MECHANISMS OF AUTORECEPTOR-MEDIATED
INHIBITION IN CENTRAL MONOAMINE NEURONS
By
NICHOLAS A. COURTNEY
Submitted in partial fulfillment for the requirements
For the degree of Doctor of Philosophy
Thesis Advisor: Christopher P. Ford, Ph.D.
Department by Physiology and Biophysics
CASE WESTERN RESERVE UNIVERSITY
January, 2016
CASE WESTERN RESERVE UNIVERISTY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
Nicholas A. Courtney
candidate for the degree of Doctor of Philosophy.
Thesis Advisor………………………………. Dr. Christopher Ford
Committee Chair……………………..………………Dr. Corey Smith
Committee Member………………..…………… Dr. Stephen Jones
Committee Member………………...…….… Dr. Ben Strowbridge
Committee Member…………………..………… Dr. Roberto Galán
Defense Date: October 30, 2015
*We also certify that written approval has been obtained
for any proprietary material contained therein.
ii TABLE OF CONTENTS
LIST OF FIGURES ………………………………………………………………………………………….....…. vi
LIST OF ABBREVIATIONS …………………………………….…………………………………………… vii
ACKNOWLEDGEMENTS ……………………………………………………………..……………………… ix
ABSTRACT …………………………………………..………………….….……………………………………… xi
CHAPTER 1
Introduction………………………………………………………………..……..…………..….. 1
Foreword………………………….……………...…………………………………..…….….………. 2
Monoamine life cycles…………….……………...………………………...……………….….… 5
G-protein coupled receptor signaling……………………………………………………. 12
Dopamine receptors………………………………………………………...…..……....……… 16
Noradrenaline receptors……………………………………………..…………..…………… 18
Serotonin receptors…………………………………………….…………………..…………… 20
G-protein coupled, inwardly rectifying potassium channels……………...…… 22
Central monoamine systems…………………..…………………………..………………… 25
Mechanisms of neurotransmission…………………………………….………….……… 37
Mechanisms of monoamine feedback inhibition.…………………..…..…………… 42
Rationale………………………………………………………………………………….………..… 45
Statement of the problem …………………………………………………….………….…… 46
CHAPTER 2
iii Species differences in somatodendritic dopamine transmission
determine D2-autoreceptor mediated inhibition of ventral
tegmental area neuron firing ……………………………………………………………. 51
Foreword……………………………………………………………………………………….……… 52
Abstract……………………………………………………………………………………..……...... 53
Introduction……………………………………………………………………………………..……. 54
Materials and methods………………………………………………………………………...... 56
Results…………………………………………………………………………………………….…..… 60
Discussion……………………………………………………………………………………….…….. 68
Figures and figure legends…………………………………………..…………………...…..… 74
CHAPTER 3
The timing of dopamine- and noradrenaline-mediated
transmission reflects underlying differences in the extent
of spillover and pooling. …………………………………………………………………… 84
Abstract……………………………………………………………………………………..…...... 85
Introduction…………………………………………………………………………………..………. 86
Materials and methods……………………………………………………………………..…….. 88
Results……………………………………………………………………………………………...….... 94
Discussion……………………………………………………………………………………….…... 108
Figures and figure legends…..…………………………………………………………...…... 114
iv CHAPTER 4
Synaptic activation of 5-HT1A receptors in dorsal raphe serotonin
neurons. …………………………………………………………………………………………. 130
Abstract……………………………………………………………………………………..…...... 131
Introduction……………………………………………………………………………………….... 132
Materials and methods…………..………….….……………………………………………..... 134
Results…………………………………..…………..…………………………………………….…... 137
Discussion………………………………..………….………………………………………….…… 145
Figures and figure legends…………..……….…………………………………………...…... 150
CHAPTER 5
Discussion.……………………………………………………………………………..…..…….162
Inhibitory timing and monoamine activity………………………………………..….. 164
Synchronous verse asynchronous neuronal activity……………...………….…….. 167
Reuptake transporters actively shape monoamine transmission ……….…... 170
Implications for monoamine transmission in terminal regions …………….…. 172
Future directions…………………………………….………………………………………..…... 175
REFERENCES…………………………………………………………………………………………….... 179
v LIST OF FIGURES
FIGURE 1.1 ……………………………………………………………………………………………………… 47
FIGURE 1.2 ……………………………………………………………………………………………………… 49
FIGURE 2.1 ……………………………………………………………………………………………………… 74
FIGURE 2.2 ……………………………………………………………………………………………………… 76
FIGURE 2.3 ……………………………………………………………………………………………………… 78
FIGURE 2.4 ……………………………………………………………………………………………………… 80
FIGURE 2.5 ……………………………………………………………………………………………………… 82
FIGURE 3.1 …………………………………………………………………………………….……………… 114
FIGURE 3.2 …………………………………………………………………………………….……………… 116
FIGURE 3.3 …………………………………………………………………………………….……………… 118
FIGURE 3.4 …………………………………………………………………………………….……………… 121
FIGURE 3.5 …………………………………………………………………………………….……………… 123
FIGURE 3.6 …………………………………………………………………………………….……………… 125
FIGURE 3.7 …………………………………………………………………………………….……………… 127
FIGURE 4.1 …………………………………………………………………………………….……………… 150
FIGURE 4.2 …………………………………………………………………………………….……………… 152
FIGURE 4.3 …………………………………………………………………………………….……………… 154
FIGURE 4.4 …………………………………………………………………………………….……………… 156
FIGURE 4.5 …………………………………………………………………………………….……………… 158
FIGURE 4.6 …………………………………………………………………………………….……………… 160
vi LIST OF ABBREVIATIONS
5-HT – Serotonin
ACSF – Artificial Cerebral Spinal Fluid
COMT - Catechol-O-Methyl Transferase
DA – Dopamine
DAT – Dopamine Reuptake Transporter
DBH – Dopamine β Hydroxylase
DRN – Dorsal Raphe Nucleus
FSCV – Fast Scan Cyclic Voltammetry
GIRK – G-protein Coupled, Inwardly Rectifying Potassium Channel
GPCR – G-protein Coupled Receptor
IPSC – Inhibitory, Post-synaptic Current
LC – Locus Coeruleus
LDCV – Large Dense Core Vesicle
MAO – Monoamine Oxidase
NA – Noradrenaline
NET – Noradrenaline Reuptake Transporter
N.S. – No Statistical Significance
SERT – Serotonin Reuptake Transporter
SSV – Small Synaptic Vesicle
TPH – Tryptophan Hydroxylase
TH – Tyrosine Hydroxylase
vii VMAT – Vesicular Monoamine Transporter
VTA – Ventral Tegmental Area
viii ACKNOWLEDGEMENTS
This work would not have been possible without the support that I have received from many people for which I am extremely grateful.
First and foremost, I would like to thank my advisor, Dr. Chris Ford. He has provided me sound guidance in both my graduate studies and my pursuit of a research career. For all that Dr. Ford has taught me about science, it is his passion for his research that I most hope to emulate. Thank you for your guidance, wisdom, and at times patience. I have enjoy my time in the Ford lab and will look back fondly on my Ph.D. experience.
I would like to thank the members of my thesis committee: Dr. Corey Smith, Dr.
Stephen Jones, Dr. Ben Strowbridge, and Dr. Roberto Galán, as well as a previous member Dr. Diana Kunze. This committee has played instrumental part in my development as a scientist. Thank you very much for dedicating time and effort to assisting me throughout my graduate studies.
I would like to thank the members of the Ford lab: Didi Mamaligas and Pam Marcott.
I cannot count the number of times you have helped me think through experiments, evaluate drafts, practice talks and posters, and generally be supportive during my graduate studies. Thank you very much and I wish you the best of luck with your own studies.
ix
I would like to thank Dr. Joe LaManna and the Neurodegenerative Training Grant staff here at Case Western. Thank you for the support demonstrated by funding my research, and more importantly, my training and future career.
I would like to thank the entire Department of Physiology and Biophysics, especially the front office staff and fellow graduate trainees.
I would like to thank my family members and friends that have supported me through my graduate career. To my parents, thank you for patience and encouragement while I was searching for a career that I could be happy pursuing.
Finally, I would especially like to thank my wife and fellow graduate student, Becki.
Your unwavering support has been instrumental in my Ph.D. journey. Thank you for everything.
x Mechanisms of Autoreceptor-Mediated Inhibition in Central Monoamine Neurons
Abstract
By
NICHOLAS A. COURTNEY
This thesis examined the mechanisms of neurotransmission underlying
feedback inhibition mediated by somatodendritic G-protein coupled dopamine D2-
autoreceptors in the ventral tegmental area (VTA), noradrenaline α2-autoreceptors
in the locus coeruleus (LC), and serotonin 5-HT1A autoreceptors in the dorsal raphe
nucleus (DRN). Local feedback inhibition mediated by these GI-coupled monoamine receptors had been speculated to ubiquitously occur via extended transmitter spillover and obligatory extracellular transmitter pooling. Collectively termed volume transmission, these mechanisms of transmission are presumed to result in a tonic, inhibitory tone that modulates the firing rates of monoaminergic neurons in response to slowly changing extracellular concentrations of monoamines. However, evidence for the volume transmission hypothesis is indirect and relies on measuring bulk extracellular monoamine concentrations and predicting receptor activation
based on steady-state affinities and mathematical diffusion modeling.
In monoamine neurons, autoreceptors couple to G-protein coupled, inwardly rectifying potassium channels (GIRKs) through the βγ-subunits of trimetric G- proteins. Activation of somatodendritic autoreceptors by locally released monoamines generates potassium currents that inhibit neuronal excitability. In this
xi thesis, I directly investigated the synaptic mechanisms controlling autoreceptor activity by using electrophysiological methods to measure GIRK-mediated currents in response to evoked transmitter release in rodent brain slices. After characterizing the calcium dependence and clearance of midbrain dopamine transmission to demonstrate species-dependent differences in dopamine transmission, I tested the contributions of spillover and pooling to shaping autoreceptor activation in the dopamine, noradrenaline, and serotonin systems.
While both spillover and transmitter pooling contributed to noradrenergic α2- autoreceptor activation, dopamine reuptake transporters prevented both spillover and pooling in D2-autoreceptor mediated dopamine transmission. In the dorsal raphe nucleus, transmitter spillover contributed to 5-HT1A autoreceptor activation while serotonin reuptake transporters prevented serotonin pooling and synaptic crosstalk.
This thesis concludes that autoreceptor-mediated inhibitory feedback transmission encoded by locally released dopamine, noradrenaline, and serotonin occurs via distinct synaptic mechanisms. While the volume transmission hypothesis may accurately describe noradrenergic transmission in the LC, dopaminergic transmission in the VTA and serotonergic transmission in the DRN likely occur in a point-to-point fashion that is inconsistent with volume transmission. These findings provide insight into how these regions may differentially encode and influence behavior.
xii CHAPTER 1
INTRODUCTION
1 Foreword
In the mammalian central nervous system, monoamine neurotransmitters
are thought to act as neuromodulators that broadly influence cognitive and
emotional processing. The primary central monoamine neurotransmitters,
including dopamine, noradrenaline, and serotonin, are synthesized and released by
a relatively small number of neurons whose somas are clustered into nuclei regions.
From these nuclei regions, widespread axonal projections provide brain-wide
monoaminergic innervation that acts on post-synaptic heteroreceptors, located on
non-monoaminergic neurons, to influence brain function and behavior. While a detailed, mechanistic understanding of how axonal monoamine release activates these heteroreceptors is undoubtedly important to advancing our understanding of the overall function of monoamine neurotransmitter systems, the slow signaling of these metabotropic heteroreceptors, often occurring over many seconds to minutes, makes it difficult to study the synaptic mechanisms regulating heteroreceptor activation.
In addition to release in projection regions, vesicular monoamine release also occurs locally within nuclei regions from somatic, dendritic, and sometimes axonal sites. This local release activates GI-coupled autoreceptors located on the somas and
dendrites of monoamine neurons to mediate feedback inhibition via the opening of
G-protein coupled, inwardly rectifying potassium channels (GIRKs). The resulting efflux of potassium ions hyperpolarizes monoamine neurons to modulate local firing
rates, patterns, and global impulse activity and ultimately influences terminal
2 monoamine release. Because of the relatively fast coupling between autoreceptors and GIRK channels, autoreceptor-mediated inhibition occurs with a second to sub- second time course that is more amiable to studying how the release of monoamines activates receptors on the synaptic level. A mechanistic understanding of how local release activates autoreceptor would advance our knowledge on the role of feedback inhibition in central monoamine neurotransmitter systems and furthermore could serve as an informative model for heteroreceptor activation in projection regions.
Numerous studies have examined how local monoamine release activates dopamine, noradrenaline, and serotonin autoreceptors. Classically, autoreceptor- mediated transmission has been studied by measuring extracellular monoamine concentrations using dialysis or electrochemistry and then predicting receptor activation, and thus transmission, using mathematical models based on receptor affinities. These studies conclude that monoamine autoreceptor-mediated transmission occurs by paracrine mechanisms similar to the canonical mechanisms of metabotropic-receptor mediated transmission. Specifically, these studies find that autoreceptor-mediated transmission occurs over long synaptic distances resulting in low monoamine concentrations acting on high-affinity GPCR autoreceptors. Loose regulation by clearance mechanisms such as reuptake transporters results in long-lasting signaling and an ambient tone of autoreceptor- mediated inhibition. Together, this is thought to result in multiple pre-synaptic release sites converging and activating overlapping pools of post-synaptic autoreceptors, termed transmitter pooling or synaptic crosstalk, and a lack of the
3 paired, independent sites of linear transmission that defined classical synaptic
transmission. In the monoamine field, this loosely regulated form of transmission is
generally referred to as volume transmission. However, it is important to note that
these paracrine mechanisms of transmission have only been predicted by modeling
approaches and have never been directly tested from the perspective of post-
synaptic autoreceptor activation.
The primary objective of this thesis work was to directly measure post-
synaptic autoreceptor activity to explicitly test the proposed paracrine mechanisms
regulating monoamine autoreceptor-mediated transmission. Nearly a decade ago, studies demonstrated that electrophysiological recordings of G-protein coupled, inwardly rectifying potassium channel (GIRK) mediated currents could be used to directly assess dopamine autoreceptor activity in response to local dopamine release. By utilized GIRK-mediated currents generated by dopamine D2- autoreceptor activity in the ventral tegmental area, noradrenaline α2-autoreceptor activity in the locus coeruleus, and serotonin 5-HT1A-autoreceptor activity in the dorsal raphe nucleus to directly assess the synaptic mechanisms by which local monoamine release activates autoreceptors, this thesis tests how synaptic distances, transmitter diffusion, monoamine clearance, and transmitter pooling shape the activity of monoamine autoreceptors. Surprisingly, these experiments found that transmission mediated by dopamine, noradrenaline, and serotonin autoreceptors was all differentially regulated at the level of synaptic transmission and in certain cases deviated significantly from canonical metabotropic, paracrine mechanisms.
4 Before discussing the experiments and results supporting this conclusion,
this thesis begins with a general introduction to monoamines and monoamine
neurotransmission. First, the synthesis, packaging, release, and degradation of
monoamine, general mechanisms of GPCR signaling and the types of monoamine
receptors, and the neuroanatomy of monoamine neurotransmitter systems will be
discussed. Next, this introduction will discuss general mechanisms of synaptic
transmission as evidenced by classical fast, ligand-gated synapses and provide a
more detailed account of the electrochemical studies used to generate the paracrine
model of monoamine transmission. Finally, this introduction will conclude by
discussing some of the relevant findings of GIRK-recordings of dopamine
autoreceptor-mediated transmission.
Monoamine life cycles
Synthesis
The biosynthesis of dopamine and noradrenaline occurs via a shared metabolic pathway that begins with the non-essential amino acid tyrosine. Located
in the cytosol of catecholaminergic neurons, tyrosine hydroxylase (TH) catalyzes the
formation of L-DOPA (L-3,4-dihydroxyphenylalanine) from tyrosine. Non-primate
mammals express a single isoform of TH (Nagatsu, 1995) and hydroxylation of tyrosine by TH is the rate-limiting step in the formation of catecholamines (Pothos et al., 1996). The catalytic rate of TH is highly regulated to control catecholamine availability, such as by competitive inhibition by dopamine and noradrenaline
5 (Daubner et al., 2011), phosphorylation of its regulatory domain (Dunkley et al.,
2004), protein-protein interactions (Perez et al., 2002), and adenylate-cyclase
mediated inhibition resulting from catecholaminergic GI-coupled GPCR activity
(Schmitz et al., 2003). Dopamine is synthesized from L-DOPA by the enzyme
aromatic amino acid decarboxylase (DDC). In noradrenergic neurons, dopamine is
hydroxylated to form noradrenaline by the enzyme dopamine β-hydroxylase (DBH).
Whereas the rest of metabolic pathway occurs in the cytosol, DBH is located within synaptic vesicles (De Potter et al., 1970) and thus requires dopamine to be loaded into vesicles before being converted into noradrenaline.
Serotonin biosynthesis begins with the essential amino acid tryptophan.
Tryptophan is converted into 5-hydroxy-L-tryptophan (5-HTP) by the enzyme tryptophan hydroxylase (TPH) with TPH2 being the predominate isoform in the
central nervous system (Invernizzi, 2007). This is the rate-limiting step of serotonin
synthesis. 5-HTP is then converted into serotonin also by DDC.
The synthesis enzymes for dopamine (TH), noradrenaline (DBH), and
serotonin (TPH) are found in the cell bodies, dendrites, and axons of monoamine
neurons (Pickel et al., 1976). Monoamine synthesis likely occurs at both
somatodendritic sites and axon terminals (Demarest and Moore, 1979)(Demarest
and Moore, 1979)(Demarest and Moore, 1979).
VMAT and vesicular release
Dopamine, noradrenaline, and serotonin are packaged into vesicles via the
vesicular monoamine transporter (VMAT). VMAT is a twelve transmembrane
6 domain protein that exchanges a single monoamine molecule for two molecules of
hydrogen using the hydrogen electrochemical gradient generated by vacuolar-type
H+-ATPases (Chaudhry et al., 2008). VMAT has two isoforms of which primarily
VMAT2 is expressed in central dopaminergic, noradrenergic, and serotoninergic
neurons (Erickson et al., 1996). While VMAT2 has relatively similar affinities and
maximum turnover rates for NE, DA, and 5HT (Wimalasena, 2011), the relatively
low maximum turnover rate of VMAT (Peter et al., 1994) sometimes results in
vesicle loading being the rate-limiting step of transmission (Colgan et al., 2009).
Similar to the distribution of synthesis enzymes, VMAT is found at both
somatodendritic and axonal sites (Nirenberg et al., 1995, 1996) suggesting that the
synthesis and vesicular packaging of monoamines likely occurs at both sites.
Within monoamine neurons, VMAT2 localizes to both small synaptic vesicles
(SSVs) and more predominately to large, dense-core vesicles (LDCVs) (Nirenberg et al., 1995, 1996). Both SSVs and LDCVs are thought to be involved in the storage and exocytosis of serotonin (Bruns et al., 2000; Colgan et al., 2009, 2012), dopamine
(Nirenberg et al., 1996), and noradrenaline (Groves and Wilson, 1980). SSVs and
LDCVs are both present in the somas and dendrites of monoamine neurons (Colgan et al., 2012; Nirenberg et al., 1996). Serotonin terminals may preferentially contain
SSVs (Bruns et al., 2000) while dopamine terminals are reported to contain both
SSVs and LDCVs (Nirenberg et al., 1995). In addition to SSVs and LDCVs, VMAT localizes to tubulovesicles in the dendrites of midbrain dopamine neurons
(Nirenberg et al., 1996). These vesicles were reported to resemble the morphology of smooth endoplasmic reticulum (Nirenberg et al., 1996). While it is unclear to
7 what degree tubulovesicles, SSVs, or LDCVs contribute to monoamine storage and
release, some of the general mechanisms underlying release have been elucidated.
The mechanisms of monoamine release have been most studied in the
dopamine system. Both axonal and somatodendritic dopamine release require
VMAT-dependent vesicle loading (Millar et al., 1985; Rice et al., 1994) and
functional SNARE complex proteins (Bergquist et al., 2002; Fortin et al., 2006).
While SNAP-25 is likely required at both sites, the specific isoforms of other SNARE proteins may differ for axonal and dendritic release (Bergquist et al., 2002). Axonal
dopamine release clearly requires influx of calcium through voltage-gated calcium
channels (Herdon and Nahorski, 1989; Moghaddam and Bunney, 1989). However,
the calcium dependence of somatodendritic release is debated. One series of
studies measuring extracellular dopamine is response to evoked release in guinea
pigs concluded that somatodendritic dopamine release is only weakly facilitated by
extracellular calcium concentrations (Chen and Rice, 2001; Chen et al., 2011). In
this study, peak dopamine release was achieved with as little as 1.5 mM Ca2+ extracellular calcium and nearly half maximal release was still observed in 0 mM
Ca2+ (Chen and Rice, 2001). Mobilization of calcium from intracellular stores may have accounted for this weak dependence on extracellular calcium (Patel et al.,
2009). In contrast, studies performed in mice found that both extracellular dopamine concentrations and post-synaptic dopamine receptor activation in response to evoked release was strongly calcium dependent (Ford et al., 2010). This study found that dopamine release was abolished when extracellular calcium was removed, that maximum dopamine release required greater than 2.4 mM Ca2+, and
8 that depleting intracellular calcium with cyclopiazonic acid (CPA) had no effect on
evoked dopamine receptor activation (Ford et al., 2010).
Release of serotonin from the somas and dendrites of DRN neurons has been
studied using serotonin optical imaging. While both somatic and dendritic release
are dependent on extracellular calcium and VMAT (Colgan et al., 2009, 2012), only
somatic release was found to be dependent on action potentials (Colgan et al.,
2009). Because serotonin neuron dendrites are electrically isolated from somatic action potentials (Colgan et al., 2012), activation of local NMDA receptors instead
drives calcium influx and ensuing serotonin vesicle fusion (Colgan et al., 2012).
Release of serotonin from cultured DRN neurons is sensitive to tetanus toxin
(Lautenschlager et al., 2000) suggesting involvement of SNARE complexes; however, this has not been test either in slice or in vivo. Additionally, non-vesicular serotonin release has also been suggested in a carrier-mediated fashion that can be driven by
ρ-chloroamphetamine (Adell et al., 1989, 2002; Kuhn et al., 1985; Levi and Raiteri,
1993). Whether this occurs under normal physiology, however, remains unclear.
Monoamine reuptake transporters
Following transmission, monoamines are cleared from the extrasynaptic space in a Na+/Cl- dependent manner by the actions of three reuptake transporters: noradrenaline reuptake transporters (NETs), dopamine reuptake transporters
(DATs), and serotonin reuptake transporters (SERTs) (Ramamoorthy et al., 2011).
DATs are primarily expressed on the somas, dendrites, and axon terminals of
dopamine neurons in the central nervous system (Ramamoorthy et al., 2011) and
9 mediate the clearance of dopamine. DAT knockout mice have increased
extracellular dopamine levels (Giros et al., 1996) and are notably hyperactive, indifferent to cocaine, and have cognitive defects (Gainetdinov and Caron, 2003;
Giros et al., 1996). The primary actions of psychostimulants, such as cocaine and amphetamine, are mediated by DATs. Whereas cocaine blocks DAT activity to inhibit dopamine uptake, amphetamine and its derivatives trigger the efflux of
dopamine through DATs (Ramamoorthy et al., 2011).
NETs are expressed exclusively in noradrenaline neurons in the central
nervous system; noradrenaline reuptake in terminal regions relies on pre-synaptic
NETs (Foote et al., 1983) and mediates the clearance of noradrenaline. While NET is
encoded by a single gene, alternative splicing of the 3’ carboxy terminal may
regulate its expression and function (Kitayama et al., 2001). Mice lacking NET
activity, due either to genetic knockout or pharmacological inhibition, are prone to
seizures, sensitive to opioids and cocaine, and have altered response to social stress
(Ahern et al., 2005; Bohn et al., 2000; Haller et al., 2002). In addition to
noradrenaline, NETs can also mediate the clearance of dopamine in brain regions
lacking significant DAT expression (Madras et al., 2005; Morón et al., 2002).
SERTs are also encoded by a single gene (Ramamoorthy et al., 1993) and are expressed only in serotonin neurons in the central nervous system (Ramamoorthy et al., 2011). On serotonin neurons, SERTs localize both to somatodendritic regions as well as projection axon terminals. SERT knock-out mice have impaired serotonin clearance resulting in an altered response to cocaine and MDMA, increased anxiety- like behaviors, and altered stress responses (Bengel et al., 1998; Holmes et al., 2003;
10 Li et al., 1999; Sora et al., 1998). SERTs are the primary target of selective serotonin
reuptake inhibitors (SSRIs) used in the treatment of depression.
Monoamine degradation
After monoamines are transported into the cytosol by reuptake transporters,
they are either recycled back into synaptic vesicles for future use or degraded by
either monoamine oxidase (MAO) or catechol-O-methyl transferase (COMT).
Serotonin is degraded to 5-HIAA in one step catalyzed by MAO. The degradation of
dopamine to HVA and noradrenaline to MHPG is a two-step process with one step
catalyzed by MAO and the other by COMT; these steps can occur in either order.
Monoamine oxidase catalyzes the removal of an amino group (Hare, 1928).
Two separate genes encode two isoforms of monoamine oxidase, MAO-A and MAO-
B, that have different affinities for their substrates (Finberg, 2014). While MAO-A primarily degrades serotonin and noradrenaline, both MAO-A and MAO-B can equally degrade dopamine (Kalgutkar et al., 2001). Consistent with this, noradrenergic neurons in the locus coeruleus primarily express MAO-A (Fagervall and Ross, 1986; Finberg, 2014). MAO activity in midbrain dopamine neurons is more complex; striatal dopamine degradation is achieved mainly by MAO-A
(Wachtel and Abercrombie, 1994) while midbrain cell bodies display little MAO-A expression or activity (Arai et al., 1998; Westlund et al., 1993). Finally despite serotonin being a selective substrate of MAO-A, serotonin neurons in the DRN primarily express MAO-B (Finberg, 2014), though their projection fields may have
MAO-A expression (Fagervall and Ross, 1986). At the subcellular level, both MAO
11 isoforms localizes to mitochondrial outer membranes (Denney and Denney, 1985;
Edmondson et al., 2009).
Catechol-O-methyl transferase (COMT) catalyzes the addition of a methyl
group to catecholamines. Both a membrane-bound and a soluble form of COMT are
encoded by a single gene; a truncation of ~50 amino acids from the amino terminal
of COMT results in the loss of the signal-anchor peptide to form the soluble isoform
(Ulmanen et al., 1997). In the central nervous system, COMT is primarily located
intracellularly in neurons (Schott et al., 2010; Ulmanen et al., 1997), though
extracellular COMT has been reported in the periphery.
G-protein coupled receptor signaling
Because this thesis deals extensively with actions mediated by G-protein coupled receptors (GPCRs), their basic properties will be detailed along with specific information on monoaminergic receptors.
GPCRs and the G-protein cycle
G-protein coupled receptors (GPCRs) are the largest family of surface- expressed receptors and are involved in almost every physiological process and behavior. The larger superfamily of GPCRs can be subdivided into five classes based on sequence alignment using the recent GRAFS classification system: Glutamate,
Rhodopsin, Adhesion, Frizzled/Taste2, and Secretin (Schiöth and Fredriksson,
2005). Despite having multiple subfamilies, all GPCRs share a common structure of
12 seven transmembrane alpha helixes joined by intra- and extracellular loops
(Rosenbaum et al., 2009; Trzaskowski et al., 2012). Diversity in the sequence and
length of the extracellular loops, the extracellular N-terminus, and in some cases the
transmembrane helixes themselves leads to specificity of ligand binding. Binding of
ligand to the extracellular face of the GPCR is thought to induce a membrane-
spanning conformational change in the spatial arrangement of the helixes that
exposes the G-protein binding domain (Rasmussen et al., 2011; Yeagle and Albert,
2003). Subsequent to G-proteins binding, GPCR GEF domains catalyze the release of
guanosine diphosphate (GDP) from the Gα subunit of heterotrimeric G-proteins.
The subsequent binding of guanosine triphosphate (GTP) triggers the dissociation of the βγ subunit dimer (Neer, 1995). This activates the G-protein and allows free Gα
and βγ dimers to interact with downstream intracellular effectors. GPCR-induced G-
protein signaling cascades are terminated by the hydrolysis of the Gα-bound GTP.
Hydrolysis occurs due to intrinsic GTPase activity of Gα subunits (Hollinger and
Hepler, 2002; Kach et al., 2012). However, this process is slow and is often accelerated by the action of regulators of G-protein signaling (RGS) proteins.
Hydrolysis of GTP to GDP causes the reassociation of α and βγ subunits and thereby inactivates G-proteins, ceasing GPCR signaling.
Heterotrimeric G-proteins are the primary effectors of GPCRs. Specific associations between GPCRs and G-protein isoforms, determined by the Gα subunit isoform, allow for discriminate activation of signaling pathways and effectors.
Currently, at least 20 isoforms of Gα have been described that divide into four families: GαS, GαI/O, GαQ, and Gα12/13.
13
GαS
Activation of GαS increases intracellular concentration of cyclic adenosine
monophosphate (cAMP) by stimulating adenylyl cyclase (AC) (Rall and Sutherland,
1962). While signaling can also occur through cyclic-nucleotide-gated (CNG)
channels and guanine-nucleotide exchanging proteins activated by cAMP (Epac)
(Calebiro and Maiellaro, 2014), cAMP classically activates protein kinase A (PKA) as
the primary effector of GαS signaling. GαS signaling is terminated by the
degradation of cAMP by cyclic nucleotide phosphodiesterases.
GαI/O
Activation of GαI/O decreases the production of cAMP by inhibiting adenylyl
cyclase (Birnbaumer, 1992). Because of ongoing degradation by
phosphodiesterases, this results in lower concentrations of cAMP and reduced PKA
activity. Thus, the GαS and GαI/O signaling pathways ultimately oppose one another
in their cAMP-mediated effects. Dimers composed of a β and γ subunit that disassociate from GαI/O can also independently activate effectors including G-
protein coupled inwardly rectifying potassium channels (GIRKs) (Kofuji et al.,
1995). Because GIRKs are the major effectors studied in this thesis, they will be covered in detail in a later section.
GαQ
14 GαQ activates phospholipase C (PLC) to cleave phosphatidylinositol 4,5- bisphosphate (PIP2) into membrane-bound diacyl glycerol (DAG) and freely diffusible inositol trisphosphate (IP3). One major effect of IP3 is the activation of IP3-
sensitive calcium channels located on the endoplasmic reticulum (ER) causing
calcium release from ER stores resulting in a rise in cytosolic calcium. In
conjunction with intracellular calcium, DAG then activates protein kinase C (PKC)
(Sánchez-Fernández et al., 2014).
Gα12/13
Gα12/13 proteins are the most recently identified family of Gα proteins and
their signaling pathway is yet to be fully understood (Neves et al., 2002). The first
proposed direct effectors of Gα12/13 signaling are Bruton’s tyrosine kinase (BtK) and
Gap1m, a Ras GTPase-activating protein (Jiang et al., 1998). Gα12/13 proteins are
also thought to lead to downstream activation of certain RhoGEF proteins, though
this mechanistic pathway is not understood (Siehler, 2009). Ultimately, the net
effect of Gα12/13 activation is to stimulate phospholipase D, c-SC, and PKC though
this signaling is achieved remains unclear (Neves et al., 2002).
Arrestin signaling and GPCR desensitization
When exposed to agonist, GPCRs are dynamically regulated to reduce their
sensitivity (Ferguson, 2001). This rapid attenuation of responsiveness is termed
desensitization. During homologous desensitization, activated GPCRs are
phosphorylated by G-protein coupled receptor kinases (GRKs) that recruit arrestin
15 adaptor proteins (Gainetdinov et al., 2004; Lohse et al., 1990; Pitcher et al., 1998).
Binding of arrestins prevents the further activation of G-proteins and can additionally recruit clathrin (Laporte et al., 2002) to internalize the receptor. When agonist is removed from the receptor, GPCRs resensitize via dephosphorylation and endosome recycling (L. Mohan et al., 2012). In addition to inducing desensitization, arrestin binding can also trigger GPCR signaling pathways independent of trimeric
G-proteins (Hall et al., 1999; Luttrell et al., 1999).
Dopamine receptors
Dopamine acts on five known classes of receptors, all of which are GPCRs,
that subdivide into two families: D1-like receptors that increase intracellular cAMP and D2-like receptors that reduce cAMP (Spano et al., 1978). Genetic cloning has
identified multiple receptor subtypes in both of these families (Dearry et al., 1990;
Monsma et al., 1990; Sokoloff et al., 1990; Sunahara et al., 1991; Tiberi et al., 1991;
Van Tol et al., 1991; Zhou et al., 1990).
D1-like receptors include the D1 and D5 receptor. These receptors are exclusively found on post-synaptic, non-dopaminergic neurons and stimulate cAMP production through GαS signaling. The D1-receptor is the most abundantly expressed family member and is found with high expression density in areas such as the striatum, substantia nigra, olfactory bulb, amygdala, and frontal cortex. It is also expressed at lower densities in the hippocampus, cerebellum, and thalamic and hypothalamic areas (Beaulieu and Gainetdinov, 2011). Compared to the D1-
16 receptors, the D5-receptor has a much higher affinity for dopamine (Grandy et al.,
1991) but is expressed far less frequently. D5-receptors are expressed at low levels
in prefrontal cortex pyramidal neurons, the substantia nigra, the hypothalamus, the
hippocampus, the dentate gyrus, and the premotor, cingulated, and entorhinal
cortex (Beaulieu and Gainetdinov, 2011). D5-receptors are also reportedly
expressed in specific features of the nucleus accumbens (Muly et al., 2010).
Additionally, two genes have been described that encode non-functional, truncated
D5-receptors (Grandy et al., 1991).
D2-like receptors include the D2, D3, and D4 receptor. Unlike D1-like receptors, D2-like receptors are expressed dopamine neurons themselves as well as in non-dopaminergic post-synaptic targets. In dopamine neurons, these receptors
act both at pre-synaptic terminals and somatodendritic sites. D2-like receptors
couple to GαI; thus, activation of these receptors inhibits cAMP production and in
some neurons, such as midbrain dopamine neurons, activates GI coupled GIRK
channels. D2-receptors are the most abundantly expressed receptor in this family,
with high levels of expression found in the striatum, nucleus accumbens, and the
olfactory tubercle. D2-receptors are also expressed at moderate densities in the
substantia nigra, ventral tegmental area, hypothalamus, septum, amygdala,
hippocampus, and cortical areas (Beaulieu and Gainetdinov, 2011). D3- and D4-
receptors have significantly lower levels of expression than D2-receptors. D3 receptors are expressed in the nucleus accumbens shell and olfactory tubercle and at lower levels in the striatum, substantia nigra pars compacta, ventral tegmental area, the hippocampus, and some cortical regions. D4-receptors have been found in
17 the hippocampus, hypothalamus, globus pallidus, substantia nigra pars reticulate,
and thalamus (Beaulieu and Gainetdinov, 2011). While the genes encoding D1-like receptors contain no introns in their coding sequence, genes encoding D2-like receptors contain introns and thus can have multiple splice variants (Gingrich and
Caron, 1993). Specific to D2-receptors, alternate slicing can lead to short- and long- form D2-receptors (Dal Toso et al., 1989). These isoforms have different agonist affinities and effector coupling properties (Castro and Strange, 1993; Ford, 2014) suggesting a physiological relevance for alternate slicing.
Dopamine neurons in the midbrain express D2- and D3-receptors (Diaz et al.,
2000). Activation of D3 receptors by specific agonists has been reported to inhibit
VTA dopamine neuron firing rates (Lejeune and Millan, 1995). However, in D2- receptor knockout mice the D2/D3 receptor agonist quinpirole was unable to generate GIRK currents in substantia nigra dopamine neurons (Davila et al., 2003) suggesting that D3 receptors alone are insufficient to activate GIRK channels.
Noradrenaline receptors
Noradrenaline acts through at least nine known adrenergic receptors, all of which are GPCRs (Gannon et al., 2015). Adrenergic receptors can also be activated by adrenaline in addition to noradrenaline, though this primarily occurs in the periphery. There are two major groups of adrenergic receptors: α and β receptors.
Alpha-receptors include Gq-coupled α1-receptors and GI-coupled α2-receptors; β-
receptors are all GS-coupled and include β1, β2, and β3 subtypes. β-receptors are
18 largely peripheral receptors, though they do have some expression in the central
nervous system.
Three subtypes of α1-receptors have been cloned: α1A, α1B, and α1D. Being
GαQ-coupled, these receptors signal through IP3 and cytosolic calcium (Chen and
Minneman, 2005; Perez, 2007). In the central nervous system, all subtypes of α1-
receptors are exclusively expressed on non-noradrenergic, post-synaptic neurons
(Gannon et al., 2015). Holistically, α1-receptors are important for fear conditioning and spatial learning. Application of α1-receptor antagonists hamper olfactory fear conditioning (Do Monte et al., 2013) and impair spatial memory measured by a
Morris water maze (Torkaman-Boutorabi et al., 2014). Consistent with this, treatment with α1-receptor agonists improves performance in Morris water maze tests (Puumala et al., 1998). Finally, α1-receptors are an important excitatory driver of tonic firing of serotonergic neurons in the DRN (Baraban and Aghajanian,
1981).
Three subtypes of α2-receptors have also been identified that are encoded by separate genes: α2A, α2B, and α2C (Harrison et al., 1991). While α2-receptors are
expressed post-synaptically on non-noradrenergic neurons, they are unique among
adrenergic receptors in that they are also expressed as autoreceptors both on
axonal terminals and somatodendritically on noradrenergic neurons (Lee et al.,
1998a, 1998b). These α2-receptors are GI/O-coupled and thus inhibit cAMP
production and PKA activity. In the locus coeruleus α2-receptors couple to GIRK
channels. Both α2A and α2C receptor subtypes have been identified by electron
19 microscopy studies as residing on the somas and dendrites of noradrenergic
neurons (Lee et al., 1998b, 1998c). However, in situ hybridization assays only
detected α2A mRNA in the locus coeruleus (Scheinin et al., 1994). mRNA for α2A
receptors is expressed in almost all brain regions, including the locus coeruleus and
other brain stem regions, dorsal and medial raphe nuclei, hypothalamic regions,
amygdala, hippocampus, and cerebral cortex (Scheinin et al., 1994). In contrast, α2B and α2C receptors have much more restricted expression patterns, with α2B receptor mRNA found only in thalamic regions and α2C receptor mRNA found primarily in the hippocampus, cortical, and olfactory regions (Scheinin et al., 1994). It is unclear how these receptor subtypes differ either mechanistically or in their physiological relevance.
Serotonin receptors
In the central nervous system, serotonin signaling is mediated by 14 distinct serotonin receptors that are subdivided into seven subtypes: 5-HT1-7. Almost all
serotonin receptors are metabotropic GPCRs; the only ionotropic receptors are 5-
HT3 receptors which are pentameric, cation-selective, ligand-gated ion channels
(Hoyer et al., 1994). Depending on the subtype, serotoninergic GPCRs can signal
through GS, GI/O, or GQ; 5-HT1 and 5-HT5 receptors couple to GI/O, 5-HT2 receptors
couple to GQ, and 5-HT4, 5-HT6, and 5-HT7 receptors couple to GS (McCorvy and
Roth, 2015).
20 Subtypes of GI-coupled 5-HT1 receptors include 5-HT1A, 5-HT1B, 5-HT1D, 5-
HT1E, and 5-HT1F receptors. In addition to being located on non-serotonergic post-
synaptic neurons, 5-HT1A receptors are expressed as autoreceptors on the somas and dendrites of serotonin neurons. Consistent with this role, 5-HT1A receptors are
most highly expressed in the dorsal raphe nucleus, hippocampus, and cortical
regions (Hoyer et al., 1994). However, unlike dopamine and noradrenaline
autoreceptors, 5-HT1A receptors are not expressed on serotonergic terminals.
Instead, either 5-HT1B receptors in rodents or 5-HT1D receptors in non-rodent mammals (Hoyer et al., 1994) are located on both serotonergic and non- serotonergic axon terminals where they serve to inhibit transmitter release (Sari et al., 1999). GI-coupled 5-HT5 receptors include 5-HT5A and 5-HT5B receptors in
rodents (Matthes et al., 1993) with only 5-HT5A receptors being functional in
humans. Little is known about the function and expression patterns of these
receptors.
5-HT2 receptors are divided into 3 subtypes: 5-HT2A, 5-HT2B, and 5-HT2C. 5-
HT2A and 5-HT2C receptors are expressed on DRN serotonin neurons (Boothman et
al., 2003), though to a lesser extent than 5-HT1A receptors (Marinelli et al., 2004).
Because these receptors are GQ-coupled, their activation increases the excitability of
DRN serotonin neurons (Marinelli et al., 2004). Additionally, these receptors are
found on non-serotonergic neurons in the DRN as well as in the substantia nigra,
cortical regions, globus pallidus, choroid plexus, claustrum, and olfactory tubercle
(Hoyer et al., 1994).
21 Of the GS-coupled serotonin GPCRs, 5-HT4 receptors are primarily found in the colliculus and the hippocampus (Hoyer et al., 1994). 5-HT6 receptor mRNA is present at the highest levels in the striatum, followed by the olfactory tubercle, the cerebral cortex, and the hippocampus (Hoyer et al., 1994). 5-HT6 receptors typically localize to GABA neurons so that their activation produces an overall inhibitory effect on brain circuitry (Schechter et al., 2007). 5-HT7 receptors are mainly expressed in the thalamus and other forebrain regions (Lovenberg et al., 1993) and are implicated in circadian rhythms.
G-protein coupled, inwardly rectifying potassium channels
G-protein coupled, inwardly rectifying potassium channels are the primary effector of GPCR-mediated signaling studied in this thesis. As such, their basic properties and expression profiles are detailed.
GIRK channel isoforms and expression patterns.
Perhaps the fastest known effectors of inhibitory monoamine signaling are G- protein coupled, inwardly rectifying potassium channels (GIRKs). GIRK channels are members of the inwardly rectifying potassium channel superfamily (Kir3) and are functionally tetramers (Liao et al., 1996; Tucker et al., 1996; Yang et al., 1995).
In mammals there are four GIRK subunits (labeled either GIRK1 – GIRK4 or Kir3.1-
Kir3.4), though GIRK4 is predominantly cardiac and has little brain expression
(Wickman et al., 2000). Of these subunits, GIRK2 alone is thought to be able to form
22 functional homotetramers (Lesage et al., 1995; Slesinger et al., 1996), while the
other subunits combine pairwise with each other or with GIRK2 to form functional
heterotetramers (Jelacic et al., 1999, 2000; Lesage et al., 1995; Lüscher and
Slesinger, 2010). Different subunit compositions are expressed in various neurons
and have different single channel properties resulting in different macroscopic
currents and effects on excitability.
Noradrenergic neurons of the locus coeruleus can express mRNA for all four
GIRK isoforms with GIRK1 and GIRK2 being the most ubiquitously expressed
(Kawano et al., 2004). While it is unknown which GIRK isoforms couple to α2-
receptor signaling, currents generated by GI-coupled opioid receptors in the LC are
reduced by > 80% in GIRK2/3 knockout mice (Torrecilla et al., 2002).
Protein expression of GIRK1, GIRK2, and GIRK3 can all be detected to some degree in the ventral tegmental area (del Burgo et al., 2008). Midbrain dopamine neurons specifically all express some degree of GIRK2; however, while all substantia nigra neurons strongly express GIRK2, only 50-60% of VTA neurons have strong expression (Reyes et al., 2012). GIRK2 knockout mice lack evoked D2-IPSCs while
GIRK3 knockout mice are similar to wild type (Beckstead et al., 2004). This suggests that GIRK2 subunits are at least necessary and perhaps sufficient for D2-receptor-
GIRK coupling in the midbrain.
Neurons in the dorsal raphe nucleus express both the mRNA and protein for
GIRK1, GIRK2, and GIRK3 (del Burgo et al., 2008). Mice lacking GIRK2 subunits specifically display blunted effects of the 5-HT1A agonist, 8-OH-DPAT, and the
23 serotonin reuptake inhibitor, citalopram, on inhibiting DRN firing rates (Llamosas et
al., 2015) implicating GIRK2 subunits in 5-HT1A receptor signaling.
GPCR-GIRK signaling
Historically, the mechanisms of GIRK channel activation were first described
in relation to cardiac muscarinic acetylcholine receptors (mAChRs). While mAChRs
required GTP to produce GIRK currents (Pfaffinger et al., 1985), it was canonically
thought that isoforms of α subunits alone could mediate G-protein signaling. The observation that the application of βγ subunits, but not α subunits, to pulled patches
containing GIRK-channels was sufficient to induce a potassium current both
identified the molecular pathway leading to GIRK activation as well as
revolutionized theories of G-protein signaling to include non-α subunit mediated
pathways (Logothetis et al., 1987).
GIRK channels are activated directly by the binding of the βγ subunit of GI/O
proteins (Huang et al., 1995; Logothetis et al., 1987; Wickman et al., 1994). There is
strong interest in whether GIRK channels reside nearby GPCRs or if βγ subunits
undergo extended membrane-delimited diffusion before activating GIRKs. Recent studies have provided evidence that GPCRs, G-proteins, and GIRKs might be pre- associated in macromolecular signaling complexes (David et al., 2006; Lavine et al.,
2002; Lober et al., 2006; Nobles et al., 2005). As Gα subunits have been reported to associate with GIRKs, specific signaling between certain GPCRs and GIRKs may be facilitated by these complexes (Clancy et al., 2005). Additionally, preformed
24 complexes may result in faster kinetics of both GIRK activation and deactivation
(Bünemann et al., 2003) in response to GPCR signaling.
Plasticity of GIRK channels
Recent studies have begun illustrating GIRK channel plasticity that can either
potentiate or depress currents likely via relocation of GIRK channels to and from the
cell surface membrane (Luján et al., 2014). Long term potentiation of GIRK currents, first demonstrated studying GABAB mediated GIRK currents in
hippocampal pyramidal neurons, can result from calcium entry via NMDA receptors
activated calcium-calmodulin dependent kinase II (CaMKII) (Huang et al., 2005)
leading to insertion of GIRK channels into the surface membrane (Chung et al.,
2009). In dopamine neurons, classical long-term potentiation and long-term depression protocols result in GIRK current plasticity. Bursts of stimulation potentiate GIRK currents while stimulation mimicking tonic firing depresses GIRK currents (Lalive et al., 2014). Both of these processes are GPCR independent, instead occurring by glutamate receptors, CaMKII activity, and increased GIRK trafficking to the surface membrane (Lalive et al., 2014). Prolonged morphine exposure (~ 20 hours) has also been observed to increase the localization of GIRK channels to dendritic spines in cultured hippocampal neurons through the CamKII pathway (Nassirpour et al., 2010).
Separately, acute D2-receptor activation induces a long-term depression of
D2-receptor mediated GIRK currents that is dependent on post-synaptic intracellular calcium and prevented by intracellular calcium chelators (Beckstead
25 and Williams, 2007). While this depression is likely due in part to internalization of
D2-receptors via the β-arrestin pathway (Beckstead and Williams, 2007), depression of other GI-coupled receptors has resulted from co-internalization of receptors and GIRK channels (Clancy et al., 2007).
Central monoamine systems
Central dopamine system
The activity of dopamine neurons has been associated with a wide range of behaviors, including: reward encoding, reward-based learning, aversion, salience, uncertainty, novelty, and initiating coordinated movements (Lammel et al., 2014;
Schultz, 2007; Ungless, 2004; Ungless et al., 2010). Globally knocking out dopamine signaling in mice most notably results in a lack of goal-directed movements. For example, knockout mice will right themselves when flipped and maintain balance but will not move to seek out food even when faced with starvation (Zhou and
Palmiter, 1995).
Dopamine neurons in the central nervous system are primarily clustered into two bilateral midbrain nuclei: the ventral tegmental area (VTA) and the substantia nigra pars compacta (SNc). Despite the relatively small number of dopamine neurons, ~450,000 in humans and ~25,000 in mice (German et al., 1983), wide ranging, highly branched projections from these neurons provide substantial innervation brain-wide (German et al., 1983)(German et al., 1983)(German et al.,
1983)(German et al., 1983)(German, Schlusselberg, and Woodward 1983). The
26 largest recipient of dopamine innervation is the striatum, with the VTA
preferentially innervating the nucleus accumbens and the SNc preferentially
innervating the dorsal striatum. Individual medium spiny neurons in the striatum
receive input from ~1,000 dopamine synapses (Arbuthnott and Wickens, 2007) and
the axonal arborization from a single midbrain dopamine neuron can cover ~3% of
the volume of the entire striatum (Matsuda et al., 2009). Dopamine axons primarily
synapse onto the necks of dendritic spines (~70% of total synapses), though they
sometimes synapse on dendritic spine heads (Descarries et al., 1996; Freund et al.,
1984). Whether these synaptic contacts are true, membrane-specialized synapses
remains debated and it has been proposed that axonal DA release in the striatum
can occur by both synaptic and non-synaptic mechanisms (Bérubé-Carrière et al.,
2012; Descarries et al., 2008). Dopamine projections from the substantia nigra pars
compacta to the dorsal striatum are specifically thought to be important for
initiating coordinated movements, and dysfunctions in this pathway are thought to
underlie the neurodegenerative disorder Parkinson’s disease (Burns et al., 1983;
Schultz et al., 1983, 1989). Dopamine projections from the ventral tegmental area to
the nucleus accumbens, in contrast, are implicated in reward-based learning and addiction (Koob, 1992; Schultz, 1997). In addition to the striatum, dopamine neurons also send projections to the olfactory bulb, amygdala, pre-frontal cortex,
and piriform cortex (Sara, 2009) (Figure 1.1).
Both the VTA and SNc have heterogeneous cell populations. In the VTA,
~2/3 of neurons are dopaminergic. Of the remaining neurons, the vast majority are
GABAergic interneurons that regulate the activity of dopamine neurons (~1/3 of
27 total neurons) with a trace amount of glutamatergic neurons also being observed
(Nair-Roberts et al., 2008). Markers of dopamine and glutamate vesicles co-localize in some VTA neurons, and co-release of dopamine and glutamate has been observed in the nucleus accumbens (Stuber et al., 2010). The SNc has a similar distribution of dopaminergic and GABAergic interneurons (~70:30) but lacks glutamatergic neurons and glutamate co-release (Nair-Roberts et al., 2008). The relative distribution of dopamine and GABA neurons is non-uniform across spatially defined subregions in the midbrain (Nair-Roberts et al., 2008), and midbrain dopamine neurons in general can co-release GABA in the absence of GABA synthesis by internalizing and repackaging ambient GABA (Tritsch et al., 2014). For both regions, dopamine neuron markers include tyrosine hydroxylase (TH) and dopamine reuptake transporters (DATs). Electrophysiologically, dopamine neurons can be identified by the presence of D2-receptor mediated dopamine conductance, pacemaker firing of 1-5 Hz, broad action potential waveforms (> 1.2 ms), high membrane resistance (>400 MΩ), and comparatively larger cell bodies (>25 pF capacitance) (Courtney et al., 2012; Ford et al., 2009; Lammel et al., 2008).
Whole-brain inputs to midbrain dopamine neurons have been mapped by
utilizing modified rabies virus. Dopamine neurons in the SNc receive mono-synaptic
inputs primarily from the dorsal striatum, globus pallidus, superior colliculus,
substantia nigra pars reticulate, pedunculopontine tegmental nucleus, parabrachial
nucleus, subthalmic nucleus, and parts of the motor cortex (Watabe-Uchida et al.,
2012). Mono-synaptic inputs to the VTA are slightly different, with the primary sources of innervation originating from the dorsal striatum, dorsal raphe nucleus,
28 lateral hypothalamic area, nucleus accumbens shell and core, ventral pallidum, globus pallidus, and the amgydala (Watabe-Uchida et al., 2012). Local dopamine release also influences the excitability of midbrain dopamine neurons. In both the
VTA and SNc, dopamine is thought to be released from somatodendritic locations
(Cheramy et al., 1981; Courtney et al., 2012; Ford et al., 2010; Geffen et al., 1976;
Groves and Linder, 1983; Jaffe et al., 1998; Wilson et al., 1977), while dopamine released in the VTA can also originate from axon collaterals (Nirenberg et al., 1996).
GI-coupled D2-autoreceptors are the only dopamine receptors expressed on midbrain dopamine neurons. Local dopamine release acts on these inhibitory autoreceptors to modulate the firing patterns of dopamine neurons. Most notably, transient activation of these receptors following bursts of dopamine activity lead to pauses in neuron firing that are thought to be important in reward encoding
(Schultz et al., 1997). Furthermore, mice lacking dopamine autoreceptors in the midbrain are hyperactive and have increased sensitivity to cocaine (Anzalone et al.,
2012; Bello et al., 2011) indicating a behavioral relevance for midbrain dopamine transmission.
Electron microscopy studies have attempted to understand midbrain dopamine transmission by examining the ultrastructural characteristics of dopamine transmission. VTA dopamine neurons, labeled by TH immunoreactivity, can form dendrodendritic and axodendritic synapses with other TH+ membranes
(Bayer and Pickel, 1990); however, only 66% of all labeled somas and 15% of all labeled dendrites were found in apposition to other TH-labeled dendrites or somas
(Bayer and Pickel, 1990). Pre-synaptic dopamine release sites, labeled by the
29 vesicular monoamine transporter (VMAT), were rarely observed with TH+ apposing membranes in the SNc while clusters of VMAT labeled vesicles could sometimes be observed near pre-synaptic membranes that were apposed to a TH+ post-synaptic membrane in the VTA (Nirenberg et al., 1996). In the SNc these pre-synaptic release sites are located only on somas and dendrites of TH+ neurons while in the VTA release sites are also located on axons (Juraska et al., 1977; Wassef et al., 1981).
Thus, dopamine release in the SNc is primarily somatodendritic while release in the
VTA may be from both somatodendritic and axon collateral sites. Midbrain post- synaptic dopamine reception sites, labeled by the dopamine D2-receptor, were found both apposed to axon terminals and widespread throughout non-synaptic portions of plasma membranes (Sesack et al., 1994). Thus, ultrastructural studies of midbrain dopamine neurons have not illuminated the mechanisms or structural features by which transmission occurs. This is partially due to a lack of serial, quantitative electron microscope imaging to clearly define the prevalence of synaptic and non-synaptic structures.
Central noradrenaline system
Noradrenergic transmission has been implicated in attention and arousal, stress, anxiety, and depression (Aston-Jones et al., 1999; Itoi and Sugimoto, 2010).
Targeted ablation of noradrenaline neurons in adult mice results in mice that display reduced anxiety behaviors as measured by an elevated plus-maze and open- field test (Itoi, 2008).
30 Noradrenaline neurons are clustered into hindbrain nuclei in the central
nervous system. Historically, seven noradrenergic nuclei (A1 - A7) have been identified in rats (Dahlström and Fuxe, 1964), with most of these nuclei also found in primates and humans (Bogerts, 1981; Felten and Sladek, 1983; Jacobowitz and
MacLean, 1978). These nuclei can be subdivided into caudal, central, and rostral
groups (Szabadi, 2013). The locus coeruleus (LC; A6), the only member in the
rostral group, is the largest of all the noradrenergic nuclei and it is located
bilaterally on the corner of the fourth ventricle. The LC contains almost 50% of all
central noradrenergic neurons, ~20-50,000 neurons in humans and ~1,500
neurons in rats (Mouton et al., 1994; Singewald and Philippu, 1998). Noradrenergic
projections are primarily organized into three pathways: the ascending, cerebellar,
and descending pathway (Szabadi, 2013). The ascending pathway innervates the
midbrain, thalamus, amygdala, and the neocortex; the cerebellar pathways
innervates the cerebellar nuclei and cortex; and the descending pathway projects to
motor nuclei in the lower brainstem to descend down the spinal cord (Szabadi,
2013) (Figure 1.1). Reciprocal connections between the locus coeruleus and the
basolateral amygdala have been implicated in the stress response (Buffalari and
Grace, 2007; Galvez et al., 1996; Hatfield et al., 1999) and noradrenergic innervation
of corticotrophin-releasing factor neurons in the paraventricular nucleus modulate
the hypothalamic-pituitary-adrenal (HPA) axis to influence the stress response (Itoi
and Sugimoto, 2010).
Interestingly, noradrenergic projection fibers contain varicosities, or non-
terminal enlargements that have the structure of pre-synaptic terminal boutons and
31 are capable of neurotransmitter release. These structures were initially observed in
the cortex, amygdala, and hypothalamus (Beaudet and Descarries, 1978; Farb et al.,
2010; Marotte and Raisman, 1974) and release from these varicosities is now
thought to be a general feature of terminal noradrenaline release (Chiti and
Teschemacher, 2007; Kasparov and Teschemacher, 2008). Noradrenergic axon
terminals can contact post-synaptic neurons or end in the extracellular space in the
absence of apposed membranes (Szabadi, 2013). Together, these observations
suggest that noradrenaline release is likely to have a diffuse, non-synaptic effect in
terminal regions.
In rodents, the locus coeruleus is comprised entirely of noradrenergic
neurons (Singewald and Philippu, 1998). Noradrenergic neuron dendrites primarily
extend laterally from the LC into two pericoerulear zones: the potine tegmentum
medial rostral to the LC and the caudal juxtaependymal periocoerulear region
adjacent to the 4th ventricle (Shipley et al., 1996). These dendrites connect extensively via gap junctions, which induces synchrony in LC neuron activity
(Ishimatsu and Williams, 1996). Electophysiologically, noradrenaline neurons in the LC have α2-receptor mediated noradrenaline conductances, large cell bodies (>
40 pF capacitance), and tonic pacemaker firing of 1 - 5 Hz (Williams et al., 1984).
Major inputs to the locus coeruleus originate from the neocortex, amygdala,
hypothalamus, ventral tegmental area, dorsal raphe nucleus, medulla, and sensory
neurons in the spinal cord (Luppi et al., 1995; Szabadi, 2013). Noradrenaline is also
released within the LC, originating either from projection terminals from other
noradrenergic nuclei (Singewald and Philippu, 1998) including the contralateral LC
32 (Cedarbaum and Aghajanian, 1978) or locally from somatodendritic (Huang et al.,
2007, 2012) and axon collateral sites (Aghajanian et al., 1977).
GI-coupled α2-autoreceptors are the only adrenergic receptors expressed by
noradrenergic neurons in the LC. Noradrenaline release within the LC is thought to
activate these receptors to regulate the tonic firing rates of LC noradrenergic
neurons especially during different modes of attention and arousal (Aston-Jones and
Cohen, 2005a; Aston-Jones et al., 1999). Additionally, these autoreceptors are thought to enable firing rates to be reset to baseline levels by generating a pause in firing following a phasic burst of LC activity (Aston-Jones and Cohen, 2005a; Aston-
Jones et al., 1999).
Electron microscopy studies aiming to understand the underlying mechanisms of noradrenergic feedback inhibition have examined the ultrastructure location of post-synaptic α2-autoreceptors in the LC. While these receptors clearly localize to the dendrites of TH+ noradrenaline neurons, dendritic receptor sites were rarely observed to be apposed by TH+ noradrenergic pre-synaptic membrane
(Lee et al., 1998c). Furthermore, when dendrites did receive contacts from TH+ pre- synaptic membranes, noradrenergic receptors were located on non-junctional regions of the post-synaptic membrane outside of synaptic densities (Lee et al.,
1998b, 1998c). Finally, these receptors could be found portions of the plasma membrane not associated with any synapses (Lee et al., 1998b). Taken together, electron microscopy studies indicate a lack of distinct synaptic structures by which noradrenergic transmission might occur.
33 Central serotonin system
The activity of serotonin neurons in the dorsal raphe nucleus is associated
with sleep/wake cycles, mode, stress response, and reward encoding (Adell et al.,
2002; Inaba et al., 2013; Maier et al., 2006; McDevitt et al., 2014). Central serotonin
neurons are clustered within seven raphe nuclei located in the midbrain, pons, and
medulla (Carlsson et al., 1962; Dahlström and Fuxe, 1964; Palkovits et al., 1974;
Vasudeva et al., 2011). The largest of these nuclei is the dorsal raphe nucleus (DRN;
B6 and B7), which is located on the midline of the brainstem just ventral of the
cerebral aqueduct. The dorsal raphe is the primary source of serotonergic
innervation in the forebrain. Projection targets of the DRN serotonergic neurons
(Figure 1.1) are topographically organized. Neurons located rostrally within the
DRN project to the caudate putamen, substantia nigra, and neocortical regions
(Abrams et al., 2004; Imai et al., 1986; Vertes, 1991). In contrast, neurons located
caudally within the DRN project to the septum, hippocampus, and entorhinal cortex
(Abrams et al., 2004; Köhler and Steinbusch, 1982; Vertes, 1991). While these rostral-caudal projection fields do not overlap for given serotonergic neurons (Imai et al., 1986), branched axons arising from a single neuron can innervate multiple, functionally related brain regions (Lowry, 2002).
In rodents, only ~50% of the neurons present in the DRN are serotonergic
(Vasudeva et al., 2011). In addition to having rostral-caudal topography, serotonin neurons within the dorsal raphe are typically divided into four anatomical groups: dorsomedial, ventromedial, and two bilateral wings (Abrams et al., 2004; Agnati et al., 1982; Steinbusch et al., 1981). In addition to serotonin neurons, neurons that
34 display markers for dopamine, GABA, and glutamate synthesis or vesicular
packaging are also present in DRN. Dopamine neurons in the DRN are sometimes
referred to as a dorso-caudal extension of the VTA. These neurons project to the
neostriatum, lateral septum, and frontal cortex (Descarries et al., 1986; Stratford
and Wirtshafter, 1990) and are thought to be involved in the rewarding properties
of opiates (Dougalis et al., 2012; Flores et al., 2004; Meyer et al., 2009). Glutamate
neurons in the DRN are thought to be involved in reward encoding by projection
onto dopamine neurons located in the VTA (McDevitt et al., 2014; Qi et al., 2014).
Additionally, it is thought that GABA and 5-HT markers do not co-express in DRN neurons suggesting a lack of GABA/serotonin co-release (Day et al., 2004; Stamp and Semba, 1995). However, co-release of glutamate and serotonin does occur in a small number of medial DRN neurons as demonstrated by the co-expression of markers for the synthesis or vesicular packaging of glutamate and serotonin
(Commons, 2009; Hioki et al., 2010). Despite earlier criteria using firing rates and action potential width (Aghajanian and Vandermaelen, 1982), recent studies have shown that DRN serotonin neurons cannot be positively identified on the basis of electrophysiology alone (Marinelli et al., 2004). Instead, TPH, SERT, or the differentiation factor PET1 (Scott et al., 2005) can be used to genetically label serotonergic DRN neurons.
The major inputs to the DRN originate from the lateral and medial habenula, locus coeruleus, superior vestibular nucleus, substantia nigra, hypothalamus, nucleus of the solitary tract, and the prefrontal cortex (Jacobs and Azmitia, 1992;
Vasudeva et al., 2011). Noradrenergic axon inputs from the locus coeruleus act on
35 α1-receptors to provide the major excitatory drive underlying rhythmic DRN serotonin neuron firing (0.5 – 3 Hz) (Baraban and Aghajanian, 1981). Similar to
DRN projections, inputs to the DRN also display some topographical organization
(Peyron et al., 1998). Serotonin is released locally within the DRN from somatic
(Colgan et al., 2009), dendritic (Colgan et al., 2012), and axon terminals originating both locally and from other raphe nuclei (Bruns et al., 2000) to provide feedback control over DRN activity.
Local serotonin release within the dorsal raphe nuclei can be encoded by three types of serotonin receptors: 5-HT1A, 5-HT1B, and 5-HT2 receptors (Marinelli et
al., 2004). While 5-HT1B receptors are located exclusively on axon terminals (Sari et
al., 1999) and 5-HT2 receptors primarily locate to DRN GABAergic neurons
(Marinelli et al., 2004), serotonin neurons in the DRN mainly express 5-HT1A
autoreceptors (Marinelli et al., 2004). These 5-HT1A-autoreceptors are thought to
primarily modulate the firing rates of serotonin neurons in response to ambient
levels of serotonin released within the DRN (Haddjeri et al., 2004; Hajos et al., 2001;
Judge and Gartside, 2006). However, evidence of transient bursts and pauses during
reward encoding suggests that these receptors might also act on a more phasic time
scale (Cohen et al., 2015). To assess the behavioral relevance of 5-HT1A autoreceptor mediated transmission, mice were generated with either an over- or under-expression of 5-HT receptors only in serotonin neurons. Compared to under-
expression, increased autoreceptor expression was associated with a blunted
response to stress, increased behavioral despair, and a lack of a response to
36 antidepressants (Richardson-Jones et al., 2010) demonstrating a behavioral function
of serotonin feedback in the DRN.
Ultrastructurally, 5-HT1A receptors are located on the somas and dendrites of
DRN serotonin neurons (Kia et al., 1996; Riad et al., 2000). Whether or not these sites display a distinct, synaptic architecture remains unclear. Within the DRN, markers of post-synaptic transmission sites were found to lack any associated pre- synaptic membranes (Kia et al., 1996; Riad et al., 2000). However, in other brain regions these same markers were found within defined synapses (Kia et al., 1996).
Due to a lack of serial quantitative electron microscopy, it is difficult to say
definitively that these synaptic localizations do not also occur within the DRN.
Mechanisms of neurotransmission
Many insights about synaptic function and regulation were first described in
fast, ionotropic systems in peripheral neuromuscular junctions and central
synapses. Thus, this section will begin by examining the mechanisms that impact
transmission mediated by ionotropic receptors before moving on to discuss general
mechanisms of metabotropic signaling.
Mechanisms of chemical synaptic transmission were first described at the
junction between peripheral nerves and muscle fibers. At these neuromuscular
junctions, acetylcholine released from the pre-synaptic nerve acts on ionotropic
nicotinic receptors located post-synaptically on the muscle fiber directly opposed to
the nerve terminal, a region of the muscle dubbed the motor endplate, to initiate a
37 muscle contraction. The clustering of nicotinic receptors into motor endplates was
first observed by using the iontophoretic application of acetylcholine to map post-
synaptic sensitivity to acetylcholine at the micron level (Kuffler and Yoshikami,
1975). These experiments found that the strongest post-synaptic response to
acetylcholine application always occurred at the motor endplate directly under
nerve terminals and that the sensitivity of the post-synaptic membrane to
acetylcholine dropped off near 50-fold merely 2 µm away from these synaptic
endplate regions (Kuffler and Yoshikami, 1975). This, along with the close proximity of nerve terminals and motor endplates and the rapid degradation of acetylcholine by acetylcholinesterase in the extracellular space, resulted in acetylcholine released from a pre-synaptic nerve terminal acting only on directly apposed nicotinic receptors on the motor endplate. Thus, communication by nicotinic receptors at the neuromuscular junction occurs by linear, point-to-point transmission via independent synapses between paired nerve terminals and motor endplates.
Until the mid to late 1990s, fast ionotropic neurotransmission in the central nervous system was also thought to occur only via independent synapses. As such, transmitter released from pre-synaptic sites was thought to exclusively act on immediately apposed post-synaptic receptors forming direct pathways of communication (Barbour and Hausser, 1997; Huang, 1998). However, beginning in the mid 1990s a series of papers began demonstrating that neurotransmitters could escape from synaptic clefts at some synapses to activate more distal receptors not immediately apposed to release sites which were located on either pre- or post-
38 synaptic membranes (Kullmann et al., 1996; Min et al., 1998a; Rossi and Hamann,
1998). This long-range form of transmission was termed spillover because of the
necessity for transmitter to ‘spill’ out of the synaptic cleft (Huang, 1998). In some
cases, spillover transmission was sufficient to activate receptors located at
neighboring synapses, resulting in crosstalk between synaptic sites and a
corresponding loss of synaptic independence (Rusakov and Kullmann, 1998a).
While completely independent synapses maximize the computational and storage
capacity of a neuronal network, crosstalk between functionally related synapses
improves the reliability of transmission and provides an efficient means of setting
background activity levels and synchronizing entire networks (Barbour and
Hausser, 1997; Isaacson, 1999). Thus, both forms of transmission could be
advantageous for a given neuronal network depending on the structure and the
information it is intended to convey.
Point-to-point activation of synaptic receptors
Classical, point-to-point neurotransmission is mediated by post-synaptic receptors that reside immediately apposed to release sites in the post-synaptic density. Release sites and receptors are directly apposed at the ultrastructure level and separated only by a synaptic cleft which is ~20 nm in width and ~2 attoliters in volume (Bergles et al., 1999). Receptors mediating classical synaptic transmission typically have low affinities and include AMPA, GABAA, nACh and NMDA receptors.
Partially because of their low affinities, these receptors generally respond to
39 transmitter released from the associated pre-synaptic membrane resulting in point- to-point transmission.
In the case of synaptic glutamate transmission, studies using competitive antagonists have determined the concentration and time course of glutamate in the synaptic cleft that interacts with post-synaptic receptors. In the synaptic cleft, glutamate concentration peaks around 1.1 mM and rapidly decays with a time constant of 1.2 milliseconds (Clements et al., 1992). Because of this rapid clearance of glutamate from the synaptic cleft, the time course of transmission at these synapses is governed by channel kinetics (Jahr and Lester, 1992). For some synaptic receptors, these channel kinetics are determined by the relative rates of transmitter-receptor dissociation. For example, slower dissociation rates for glutamate from NMDA receptors can result in a longer time course of transmission
(Lester et al., 1990). Alternatively, high agonist concentrations found in a synaptic cleft can result in receptor desensitization that significantly curtails the duration of signaling such as is the case with AMPA receptors (Jones and Westbrook, 1996).
Glutamate clearance from the synaptic cleft is primarily achieved by diffusion. However, the driving force for the rapid diffusion of glutamate out of the synaptic cleft requires low ambient concentrations of glutamate. This low ambient glutamate concentration is maintained by glutamate reuptake transporters located on both neurons and glial cells. As a result, acutely inhibiting glutamate reuptake transporters has no effect on the time course of AMPA or NMDA mediated transmission at many synapses (Hestrin et al., 1990; Isaacson and Nicoll, 1993;
Sarantis et al., 1993). However, inhibiting reuptake transporters at some glutamate
40 synapses prolongs a slow phase of signaling that presumably results from the
entrapment of glutamate by a specialized synaptic cleft, necessitating clearance by
transporters (Barbour et al., 1994; Kinney et al., 1997; Otis et al., 1996).
Spillover activation of extrasynaptic receptors
The effective concentration of neurotransmitter falls drastically as it leaves
the synaptic cleft and diffuses in all directions (Barbour and Hausser, 1997).
Reflecting this fact, the extrasynaptic receptors that respond to transmitter spillover
typically have higher affinities for neurotransmitters compared to synaptic
receptors (Huang, 1998). Activation of extrasynaptic receptors is inherently
dependent on the extracellular concentration profile of neurotransmitter; thus,
clearance mechanisms such as the rate of diffusion (Barbour and Hausser, 1997)
and reuptake (Asztely et al., 1997; Carter and Regehr, 2000; Diamond, 2001;
Wadiche and Jahr, 2005) can strongly govern or even prevent the activity of
extrasynaptic receptors. Consistent with these functional studies, ultrastructural
studies of spillover-activate metabotropic GABAB receptors find that these receptors
are primarily located either just outside of closely-apposed synaptic contacts
(perisynaptic) or in regions of the plasma membrane completely unassociated with
apposed synaptic structures (Charara et al., 2005). Furthermore, these GABAB
receptors are not observable within closely apposed synaptic structures unlike
point-to-point, synaptic GABAA receptors (Charara et al., 2005). Because these
receptors are unopposed by pre-synaptic GABA release site, they require extended
GABA diffusion if they are to be activated.
41 Whether or not transmitter spillover from a single release event is sufficient to activate extrasynaptic receptors varies across synapses. In many cases, transmitter release from a single synapse is often insufficient to activate extrasynaptic receptors. Instead, multiple release events generated by stimulation trains or the synchronous activation of multiple release sites are necessary to elicit a response from extrasynaptic receptors (Isaacson et al., 1993; Scanziani, 2000). The transmitter released during these stimulation paradigms presumably pools in the extracellular space summating to reach the concentrations necessary to activate extrasynaptic receptors. As a result, both the reliability and kinetics of spillover transmission resulting in transmitter pooling are dependent on the number of active release sites and the probability of vesicular release (Balakrishnan et al., 2009;
Carter and Regehr, 2000).
Mechanisms of monoamine feedback inhibition
Feedback inhibition encoded by monoaminergic autoreceptors is thought to occur via an extended form of spillover and obligate pooling that is termed ‘volume transmission.’ As a consequence of volume transmission, monoamine release is thought to contribute to a general receptor tone that tonically inhibits neurons based solely on extrasynaptic concentrations rather than any synaptic associations.
While volume transmission has been predicted for noradrenergic autoreceptor- mediated transmission in the LC, dopaminergic autoreceptor-mediated transmission in the VTA, and serotoninergic autoreceptor-mediated transmission in
42 the DRN, the evidence supporting this hypothesis is indirect and heavily relies on
modeling predictions rather than tested hypotheses.
Monoamine-mediated transmission is typically studied using electrochemical measurements of transmitter release. Dopamine, noradrenaline, and serotonin all undergo redox chemical reactions at specific voltage ranges. This chemical property can be leveraged to measure the concentrations of monoamine in the bulk
extracellular space using either fast-scan cyclic voltammetry (FSCV) or constant
potential amperometry (Ewing et al., 1982; Jackson et al., 1995). In both techniques,
a large carbon-fiber electrode placed either into brain slices or in vivo detects the
average monoamine concentration along its entire surface area (Kelly and
Wightman, 1987; Kuhr and Wightman, 1986). This results in relatively poor spatial
resolution for monoamine concentrations that typically cannot discriminate
individual release sites or events in brain slices or in vivo. In generally, FSCV has
significantly better signal to noise ratios while amperometry has improved temporal precision.
Using electrochemistry, multiple studies have demonstrated that electrically evoking release generates extracellular transients of dopamine in the midbrain, noradrenaline in the LC, and serotonin in the DRN that can last for many seconds
(O’Connor and Kruk, 1991; Palij and Stamford, 1994; Rice et al., 1997). These evoked transients reach a peak concentration of ~20-100 nM, which is relatively similar to reported EC50 concentrations of monoamine autoreceptors (Cragg and Rice, 2004;
Jennings, 2013). Diffusional modeling that utilizes both voltammetry data and
receptor EC50 values predicts that the vesicular release of dopamine, noradrenaline,
43 and serotonin can all act on receptors that are located several microns from release
sites and continually activate receptors for many seconds following release (Cragg
and Rice, 2004; Jennings, 2013). These results, in combination with the
extrasynaptic localization of autoreceptors suggested by electron microscopy
studies, provide the primary evidence supporting the volume transmission
hypothesis. However, D2-autoreceptors more likely exist in low-affinity states
under physiological conditions that are orders of magnitude less sensitive to
dopamine than the high-affinity states used in modeling (Ford, 2014; Schoffelmeer
et al., 1994; Sibley et al., 1983) and the rapidly-changing neurotransmitter concentrations seen by receptors during synaptic transmission significantly complicates the application of steady-state measurements such as EC50 values.
Furthermore, ultrastructural studies in the dopamine system have also reported
synaptically localized D2-autoreceptors (Sesack et al., 1994) suggesting that
transmission may also be occurring over shorter distances.
In the mid 2000s, studies began using GIRK channel mediated currents to
examine the activity of midbrain D2-autoreceptors (Beckstead et al., 2004) (Figure
1.2). These studies found that evoked dopamine release only transiently activated
D2-autoreceptors to produce GIRK currents that rose and fell within 1-2 seconds
(Beckstead et al., 2004) and poorly correlated to extracellular dopamine transients measured by voltammetry (Ford et al., 2009). These currents could only be replicated by applying greater than 10 µM dopamine to receptors (Ford et al., 2009), which is significantly higher than both bulk dopamine concentrations measured by electrochemistry and reported EC50 values. Furthermore, slowing diffusion with the
44 macromolecule dextran (Min et al., 1998b) had no effect on evoked D2-receptor mediated GIRK currents, suggesting that dopamine spillover did not activate D2-
autoreceptors (Ford et al., 2010). Together, these studies suggest that the volume
transmission hypothesis may not fully explain midbrain dopamine feedback
transmission. This methodology had not yet been systematically applied to either
the serotonergic or noradrenergic system to address questions of spillover, pooling,
or general mechanisms of transmission.
Rationale
Volume transmission has been proposed as the key mechanisms of autoreceptor activation for the monoamines dopamine, noradrenaline, and serotonin. However in each system, the case supporting volume transmission relies on indirect measurements of transmission. While electrochemical methods do measure extracellular transients of monoamines for seconds following evoked release, receptor activation has only be inferred based on affinity measurements and diffusion modeling. Importantly, these key affinity measurements are often
EC50 values that are measured at steady state and may not prove insightful in the
rapidly changing environment at sites of synaptic transmission.
Recent advances using GIRK-channel currents as a readout of
monoaminergic receptor activation offer a unique opportunity to directly test the
volume transmission hypothesis. By applying techniques developed to study
transmitter spillover and pooling at ionotropic synapses to GPCR-mediated GIRK
45 currents, this thesis aims to evaluate the contributions of spillover, pooling, and
transmitter clearance in shaping the timing of transmission. Knowing the
mechanisms governing the spatial and temporal limitations of signaling is necessary
to understand the role of autoreceptor-mediated feedback in shaping global monoamine signaling and monoamine-influenced behaviors.
Statement of the problem
This thesis begins with the hypothesis that dopamine-mediated transmission in the mouse midbrain may occur via independent, point-to-point synaptic transmission. This hypothesis will be tested by recording dopamine-mediated GIRK currents in response to evoked transmitter release in midbrain dopamine neurons.
Because the methodology for evaluating spillover and pooling were developed for fast, ionotropic receptor-mediated transmission, noradrenaline-mediated GIRK currents in the locus coeruleus and serotonin-mediated GIRK currents in the dorsal raphe will be recorded and tested with respected to the volume transmission hypothesis. Comparing the transmission mechanisms underlying various monoamines provides some validation of the application of ionotropic methodology to metabotropic-receptor mediated currents. Perhaps more importantly, this comparison also addresses the question of whether neuromodulation by monoamines is governed by similar transmission mechanisms.
46 Figure 1.1: Projection targets of monoamine nuclei regions. A. Cartoon diagram of the major axonal projection targets of dopamine neurons located in the midbrain ventral tegmental area (VTA) and the substantia nigra pars compacta (SNc). B.
Cartoon diagram of the major axonal projection targets of noradrenaline neurons located in the hindbrain locus coeruleus (LC). B. Cartoon diagram of the major axonal projection targets of serotonin neurons located in the hindbrain dorsal raphe nucleus (DRN). Labels for all panels are as follows: Am: amygdala; CB: cerebellum;
CP: caudate putamen; HC: hippocampus; OB: olfactory bulb; PFC: pre-frontal cortex;
Pir: piriform cortex; Sp: septum; SNr: substantia nigra pars reticulate; St: striatum;
Th: thalamus
47
Figure 1.1
48 Figure 1.2: Autoreceptor mediate inhibition in monoamine neurons. Left:
Within nuclei regions, inhibitory transmission between monoamine neurons
regulates their activity. Right: GI-coupled autoreceptors located on the somas and dendrites of monoamine neurons respond to locally released monoamines by activating G-protein coupled, inwardly recertifying potassium channels (GIRKs) via
βγ signaling. The resulting efflux of potassium hyperpolarizes monoamine neurons to inhibit their activity. In the locus coeruleus, local noradrenaline release activates
α2-auotreceptors located on noradrenergic neurons. In the ventral tegmental area, local dopamine release activates D2-autoreceptors located on dopaminergic neurons. In the dorsal raphe nucleus, local serotonin release activates 5-HT1A-
autoreceptors located on serotonergic neurons.
49
Figure 1.2
50
Chapter 2
Species differences in somatodendritic
dopamine transmission determine D2-
autoreceptor mediated inhibition of ventral
tegmental area neuron firing.
Courtney NA, Mamaligas AA, and Ford CP (2012) Species Difference in
Somatodendritic Dopamine Transmission Determine D2-Autoreceptor-Mediated
Inhibition of Ventral Tegmental Area Neuron Firing. J Neurosci 32: 13520-13528.
51 FOREWORD
This chapter was published with myself as a co-first author. For this publication, I personally conducted all of the electrochemical experiments and contributed to the analysis, figure construction, and writing of the entire work.
52 ABSTRACT
The somatodendritic release of dopamine within the ventral tegmental area
(VTA) and substantia nigra pars compacta (SNc) activates inhibitory post-synaptic
D2-receptors on dopaminergic neurons. The proposed mechanisms that regulate
this form of transmission differ between electrochemical studies using rats and
guinea pigs and electrophysiological studies using mice. This study examines the
release and resulting dopamine D2-autoreceptor mediated inhibitory post-synaptic
currents (D2-IPSCs) in the VTA of mouse, rat and guinea pig. Robust D2-IPSCs were observed in all recordings from neurons in slices taken from mouse, whereas in rat and guinea pig D2-IPSCs were observed less frequently and were significantly smaller in amplitude. In slices taken from guinea pig, dopamine release was more persistent under conditions of reduced extracellular calcium. The decline in the concentration of dopamine was also prolonged and not as sensitive to inhibition of reuptake by cocaine. This resulted in an increased duration of D2-IPSCs in the guinea pig. Therefore, unlike the mouse or the rat, the time course of dopamine in the extracellular space of the guinea pig determined the duration the D2-IPSC.
Functionally, differences in D2-IPSCs resulted in inhibition of dopamine neuron firing only in slices from mouse. The results suggest that the mechanisms and functional consequences of somatodendritic dopamine transmission in the VTA vary among species. This highlights the complexity that underlies dopamine dependent transmission in one brain area. Differences in somatodendritic transmission would be expected in vivo to affect the downstream activity of the mesocorticolimbic dopamine system and subsequent terminal release.
53
INTRODUCTION
Midbrain dopamine neurons play an essential role in goal-orientated motor and cognitive functions that include planning and initiating movements, and encoding the associations between rewards and salient environmental stimuli
(Schultz, 2007; Wise, 2004). Disruptions in the mesocorticolimbic dopamine system may contribute to psychiatric disorders such as schizophrenia, attention-deficit and hyperactivity disorder (ADHD) and drug addiction (Madras et al., 2005; Meisenzahl et al., 2007; Volkow et al., 2009).
Dopamine release in terminal regions is facilitated in vivo during bursts of activity that are driven by the appearance of unexpected rewards or by cues predicting rewards (Bromberg-Martin and Hikosaka, 2009; Phillips et al., 2003;
Tobler et al., 2005). In addition to axonal release, dopamine neuron firing also drives the local release of dopamine from somatodendritic terminals in the midbrain (Beckstead et al., 2004; Cheramy et al., 1981; Geffen et al., 1976; Groves and Linder, 1983; Jaffe et al., 1998; Nirenberg et al., 1996; Rice et al., 1994; Wilson et al., 1977). Somatodendritic dopamine release in both the VTA and SNc activates D2- autoreceptors on post-synaptic dopamine neurons leading to an inhibition via the activation of G-protein coupled inwardly rectifying potassium (GIRK) channels
(Lacey et al., 1987). Local release of dopamine in the VTA is therefore a form of feedback or lateral inhibition that can regulate dopamine neuron activity.
Somatodendritic release has often been examined using electrochemical or dialysis techniques to measure the bulk concentration of dopamine released into the
54 extracellular space. Using these techniques in the rat and guinea pig,
somatodendritic release has been found to differ from axonal release in several
ways, including a prolonged duration lasting several seconds and a weak
dependence upon extracellular calcium entry (Chen and Rice, 2001; Chen et al.,
2011; Cragg and Greenfield, 1997; Hoffman and Gerhardt, 1999; Rice et al., 1997). In
contrast, when measured using electrophysiology in the mouse, somatodendritic
transmission in the VTA is short in duration and exhibits a steep dependence upon
calcium entry (Beckstead et al., 2004; Ford et al., 2007, 2010). Differences in release
could lead to differential activation of post-synaptic D2-receptors during
somatodendritic transmission. Because D2-autoreceptors regulate dopaminergic excitability (Beckstead et al., 2004) and terminal release (Benoit-Marand et al.,
2001; Phillips et al., 2002; Schmitz et al., 2002), alterations in the strength of somatodendritic transmission may have functional consequences for downstream dopamine neuron activity. Underlying this fact is recent work showing that D2- autoreceptor dysfunctions disrupt several dopamine-dependent behaviors that include locomotor activity, impulsivity and motivation for rewards in both mice and humans (Bello et al., 2011; Buckholtz et al., 2010).
To examine how differences in somatodendritic release regulate VTA neuron activity, the present study examined dopamine transmission in brain slices from the mouse, rat and guinea pig. Using a combination of electrochemical and electrophysiological techniques, the results indicate that release across species varied dramatically in calcium dependence, strength and time course. The functional
55 consequences of these differences were apparent in the ability of phasic
somatodendritic release to regulate dopamine neuron firing in the VTA.
METHODS
Slice preparation and solutions. All procedures were in accordance with
CWRU IACUC guidelines. Following anesthesia, horizontal midbrain slices
containing the VTA (220 m) or coronal forebrain slices containing the dorsal
striatum (240 m) were madeμ from 3-5 week old male and female C57Bl6 mice
(Jackson Laboratories),μ Sprague Dawley rats (Charles River) or Hartley guinea pigs
(Charles River). Brain slices were cut in ice-cold sucrose cutting solution that
contained 75 mM NaCl, 2.5 mM KCl, 6 mM MgCl2, 0.1 mM CaCl2, 1.2 mM NaH2PO4, 25
mM NaHC03, 2.5 mM D-glucose, 50 mM sucrose; bubbled with 95 % O2/ 5% CO2.
Slices were incubated post-cutting at 35°C in oxygenated 95% O2/ 5% CO2 ACSF
solution that contained 126 mM NaCl, 2.5 mM KCl, 1.2 mM MgCl2, 2.5 mM CaCl2, 1.2
mM NaH2PO4, 21.4 mM NaHCO3, 11.1 mM D-glucose for 45 minutes before
recording. During incubation, 10 M MK-801 was included to reduce excitotoxicity and increase slice viability. Followingμ incubation, slices were placed in a recording chamber and constantly perfused with warm ACSF (34 ± 2°C) containing 100 M
picrotoxin, 10 M DNQX and 200 nM CGP 55845 at 2 ml/min. For experimentsμ testing the calciumμ dependence of dopamine release, the reduced Ca2+ was substituted for an equal molar amount of Mg2+ in order to maintain a constant
divalent cation concentration (3.7 mM). Dopamine neurons were visualized with a
BXWI51 microscope (Olympus) with infrared custom-built gradient contrast optics.
56 The VTA was defined as the area within and medial to the fibers of the medial
lemniscus (Ford et al., 2006, 2007). As dopamine neurons projecting to the
prefrontal cortex are insensitive to the inhibitory actions of dopamine due to a low
expression of D2-receptors and GIRK channels (Chiodo et al., 1984; Lammel et al.,
2008), we avoided recording from dopamine neurons in the most medial portions of
the VTA.
Electrophysiology. Whole-cell current-clamp or voltage-clamp recordings were made from VTA dopamine neurons using an Axopatch 200B amplifier
(Molecular Devices). Patch pipettes (1.5-
(World Precision Instruments). The intracellular2 MΩ) were pipette pulled solution from borosilicatecontained 115 glass mM
K-methylsulphate, 20 mM NaCl, 1.5 mM MgCl2, 10 mM HEPES(K), 10 mM BAPTA-
tetrapotassium, 1mg/ml ATP, 0.1 mg/ml GTP, and 1.5 mg/ml phosphocreatine (pH
7.4, 275 mOsm). Data was acquired using an ITC-18 interface (Instrutech) and
Axograph X (Axograph Scientific) at 10 KHz and filtered to 2KHz for voltage-clamp
recordings. For voltage-clamp experiments, neurons were held at -60 mV. Series
resistance was not compensated and cells were discarded if the access resistance
exceeded 15
membrane potentialMΩ. Action set withpotentials constant were current recorded injection in current to ensure clamp tonic mode pacemaker at a
firing. VTA dopamine neurons were identified by the presence of a D2-receptor
sensitive dopamine conductance, pacemaker firing (0.5 - 5 Hz), presence of an h-
current (Ih
(Chieng et ),al., an 2011; input Ford resistance et al., 2006) of < 300. To MΩ pharmacologically and an input capacitance isolate evoked > 25 pFD2 -
receptor mediated GIRK currents, 100 M picrotoxin, 10 M DNQX and 200 nM CGP
μ μ
57 55845 were added to the ACSF solution. The high concentration (10 mM) of BAPTA
in the pipette solution and MK-801 in the incubation solution were used to block
mGluR signaling and NMDA receptors. The D2-receptor antagonist sulpiride (200 nM) was used to confirm that dopamine currents were mediated through the D2- receptor.
Dopamine release was elicited with an ACSF-filled extracellular glass mono- polar stimulating electrode placed ~50 - 100 m from the neuron recorded. A train of stimuli (5 pulses, 0.6 ms, 40 Hz, 40-60 A) wasμ used to evoke release. All drugs were applied by bath perfusion except dopamine,μ which was applied by iontophoresis. Dopamine (1 M) was ejected as a cation (160 nA) for 1s using an Ion-
100 Iontophoresis Generator (Dagan) from thin walled iontophoretic electrodes placed ~10 m from the cell. A retention current of 3-20 nA was applied to prevent the leakage μof dopamine.
Electrochemistry. Fast scan cyclic voltammetry (FSCV) recordings were made with glass encased carbon fiber electrodes as previously described (Ford et al.,
2009) using custom-built hardware (Electronics and Materials Engineering Shop,
University of Washington, Seattle, WA) and software (Tarheel CV, Labview). Carbon fibers (34-700; 7 m diameter; Goodfellow) were cut to a final length of 50 – 100
m. Once cut, the μtip was placed in isopropanol with activated carbon for 10 min priorμ to use. The exposed tip of the electrode was placed in the VTA ~50 m below the surface of the slice. A triangular waveform from -0.4 V to +1.3 V versusμ Ag/AgCl at changing voltage of 400V/s at 10 Hz was used for all recordings. Between scans, the electrode was maintained at -0.4 V (versus Ag/AgCl). Background subtracted
58 cyclic voltammogram currents were obtained by subtracting the average of 10 cyclic
voltammograms obtained before stimulation. The time course of dopamine release
was obtained by plotting the current at the peak of oxidation (0.6 to 0.7 V) against
time. The chemical identity of the FSCV recordings was examined by comparing
cyclic voltammograms produced from the evoked release of dopamine to
voltammograms produced by the exogenous iontophoretic application of dopamine
hydrochloride (1M) or serotonin hydrochloride (0.5 M). The peak reduction values
of evoked voltammograms (- 0.2 to - 0.3 V) were similar across VTA of mice, rats
and guinea pigs (p > 0.7, one way ANOVA) and were similar to the peak reduction
value produced by exogenous dopamine (p > 0.5, one way ANOVA) but not
serotonin (p < 0.0001, one way ANOVA) indicating that serotonin release did not
contribute significantly to FSCV signals in the VTA. Currents were calibrated against
dopamine standards ranging from 0.1 to 10 M.
As serotonergic terminals are also presentμ in the SNc and the release of
serotonin contributes to the FSCV signal detected in the SNc (Cragg and Greenfield,
1997; Ford et al., 2010; John et al., 2006), only the VTA was examined in the present
study. This prevented the release of serotonin from axon terminals from interfering
with FSCV measurements of dopamine release.
Chemicals. Picrotoxin, DNQX, and MK-801 were from Ascent Scientific.
CGP55845 and S-(-)-sulpiride were from Tocris Bioscience. K-methylsulphate was
from Acros Organics. BAPTA was from Invitrogen. Cocaine hydrochloride was from
the National Institute of Drug Abuse. All other chemicals were from Sigma-Aldrich.
59 Statistics and analysis. All data are shown as mean ± standard error (SEM).
Statistical significance (p < 0.05) was assessed by a one-way ANOVA or a Students
unpaired t-test unless as noted (InStat 3.0, Graphpad). If ANOVA showed statistical
significance (p < 0.05), all pairwise post-hoc analysis stated was performed using a
Tukey post hoc test. Decay kinetics of D2-IPSCs and extracellular dopamine
transients were fit with a single or double exponential using a Simplex algorithm
optimized by the sum of squared errors in Axograph X (Axograph Scientific).
RESULTS
The kinetics of dopamine release and transmission in the VTA
Carbon fiber electrodes were used with fast-scan cyclic voltammetry (FSCV)
to compare the kinetics and the amount of dopamine released in VTA brain slices
from the mouse, rat and guinea pig. A train of 5 stimuli (40 Hz) evoked a rapid
increase in the concentration of dopamine in the extracellular space ([DA]o). Nearly
twice as much dopamine could be detected within the VTA of the mouse (145 ± 15
nM, n = 11) and guinea pig (153 ± 18 nM, n = 11) as in the VTA of the rat (86 ± 8 nM,
n = 10, p < 0.05 versus mouse; p < 0.01 versus rat) (Fig. 1A). Despite the range in
[DA]o that was evoked, the rise time and time to peak of the dopamine transients
were similar among rats, mice and guinea pigs (time to peak mouse: 536 ± 74 ms, n
= 11; rat: 570 ± 73, n = 10; guinea pig: 545 ± 67 ms, n = 11; p > 0.9, one-way ANOVA)
(Fig. 1A). Following release, however, the [DA]o remained elevated for longer in the
guinea pig VTA than in the mouse or rat. The [DA]o had a decay of 2.3 ± 0.2 s (n = 11) in the guinea pig VTA, which was slower than in the VTA ofτ the rat (1.3 ± 0.2 s, n =
60 10) or mouse (1.4 ± 0.2 s, n = 11; p < 0.001) (Fig. 1B). Thus, despite a similarity in
the rate of release, the amount and duration over which dopamine was present in
the extracellular space varied between species with the concentration of dopamine
being elevated for the longest in the VTA of the guinea pig.
The extended extracellular presence of dopamine has often been implicated
to indicate that dopamine is not constrained within the synapse following release.
This has led to the hypothesis that spillover of dopamine or volume transmission
may play a major role in shaping the time course of somatodendritic signaling.
However, this may not be the case in the VTA of the mouse as the time course of the
dopamine transient underlying somatodendritic transmission may be brief (Ford et
al., 2009). As the amount and duration of dopamine released into the extracellular
space varied between species, we next examined what effect this difference in [DA]o had on synaptic transmission. Whole-cell recordings were made from VTA dopamine neurons in brain-slices from mice, rats and guinea pigs. Dopamine D2- receptor mediated inhibitory post-synaptic currents (D2-IPSCs) were evoked by stimulating somatodendritic dopamine release (train of 5 pulses at 40 Hz) from adjacent presynaptic dopamine neurons with an extracellular stimulating electrode placed in the VTA. Only dopamine neurons in the central and lateral portions of the
VTA were recorded as these neurons do not project to the prefrontal cortex and are known to exhibit autoreceptor mediated inhibitory responses (Ford et al., 2006;
Lammel et al., 2008). Robust D2-IPSCs were observed in all experiments from mouse VTA slices (n = 38). D2-IPSCs in slices from rat and guinea pig in contrast, were smaller and observed less frequently (Fig. 1D). The average amplitude of D2-
61 IPSCs in mouse dopamine cells was 115 ± 9 pA (n = 38) (Fig. 1 D). The amplitude of
the synaptic currents in rat VTA neurons was less than a fifth of that seen in the
mouse (24 ± 2 pA, n = 26, p < 0.001). Surprisingly, the high [DA]o that could be
detected in the guinea pig VTA did not reliably produce D2-IPSCs. IPSCs were observed in only 23 out of 48 neurons (Fig. 1D), despite the exogenous application of dopamine evoking a D2-receptor mediated outward current in all cells examined.
Of the D2-IPSCs that were recorded, the amplitude was also small relative to those in the mouse (24 ± 3 pA, n = 23, p < 0.001) (Fig. 1 C, D). Thus between species, the
[DA]o detected by FSCV did not correlate with the amplitude of D2-IPSCs.
Despite the variation in amplitude, the time course of D2-IPSCs was remarkably similar between the rat and mouse (Fig 1E). In agreement with previous results, D2-IPSCs from the mouse VTA peaked in 362 ± 7 ms (n = 38) (Fig 1E)(Ford et al., 2009). D2-IPSCs from the rat had similar rise times to those in mice (time to
peak: 349 ± 9 ms, n = 26, p > 0.2) (Fig 1E). The time constant of decay was also
similar between D2-IPSCs from rat and mouse VTA cells (mouse: decay = 327 ± 14
ms, n = 37; rat: decay = 396 ± 21 ms, n = 26; p > 0.05 one-way ANOVA)τ (Fig 1F).
These results indicateτ that while large differences occur in the amount of dopamine released and the amplitude of D2-IPSCs between rats and mice, the mechanisms controlling the time course of transmission are similar; in both species the duration of D2-receptor activation appears to be tightly regulated. The reduction in both the
[DA]o and amplitude of D2-IPSCs in the rat compared to the mouse may result from
a difference in the number of release sites and/or points of synaptic contact in the
rat.
62 D2-IPSCs in guinea pig VTA dopamine neurons were slower. The time to peak
was 547 ± 22 ms (n = 23; p < 0.001) (Fig. 1E). Likewise the decay of D2-IPSCs was slower in dopamine cells from the guinea pig VTA than rat orτ mouse (889 ± 73 ms; n
= 22; p < 0.0001). Thus when scaled to their peak amplitude, D2-IPSCs from guinea pig VTA neurons did not overlap with those in rat and mouse dopamine cells (Fig.
1E). These results suggest dopamine is more loosely regulated within guinea pig
VTA somatodendritic synapses than rat and mouse. These observations are consistent with the idea that the extent of transmitter spillover or volume transmission varies widely at different somatodendritic synapses among species.
The difference in time course in D2-IPSCs may result from post-synaptic differences between guinea pigs and rats and mice. This possibility may be unlikely however as the dopamine transient measured electrochemically was also prolonged in the guinea pig VTA relative to rats and mice (Fig 1A, B).
Calcium dependence of dopamine release in the VTA and Striatum
Somatodendritic dopamine release in the guinea pig persists under conditions where extracellular calcium is reduced (Chen and Rice, 2001; Chen et al.,
2011; Rice et al., 1997). As the dopamine transmission in the mouse is inhibited in low extracellular calcium (Ford et al., 2010), we next asked if lowering calcium would similarly affect somatodendritic release in the guinea pig, rat and mouse.
Dopamine release in both the mouse and rat VTA was strongly inhibited in 0.5 mM extracellular calcium. Lowering the extracellular calcium concentration ([Ca2+]o)
from control values (2.5 mM) to 0.5 mM led to an inhibition of 91 ± 4% (n = 7) and
63 94 ± 2% (n = 11) of [DA]o and D2-IPSCs respectively in the mouse VTA (Fig 2A, D).
The inhibition was similar to that observed in the mouse SNc (% inhibition D2-IPSC
: 91 ± 2%, n = 7; p > 0.2; data not shown). Lowering [Ca2+]o to 0.5 mM led to a similar
reduction of 90 ± 3% (n = 10) and 94 ± 2% (n = 11) of [DA]o and D2-IPSCs in the rat
VTA (Fig 2B, D). Thus, dopamine release in the rat and mouse midbrain exhibits a strong dependence on extracellular calcium.
In the guinea pig VTA, a significant amount of extracellular dopamine remained when calcium levels were lowered (Fig 2C, D). Lowering [Ca2+]o to 0.5 mM
led to a reduction of only 70 ± 4% (n = 8) in the [DA]o. D2-IPSCs in the VTA were also less sensitive to lowering the [Ca2+]o, as in the presence of 0.5 mM [Ca2+]o, D2-
IPSCs were reduced by 83 ± 3% of control levels (n = 9) (p < 0.01) (Fig 2C, D). These results support previous findings that somatodendritic dopamine release in the guinea pig shows weak dependence on extracellular calcium (Chen et al., 2011). The
calcium dependence of somatodendritic release of dopamine therefore also varies
between guinea pigs and other species.
Recurrent axon collaterals are also present within the VTA (Bayer and Pickel,
1990; Deutch et al., 1988). These terminals may also contribute to dopamine release
in the VTA (Chen et al., 2011). We next compared release in the VTA to the dorsal
striatum where release is purely axonal. Lowering [Ca2+]o to 0.5 mM reduced the
amount of dopamine detected in the striatum by 95 ± 1% (n = 5, Fig. 2D) in the
mouse and likewise by 98 ± 4% (n = 5, Fig 2D) in the rat, indicating that in the rat
and mouse, dopamine exhibits a strong dependence on [Ca2+]o at both axonal and
dendritic release sites. In the guinea pig striatum, dopamine release was also
64 strongly reduced in 0.5 mM [Ca2+]o (96 ± 4%, n = 5, Fig 2D). Thus, the weak dependence of dopamine release on extracellular calcium entry was selective for only somatodendritic terminals of the guinea pig.
Dopamine D2-autoreceptor currents vary among species
The variation in D2-IPSCs among species and the lack of correlation with the measured [DA]o may reflect differences in D2-receptor/GIRK channel density, D2- receptor signaling efficiency, and/or the concentration of dopamine at post-synaptic
D2-receptors. To examine these possibilities, we first examined D2-receptor mediated outward potassium currents evoked by the exogenous iontophoretic application of dopamine. Application of a saturating concentration of dopamine by iontophoresis evoked a maximal outward current of 309 ± 26 pA in mouse VTA cells
(n = 22), which was greater than the outward currents observed in rat (221 ± 18 pA
, n = 18) or guinea pig dopamine neurons (143 ± 12, n = 46; p < 0.05) (Fig 3A).
Likewise, a similar difference among species was present when cell size was taken into account by normalizing the outward currents to the input capacitance (mouse:
7 ± 0.6 pA/pF, n = 22; rat: 5.3 ± 0.4 pA/pF, n = 18; guinea pig: 3.7 ± 0.3 pA/pF, n =
46; p < 0.05, Fig 3B). Thus, D2-receptor mediated GIRK-mediated currents are largest in mouse VTA dopamine cells and smaller in rat and guinea pig neurons. As the density of D2-receptors in the midbrain is greater in the mouse than the rat than the guinea pig (Camps et al., 1990), this difference may likely be due to different levels of receptor expression among species (Werkman et al., 2011).
65 Dopamine transporter regulation of extracellular dopamine in the VTA
Dopamine transporters regulate the amplitude and duration of D2-IPSCs following evoked release. Blocking transporters enhances spillover and prolongs the time course of transmission (Beckstead et al., 2004; Christie and Jahr, 2006; Ford et al., 2009; Isaacson et al., 1993). As the duration of [DA]o and D2-IPSCs were longer
in the guinea pig VTA than the VTA of rats or mice (Fig 1A, C) we next asked if
transport function differed within the VTA. Blocking dopamine reuptake with the
monoamine transport blocker cocaine (1 M) led to a similar increase in the amount and duration of [DA]o within the VTA of allμ species. Cocaine (1 M) increased the
peak [DA]o detected by 225 ± 19% (n = 6) in the mouse, 227 ± 32%μ in the rat (n = 6)
and by 171 ± 8% (n = 4) in the guinea pig (p > 0.2; one way ANOVA). Blocking
reuptake, however, increased the amplitude of D2-IPSCs in the rat and mouse VTA
more than in the guinea pig VTA. Cocaine (1 M) enhanced the peak of the D2-IPSC in the mouse and rat by 264 ± 31% (n = 7) andμ 336 ± 29% (n = 8) respectively, which was greater than the increase in the D2-IPSCs in guinea pig (167 ± 9%; n = 7; p < 0.05) (Fig 4 A, B). The smaller effect on D2-IPSCs from the guinea pig suggests that dopamine transporters are less effective in regulating the time course of dopamine at somatodendritic synapses in guinea pig VTA dopamine neurons than in the rat or mouse. These observations indicate that D2-autoreceptor mediated transmission varies in both strength and transporter-dependent regulation among species.
66 Extent of somatodendritic dopamine transmission correlates with evoked
pauses in pacemaker firing
We next asked what functional consequences somatodendritic dopamine transmission would have on the firing patterns of dopamine neurons. Evoked somatodendritic release was hypothesized to have the greatest effect on mouse
dopaminergic activity and less of an effect on firing in guinea pig or rat neurons.
Initially, the predicted change in voltage in VTA dopamine neurons was calculated
based upon the observed amplitude of D2-IPSCs from all cells recorded (including
failures) and the species-specific cellular input resistance of VTA dopamine neurons
(VD2-IPSP = ID2-IPSC × Rinput). The estimated hyperpolarization in mouse VTA cells was
~14.8 mV (115 pA * 129 MΩ), which was greater than that predicted for rat or
guinea pig VTA neurons (rat: ~2.1 mV = 21 pA * 101 MΩ; guinea pig: ~0.7 mV = 11
pA * 68To MΩ). test whether the D2-IPSC could regulate the firing patterns of VTA neurons, whole cell current clamp recordings were made from dopamine cells in the mouse, rat and guinea pig. Dopamine neurons typically fire in vitro in a tonic pacemaker pattern at ~1-5 Hz. A train of stimuli, similar to that used to evoke D2-
IPSCs (5 pulses at 40 Hz) evoked large hyperpolarizations in mouse VTA dopamine neurons, sufficient to induce a pause in pacemaker firing (Fig 5A). The hyperpolarization-induced pause lasted 1.5 ± 0.1 s (n = 18), which was similar to duration of D2-IPSCs recorded in voltage-clamp (Fig 5A). The pause in pacemaker firing was blocked by bath application of the D2-receptor antagonist sulpiride (200 nM) (0.5 ± 0.1 s; n = 13) (Fig 5A). When normalized to the baseline pacemaker firing
67 frequency of each neuron, stimulation led to a 364 ± 23% inhibition in firing frequency (n = 18). Sulpiride (200 nM) reduced the pause in firing to 134 ± 11% of control levels (n = 13; p < 0.0001). In contrast, trains of stimuli failed to evoke large dopamine-mediated hyperpolarizations in dopamine cells from rat or guinea pig
VTA slices (Fig 5 B, C). Thus the stimulation induced change in firing rate was not different in rat or guinea pig VTA dopamine neurons between control slices and slices treated with sulpiride (200 nM) (rat control: 164 ± 16%, n = 19 sulpiride: 145
± 11%, n = 12; guinea pig control: 164 ± 22%, n = 18, sulpiride: 177 ± 25%, n = 16%; p > 0.7; one-way ANOVA) (Fig 5C). Together these data suggest that the extent of somatodendritic transmission determines the role that D2-autoreceptors play in regulating the phasic actions of dopamine on VTA pacemaker firing.
DISCUSSION
The results show that the mechanisms controlling release, time course and strength of somatodendritic dopamine transmission vary widely across the VTA of different species. While much of the past work examining somatodendritic release has been done in rats and guinea pigs (Chen and Rice, 2001; Chen et al., 2011; Cragg and Greenfield, 1997; Hoffman and Gerhardt, 1999; Rice et al., 1997), the physiological actions of dopamine as a synaptic transmitter have been examined only in mice (Beckstead et al., 2004; Bello et al., 2011; Ford et al., 2009, 2010). In contrast to the guinea pig, somatodendritic dopamine transmission in rats and mice exhibited a strong dependence on extracellular calcium. The time course of dopamine within somatodendritic synapses was also more strongly regulated by
68 dopamine transporters in rats and mice than guinea pigs. This limited dopamine spillover and prevented prolonged activation of post-synaptic D2-receptors. Thus somatodendritic dopamine transmission in the guinea pig varied from other species in time course and the dependence of release on extracellular calcium entry. This work suggests that species-specific differences may be one source accounting for the differences in the mechanisms underlying somatodendritic release and transmission reported in previous publications.
Calcium dependence of dopamine release in the VTA
Dopamine release from dendritic terminals has often been reported to persist in low extracellular calcium, leading to the conclusion that somatodendritic release exhibits a weak dependence on calcium entry not seen at conventional CNS synapses (Chen and Rice, 2001; Chen et al., 2011; Fortin et al., 2006; Hoffman and
Gerhardt, 1999). Here we found that in the rat and the mouse, both axonal and dendritic dopamine release showed an equally strong dependence on extracellular
incalcium. the VTA Lowering and dorsal calcium striatum. to 0.5 The mM VTA led containsto a ≥ 90% both reduction axon collaterals in dopamine and release somatodendritic terminals (Bayer and Pickel, 1990; Deutch et al., 1988) and the release of dopamine from both sets of terminals can contribute to the total detected increase in [DA]o with FSCV (Chen et al., 2011; Ford et al., 2010). However, as D2-
IPSCs from VTA and SNc dopamine cells are similar in amplitude, kinetics, duration and pharmacological regulation (Beckstead et al., 2004; Ford et al., 2007, 2010), the physiological activation of D2-receptors during phasic release may occur primarily
69 at somatodendritic synapses. Like the reduction in the [DA]o, D2-IPSCs in the VTA
were also similarly reduced by > 90% in the presence of 0.5 mM calcium in rats and
mice. Thus, whether measured electrochemically or electrophysiologically,
somatodendritic dopamine release in both the rat and mouse shows a strong
dependence on calcium entry from the extracellular space.
Dopamine transmission in the VTA of the guinea pig however showed a
marked difference in regulation. As previously reported, lowering extracellular
calcium (0.5 mM) did not inhibit DA release to the same extent in the VTA compared
to the striatum (Chen et al., 2011). In low calcium ~30% of total dopamine release
remained in the VTA in 0.5 mM extracellular calcium. Likewise, D2-IPSCs in VTA dopamine neurons from the guinea pig were also less sensitive to reductions in extracellular calcium. As dopamine neurons of the guinea pig do not express many of the vesicular fusion release proteins commonly expressed at fast synapses
(Witkovsky et al., 2009), the results suggest that dopamine transmission in the guinea pig may occur via a different set of mechanisms than at other central synapses.
Time course of transmission
The duration of synaptic transmission is dependent upon both the concentration time course of neurotransmitter and the kinetics of the neurotransmitter-receptor interactions (Barbour et al., 1994; Katz and Miledi, 1973;
Lester et al., 1990). Many G-protein coupled receptors (GPCRs) that mediate slow metabotropic transmission, such as GABAB receptors, are located at extrasynaptic
70 sites and are activated by transmitter spillover away from the synaptic cleft
(Beenhakker and Huguenard, 2010; Isaacson et al., 1993; Kulik et al., 2002; Otis and
Mody, 1992; Scanziani, 2000). The long lasting presence of dopamine that can be detected in the extracellular space of rats and guinea pigs has often thus been interpreted to indicate that dopamine transmission also occurs by a paracrine spillover type mechanism (volume transmission). In the mouse VTA, somatodendritic dopamine transmission is thought to be mediated by a high concentration of dopamine that activates a pool of receptors located within 1 m from the site of release (Ford et al., 2009). Somatodendritic dopamine transmissionμ may therefore be more localized than previously assumed.
Dopamine D2-IPSCs were significantly larger in the mouse than in the rat.
The iontophoresis of exogenous dopamine evoked smaller maximal outward currents in rat dopamine cells, suggesting that D2-receptor/GIRK channel density, number of release sites and/or the number of points of synaptic contact may be reduced in the rat compared to the mouse. However despite the smaller amplitude, the kinetics and duration of synaptic transmission was similar. Because the concentration of dopamine determines the rate of activation of the GIRK conductance (Ford et al., 2009), the similarity in the rate of rise of D2-IPSCs and time to peak of synaptic currents suggests that during transmission D2- autoreceptors are exposed to similar concentrations of dopamine in both the rat and mouse.
Dopamine D2-IPSCs were significantly slower in the guinea pig. In guinea pig dopamine cells, IPSCs had a slower rise time, time to peak and decay kinetics than
71 either the rat or mouse. As blocking dopamine reuptake with cocaine had a limited
effect on the amplitude of D2-IPSCs in the guinea pig, the reuptake of dopamine in
the VTA of the guinea pig may be less efficient. The slow kinetics of D2-IPSCs in the
guinea pig most likely results from this decreased rate of dopamine uptake. The
effect of this in the guinea pig would be an increase in dopamine spillover to
extrasynaptic sites and a prolonged activation of post-synaptic receptors.
Functional Implications
The functional consequence of D2-IPSCs in dopamine cells from the VTA of
the mouse was a pause in pacemaker firing. In rats and guinea pigs electrical
stimulation often led to a slowing of firing. However, the effect was not altered in the
presence of the D2-recptor antagonist, sulpiride. Multiple metabotropic
neurotransmitter receptors including GABAB, mGluR, and 1-noradrenergic
receptors inhibit dopamine cells (Fiorillo and Williams, 1998;α Morikawa and
Paladini, 2011; Morikawa et al., 2003; Paladini and Tepper, 1999; Paladini and
Williams, 2004). The pause was unlikely to be mediated by GABAB and mGluR
receptors as receptor signaling was prevented by antagonists (CGP55845) or
blockers of the receptor-effector signaling cascade (BAPTA), suggesting that
activation or inactivation of other conductances may have been engaged as a result
of the train of stimuli. This species difference in ability of D2-autoreceptors to induce a pause in firing may result from a higher level of expression of D2-receptors in the mouse than either the rat or guinea pig (Camps et al., 1990; Werkman et al.,
2011). While lower coupling efficiency or GIRK channel expression in rats or guinea
72 pigs may also have contributed to the observed differences, this possibility may be
less likely, as robust GABAB mediated synaptic potentials that couple via GIRK
channels have been reported previously in guinea pig dopamine neurons (Bonci and
Williams, 1996; Cameron and Williams, 1993).
The activation of D2-autoreceptors has long been known to regulate
dopaminergic neuron firing (Aghajanian and Bunney, 1977). The present results
indicate that strong activation of these receptors during robust somatodendritic
synaptic events can be an efficient form of lateral inhibition that regulates the firing
patterns of VTA dopamine neurons. The species dependent variations in the
strength of D2-IPSCs that we observed would therefore be expected to differentially
control the firing pattern of dopamine neurons in vivo and subsequently the
forebrain terminal release of dopamine during behaviorally relevant tasks. As
multiple mechanisms can underlie dopamine transmission, subtle alterations in any
of these mechanisms would be expected to have significant effects on the efficiency
of transmission. Recent work in humans has shown that reduced midbrain D2-
autoreceptor levels correlated with highly impulsive individuals (Buckholtz et al.,
2010). Disruptions in somatodendritic transmission may therefore lead to widespread dysfunctions in this system at the behavioral and clinical level.
Differences in this system highlight the need to consider species-specific aspects of
dopamine signaling in the future when assessing the pre-clinical efficacy of
pharmacological treatments for a variety of psychiatric diseases.
73 Figure 2.1: Inter-species characterization of dopamine release and D2-IPSCs in the VTA. (A) FSCV measurements of the average evoked [DA]o by a train of stimuli
(5 pulses, 0.5 ms, 40 Hz) in the mouse (black, n = 11), rat (blue, n = 10) and guinea pig (red, n = 11). Stimulation evoked less dopamine release in the rat than the guinea pig or mouse. (B) Summary data illustrating that the decay time constants for the [DA]o was longer in the guinea pig than the rat or mouse VTA. (C)
Representative traces of D2-IPSCs evoked by a train of stimuli (5 pulses, 0.5 ms, 40
Hz) in the mouse (black), rat (blue) and guinea pig (red). Synaptic currents were larger in amplitude in the mouse than guinea pig or rat VTA. (D) Summary distribution of D2-IPSC amplitude of all neurons recorded in the VTA of the mouse, rat and guinea pig. Numbers in parenthesis refer to the proportion of neurons recorded in the VTA of each species that expressed a measurable D2-IPSC. Large filled circle illustrates the average synaptic currents of responding neurons. (E)
Normalized D2-IPSCs illustrated in previous panel. D2-IPSCs have been scaled to the peak of their amplitude. D2-IPSCs from the mouse and rat exhibited similar kinetics while D2-IPSCs in the guinea pig had a slower rise time, time to peak and time of decay. (F) Summary data illustrating the slower time constant of decay of D2-IPSCs from the guinea pig (n = 23) than the mouse (n = 37) or rat (n = 26). Asterisk: ** = p
< 0.01; *** = p < 0.001.
74
Figure 2.1
75 Figure 2.2: Dopamine release in the VTA shows a weaker dependence on extracellular calcium in guinea pigs than rats or mice. (A-C) Averaged [DA]o (n =
7 - 10) and representative traces of D2-IPSCs from the mouse, rat and guinea pig
under control conditions (2.5 mM [Ca2+]o; black) and low calcium (0.5 mM [Ca2+]o;
grey). (D) Summary data illustrating the [DA]o remaining in 0.5 mM [Ca2+]o in the
VTA and striatum and the amplitude of D2-IPSCs in the VTA in 0.5mM Ca2+as
compared to control (2.5 mM [Ca2+]o). Asterisk: ** = p < 0.01; *** = p < 0.001
76
Figure 2.2
77 Figure 2.3: Dopamine mediated D2-autoreceptor currents are smallest in guinea pig VTA dopamine neurons. (A) Representative traces of outward currents evoked by the exogenous iontophoretic application of dopamine (160 nA, 1 s). (B)
Current density (pA / pF) showing that a maximal application of dopamine evoked the largest currents in the mouse and the smallest currents in the guinea pig.
Asterisk: * = p < 0.05; *** = p < 0.001
78
Figure 2.3
79 Figure 2.4: Dopamine uptake in the VTA regulates D2-IPSCs to a greater extent in mice and rats than guinea pigs. (A) Representative traces illustrating evoked
D2-IPSCs under control conditions (black) and in the presence of cocaine (1 M, grey) from VTA dopamine neurons in mice, rats and guinea pigs. Note the greaterμ increase in amplitude of D2-IPSCs recorded from rats and mice than guinea pigs in the presence of cocaine. D2-IPSCs under control conditions have been sized to the same initial amplitude in order to illustrate the relative increase induced by blocking re-uptake. (B) Summarized data illustrating the increase in amplitude of
D2-IPSCs in the presence of cocaine (1 M). Asterisk: * = p < 0.05; *** = p < 0.001
μ
80
Figure 2.4
81 Figure 2.5: Evoking D2-IPSCs does not induce a pause in the firing of VTA dopamine neurons from the rat or guinea pig. (A) Voltage clamp recording of a
D2-IPSC (top trace) from a mouse VTA dopamine neuron and current clamp recordings of tonic pacemaker firing of the same neuron (lower traces). The time course of the evoked D2-IPSCs was similar to that of the evoked hyperpolarization and the pause in firing when recorded in current clamp. The D2-receptor antagonist, sulpiride (200 nM) blocks the evoked pause in firing suggesting that D2-receptor activation during phasic release underlies the pause in firing. (B – C) Current-clamp recordings of tonic pacemaker firing of rat and guinea pig VTA dopamine neurons under control conditions and in the presence of sulpiride (200 nM). Stimulation does not induce an extended pause in firing. Sulpiride does not change the firing rate following stimulation suggesting that D2-receptor activation by evoked release in the rat and guinea pig is not sufficient to induce a phasic pause in firing. (D)
Summary data of the above conditions. Data is illustrated as the time from the start of stimulation to the first action potential following the stimulation as a percent of the average interspike interval recorded from each cell during 3s of baseline tonic firing. Asterisk: *** = p < 0.001
82
Figure 2.5
83 Chapter 3
The timing of dopamine- and noradrenaline-
mediated transmission reflects underlying
differences in the extent of spillover and
pooling.
Courtney NA and Ford CP (2014) The Timing of Dopamine- and Noradrenaline-
Mediated Transmission Reflects Differences in the Extent of Spillover and Pooling. J
Neurosci 34: 7645-7656.
84 ABSTRACT
Metabotropic transmission occurs at many sites through the spillover activation of extrasynaptic receptors. This study examined the mechanisms underlying somatodendritic dopamine and noradrenaline transmission and found that the extent of transmitter spillover and pooling varied dramatically between these two synapses. In the mouse ventral tegmental area, the time course of D2- receptor mediated inhibitory post-synaptic currents (D2-IPSCs) was consistent between cells and unaffected by altering stimulation intensity, probability of release or the extent of diffusion. Blocking dopamine reuptake with cocaine extended the time course of D2-IPSCs and suggested that transporters strongly limited spillover.
As a result, individual release sites contributed independently to the duration of D2-
IPSCs. In contrast, increasing the release of noradrenaline in the rat locus coeruleus prolonged the duration of 2-receptor mediated IPSCs even when reuptake was intact. Spillover and subsequentα pooling of noradrenaline activated distal α2- receptors which prolonged the duration of 2-IPSCs when multiple release sites were synchronously activated. By using theα rapid application of agonists onto large macro patches, we determined the concentration profile of agonists underlying the two IPSCs. Incorporating the results into a model simulating extracellular diffusion predicted that the functional range of noradrenaline diffusion was nearly five-fold greater in the locus coeruleus than dopamine in the midbrain. This study demonstrates that catecholamine synapses differentially regulate the extent of spillover and pooling to control the timing of local inhibition and suggests a
85 diversity in the roles of uptake and diffusion in governing metabotropic
transmission.
INTRODUCTION
Synaptic potentials mediated by G-protein coupled receptors (GPCRs)
typically occur through the activation of extrasynaptic receptors by transmitter
spillover from nearby synapses (Beenhakker and Huguenard, 2010; Isaacson et al.,
1993; Kulik et al., 2002; Otis and Mody, 1992; Scanziani, 2000). These slow events
differ from fast ligand-gated ion channel mediated events in which a high
concentration of transmitter rapidly rises and falls within the synapse (Barbour et
al., 1994; Beato, 2008; Clements et al., 1992). As GPCRs are primarily located
outside the synapse (Sesack et al., 1994), reuptake by transporters tightly regulates
the spatial range of receptor activation (Beenhakker and Huguenard, 2010; Isaacson
et al., 1993; Scanziani, 2000). Bursts of stimulation are therefore often required to
overcome uptake in order to facilitate transmitter pooling and evoke GPCR-
mediated synaptic events (Isaacson et al., 1993; Otis and Mody, 1992; Scanziani,
2000).
Transmission mediated by monoamines like dopamine and noradrenaline is classically thought to occur via an extended form of spillover known as volume transmission (Agnati et al., 2010; Zoli et al., 1998). Catecholamine transients measured electrochemically in the extracellular space can be detected for several seconds following release (Cragg and Rice, 2004; Garris et al., 1994; Kawagoe et al.,
1992; Palij and Stamford, 1994; Rice and Cragg, 2008). Modeling studies have
86 predicted that these low concentration transients may diffuse many microns away
from the site of release before activating distant extrasynaptic receptors (Agnati et
al., 2010; Cragg and Rice, 2004; Dreyer et al., 2010; Rice and Cragg, 2008). However,
functional studies in the ventral tegmental area (VTA) have found that at
somatodendritic synapses, saturating concentrations of dopamine are required to
evoke D2-receptor mediated IPSCs (Ford et al., 2009). It therefore remains unclear
to what extent the prolonged diffusion of catecholamines within the extracellular
space contributes to the timing of transmission.
Within the locus coeruleus (LC), somatodendritic noradrenaline release
evokes α2-receptor mediated IPSCs through an analogous G-protein inwardly rectifying potassium (GIRK) conductance (Egan et al., 1983) to that which mediates somatodendritic dopamine D2-IPSCs in the VTA (Beckstead et al., 2004; Cheramy et al., 1981; Geffen et al., 1976; Jaffe et al., 1998; Lacey et al., 1987; Palij and Stamford,
1994; Rice et al., 1994). The present work compares dopamine and noradrenaline transmission mediated by autoreceptors in the VTA and LC in brain slices. The results show that diffusion and reuptake of catecholamines play different roles at these two synapses. Whereas the kinetics of transmission mediated by noradrenaline is dependent on both diffusion and reuptake, dopamine-dependent transmission is regulated primarily by reuptake. This difference maintains independence between dopamine synapses in the VTA, yet allows for pooling of noradrenaline in the extracellular space of the LC that prolongs transmission when multiple terminals are synchronously active. These results show that by regulating
87 the extent of spillover, transporters control the timing and the extent of cross talk
that occurs between dopamine synapses.
METHODS
Slice Preparation and visualization: All procedures were approved and performed in accordance with CWRU IACUC guidelines. Horizontal brain slices were obtained from deeply anesthetized 3-6 week old male and female Sprague
Dawley rats (Charles River) containing the locus coeruleus (240 µM) or 3-6 week
old male and female C57BL6 mice (Jackson Laboratories) containing the VTA (220
µM). Brain slices were cut using a Vibratome (Leica) in ice-cold cutting solution
containing (in mM): 75 NaCl, 2.5 KCl, 6 MgCl2, 0.1 CaCl2, 1.2 NaH2PO4, 25 NaHCO3,
2.5 D-glucose and 50 sucrose and continuously bubbled with 95% O2 and 5% CO2.
Slices were transferred to artificial cerebral-spinal fluid (ACSF) containing (in mM):
126 NaCl, 2.5 KCl, 1.2 MgCl2, 2.5 CaCl2, 1.2 NaH2PO4, 21.4 NaHCO3, and 11.1 D- glucose bubbled with 95% O2 and 5% CO2 and incubated at 35°C for at least 45
minutes prior to use. MK-801 (10 µM) was included during the incubation to block
NMDA receptors. Following incubation, slices were transferred to a recording
chamber and constantly perfused at 2 ml/min with oxygenated ACSF warmed to 34
± 2°C. Slices were visualized with a BX51WI microscope (Olympus) with custom-
built infrared gradient contrast optics.
Electrophysiology: Whole-cell voltage clamp recordings were made using an
Axopatch 200B amplifier (Molecular Devices). Patch pipettes (2.0-
2.5 MΩ) were
88 pulled from borosilicate glass (World Precision Instruments). The pipette internal
solution contained (in mM): 115 K-methylsulphate, 20 NaCl, 1.5 MgCl2, 10 K-HEPES,
10 BAPTA-tetrapotassium, 2 ATP, 0.3 GTP, and 6 sodium phosphocreatine; pH 7.4;
275 mOsm. Data were acquired using an ITC-18 interface (Instrutech) and Axograph
X (Axograph Scientific) at 10 kHz and filtered to 2 kHz. Series resistance was not
compensated and recordings were discarded if the access resistance rose above 15
2 was
substitutedMΩ. In the caseto maintain of experiments a constant with divalent reduced ion extracellular concentration. calcium, All drugs, MgCl except
where otherwise noted, were applied by bath perfusion. The VTA was defined as
the area within and medial to the fibers of the medial lemniscus (Ford et al., 2006).
Dopamine neurons within the VTA were identified by the presence of a dopamine
D2 receptor-mediated dopamine conductance, a capacitance of 25 - 40 pF, an input resistance < 300 M -current, and a tonic firing rate of 0.5 – 2 Hz as previous described (Ford et Ω,al., an 2006) h . The locus coeruleus was identified by its location between the corner of the 4th ventricle and the mesencephalic tract motor neurons.
Noradrenergic neurons within the LC were identified by having an α2-sensitive
andnoradrenaline a tonic firing conductance, rate of 0.5 –a 2capacitance Hz. > 40 pF, an input resistance < 100 MΩ,
Catecholamine release was elicited with an extracellular, ACSF-filled
lices containing the VTA, the stimulating
electrodemonopolar was glass placed electrode 50-100 (~5 µm MΩ). away In from s the neuron being recorded (or the tip of
the carbon fiber where applicable). In slices containing the LC, the stimulating
electrode was placed into the caudal side of the LC. Either single pulses (0.5 ms) or
89 trains of stimuli (0.5 ms at 40 Hz) were used to drive catecholamine release (~40-
50 µA unless otherwise noted). D2- and α2-receptors were pharmacologically
isolated with DNQX (10 µM), MK801 (10 µM), picrotoxin (100 µM) and CGP55845
(200 nM) to block AMPA, NMDA, GABAA, and GABAB receptors. BAPTA (10 mM) in
the internal pipette solution was used to block intracellular Ca2+ signaling and
associated mGluR responses. For experiments in the LC in which the rising kinetics
were analyzed, the stimulation artifact was removed by antagonist subtraction
(idazoxan, 2 M). The effect of cocaine on transmission was verified to be due to inhibition of μreuptake by comparisons to a DAT-specific antagonist (GBR-12909, 1
M) and a NET-specific antagonist (nisoxetine, 1 M).
μ Electrochemistry: Fast-scan cyclic voltammetryμ (FSCV) recordings were made using glass-encased carbon fiber electrodes (34-700, Goodfellow) with an exposed diameter of 7 µM and an exposed length of 50-100 µm. Before use, the exposed carbon fiber tip was soaked in isopropanol purified with activated carbon for at least 20 minutes. The tip of the fiber was placed directly into either the LC or VTA
~50 M below the surface of the slice. While holding the fiber at -0.4 V, either triangularμ waveforms (-0.4 to 1.3 V vs. Ag/AgCl at 400 V/S) or sinusoidal waveforms
(-0.4 to 1.0 V vs. Ag/AgCl over 10 ms) were applied at 10 Hz. Background subtracted cyclic voltammogram currents were obtained by subtracting the average of 10 voltammograms obtained prior to stimulation from each voltammogram obtained after stimulation. The time course of dopamine and noradrenaline were determined by plotting their peak oxidation potentials vs. time. After
90 experimentation, each electrode was calibrated against known concentrations of
either dopamine or noradrenaline.
To confirm the chemical identify of the signal, we compared the
voltammogram of evoked transmitter release to voltammograms obtained by
applying serotonin, dopamine, and noradrenaline directly onto the carbon fiber via
iontophoresis. The single-stimulation evoked signal had a peak reduction at – 257
or -223 mV in the LC or VTA, respectively, which was consistent with the
voltammograms of noradrenaline and dopamine. Serotonin did not contribute to
either evoked signal as evidenced by the lack of characteristic twin reduction peaks
(Rice et al., 1997). Inhibiting dopamine reuptake with the DAT inhibitor GBR12909
(1 M) had no effect on evoked catecholamine signal in the LC (p > 0.4, data not shown,μ n = 5, Student’s paired t-test) confirming that dopamine likely does not contribute to the evoked signal in the LC (Palij and Stamford, 1994).
Isolated macro patches: Large nucleated outside-out macro patches were obtained from either dopamine cells from the VTA or noradrenaline cells from the
LC, as previously described (Ford et al., 2009). Patches were pulled out of the slice and into the free solution above the tissue. In some experiments, the patches were lowered onto the surface of the slice to attempt to detect the evoked release of noradrenaline or dopamine. For these experiments, mono-polar electrode stimulation was used, similar for evoking IPSCs. Other experiments used fast flowing solutions through a theta tube flow pipe to rapidly apply known concentrations of dopamine or noradrenaline. Macro patches were raised ~500 m above the surface of the slice and placed in front of the theta tube. Once in place, μ
91 solution was allowed to flow through both barrels of the flow pipette at a rate of
~20 l per min. The bath volume was adjusted such that 1 cm of the flow pipette
was immersedμ in the bath allowing the temperature of the flow solution to reach
equilibrium with the temperature of the bath (34 ± 2°C). One barrel of the flow
pipette perfused a control solution containing (in mM): 121 NaCl, 2.5 KCl, 1.2 MgCl2,
2.4 CaCl2, 1.4 NaH2PO4, 0.6 ascorbate-NA, and 5 HEPES-Na and pH 7.3, while a similar solution with the addition of a known concentration of dopamine or noradrenaline was perfused through the other barrel. The flow pipette was attached to a piezoelectric bimorph. Passing 25 V across the bimorph changed the location of the flow pipette tip such that the solution bathing the macro patch was rapidly exchanged. To determine the kinetics of this exchange, open tip junction potentials were measured with the flow pipette perfusing solutions with different concentrations of KCl. The average 10-90% exchange time, as measured by changes in junction potential, was 2 ± 1 milliseconds.
Point-Release Diffusion Modeling: The spatiotemporal profile of transmitter
diffusing away from a single vesicle of release was approximated using a point
diffusion model (Cragg and Rice, 2004; Cragg et al., 2001; Rice and Cragg, 2008). In
brief, we solved for the concentration (C) of dopamine or noradrenaline as a
function of radius (r) from the release site and time following release (t) using the
equation:
( , ) = 𝑄𝑄 ( ), 2 ( 𝑁𝑁𝑎𝑎 ) −𝑟𝑟 3 ∗ ∗ 2 4𝐷𝐷 𝑡𝑡 𝐶𝐶 𝑟𝑟 𝑡𝑡 𝛼𝛼 4𝐷𝐷 𝑡𝑡𝑡𝑡 𝐸𝐸𝐸𝐸𝐸𝐸
92 where Q is the quantal content of a vesicle in molecules, Na is Avogadro’s number,
is the volume fraction of the tissue, and D* is the tissue corrected diffusion constantα
2
tortuosityequal to D/λ of thewhere tissue. D is For the the diffusion diffusion constant of dopamine of the neurotransmitter in the VTA, we used and the λ is the
following values obtained from the literature (Cragg et al., 2001):
Q = 14,000 (Jaffe et al., 1998), and D = 7.63 x 10-6 cm2 sec-1. For theα diffusion= 0.30, λ =of 1.62,
noradrenaline in the LC, we used: Q = 37,000 (Huang et al., 2007), D = 7.63 x 10-6
cm2 sec-1 (adapted from: (Bennett et al., 2004). The values of were used in the LC. α = 0.30 and λ = 1.58
Statistics and Data Analysis: All data are shown as mean ± standard error of
the mean (SEM). Statistical significance (p < 0.05) was assessed by Student’s paired
t-test, Student’s unpaired t-test, ANOVA, repeated measures ANOVA, or Pearson correlation where applicable (InStat 3.0). Decay kinetics of D2- and 2-IPSCs and extracellular catecholamine transients were fit with a single exponentialα using a
Simplex algorithm optimized by the sum of squared errors in Axograph X (Axograph
Scientific). Single exponential decays provided accurate fits to the decay kinetics and were used to indicate the duration of the IPSC decay phase. Multiple exponential fits were not considered during analysis. For all figures, * denotes p <
0.05, ** denotes p < 0.01, *** denotes p < 0.001, and n.s. denotes p > 0.05.
Materials: Picrotoxin, DNQX, CGP 55845, and MK-801 were from Ascent
Scientific. S-(-)-sulpiride and idazoxan hydrochloride were from Tocris Bioscience.
Dextran (35,000-50,000 kD) was from MP Biomedical. K-methylsulphate was from
Acros Organics. BAPTA was from Invitrogen. Cocaine hydrochloride was obtained
93 from the National Institute of Drug Abuse. Dopamine hydrochloride, noradrenaline
hydrochloride, D-glucose, sodium L-ascorbate, and HEPES sodium salt were from
Sigma-Aldrich. All other chemicals were from Fischer Scientific.
RESULTS
The time courses of dopamine and noradrenaline transmission are different
To examine the mechanisms regulating catecholamine synapses, voltage
clamp recordings were made from either noradrenaline neurons in the LC or
dopamine neurons in the VTA. A single stimulation in the LC from brain slices from
the rat evoked a slow IPSC (67.3 ± 6.9 pA, n = 26) that was blocked by the
noradrenaline 2-receptor antagonist idazoxan (2 M) (Fig. 1A). Bath application of
noradrenaline (30α µM) also evoked large outward currentsμ in these LC neurons
(298 ± 34 pA, n = 3). As the amplitude of α2-receptor mediated outward currents in
LC neurons was significantly smaller in slices from the mouse (bath application 30
µM: 83 ± 14 pA, n = 3; p < 0.01, Student’s t-test), all noradrenergic α2-receptor
mediated transmission was subsequently examined in LC slices from the rat LC.
Previous work has shown that D2-IPSCs recorded from the VTA of rats and mice have similar kinetics though the amplitudes are significant larger in mice
(Courtney et al., 2012). As bath application of dopamine (30 µM) evoked
significantly larger outward currents in VTA neurons from brain slices from mice
than from rats (mouse: 223 ± 8 pA, n = 3, rat: 126 ± 22 pA, n = 4, p < 0.05, Student’s
t-test), dopamine D2-receptor mediated transmission was examined in mouse VTA
94 brain slices. A single stimulation in the VTA evoked a dopamine-mediated IPSC (29.9
± 3.5 pA, n = 14) that was abolished by the D2-receptor antagonist sulpiride (200 nM) (Fig. 1B). Both IPSCs reversed near the potassium equilibrium potential and were inhibited by BaCl2 (300 µM), suggesting that both currents resulted from GIRK
channel activation (Egan et al., 1983; Lacey et al., 1987). A comparison of the time
courses of D2-IPSCs in the VTA with those of 2-IPSCs in the LC revealed that D2-
IPSCs had markedly faster kinetics. When evokedα with similar stimulation intensity,
dopamine D2-IPSCs occurred following a latency of 64 ± 3 ms (10% onset) and had
10 - 90% rise times of 163 ± 6 ms (Fig. 1C). This led to D2-IPSCs reaching their peak
in less than 300 ms (n = 17) (Fig. 1C). In contrast, the kinetics of noradrenaline α2-
IPSCs from the LC were slower (latency to 10% onset: 154 ± 6 ms; 10 - 90% rise
time: 388 ± 11 ms; n = 26, p < 0.001 versus D2-IPSC, Student’s t-test). Thus α2-
IPSCs in the LC did not peak until nearly 900 ms (Fig. 1C). As the rate of GIRK
current activation is determined by the concentration of agonist at the receptor
(Ford et al., 2009; Ingram et al., 1997; Sodickson and Bean, 1996), the faster
activation of D2-IPSCs suggests that a higher effective concentration of dopamine
may underlie transmission at D2-receptors in the VTA than noradrenaline at 2-
receptors in the LC. α
In addition to activating more slowly, 2-IPSCs were longer than D2-IPSCs
and slower to decay (τdecay 2-IPSC: 1.1 ± 0.1 αsec, n = 26; τdecay D2-IPSC: 0.4 ± 0.1 sec,
n = 17; p < 0.001; half-widthα α2-IPSC: 1.2 ± 0.1 sec, n = 26; half-width D2-IPSC: 0.5 ±
0.02 sec, n = 17; p < 0.001) (Fig. 1D, F, G). Noradrenaline-mediated α2-IPSCs also
varied widely in decay time (τDecay standard deviation: 0.56 sec, n = 26) (Fig. 1D) and
95 the amplitude of larger events correlated with longer decay times (R2 = 0.176, n =
26, p < 0.05, Pearson correlation) (Fig. 1E). Surprisingly, the fast decay kinetics of
dopamine IPSCs were remarkably similar among neurons (τDecay standard deviation:
0.09 sec, n = 17) (Fig. 1G) such that there was no correlation with amplitude (Fig.
1H). As the rise time (10 - 90%) of neither α2-IPSCs nor D2-IPSCs correlated with the amplitude of each event (Pearson correlation; 2-IPSCs: R2 = -0.1, p = 0.8; n = 26;
D2-IPSCs: R2 = 0.04, p = 0.4; n = 17) (Fig. 1I), the slowα time course of α2-IPSCs is
unlikely to result from the asynchronous release of noradrenaline in the LC. These
results are consistent with the idea that an increase in the amount of noradrenaline
released leads to increased exposure of α2-receptors. The consistent time course of
D2-IPSCs, in contrast, suggests that dopamine is regulated such that the duration of transmission remains independent of the number of active release sites.
Pooling of transmitter prolongs the duration of noradrenaline transmission
Spillover of transmitter from within the synapse to distal extrasynaptic sites
can prolong of the duration of synaptic events (Beenhakker and Huguenard, 2010;
Carter and Regehr, 2000; Cathala et al., 2005; DiGregorio et al., 2002; Isaacson et al.,
1993; Oláh et al., 2009; Scanziani, 2000; Szabadics et al., 2007). In several cases, spillover can allow for pooling in the extracellular space of transmitter that originated from multiple pre-synaptic release sites (Balakrishnan et al., 2009;
Barbour et al., 1994; Otis et al., 1996; Overstreet and Westbrook, 2003; Silver et al.,
1996). To examine whether pooling contributed to the time course of transmission at these two synapses, we next varied the release of dopamine and noradrenaline to
96 determine whether increased transmitter release could result in longer duration
IPSCs.
The stimulus of single pulse was varied in order to determine the intensity
that produced ~85% of the maximal IPSC amplitude (high stimulation). The
stimulus intensity was then lowered to reduce the IPSC amplitude by 62 ± 4% for
D2-IPSCs (n = 8) and 59 ± 2% for 2-IPSCs (n = 9, p = 0.5. vs. D2-IPSCs, Student’s t-
test) (Fig. 2A). Despite the reductionα in amplitude, the decay kinetics of dopamine
D2-IPSCs remained constant (0.4 ± 0.1 sec to 0.4 ± 0.1 sec, high to low stimulation intensity, p = 0.9, Student’s paired t-test) (Fig. 2A). In contrast, reducing the
stimulation intensity significantly accelerated the decay kinetics of 2-IPSCs (0.8 ±
0.1 sec to 0.4 ± 0.1 sec, high to low stimulation intensity, p < 0.001, Student’sα paired
t-test) (Fig. 2A).
Next, we lowered the release probability (Pr) by decreasing the extracellular
concentration of calcium. The amplitude of D2-IPSCs and α2-IPSCs were reduced to
48 ± 3% and 53 ± 3% of control levels when Ca2+ was reduced from 2.5 mM to 1.0
mM. Lowering extracellular Ca2+ had no effect on the decay time of D2-IPSCs in the
VTA (τdecay: 106 ± 8% of 2.5 mM Ca, n = 5, p > 0.1, Student’s paired t-test), yet it accelerated the decay of α2-IPSCs in the LC by 48 ± 5 % (n = 8, p < 0.001, Student’s paired t-test) (Fig. 2B).
Finally, the intensity (ranging from 10 to 80 µA) of bursts of stimuli (5 pulses at 40 Hz) was varied. Despite increasing the amplitude 4-fold over the range of stimulation, the kinetics of D2-IPSC decay remained constant (0.4 ± 0.1 seconds to
0.4 ± 0.1 seconds, lowest versus highest, n = 12, Pearson Correlation p > 0.05)
97 (Beckstead et al., 2004). Increasing the intensity of stimulation in the LC, however,
resulted in larger amplitude 2-IPSCs that also had longer decay times (0.7 ± 0.1
seconds to 1.5 ± 0.2 seconds,α lowest versus highest; n = 15 cells; Pearson Correlation
p < 0.001) (Fig. 2C). Similar results were obtained when the frequency of
stimulations during the train was varied (Fig. 2D). IPSCs evoked at low frequency
(12.5 Hz) in both the VTA and LC had smaller amplitudes (Fig. 2D). This reduced the
decay time of noradrenaline IPSCs in the LC by 23% (n = 5, p < 0.01, Student’s
paired t-test) but did not alter rate of decay of the D2-IPSC (n = 5, n.s., Student’s
paired t-test) (Fig. 2D). Thus in all cases the time course of dopamine transmission
remained constant regardless of the size of D2-IPSC.
As spillover of transmitter from neighboring sites is limited when the
probability of release is lowered (Balakrishnan et al., 2009; DiGregorio et al., 2002;
Hartzell et al., 1975; Silver et al., 1996; Trussell et al., 1993), the acceleration in the
decay of α2-IPSCs under conditions of low release suggests that pooling of
noradrenaline from adjacent release sites contributed to the extended time course
of the α2-IPSC. In contrast, the consistent duration of dopamine D2-IPSCs suggests
that limited dopamine spillover occurs during phasic transmission.
Differences in receptor signaling and dendritic filtering cannot account for the variation in kinetics of catecholamine transmission
We next tested whether differences in receptor signaling or the rate of
release could alternatively explain the different time courses between the two
IPSCs. First, the efficiency of α2-/D2-receptor signaling to GIRK channels was
98 examined to see if the kinetics of dopamine IPSCs could result from faster coupling of D2-receptors to potassium channels. Diffusion and reuptake severely slow the local application of exogenous agonists (Barbour and Hausser, 1997). To avoid this, outside-out macro patches were pulled from either noradrenergic neurons of the LC or dopaminergic neurons of the VTA and moved out of the slice into free solution
(Ford et al., 2009). Currents were evoked from macro patches by rapid-flow application of either noradrenaline or dopamine via a two-barrel theta tube flow pipe. One barrel of the flow pipette contained a control solution while the second barrel contained a known concentration of dopamine or noradrenaline. The flow pipe was attached to a piezoelectric bimorph to allow for rapid exchange between the two the solutions (Fig. 3A). The application of noradrenaline (100 M, 100 ms) onto LC macro patches resulted in idazoxan-sensitive outward currentsμ that initiated within 58 ± 8 ms (10% onset, n = 6) and had a time to peak of 202 ± 14 ms
(n = 6) (Fig. 3A, B). The kinetics of these currents were indistinguishable from the kinetics of sulpiride-sensitive currents evoked from patches from VTA neurons when dopamine was similarly applied (100 M, 100 ms) (10% onset: 66 ± 4 ms; time to peak: 217 ± 22; n = 13, p = n.s. comparedμ to NA onto LC patches, Student’s t- test) (Fig. 3A, B). This indicates that the kinetics of both the 2- and D2-receptor mediated currents are similar and that coupling efficiency cannotα explain the difference in time course between D2- and 2-IPSCs.
Next, the rise and fall of the concentrationα of dopamine and noradrenaline in the extracellular space was measured with fast-scan cyclic voltammetry. A carbon- fiber electrode was placed into either the LC or VTA and cycled through a triangular
99 waveform to detect either noradrenaline or dopamine (10Hz, Fig. 3C, D). The rise as well as the duration of catecholamines in the extracellular space was similar between the VTA and LC (Fig. 3C, D). Using a triangular voltage protocol of -0.4 V to +1.3 V at 400V/sec to maximize sensitivity (Heien et al., 2003), both dopamine and noradrenaline release peaked in less than 500 ms following a single stimulation
(0.5 ± 0.1 sec, n = 9, VTA; 0.3 ± 0.1 sec, n = 13, LC; n.s.) and decayed with time constants of ~1.5 sec (1.4 ± 0.2 sec, n = 9, VTA; 1.7 ± 0.2 sec, n = 13, LC, n.s.).
Similarly, the kinetics of both release and decay were comparable for both dopamine and noradrenaline if a sine-wave voltage protocol of -0.4 V to +1.0 V at
300 V/sec was used to more accurately measure the kinetics of release (Heien et al.,
2003). Thus the release of catecholamines into the extracellular space had similar time courses in both regions.
As the dendritic location of inputs could differ between dopamine and noradrenaline neurons, the slow decay of α2-IPSCs may result from filtering of faster events at distal dendritic sites within the LC. To test whether the noradrenaline IPSC extended beyond the time when synaptic receptor signaling was active, we applied a voltage jump during the IPSC (Barbour et al., 1994; Häusser and
Roth, 1997; Pearce, 1993). Voltage steps from near the potassium equilibrium potential to depolarized potentials increased the synaptic current to levels equivalent to that under control conditions for both D2-IPSCs and α2-IPSCs when given after the peak of the IPSC (Fig. 3E). As measured by the average current for
500 ms after the depolarizing voltage step, there was no difference between IPSCs with or without the voltage steps for either D2-IPSCs (without voltage vs. with
100 voltage step; 500 ms post stimulation step: 161 pA vs. 162 pA; 1000 ms post stimulation step: 28 pA vs. 28 pA; 2000 ms post stimulation step: 6 pA vs. 7 pA; n =
6; p = n.s. for all comparisons, Student’s paired t-test) or 2-IPSCs (without voltage vs. with voltage step; 1000 ms post stimulation step: 109α pA vs. 113 pA; 2000 ms post stimulation step: 50 pA vs. 56 pA; 3000 ms post stimulation step: 24 pA vs. 24 pA; n = 3; p = n.s. for all comparisons, Student’s paired t-test). This indicates that synaptic conductances were active even at late times and that the slow decay of α2-
IPSCs in the LC is due to the intrinsic slow kinetics of a receptor-mediated event.
Together these results suggest that the difference in synaptic timing between dopamine and noradrenaline is unlikely to be explained by differences in dendritic filtering, receptor signaling kinetics, or the kinetics of release.
Finally, we tested whether the differences in kinetics could be explained by differences in transmission between species. First, we compared D2-IPSCs evoked from slices from the rat to those from the mouse. Due the small amplitude of D2- receptor mediated GIRK currents in the rat VTA, trains of 5 stimulations (40 Hz) were used to evoke dopamine release. The amplitude of D2-IPSCs from brain slices from the mouse was approximately 6-fold greater than D2-IPSCs from brain slices from the rat (mouse: 133 ± 21 pA, n = 11; rat: 21 ± 4 pA, n = 7; p < 0.001, Student’s t- test) (Fig. 3F). However despite the difference in amplitude, D2-IPSCs in both species had similar decay kinetics ( decay; mice: 0.34 ± 0.02 sec, n = 11; rats: 0.39 ±
0.05 sec, n = 7; p = n.s., Student’s t-test).τ Similar to brain slices from the mouse, lowering the stimulation intensity had no effect on the kinetics of rat D2-IPSCs despite a 47 ± 4% reduction in amplitude ( decay: 0.44 ± 0.04 sec, n = 7, p = n.s. vs.
τ 101 maximal stimulation, Student’s paired t-test) (Fig. 3G, H). We next tested whether
the kinetics of 2-IPSCs recorded in the LC of mice were similar to those recorded in
the LC of rats. αAgain, due the small amplitude of α2-receptor mediated GIRK
currents in mouse LC slices, noradrenaline was evoked by a burst of 5 stimulations
(40 Hz). The amplitude of α2-IPSCs recorded in the rat LC were more than 6-fold
greater than α2-receptor mediated IPSCs recorded in the mouse LC (rats: 226 ± 30
pA, n = 10; mice: 35 ± 3 pA, n = 9; p < 0.001, Student’s t-test) (Fig. 3F). The decay
kinetics of α2-IPSCs however were indistinguishable in slices obtained from the
mouse and the rat ( decay; rats: 1.3 ± 0.2 sec, n = 10; mice: 1.6 ± 0.2 sec, n = 9; p = n.s.,
Student’s t-test). Likeτ in the rat, lowering the stimulation intensity reduced both the amplitude (58 ± 7% reduction) (Fig. 3G) and time course of decay of α2-IPSCs in the
LC of slices from the mouse (50 ± 5% reduction, n = 9, p < 0.001, Student’s paired t-
test) (Fig. 3H). Together the results suggest that in both rats and mice, different
mechanisms regulate the timing of dopamine and noradrenaline transmission.
Reuptake transporters maintain the temporal fidelity of dopamine transmission in the VTA
Because the diffusion of transmitter is regulated by neurotransmitter transporters (Barbour et al., 1994; Beenhakker and Huguenard, 2010; Brasnjo and
Otis, 2001; Isaacson et al., 1993; Overstreet and Westbrook, 2003), we next examined how D2- and α2-IPSCs were affected after blocking dopamine and noradrenaline uptake transporters (DAT, NET) with cocaine (2 µM). Cocaine potentiated both the amplitude and duration of D2- and α2-IPSCs (Fig. 4A) and
102 prolonged their rising phases as indicated by a delayed time to peak. Cocaine blocks
DAT and NET with similar potency (Han and Gu, 2006) yet consistently had a
greater effect on the amplitude of D2-IPSCs than α2-IPSCs. Cocaine also prolonged
the rising phase of D2-IPSCs more than 2-IPSCs (time to peak; D2-IPSC: 76 ± 6%
increase in cocaine, n = 11; 2-IPSC: 36 α± 8% increase in cocaine, n = 10; p < 0.001,
Student’s t-test). A similar αeffect of cocaine was observed between recordings from
the rat and mouse LC and the rat and mouse VTA (D2-IPSCs, rat VTA: 360 + 44%
increase in amplitude, n = 4; α2-IPSCs, mouse LC: 114 + 11% increase in amplitude,
n = 4; p < 0.01 for change in D2-IPSCs vs. α2-IPSCs; Student’s t-test). Taken
together, these results suggest that reuptake plays a more dominant role in
regulating dopamine transmission in the VTA than noradrenaline transmission in
the LC.
We next applied brief bursts of stimuli to recruit multiple release sites.
Again, bursts facilitated the amplitude of D2-IPSCs without altering their decay
times but increased both the amplitude and duration of α2-IPSCs (D2-IPSCs: single
stimulus: 32 ± 5 pA, 0.4 ± 0.1 sec; 5 stimuli: 133 ± 21 pA, 0.3 ± 0.1 sec; n = 11; p <
0.001 for amplitude, p = n.s. for decay time. α2-IPSCs: single stimulus: 64 ± 10 pA,
0.8 ± 0.1 sec; 5 stimuli: 160 ± 22 pA, 1.3 ± 0.2 sec; n = 11; p < 0.001 for amplitude, p
< 0.01 for decay time) (Fig. 4B). In the presence of cocaine however, the decay times
of D2-IPSCs were increased as a result of increased stimulation (Fig. 4C, D). Thus
when scaled to peak amplitude, the D2-IPSC evoked by a burst was no longer super imposable on D2-IPSCs evoked by a single stimulus (Fig. 4C inset). A similar effect was also observed in the VTA of the rat in cocaine (n = 6, p < 0.05, Student’s paired t-
103 test). Thus, blocking reuptake transporters in the VTA extended the diffusion of
dopamine such that the duration of the D2-IPSC was now dependent on the amount
of transmitter released. These results suggest that in the VTA, DATs limit spillover
to prevent dopamine pooling and activation of distal extrasynaptic D2-receptors.
Dopamine transmission in the VTA does not occur via volume transmission
As both reuptake and diffusion govern the spatial limit over which spillover activates post-synaptic receptors (Barbour and Hausser, 1997; Rusakov and
Kullmann, 1998b), the time course of IPSCs were examined after slowing diffusion through the addition of dextran to the extracellular solution (5%, M.W. = 35,000 –
50,000) (Ford et al., 2010; Min et al., 1998b). Slowing diffusion with dextran alters the amplitude and time course of spillover events (Ford et al., 2010; Markwardt et al., 2009; Min et al., 1998b; Nielsen et al., 2004; Szabadics et al., 2007) thus dextran was predicted to have a greater effect on α2-IPSCs than D2-IPSCs. Following application of dextran, the amplitude of noradrenaline α2-IPSCs increased by 137 ±
16% (n = 10, p < 0.05, Student’s paired t-test) (Fig. 5A, D), suggesting that noradrenaline does not saturate receptors mediating the IPSC. Dextran also increased the decay time of α2-IPSCs by 157 ± 16% (n = 10, p < 0.05, Student’s paired t-test) (Fig. 5A, E). Because dextran altered the kinetics of the α2-IPSC, diffusion is likely a key clearance mechanism regulating noradrenaline transmission in the LC. In contrast, dextran had no effect on the kinetics or amplitude of dopamine IPSCs from the VTA (amplitude: 88 ± 7%; τDecay: 112 ± 9%; n = 9; n.s.)
(Fig. 5B, D, E). The lack of an effect of dextran on D2-IPSCs implies that under
104 control conditions extended diffusion plays little role in determining the kinetics of
the D2-IPSC. However in the presence of a sub-saturating concentration of cocaine
(500 nM), dextran slowed the kinetics of D2-IPSCs (Amplitude: 48 ± 5%; Decay: 148
± 16%; n = 6; p < 0.05, Student’s paired t-test) (Fig. 5C, E). As hindering diffusionτ reduced the amplitude of the D2-IPSC in the presence of cocaine, dextran likely prevented the activation of receptors away from the release site. Thus, diffusion of dopamine only contributes to D2-IPSCs when reuptake is blocked confirming that transporters prevent the long-range spillover activation of D2-receptors.
Additionally, we examined the effects of enzymatic degradation on transmitter clearance using the monoamine oxidase inhibitor (MAO-I) pirlindole (10
M). Acute application (10 minutes) of pirlindole had no effect on the amplitude or decayμ kinetics of either 2- (n = 3, p = n.s. for both amplitude and Decay, Student’s
paired t-test) or D2-IPSCsα (n = 3, p = n.s. for both amplitude and Decayτ , Student’s
paired t-test). Thus, enzymatic degradation by monoamine oxidaseτ is unlikely to
acutely influence the time course of dopamine or noradrenaline mediated IPSCs.
To further examine how signaling at a distance may lead to activation of catecholamine receptors in the VTA and LC, we pulled macro patches from either dopamine or noradrenaline neurons out from the slice and placed these patches back on the surface of the brain slice (Fig. 6A). Using patches as a sensor for evoked release, bursts of stimuli within the LC evoked outward currents in the majority of
LC-noradrenaline sensor patches (16 ± 3 pA; n = 12) (Fig. 6B). Currents were abolished by the α2-receptor antagonist idazoxan (2 M) (95 ± 2% reduction, n = 3, p < 0.05, Student’s paired t-test). The kinetics and timeμ course of these events were
105 similar to that of whole-cell α2-IPSCs recorded separately from LC neurons in other control experiments. When scaled to the peak of their amplitude, the evoked α2- currents from patches were indistinguishable from an average α2-IPSC (Fig. 6C).
Thus, despite patches being located at a distance from the site of release, the concentration profile of noradrenaline was similar to the α2-receptors that mediate the IPSC within the slice.
In contrast, when sensor patches were pulled from dopamine cells in the VTA or SNc and placed on the surface of the slice, bursts of stimuli failed to evoke D2- receptor mediated outward currents (Fig. 6D). Increasing the intensity of stimulation (20 stimulations, 100 Hz) also failed to produce measurable outward currents (Fig. 6 D, E). This suggests that under control conditions, the concentration of dopamine that escapes from the slice is too low to activate D2-receptors. In the presence of cocaine (5 µM), however, bursts of stimuli were able to evoke outward currents in the majority of patches (Fig. 6 D, E). Currents were abolished by sulpiride (200 nM) (93 ± 5% reduction, n = 4, p < 0.05, Student’s paired t-test), confirming they were mediated by D2-receptor activation. Thus, dopamine transporters prevented dopamine from diffusing out of the slice at a high enough concentration to activate D2-receptors. This observation is consistent with the role of dopamine reuptake in preventing transmitter pooling and synaptic crosstalk under normal conditions.
High concentrations of dopamine result in rapid activation of D2-IPSCs
106 The concentration of agonist determines the rate of activation of GPCR-
mediated GIRK currents (Ford et al., 2009; Ingram et al., 1997; Sodickson and Bean,
1996). Amperometric measurements have estimated somatic quantal content of noradrenaline vesicles in the LC to be about twice that of dopamine vesicles in the
VTA (~37,000 noradrenaline molecules vs. ~14,000 dopamine molecules per vesicle) (Huang et al., 2007; Jaffe et al., 1998). Despite the greater number of noradrenaline molecules being released per vesicle, the α2-dependent IPSC was
nearly four times slower than D2-IPSC (Fig. 1C, F). This suggests that the α2-IPSC
may be mediated by a sub-saturating concentration of noradrenaline.
To determine the concentration of catecholamines that underlies the two
IPSCs, we used rapid-flow application of either dopamine or noradrenaline via a
theta tube flow pipette to examine rising kinetics of the α2- and D2-mediated
currents. Application of high, saturating concentrations of dopamine (100 M, 100
ms) onto VTA macro patches gave rise to an outward current (Fig. 7A) thatμ
mimicked the activation kinetics of the D2-IPSC (10% onset: 66.3 ± 4.4 ms, n = 13, p
> 0.6 vs. D2-IPSCs, Student’s t-test) (Fig. 7B, C). Reducing the concentration of
dopamine resulted in slower currents that no longer matched the activation kinetics
of D2-IPSCs (10% onset; 10 M DA: 76.8 ± 5.9 ms, n = 13, p < 0.05 vs. D2-IPSCs; 3
M DA: 92.8 ± 3.7 ms, n = 6, μp < 0.001 vs. D2-IPSCs, Student’s t-test) (Fig. 7C). This
suggestsμ that at least 100 M dopamine is present at D2-receptors during the initial
phase of the D2-IPSCs, confirmingμ previous observations (Ford et al., 2009).
Next, noradrenaline was applied onto LC macro patches to determine the
concentration necessary to mimic the rise kinetics of the α2-IPSC. Again, the rate of
107 activation of α2-mediated currents was similar to D2-currents from VTA patches
when evoked with similar concentrations of agonist (100µM, 100 ms) (10% onset;
NA: 57.9 ± 8.2 ms, n = 6; p = n.s. vs. DA) (Fig. 7D top). Additionally, the α2- and D2- currents decayed at a similar rate when exposed to similar applications of agonist
( Decay; NA: 116 ± 16 ms, n = 6; DA: 117 ± 11 ms, n = 12; p = n.s., Student’s t-test)
(Fig.τ 7D top). Because α2-IPSCs peak significantly later than D2-IPSCs,
noradrenaline was applied for 1000 ms to mimic the longer rise time of the α2-IPSC
(Fig. 7D bottom). When a high concentration of noradrenaline (100 µM, 1000 ms)
was applied, α2-mediated currents rose faster than 2-IPSCs (10% onset: 66 ± 14
ms, n = 9, p < 0.01 vs. 2-IPSC, Student’s t-test) (Fig. α7G). This suggests that a lower
concentration of noradrenalineα mediates 2-IPSCs at synapses within LC. The rising
phase of the α2-IPSC was best matched byα applying 3 µM (1000 ms) noradrenaline
(3 M NA, 10% rise time: 124 ± 14 ms, n = 11, p = n.s.; 10 M NA, 10% rise time: 82
± 13μ ms, n = 9, p < 0.001 vs. 2-IPSC; 1 M, 10% rise time:μ 201 ± 30 ms, n = 8, p <
0.05, Student’s t-test) (Fig. 7F,α G). Thus,μ the concentration of noradrenaline that
interacts with 2-receptors during the rising phase of the IPSC is on the order of
thirtyfold less thanα the concentration of dopamine that mediates the D2-IPSC.
DISCUSSION
The present study examined two catecholamine-mediated IPSCs and found that
they had very different characteristics. Whole cell recordings revealed that the rise
and fall of D2-IPSCs in VTA was markedly faster than 2-IPSCs in the LC. Both GPCRs
α 108 activated GIRK conductances that displayed similar kinetics upon rapid-flow application of the respective agonist in macro patch recordings. The difference between the kinetics of the IPSCs was instead dependent on the extent of transmitter diffusion as a result of differences in the level of uptake. Within the VTA, we found that transporters strongly limited dopamine spillover. This allowed for only high, local concentrations to activate proximal D2-receptors. As the decay time of D2-IPSCs
was unaffected by changes in the extent or probability of dopamine release, our
results suggest that the time course of dopamine signaling in the VTA does not rely on
pooling of transmitter from multiple sites. This suggests that individual release sites
contribute independently to D2-receptor activation during the IPSC, which functions to maintain the temporal fidelity of dopamine transmission. In contrast, due to less efficient reuptake in the LC, noradrenaline was able to spill out away from the release site and diffuse a significant distance before activating post-synaptic α2-receptors.
Subsequent pooling of extracellular noradrenaline when multiple terminals were activated led to a prolonged duration of the α2-IPSC. As metabotropic transmission is thought to occur through the activation of extrasynaptic GPCRs by transmitter spillover and pooling in the extracellular space (Beenhakker and Huguenard, 2010;
Isaacson et al., 1993; Kulik et al., 2002; Otis and Mody, 1992; Scanziani, 2000) these results suggest that the mechanisms underlying metabotropic-receptor mediated neurotransmission are more diverse than previously believed. Differences in the temporal and spatial tuning of GPCR-mediated transmission may allow for differences in signal integration that underlies dopamine and noradrenaline mediated actions.
109 The rise time of noradrenaline α2-IPSCs was nearly four-fold slower than that of dopamine D2-IPSCs. This kinetic difference could not be explained by voltage clamp errors resulting from dendritic filtering, differences in coupling efficiency of GPCRs to GIRK channels, or differences in the kinetics of release between terminals. Instead the slower rate of activation of the α2-receptor- mediated synaptic current was likely due to lower concentrations of noradrenaline activating receptors during the IPSC (3 µM NA versus 100 µM DA, Fig. 7C, G). To understand how diffusion contributed to receptor activation we simulated the spatial profile of catecholamine diffusion from single vesicles in either the VTA or LC
(Fig. 7H). The model showed that in the VTA, the peak concentration of dopamine in the VTA was maintained at or above 100 M only within 0.4 m from the site of release (Fig. 7H left). D2-receptors locateμd further than 0.4 μm would be exposed to lower concentrations of dopamine. This would be predictedμ to significantly slow the rate of D2-receptor activation below what was observed for the D2-IPSC (Fig. 7C).
When the diffusion of noradrenaline in the LC was simulated, the peak concentration remained above 3 M until 1.7 m from the release site (Fig. 7H right). As α2-receptors located nearerμ than 1.7μ m would experience a higher peak concentration of noradrenaline, these receptorsμ would have activation kinetics faster than were observed during the α2-IPSC. This simulation suggests that α2- receptors mediating the initial phase of noradrenergic transmission in the LC may be five times further from the site of release than D2-receptors in the VTA (Fig. 7I).
Because the rate of current deactivation was similar for α2- and D2- receptors when agonist was removed (Fig. 7D top), the longer time course of α2-
110 IPSCs in the slice suggests that a longer-lasting noradrenaline transient led to a
prolonged activation of 2-receptors following release. Spillover of
neurotransmitter can oftenα prolong transmitter exposure through the activation of
extrasynaptic receptors (Beenhakker and Huguenard, 2010; Bennett et al., 2012;
Carter and Regehr, 2000; Cathala et al., 2005; DiGregorio et al., 2002; Oláh et al.,
2009; Overstreet and Westbrook, 2003; Szabadics et al., 2007; Szapiro and Barbour,
2007). To understand how spillover shapes the activation of post-synaptic
catecholamine receptors, we lowered the probability of release for both
noradrenaline and dopamine. Reducing release probability by decreasing
extracellular calcium reduced the decay time of α2-IPSCs but not D2-IPSCs. Because
transmission mediated by pooling is more sensitive to reductions in release than
point-to-point mediated transmission (Arnth-Jensen et al., 2002; DiGregorio et al.,
2002; Scanziani, 2000), pooling likely plays a significant role in shaping the decay
time of only α2-IPSCs. Support for this comes from the fact that slowing
extracellular diffusion with dextran did not alter the duration of D2-IPSCs yet
slowed the decay of α2-IPSCs. As dextran did not affect the amplitude of D2-IPSCs,
it is likely that D2-receptors are saturated during transmission by the local high
concentration of dopamine occurring near the site of release, consistent with our
finding from fast-flow experiments with macro-patches. However a potential role for enhanced activation of pre-synaptic D2-receptors modulating dopamine release under these conditions cannot be excluded.
We found that the only condition that could affect the duration of D2-IPSCs was when transporters were blocked. Reuptake or buffering of dopamine by
111 transporters is therefore the major mechanism that limits spillover or pooling in the
VTA. Cocaine had a dramatic effect on delaying the time to peak and prolonging the
duration of D2-IPSCs (Beckstead et al., 2004; Courtney et al., 2012; Ford et al.,
2010). Ultrastructure studies show that dopamine receptors in the midbrain
aggregate at both synaptic and extrasynaptic sites on dopaminergic dendrites
(Pickel et al., 2002; Sesack et al., 1994). As cocaine increases the distance over
which dopamine signals (Cragg and Rice, 2004; Rice and Cragg, 2008), blocking reuptake likely allows the concentration profile of dopamine to remain sufficiently high outside the synapse to recruit additional extrasynaptic receptors that normally do not participate in transmission. In the LC, post-synaptic 2-receptors are distributed along the plasma membrane in regions both opposedα and unopposed by other membranes (Lee et al., 1998c, 1998d). As both dextran and cocaine slowed
α2-IPSCs, the profile of 2-receptor activation is determined by both diffusion and reuptake likely becauseα of less efficient uptake in the LC. The reason for the difference in uptake between the LC and VTA is not clear but may be due to reduced levels of expression of NETs in the LC (Burchett and Bannon, 1997; Szot et al., 1996) and/or a differential distribution of transporters further away from release sites
(Beenhakker and Huguenard, 2010). Support for this comes from the observation that DATs are often associated with both the pre- and post-synaptic densities of dendro-dentritic synapses between midbrain dopamine neurons (Hersch et al.,
1997).
Through the course of these experiments, miniature α2-IPSCs were not observed while recording from LC neurons, despite the fact that evoked α2-IPSCs
112 were routinely robust in amplitude. This is consistent with volume transmission and the need for pooling of multiple noradrenaline vesicles to drive α2-receptor activation. A recent report, however, described dopamine mediated spontaneous miniature D2-IPSCs in the VTA (Gantz et al., 2013). This indicates that vesicular dopamine release must occur adjacent to a high density of D2-receptors. As the time course of miniature D2-IPSC is similar to that of IPSCs driven by evoked release
(Gantz et al., 2013), dopamine synapses likely function independently even during synchronous release from multiple terminals. The density of release sites often regulates the extent of spillover and pooling between synapses (Balakrishnan et al.,
2009; Overstreet and Westbrook, 2003). A low number of terminals in the VTA
(Ford et al., 2010; Nirenberg et al., 1996; Wilson et al., 1977) may therefore aid in maintaining the synaptic independence between sites to limit dopamine cross talk.
Autoreceptor inhibition regulates both the overall level of excitability and the patterns of firing across catecholamine neurons (Aghajanian and Bunney, 1977;
Beckstead et al., 2004; Benoit-Marand et al., 2001; Courtney et al., 2012; Egan et al.,
1983; Schmitz et al., 2002), such that changes in the duration or strength of somatodendritic autoreceptor transmission can dramatically impact catecholamine signaling throughout the brain (Bello et al., 2011; Buckholtz et al., 2010). By regulating the functional extent of spillover, different catecholamines may set the extent that inhibition occurs as result of activity in distant neurons within their local network.
113 Figure 3.1. Dopamine and noradrenaline IPSCs have different time courses.
(A) α2-IPSC from a LC neuron under control conditions and in the presence of idazoxan (2 µM). Ticks illustrate time of stimulation. The current artifact remaining in idazoxan was subtracted to allow for analysis of rise kinetics. Left: Schematic of whole-cell voltage-clamp recordings of noradrenaline neurons in the LC. (B)
Averaged D2-IPSCs from a VTA neuron under control conditions and in the presence of sulpiride (200 nM). (C) Summary of the 10% onset and time to peak of α2-IPSCs
(n = 26) or D2-IPSCs (n = 17) (Student's t-test). Artifacts remaining in the presence of antagonists (idazoxan in the LC or sulpiride in the VTA) were subtracted from control α2-IPSCs or D2-IPSCs when analyzing each cell. (D) Representative averages of α2-IPSCs grouped by amplitude. Traces in the presence of idazoxan
were subtracted to remove stimulation artifact. Bottom: Traces from above were
scaled to their peak amplitude. (E) Decay time correlates with amplitude for α2-
IPSCs. Circles represent IPSCs recorded from individual cells (Pearson’s
correlation). (F) Summary data of the half width and τdecay of α2-IPSCs (n = 26) and
D2-IPSCs (n = 17) (Student's t-test). (G) Representative averages of D2-IPSCs
grouped by amplitude. Stimulation artifacts blanked for clarity. (H) Decay time does
not correlate with amplitude for D2-IPSCs (Pearson’s correlation). Circles represent
IPSCs recorded from individual cells. (I) Lack of correlation of rise time (10-90%) with amplitude of IPSCs (Pearson’s correlation). Circles represent IPSCs recorded from individual cells. *** represents p < 0.001; error bars represent ± s.e.m.
114
Figure 3.1
115 Figure 3.2. Increasing noradrenaline release prolongs the decay time of α2-
IPSCs. (A) Averaged 2-IPSCs (n = 9) and D2-IPSCs (n = 8) evoked by high and low
intensity stimulation.α Right: Quantification of the decay times of individual IPSCs.
Changing the stimulation intensity had no effect on the decay time of D2-IPSCs
(Student's paired t-test). (B) Averaged α2-IPSCs (n = 8) and D2-IPSCs (n = 5)
evoked with a single stimulation and recorded in 1.0 mM and 2.5 mM extracellular
Ca2+ ([Ca]o). Right: Summary data of decay times in different extracellular [Ca2+]
(Student's paired t-test). (C) Summary quantification of the decay time of IPSCs evoked by bursts when divided into quartiles based upon their maximum amplitude
(α2-IPSCs: n = 15, D2-IPSCs: n = 12). For α2-IPSCs, longer decay times were significantly correlated with larger amplitude quartiles (Pearson's correlation). (D)
Averaged noradrenaline α2-IPSCs (n = 5) and D2-IPSCs (n = 5) evoked by a burst of stimuli at 12.5 and 40 Hz. Inset: Decay phase of IPSCs normalized and aligned to their peak when evoked by a burst of stimuli at 12.5 or 40 Hz. Stimulation artifacts were blanked for clarity. n.s. signifies p > 0.05, ** represents p < 0.01; error bars represent ± s.e.m.
116
Figure 3.2
117 Figure 3.3. The duration of GIRK signaling, transmitter release and the
synaptic conductance are similar for dopamine and noradrenaline. (A) Left:
schematic of experimental condition for catecholamine rapid-flow application onto nucleated patches. Right: Representative traces of currents evoked following the rapid-flow application of noradrenaline (100 M; 100 ms) onto nucleated patches of
LC cells or dopamine application (100 M; 100μ ms) onto nucleated patches of VTA cells. (B) Summary data illustrating theμ time until 10% onset and the time to peak of currents evoked in patches (Student's t-test). (C) Left: schematic of the FSCV experimental condition. Right: averaged FSCV traces measuring the extracellular concentration of noradrenaline [NA]o in the LC (n = 13) or the extracellular
concentration of dopamine [DA]o in the VTA (n = 9). Release was evoked by a single
pulse. (D) Summary data of the time to peak and decay kinetics of [NA]o and [DA]o
(Student's t-test). (E) Left: schematic of whole-cell voltage clamp recordings. Right:
current responses to a series of voltage jumps following synaptic stimulation.
Voltage jumps were given from near the potassium equilibrium potential (between -
95mV and -105 mV) to the test potential of -60mV at 500, 1000, and 1500 ms after
stimulation while recording D2-IPSCs and 1000, 2000, and 3000ms after
stimulation while recording α2-IPSCs (black traces). IPSCs were evoked by a burst
of stimuli. Each trace is the average of 3-5 events from 3 VTA dopamine cells and 6
LC noradrenaline cells. IPSCs following the jump were separated from the capacitance transients recorded with no extracellular stimulation. Following the voltage jump, the instantaneous IPSCs (black) rapidly approached control IPSCs
(color) recorded at -60mV indicating that the synaptic conductance was still active.
118 The stimulation artifact was blanked for clarity. (F) Summary data of the peak
amplitude of D2- or α2-IPSCs evoked by 5 stimulations (40 Hz) in either the mouse or rat (Student's t-test). (G) Average traces illustrating the D2- and α2-IPSCs in the mouse and rat. Top: α2-IPSCs evoked in the mouse LC have different kinetics when evoked by high or low stimulation intensity. Bottom: D2-IPSCs evoked in the rat
VTA have the same kinetics independent of stimulation intensity. (H) Quantification of the decay kinetics shown in (G) (Student's paired t-test). n.s. represents p > 0.05; error bars represent ± s.e.m.
119
Figure 3.3
120 Figure 3.4. Transporters limit dopamine spillover. (A) Left: Representative
traces of single stimulation IPSCs in control conditions and the increase in
amplitude and duration in the presence of cocaine (2 µM, grey). Right:
Quantification showing the increase in amplitude of IPSCs recorded in the presence
of cocaine. Cocaine potentiated the amplitude of D2-IPSCs more than α2-IPSCs
(Student's t-test). (B) Left: Averaged α2-IPSCs (n = 11) and D2-IPSCs (n = 11)
evoked by single stimulation or a train of 5 stimuli. Insets: Average IPSCs
normalized to peak amplitude. (C) Averaged α2-IPSCs and D2-IPSCs evoked by
either a single stimulation or a train of 5 stimuli in the presence of cocaine (2µM).
Inset: Averaged IPSCs in the presence of cocaine normalized to peak amplitude. (D)
Quantification of IPSC decay times when evoked by a single stimulation or bursts of
5 or 10 stimulations in the absence or presence of cocaine (2 µM, grey). Trains of stimuli failed to potentiate the decay time of D2-IPSCs under control conditions.
Blocking reuptake with cocaine allowed trains of stimuli to evoke longer duration dopamine IPSCs (repeated measures ANOVA, Bonferroni post-hoc test). n.s. signifies p > 0.05, * represents p < 0.05, ** represents p < 0.01, *** represents p <
0.001; error bars represent ± s.e.m.
121
Figure 3.4
122 Figure 3.5. Slowing diffusion alters α2-IPSCs but not D2-IPSCs unless reuptake is inhibited. (A) Averaged α2-IPSC (n = 10) illustrating the effect of dextran (5% w/v). (B) Averaged D2-IPSC (n = 9) illustrating the lack of effect of dextran. (C)
Averaged D2-IPSC in the presence of cocaine (500 nM) (n = 6) illustrating the effect of dextran. (D and E) Quantification of the amplitude (D) and decay time (E) of
IPSCs recorded in control solution (colored bars) and dextran (grey bars) (Student's paired t-test). n.s. signifies p > 0.05, * p < 0.05, *** p < 0.001; error bars represent ± s.e.m.
123
Figure 3.5
124 Figure 3.6. Spillover activates D2 receptors when uptake is inhibited. (A)
Schematic of experimental condition. Large sensor patches from the LC or VTA
were pulled out from the slice and re-positioned at the surface. (B) Left: Averaged
α2-receptor mediated current from noradrenaline neuron sensor patches (n = 17).
Noradrenaline release was evoked from within the LC by a train of stimuli. The stimulating electrode was positioned in the LC, in a similar location used to evoke
α2-IPSCs. Right: Distribution of current amplitudes recorded in all sensor patches
from the LC. Outward currents could be detected in the majority of LC patches. (C)
Normalized, averaged whole-cell α2-IPSC recorded within the LC (n = 15) and α2-
receptor mediated current from a sensor patch recorded at the surface of the LC (n =
12). (D) Averaged D2-receptor mediated currents from dopamine neuron sensor
patches. Left: Control recordings from VTA sensor patches did not respond to
dopamine release evoked by a burst of stimuli (5 pulses, 40 Hz; or 20 pulses, 100
Hz). Right: Averaged D2-receptor current in the presence of cocaine (5 µM; 20
pulses, 100 Hz). (E) Distribution of amplitudes of dopamine sensor patch currents
in all conditions.
125
Figure 3.6
126 Figure 3.7. D2-receptors are located near the site of dopamine release. (A)
Average current when 100 M DA was applied to VTA macro patches for 100 milliseconds (n = 13). Inset:μ Schematic of the experimental paradigm. (B) 100 M
DA mimics the rise kinetics of D2-IPSCs recorded in VTA slices. Right: Expansionμ of the rising phase (shown in the box) with the same time scale as the expansion in F.
(C) Quantification of the 10% onset of various concentrations of DA applied to VTA patches for 100 ms. The dashed line represents the 10% onset of D2-IPSCs measured in VTA slices with the grey box indicating the standard error (Student's t- test). (D) The currents resulting from 100 M NA applied to LC macro patches for
100 ms (n = 6, top) or 1000 ms (n = 9, bottom)μ have similar onset kinetics as when
100 M DA was applied to VTA patches. (E) Average current when 3 M NA was appliedμ to LC macro patches for 1000 ms (n = 11). (F) 3 M NA mimicsμ the rise kinetics of 2-IPSCs recorded in LC slices. Right: Expansionμ of the rising phase
(shown in theα box) with the same time scale as the expansion in B. (G)
Quantification of the 10% onset of various concentrations of NA applied to LC patches for 1000 ms. The dashed line represents the 10% onset of 2-IPSCs recorded in LC slices with the grey box indicating the standard errorα (Student's t- test). (H) Peak concentration versus distance of dopamine (left) and noradrenaline
(right) diffusing away from a single vesicle of transmitter release generated by a model of point-diffusion in the absence of reuptake. Orange lines indicate the distance at which DA reaches the 100 M required to mimic D2-IPSC onset kinetics.
Blue lines indicate the distance at whichμ NA reaches the 3 M required to mimic 2-
IPSC onset kinetics. (I) Scaled cartoon demonstrating the spatialμ range of dopamineα
127 and noradrenaline diffusion required to mediate the rising phase of IPSCs. n.s. represents p > 0.05, * p < 0.05, *** p < 0.001; error bars represent ± s.e.m.
128
Figure 3.7
129 Chapter 4
Synaptic activation of 5-HT1A receptors in
dorsal raphe serotonin neurons.
Courtney NA and Ford CP (2015) Synaptic activation of 5-HT1A receptors in dorsal
raphe serotonin neurons. Manuscript in preparation.
130 ABSTRACT
In the dorsal raphe nucleus (DRN), feedback activation by GI/o-coupled 5-
HT1A autoreceptors reduces the excitability of serotoninergic neurons, which
decreases serotonin release both locally within the DRN as well as in projection
regions. Serotonin transmission within the DRN is thought to occur via transmitter
spillover and paracrine activation of extrasynaptic receptors. Here, we tested this
volume transmission hypothesis in mouse DRN brain slices by recording 5-HT1A
receptor-mediated inhibitory post-synaptic currents (5-HT1A IPSCs) generated by
the activation of G-protein coupled inwardly rectifying potassium channels (GIRKs).
We found that in the DRN of ePET1-EYFP mice, the local release of serotonin
generated 5-HT1A IPSCs in serotonin neurons that rose and fell within a second. The transient activation of 5-HT1A autoreceptors resulted in brief pauses in neuron firing that did not alter the overall firing rate. The duration of 5-HT1A IPSCs was primarily
shaped by receptor deactivation due to clearance via serotonin reuptake
transporters. Slowing diffusion with dextran prolonged the rise and reduced the amplitude the IPSCs and the effects were potentiated when uptake was inhibited. By examining the decay kinetics of IPSCs, we found that while spillover may allow for the activation of extrasynaptic receptors, efficient uptake by serotonin reuptake transporters (SERTs) prevented the pooling of serotonin from prolonging the duration of transmission when multiple inputs were active. Together the results suggest that the activation of 5-HT1A receptors in the DRN results from the local
release of serotonin rather than the extended diffusion throughout the extracellular
space.
131 INTRODUCTION
The dorsal raphe nucleus (DRN) is a major source of ascending serotonergic
innervation to the forebrain and limbic regions (Jacobs and Azmitia, 1992;
Michelsen et al., 2008). Serotonin neurons of the DRN are implicated in multiple
behavioral and cognitive functions (Jacobs and Azmitia, 1992; Lucki, 1998), and dysfunction in serotonin signaling is thought to underlie mood disorders and depression (Michelsen et al., 2008). In addition to release in projection regions, vesicular serotonin (5- hydroxytryptamine; 5-HT) release occurs locally in the DRN at somatic (Colgan et al., 2009; Kaushalya et al., 2008), dendritic (Colgan et al., 2012;
De Kock et al., 2006), and axonal sites (Bruns et al., 2000) where it modulates the activity of DRN neurons (Adell et al., 2002; Andrade et al., 2015; Michelsen et al.,
2008; Piñeyro and Blier, 1999). Release of serotonin can activate inhibitory G i/o-
coupled 5-HT1A autoreceptors that inhibit serotonergic neuron impulse activity
through the opening of inwardly rectifying potassium (GIRK) channels (Aghajanian
and Lakoski, 1984; Bayliss et al., 1997; Gantz et al., 2015; Katayama et al., 1997; Pan
and Williams, 1989; Pan et al., 1989; Vivo and Maayani, 1986). By suppressing
pacemaker firing, 5-HT1A autoreceptors regulate serotonin levels both locally in the
dorsal raphe and in terminal projection regions (Adell et al., 2002; Aghajanian and
Lakoski, 1984; Hjorth and Sharp, 1991; Michelsen et al., 2008; Pan and Williams,
1989; Pan et al., 1989; Portas et al., 1996), thereby influencing behaviors such as
anxiety and stress (Richardson-Jones et al., 2010).
Ultrastructural studies have found somatodentritic 5-HT1A receptors at both
synaptic (Kia et al., 1996) and extrasynaptic sites (Kia et al., 1996; Riad et al., 2000).
132 As 5-HT1A receptors can be located at extrasynaptic sites, it has been thought that serotonergic transmission through these receptors in the DRN occurs by their
paracrine activation by low concentrations of transmitter that result from spillover
of transmitter out from the synapse (Bunin and Wightman, 1999). In support of this
extended form spillover, known as volume transmission, electrochemical studies
have found that evoked release in the DRN drives sub-micromolar increases in the concentration of 5-HT throughout the extracellular space (Bunin and Wightman,
1998; Bunin et al., 1998; Jennings, 2013). While the increase in extracellular
serotonin has been hypothesized to be sufficient to activate 5-HT1A autoreceptors up
to several microns away from release sites (Bunin and Wightman, 1999), the role of
volume transmission in gating activation of these autoreceptors in the DRN has yet
to be directly addressed.
While 5-HT1A-receptors in the DRN in vivo are thought to tonically modulate
firing rates (Fornal et al., 1996; Haddjeri et al., 2004; Hajos et al., 2001; Mundey et
al., 1996), electrophysiological studies using brain slices have found in contrast that
stimulation evokes only 5-HT1A receptor-mediated hyperpolarizations in DRN neurons that cause only brief pauses in neuron firing (Levitt et al., 2013; Morikawa et al., 2000; Pan et al., 1989; Williams et al., 1988). To examine the mechanisms that underlie the synaptic activation of 5-HT1A receptors we recorded from identified serotonergic neurons in the dorsal raphe using ePET1-eYFP mice. Through recording of 5-HT1A-receptor coupled GIRK currents, we found that evoked
serotonin release generated 5-HT1A-receptor mediated currents that rose and fell in a second. This resulted in transient pauses in firing without affecting overall firing
133 rates. While spillover likely allowed for the activation of some extrasynaptic
receptors, serotonin reuptake transporters prevented transmitter pooling from
adjacent release sites from extending the duration of transmission. These findings
suggest that serotoninergic transmission in the DRN occurs by the local activation of
of 5-HT1A autoreceptors near the sites of transmitter release with limited cross talk
between synapses.
METHODS
Slice preparation and visualization. All procedures were approved and performed in
accordance with Case Western Reserve University Institutional Animal Care and Use
Committee guidelines. Coronal brain slices (220 µM) containing the dorsal raphe
nuclei (DRN) were obtained from 4 – 8 week old male and female ePET1-eYFP mice
(Scott et al., 2005). Brain slices were cut using a vibratome (Leica) in ice-cold
cutting solution containing (in mM) 75 NaCl, 2.5 KCl, 6 MgCl2, 0.1 CaCl2, 1.2
NaH2PO4, 25 NaHCO3, 2.5 D-glucose, and 50 sucrose continuously bubbled with 95%
O2 and 5% CO2. After cutting, slices were transferred to artificial CSF (ACSF)
containing (in mM) 126 NaCl, 2.5 KCl, 1.2 MgCl2, 2.5 CaCl2, 1.2 NaH2PO4, 21.4
NaHCO3, and 11.1 D-glucose bubbled with 95% O2 and 5% CO2 and incubated at 35°
C for at least 45 minutes prior to use. MK-801 (10 µM) was included during the
incubation to block NMDA receptors. After incubation, slices were transferred to a
recording chamber and constantly perfused at 2 ml/min with oxygenated ACSF
warmed to 34 ± 2° C. Slices were visualized with a BX51WI microscope (Olympus)
with custom-built infrared gradient contrast optics.
134
Electrophysiology. Whole-cell voltage-clamp recordings (Vh = - 60 mV) were made using an Axopatch 200B amplifier (Molecular Devices). Patch pipettes (1.5 – 2.5
M ) were pulled from borosilicate glass (World Precision Instruments). The pipette intracellularΩ solution contained (in mM): 115 K-methylsulphate, 20 NaCl, 1.5 MgCl2,
10 K-HEPES, 10 BAPTA-tetrapotassium, 2 ATP, 0.3 GTP, and 6 sodium
phosphocreatine, pH 7.4, 275 mOSM. Data were acquired using an ITC-18 interface
(Instrutech) and Axograph X (Axograph Scientific) at 10 kHz and filtered to 2 kHz.
Series resistance was not compensated and recordings were discarded if the access
resistance rose above 15 M . In the case of experiments with reduced extracellular
calcium, MgCl2 was substitutedΩ to maintain a constant divalent ion concentration.
All drugs were applied by bath perfusion. Recordings were made in the
dorsomedial portion of the DRN. Serotonergic neurons within the DRN were
fluorescently identified by eYFP expression driven by the PET1 enhancer (Scott et
al., 2005). In some experiments, dextran (35,000 – 50,000 M.W.) was used to slow
diffusion via macro-molecular overcrowding. Dextran was added to ACSF (5%
weight by volume) and bath perfused after achieving stable recordings. Bath
temperature was monitored to ensure that it did not change by more than 1° C.
Cell attached recordings were made with internal solution containing (in mM): 126 NaCl, 2.5 KCl, 1.2 MgCl2, 2.5 CaCl2, 1.2 NaH2PO4. Data were acquired at 10
kHz and filtered to 5 kHz. Seals between the recording electrode and neuron ranged
from 5 – 12 M . Phenylephrine (3 µM) was included in the bath solution to mimic in
vivo excitatoryΩ drive (Vandermaelen and Aghajanian, 1983).
135
Stimulation and iontophoresis. Serotonin release was evoked with an extracellular,
ACSF-filled monopolar glass electrode. The stimulating electrode was placed 50-
100 µm away from the recorded neuron. A single stimulation pulse was used to
drive serotonin release. 5-HT1A-receptors were pharmacologically isolated with
DNQX (10 µM), MK801 (10 µM), picrotoxin (100 µM), and CGP55845 (200 nM) to block AMPA, NMDA, GABAA, and GABAB receptors, respectively. BAPTA (10 mM) in
the internal pipette solution was used to prevent intracellular calcium signaling. In
traces illustrating electric stimulation, the stimulation artifact was blanked for
presentation. For the iontophoretic application, serotonin (100 mM) was ejected as
a cation (160 nA) for 2000 ms using an Iontophoresis Generator (Dagan) from thin-
walled iontophoretic electrodes placed ~ 30 µm from the cell. A negative retention
current of 15-30 nA was applied to prevent leakage of serotonin.
Materials. Picrotoxin was from Abcam. DNQX, MK-801, CGP 55845, SB 216641 hydrochloride, WAY 100635 maleate, citalopram hydrobromide, R-(-)- phenylephrine hydrochloride, forskolin, DPCPX, and serotonin hydrochloride were from Tocris Bioscience. K-methylsulphate was from Acros Organics. BAPTA was from Invitrogen. D-glucose and HEPES sodium salt were from Sigma-Aldrich.
Dextran (35,000 – 50,000 M.W.) was from MP Biomedicals. All other chemicals
were from Fisher Scientific.
136 Statistics and Analysis. All data are shown as mean ± SEM. Statistical significance (p
< 0.05) was assessed by paired and unpaired Student’s t-tests, ANOVAs, and
Pearson’s correlations as noted (InStat 3.0). Decay kinetics of 5-HT1A IPSCs were fit
with a single exponential using a Simplex algorithm optimized by the sum of
squared errors in Axograph X (Axograph Scientific). Firing rates in cell-attached
experiments were calculated from the average inter-spike intervals.
RESULTS
Evoked 5-HT1A-IPSCs mediate a transient pause in DRN serotonergic neuron
firing.
Voltage clamp recordings were made from eYFP+ serotonergic neurons
located in the dorsomedial DRN in brain slices obtained from ePET1-eYFP mice. A single stimulation was used to elicit 5-HT1A receptor mediated inhibitory post-
synaptic currents (IPSCs). IPSCs were abolished by the 5-HT1A receptor antagonist,
WAY 100635 (200 nM, n = 5, p < 0.01, Student’s paired t-test; Fig. 1A, C), and were
reliably observed in >95% of serotonin neurons (30/31 serotonergic neurons; 85 ±
10 pA, n = 30, Fig. 1B). IPSCs were eliminated by TTX (200 nM, n = 3, p < 0.001,
Student’s paired t-test), calcium-free ACSF (n = 3, p < 0.001, Student’s paired t-test),
and barium (200 µM, n = 4 p < 0.01, Student’s paired t-test, Fig. 1B). Thus, recorded
IPSCs resulted from vesicular serotonin release acting on 5-HT1A receptors to
mediate a potassium conductance, likely via G-protein coupled, inwardly rectifying
potassium (GIRK) channels (Pan et al., 1989). Evoked IPSCs rose and fell within
approximately 1 s. IPSCs activated in 168 ± 7 ms (10-90% rise time, n = 30)
137 following a lag of 75 ± 3 ms (10% onset, n = 30), and decayed with a time constant of
0.44 ± 0.3 sec (n = 30). This resulted in an overall duration of 0.58 ± 0.4 sec
measured at half maximal amplitude (n = 30, Fig. 1C).
To examine the consequence of evoked serotonin release on excitability, cell-
attached recordings of DRN firing rates were made from serotonin neurons. In vivo,
DRN neurons fire in a regular pattern at 0.5 - 3 Hz. To mimic the excitatory
noradrenergic tone that facilitates firing (Baraban and Aghajanian, 1981, Haddjeri et
al., 2004), phenylephrine (3 µM) was added to the bath to maintain regular
pacemaker firing (Vandermaelen and Aghajanian, 1983). Single stimulation evoked
serotonin release induced a 2.56 ± 0.41 second pause in firing (n = 10) that was
blocked by the 5-HT1A receptor antagonist WAY 100635 (200 nM; n = 5; Fig 1D).
Evoked serotonin release did not cause a lasting depression of firing, as basal and
post-pause firing rates were similar (n = 5; p = 0.33; Fig 1D, E). Baseline-firing rates
did not correlate with the duration of the evoked pause (Fig. 1F). Thus, serotonin
release is capable of affecting DRN activity by generating transient pauses in
serotonin neuron firing.
Evoked 5-HT1A IPSC decay occurs by receptor deactivation and not receptor
desensitization.
At many synapses, receptor desensitization can speed the decay of receptor-
mediated currents to shorten the duration of signaling (Trussell et al., 1993). To
determine whether desensitization of 5-HT1A receptors regulates the duration of the
IPSC, a paired-pulse protocol with an interstimulus interval of 4 s was used (Fig 2A).
138 Evoking IPSCs four seconds apart resulted in a 73 ± 4% reduction in the amplitude
of the second IPSC (n = 7; p < 0.001). The paired-pulse depression was partially
attenuated in the presence of the 5-HT1B antagonist, SB 216641 (1 µM; 58 ± 8%
reduction in amplitude of the second IPSC; n = 4; p = 0.02 vs. 1st IPSC; p = 0.04 vs.
reduction in control), confirming that presynaptic activation by 5-HT1B receptors by
the first stimulation inhibited the subsequent release of serotonin (Morikawa et al.,
2000). SB 216641 (1 µM) had no effect on the amplitude of unpaired IPSCs (p = 0.78 amplitude, n = 14), indicating that pre-synaptic 5-HT1B-receptors were not tonically active in slices (Morikawa et al., 2000). To specifically examine post-synaptic depression, 5-HT was exogenously applied using paired iontophoretic applications.
The amplitude of the two resulting currents generated 4 s apart were similar (1st
iontophoretic current: 113 ± 14 pA, 2nd iontophoretic current: 123 ± 16 pA, n = 5; p
= 0.69, Fig. 2C), suggesting that the paired-pulse depression of IPSCs primarily
occurred due to pre-synaptic mechanisms. When serotonin was iontophoretically
applied during the decay phase of the evoked 5-HT1A IPSC (1 second following
stimulation) (Fig 2D), the amplitude of the resulting current was also identical to
currents that were not preceded by an IPSC (during IPSC: 120 ± 21 pA, without
IPSC: 124 ± 21 pA, n = 5, p = 0.89, Fig. 2E). Thus, 5-HT1A receptors were not desensitized during the decay of IPSCs. Because our results indicated that post- synaptic 5-HT1A receptor desensitization did not occur during IPSCs, the duration of
signaling is instead likely regulated by the deactivation of receptors..
139 5-HT1A-receptors are activated by spillover but serotonin clearance is
primarily achieved by reuptake transporters.
Clearance of monoamines in the extracellular space is achieved by a
combination of diffusion and uptake by transporters. The relative contributions of
each vary widely across monoamine synapses (Courtney and Ford, 2014). In the
DRN, it has been hypothesized that due to the limited role of uptake, diffusion of
serotonin may extend the duration of serotonin transmission by allowing activation
of distal extra synaptic 5-HT1A receptors through spillover transmission (Bunin and
Wightman, 1999; Jennings, 2013). To determine the role of serotonin reuptake
transporters (SERTs) in regulating 5-HT1A activation, the selective serotonin
reuptake inhibitor (SSRI) citalopram (Celexa, 200 nM) was bath applied while
recording IPSCs. Citalopram significantly reduced the amplitude of 5-HT1A IPSCs (39
± 8% inhibition, n = 5, p < 0.05, Fig. 3A-C). This was likely due to citalopram- induced tonic activity of pre-synaptic 5-HT1B receptors (Morikawa et al., 2000), as
combining the 5-HT1B antagonist, SB216641 (1 µM), with citalopram instead led to a
stable increase in IPSC amplitude (ctrl: 134 ± 16 pA; citalopram + SB216641: 210 ±
33 pA, n = 5, p = 0.03, Fig. 3A-C). Bath application of citalopram (200 nM) in both
the absence and presence of SB 216641 (1 µM) greatly prolonged the duration of 5-
HT1A IPSCs. Compared to controls, IPSCs recorded in citalopram had delayed peak
times (citalopram: 134 ± 22% increase vs. controls, n = 5, p = 0.003; citalopram +
SB: 189 ± 29% increase, n = 5, p = 0.047; p = 0.292 citalopram vs. citalopram + SB,
Fig. 3D) and were significantly slower to decay ( Decay; citalopram: 963 ± 120%
increase vs. controls, n = 5, p = 0.001; citalopramτ + SB: 830 ± 250% increase, n = 5, p
140 = 0.005; p = 0.6334 citalopram vs. citalopram + SB, Fig. 3D). As a result, citalopram
induced a 5-fold increase IPSC duration (half width) with or without SB 216641 (p =
0.98), indicating that clearance of serotonin clearance by reuptake transporters
limits the duration of transmission.
To examine the role of diffusion in receptor activation, dextran (5 %, 40 kDa)
was added to the bath solution to slow diffusion via macro-molecular overcrowding.
Dextran reduced the amplitude (54 ± 7% reduction; n = 6; p = 0.02), slowed the rate of rise of 5-HT1A IPSCs (38 ± 14% increase in peak time; n = 6; p = 0.02) (Fig. 4A) but
did not alter the decay kinetics of 5-HT1A IPSCs ( Decay; control: 0.60 ± 0.13 sec, 5%
dextran: 0.79 ± 0.08 sec; n = 6; p = 0.31, Student’sτ paired t-test, Fig. 4B). As impairing diffusion limits the escape of serotonin from the synaptic cleft (Min et al.,
1998b), the reduction in amplitude and delay in time to peak in dextran indicates that spillover of serotonin likely led to the activation of extrasynaptic 5-HT1A
receptors during the IPSC (Courtney and Ford, 2014; Ford et al., 2010; Markwardt et
al., 2009; Min et al., 1998b; Nielsen et al., 2004; Szabadics et al., 2007). In addition,
dextran also hinders the diffusion-mediated clearance of transmitter, which can prolong the activation of post-synaptic receptors (Markwardt et al., 2009; Min et al.,
1998b). The lack of a pronounced effect of dextran on IPSC decay suggests that clearance via reuptake, rather than diffusion, is the primary mechanism for receptor deactivation.
Next, we examined the ability of reuptake transporters to limit serotonin diffusion. When in the continuous presence of the selective serotonin reuptake inhibitor (SSRI) citalopram (Celexa, 200 nM) and the pre-synaptic 5-HT1B
141 autoreceptor antagonist SB 216641 (1 µM), bath application of 5% dextran resulted in similar changes in the amplitude (72 ± 6% reduction in 5% dextran, n = 4, p =
0.04)(p = 0.10 vs. change in absence of citalopram) and peak time (44 ± 12% increase in 5% dextran, n=4, p = 0.03; p = 0.75 vs. change in absence of citalopram) of 5-HT1A IPSCs. However, slowing diffusion while reuptake was impaired now significantly extended the duration of IPSCs ( Decay; control: 3.1 ± 0.6 sec, 5% dextran: 10.8 ± 2.9 sec; n = 4; p = 0.04; p = 0.004τ vs. change in absence of citalopram;
Fig. 4C - E). Thus when reuptake is inhibited, slowing diffusion now increased the duration of 5-HT1A receptor activation, suggesting that diffusion became the primary clearance mechanism driving receptor deactivation.
Reuptake transporters prevent pooling and crosstalk from prolonging the duration of IPSCs.
Spillover from synaptic sites can result in extrasynaptic pooling and synaptic crosstalk (Balakrishnan et al., 2009; Courtney and Ford, 2014; Isaacson et al., 1993;
Otis and Mody, 1992). As pooling allows for the duration of signaling to become dependent on the number of active release sites, greater amounts of transmitter release results in longer durations of receptor activation. While pooling likely accounts for the increases in extracellular levels of serotonin that can be detected by electrochemical approaches (Bunin and Wightman, 1998; Jennings, 2013), it is unclear if it plays a physiological role in the activation of post-synaptic 5-HT1A receptors.
142 To determine the contribution of transmitter pooling to evoked serotonin transmission, we initially compared the amplitude and duration of IPSCs recorded from serotonin neurons across the DRN (Fig. 5A). We found that across neurons, decay time did not correlate with the amplitude of evoked IPSCs (n = 30, R2 = 0.017, p = 0.54, Pearson’s correlation, Fig. 5B), suggesting that variations in the amount of transmitter release did not alter the duration of 5-HT1A receptor signaling. For a given cell, we next varied the intensity of stimulation used to evoke IPSCs.
Decreasing the stimulation intensity reduced the amplitude of events (65 ± 2 % reduction, n = 7; p < 0.001; Student’s paired t-test) but again did not alter the rate of decay (n = 7, p = 0.15, Student’s paired t-test, Fig. 5C). Likewise, decreasing the probability of transmitter release by lowering the concentration of extracellular calcium from 2.5 mM to 1.0 mM similarly reduced the amplitude of synaptic currents (n = 8, p < 0.001; Student’s paired t-test) with no effect on the decay time
(n = 8, p = 0.99, Student’s paired t-test, Fig. 5D). Because decreases in serotonin release did not result in shorter duration events, these results suggest that the time course of IPSCs under control conditions are not shaped by pooling of serotonin in the extracellular space.
In other neurotransmitter systems, spillover of transmitter between synapses can be facilitated by blocking uptake (Balakrishnan et al., 2009; Barbour et al., 1994; Courtney and Ford, 2014; Otis et al., 1996; Overstreet and Westbrook,
2003; Silver et al., 1996). As blocking reuptake transporters enabled extended diffusion to prolong 5-HT1A IPSC decay (Fig. 3), we next examined whether blocking reuptake enhanced transmitter crosstalk and pooling. In the presence of citalopram
143 (200 nM) and SB216641 (1 µM), increasing the strength of stimulation not only
increased the amplitude of IPSCs (n = 4, p = 0.01) but also prolonged the time course
of decay (80 ± 26% increase, n = 4, p = 0.006; Fig. 5E). Thus efficient uptake of
serotonin by transporters limited pooling in the extracellular space from extending
the duration of IPSCs. This suggests that the post-synaptic activation of 5-HT1A receptors may result from the local release of serotonin, rather than extended diffusion away from sites of release.
Spontaneous IPSCs occur with similar kinetics to evoked.
While making long recordings from serotonergic neurons in the DRN, we observed spontaneous IPSCs (sIPSCs) that occurred in 16/38 of neurons (42%; Fig
6A). The kinetics of sIPSCs were similar to evoked 5-HT1A IPSCs (peak time: 346 ±
23 ms; decay: 0.37 ± 0.03 sec, n = 27; Fig. 6B, D) and were prolonged in the presence
of citalopramτ (200 nM, n = 10; p < 0.001; Fig. 6C, D). The low frequency of events
that could be detected (0.2 ± 0.1 events per 10 minutes, n = 38 neurons) limited
quantitative analysis of the underlying mechanisms of individual events. To
increase the ability to detect events, phenylephrine (3 µM), forskolin (1 µM), DPCPX
(500 nM), and citalopram (50 nM) were included in the bath solution. This
pharmacological cocktail increased the frequency of spontaneous events (1.2 ± 0.3
events per 10 minutes, n = 15 neurons) with spontaneous events being observed in
13/15 neurons (87%; Fig. 6E). The apparent increase in spontaneous event
frequency may be in part due to increased amplitude of resolvable events (28 ± 3%
increase, n = 21 sIPSCs, p < 0.05; Fig. 6F). Bath application of the 5-HT1A antagonist
144 WAY 100635 (200 nM) eliminated sIPSCs (8/8 cells) and sIPSCs were never
observed in recordings made in the continuous presence of WAY 100635 (200 nM, 9
cells; Fig 6E), indicating that spontaneous events were driven by 5-HT1A receptor
activation. While it is unknown whether evoked and spontaneous IPSCs arose from
similar or separate release sites, the existence of sIPSCs implies that serotonin
release from at least some pre-synaptic release sites may be able to independently
activate post-synaptic receptors.
DISCUSSION
In the present study, we examined the mechanisms regulating serotonergic
transmission by 5-HT1A receptors in the dorsal raphe. We found that stimulation
evoked serotonin release generated 5-HT1A-receptor mediate inhibitory currents
that rose and fell within a second and paused basal firing without causing long-
lasting depressions to firing rates. The transient nature of 5-HT1A-receptor-
mediated transmission was primarily driven by transmitter clearance via serotonin
reuptake transporters and the ensuing deactivation of 5-HT1A receptors. Reuptake
transporters limited the spillover activation of distal, extrasynaptic receptors and
prevented serotonin pooling in the extracellular space from prolonging the duration
of 5-HT1A-receptor activity via synaptic crosstalk. Taken together, these results
suggest that 5-HT1A receptor-mediated serotonin transmission in the DRN occurs through functionally independent synapses despite the observation of a limited amount of spillover transmission.
145 Local feedback inhibition by inhibitory autoreceptors regulates cellular excitability at multiple monoamine synapses. In midbrain dopamine neurons, evoked and spontaneous vesicular dopamine release activates D2-receptor mediated IPSCs through GIRK channels (Beckstead et al., 2004; Gantz et al., 2013).
Like axonal synapses in the striatum (Marcott et al., 2014), somatodendritic transmission is mediated by a high concentration of dopamine (Ford et al., 2009) that in the midbrain occurs in the absence of transmitter pooling and spillover
(Courtney and Ford, 2014; Ford et al., 2010). This differs from somatodendritic transmission of noradrenaline in the locus coeruleus where less efficient uptake allows for low concentrations of noradrenaline to pool in the extracellular space when multiple inputs are synchronously active (Courtney and Ford, 2014). The lack of dopamine pooling in the ventral tegmental prevents crosstalk between synaptic sites and maintains independence between release sites (Courtney and
Ford, 2014).
In this study, we also found that the local release of serotonin in the DRN can activate synaptic 5-HT1A receptors in the absence of pooling. This was likely true for both evoked and spontaneous release due to the similarity in the rise and decay kinetics. Unlike dopamine transmission, however, spillover allowed for the activation of some pool of extrasynaptic receptors. 5-HT1A IPSCs were more variable cell to cell in their duration, and thus time course of inhibition, than was previously observed for midbrain dopamine transmission (Courtney and Ford,
2014). This increased variability may be due to the recruitment of extrasynaptic receptors in serotonin signaling. The lack of pooling at synaptic sites was attributed
146 to efficient uptake by SERT. Like somatodendritic dopamine transmission
(Courtney and Ford, 2014), blocking uptake extended the time course of IPSCs such that the duration of transmission was now dependent upon the amount of serotonin released. How serotonin reuptake transporters both allowed for the spillover activation of extrasynaptic receptors yet prevented pooling and synaptic crosstalk remains unclear. One possibility may be that a non-uniform distribution of SERTs in
DRN neurons around different release sites (Colgan et al., 2012) allows serotonin to spillover into the extrasynaptic space yet limits pooling between synapses.
Throughout these recordings we observed spontaneous IPSCs in roughly half of the neurons examined. The similarity in kinetics between these IPSCs and the finding that no events could be detected in the presence of 5-HT1A antagonists suggests that these IPSCs could be the result of spontaneous release of serotonin leading to the activation of 5-HT1A autoreceptors. Similar spontaneous release of dopamine as has been recently described at synapses in the substantia nigra (Gantz et al., 2013). It is unclear if the same pool of serotonergic vesicles mediates both evoked and spontaneous events. At ionotropic synapses, different populations of vesicles have been proposed to underlie spontaneous and evoked release (Kavalali,
2015; Ramirez and Kavalali, 2011). In the presence of phenylephrine which increases DRN firing to 2-3 Hz, the frequency of observable events was only once per ~ 10 min. While many events might be below the level of detection in our experiments, the low frequency of events suggests that these events did not result from background firing of DRN neurons. One possibility may be that these events
147 could arise from dendritic sites, where serotonin release occurs in an action-
potential independent manner (Colgan et al., 2012).
Vesicular serotonin release in the DRN occurs at axonal (Bruns et al., 2000),
somatic (Colgan et al., 2009; Kaushalya et al., 2008), and dendritic sites (Colgan et
al., 2012; De Kock et al., 2006). As evoked serotonin release underlying 5-HT1A
IPSCs is modulated by 5-HT1B receptors (Fig. 2) (Morikawa et al., 2000), which are located only on axon terminals (Sari, 2004), axonal release likely contributes to the activation of the synaptic receptors that underlie the IPSC. It is not clear whether these axons originate locally from within the DRN or arise from other brain serotoninergic nuclei (Andrade et al., 2015; Bang et al., 2012). Cell body release sites lack defined pre-synaptic active zones (Colgan et al., 2009) making it unclear
the extent to which somatic release participates in evoked IPSCs. While axonal and
somatic transmission are dependent on action potentials (Bruns et al., 2000; Colgan
et al., 2012), dendritic release is instead impulse-independent relying on local
NMDA receptors and L-type calcium channels (Colgan et al., 2012; De Kock et al.,
2006). Glutamate receptors were blocked in our experiments, making it less likely
that dendritic release sites contributed to evoked 5-HT1A IPSCs.
Recent studies have linked both tonic and transient inhibitory regulation of
DRN serotonergic neurons to behaviors such as reward encoding (Cohen et al.,
2015; Ranade and Mainen, 2009). During these behaviors, tonic and transient
inhibition independently influenced DRN neurons and could be used to identify
neuronal subpopulations (Cohen et al., 2015). There is a growing consensus that
DRN serotonergic neurons are non-homogeneous, as heterogeneity of these neurons
148 has already been suggested by anatomical, biochemical, and electrophysiological
properties (Abrams et al., 2004; Andrade and Haj-Dahmane, 2013; Calizo et al.,
2011; Marinelli et al., 2004; Vasudeva et al., 2011). Subpopulations of serotonin neurons, either within the DRN or between various raphe nuclei, are hypothesized to be interconnected, and form complex microcircuits (Altieri et al., 2013; Bang et al., 2012; Gaspar and Lillesaar, 2012). Serotonergic neurons send rich networks of recurrent axon collaterals that can span long distances inside of the DRN, often skipping their nearest neighbors to innervate more distant targets (Altieri et al.,
2013). By signaling through independent, 5-HT1A-receptor mediated synapses, these rich networks can ensure both spatial and temporal precision in how they encode mood and behavior.
149
Figure 4.1: Electrical stimulation evokes 5-HT1A IPSCs that drive pauses in
DRN serotonin neuron firing. A. Left: Averaged 5-HT1A IPSCs from a dorsal raphe
neuron evoked by a single stimulation in control conditions (black) and in the
presence of the 5-HT1A antagonist WAY 100635 (200 nM; grey). Whole-cell
recordings were voltage-clamped to -60 mV. Right: IPSCs could be evoked in 30/31
DRN serotoninergic neurons examined. B. Average traces (left) and quantification
(right) of IPSCs recorded in 200 nM TTX, calcium-free ASCF, and 200 µM barium. C.
Quantification of the rise and decay kinetics of evoked IPSCs. D. Cell-attached
recordings made in 3 µM phenylephrine to mimic in vivo excitatory drive.
Representative traces (left) and quantification (right) demonstrating that evoked
serotonin release drives a pause in firing that is absent in the 5-HT1A-receptor antagonist WAY 100635 (200 nM). E. Firing rates measured over 5 seconds before stimulation (basal) and over 5 seconds after firing had resumed (post-pause) were identical. F. Pearson correlation demonstrating that the duration of the evoked pause was independent of the baseline firing rates (n = 9). n.s. represents p > 0.05,
*** p < 0.001; error bars represent ± s.e.m.
150
Figure 4.1
151 Figure 4.2: 5-HT1A-receptor desensitization does not determine the decay time of IPSCs. A. Example trace demonstrating a paired-pulse depression when stimulating at a 4 second interval. B. Example trace demonstrating the lack of post- synaptic depression when serotonin was applied by iontophoresis twice at a 4 second interval (Ionto.). C. Quantification of amplitude ratio of evoked IPSCs in absence and presence of the 5-HT1A pre-synaptic receptor antagonist SB 216641 (1
µM) or the current generated by the iontophoresis of serotonin at a 4 second interval. D. Representative trace. Serotonin was applied by iontophoresis during the decay of evoked IPSCs (1 second post stimulation; Ionto. 1) and without preceding IPSCs (12 seconds post-stimulation; Ionto. 2). E. Quantification demonstrating the similar amplitude of currents generated by serotonin iontophoresis in D. * represents p < 0.05; error bars represent ± s.e.m.
152
Figure 4.2
153 Figure 4.3: Serotonin reuptake transporters limit the duration of IPSCs. A.
Average 5-HT1A IPSCs under control conditions and in the presence of citalopram
(200 nM) and citalopram (200 nM) + SB216641 (1 µM), illustrating that blocking
serotonin uptake increased the activation of 5-HT1A receptors and led to the tonic activation of 5-HT1B receptors. B. Quantification of the amplitude. C. Time course of
IPSC amplitude during the application of either citalopram (200 nM) or citalopram
and SB 216641 (1 µm). D. Quantification of the change in kinetics in either
citalopram or citalopram + SB 216641. * represents p < 0.05, ** p < 0.01; error bars
represent ± s.e.m.
154
Figure 4.3
155 Figure 4.4: Slowing diffusion of serotonin alters the amplitude and kinetics of
5-HT1A-IPSCs. A. Average traces of IPSCs recorded before (black) and during
(green) the bath application of 5% dextran. B. Quantification of the amplitude, peak time, and decay kinetics of IPSCs recorded in A. C. Average traces of IPSCs before
(black) and during (green) the bath application of 5% dextran recorded in the continuous presence of citalo M). When serotonin reuptake transporterspram and (200 pre nM)-synaptic and SB autoreceptors 216641 (1 μ were inhibited, slowing diffusion greatly prolonged the decay of 5-HT1A IPSCs. D. Quantification of the amplitude, peak time, and decay kinetics of IPSCs recorded in E. Relative change in IPSCs in the presence of dextran with and without citalopram (200 nM). n.s. represents p > 0.05, * p < 0.05, ** p < 0.01; error bars represent ± s.e.m.
156
Figure 4.4
157 Figure 4.5: Reuptake transporters prevent transmitter pooling and synaptic
crosstalk from prolonging the decay time of 5-HT1A IPSCs. A. Example IPSCs recorded from 4 neurons demonstrating the variation in amplitude and duration. B.
Lack of correlation between decay kinetics and amplitude of 5-HT1A IPSCs in 30 DRN
neurons from Fig. 1. C. Example IPSCs evoked with different stimulation intensities
(low: 10- - A). Inset show the traces normalized and aligned to
their peak,20 illustrating μA; high: 60 that80 responses μ to lower stimuli had similar rates of decay.
Right: quantification of amplitudes and decay times between conditions. D. Example
IPSCs evoked in 2.5 mM and 1.0 mM extracellular Ca2+. Inset show the traces
normalized and aligned to their peak, illustrating that responses in reduced calcium
had similar rates of decay. Right: quantification of amplitudes and decay times
between conditions. Scale bar (for D and E): 30 pA; 500 ms. n.s. represents p > 0.05.
E. Example traces (left) and quantification (right) of IPSCs in the presence of
SB216641 (1 µM) + citalopram (200 nM) evoked with low (10 – 20 µA) and high
(60-80 µA) stimulation intensities. Lower trace shows the recordings normalized, illustrating that in the presence of citalopram, larger stimuli prolong the decay time of 5-HT1A IPSCs. n.s. represents p > 0.05, ** p < 0.01, *** p < 0.001; error bars
represent ± s.e.m.
158
Figure 4.5
159 Figure 4.6: Spontaneous 5-HT1A–receptor mediated currents have similar
kinetics as evoked IPSCs. A. Representative trace of evoked and spontaneous
IPSCs. B. Average spontaneous IPSCs (sIPSC, left) overlaid with average sIPSCs
recorded in citalopram (200 nM, center) and average evoked 5-HT1A IPSCs (right).
C. Quantification of the amplitudes of sIPSCs recorded in control bath and citalopram vs. evoked IPSCs. (ANOVA, p < 0.001) D. Quantification of the kinetics of the spontaneous IPSCs recorded in control bath and citalopram (200 nM) compared to evoked 5-HT1A IPSCs (ANOVA: peak time p < 0.001, Decay p < 0.001). sIPSCs
recorded in control bath had similar kinetics to evokedτ IPSCs. E. Inclusion of
phenylephrine (3 µM), forskolin (1 µM), DPCPX (500 nM), and citalopram (50 nM) in
the bath increased the frequency of sIPSCs (pharm.). Further addition of WAY
100635 (200 nM) abolished sIPSCs (ANOVA p < 0.001). F. Histogram demonstrating
that sIPSC had significantly greater amplitudes (p = 0.04; n = 21 pharm.; n = 27 ctrl.) in the pharmacological bath (from E) compared to control bath. * represents p <
0.05; *** represents p < 0.001; n.s. represents p > 0.05
160
Figure 4.6
161 CHAPTER 5
DISCUSSION
162 This thesis tested the volume transmission hypothesis proposed for feedback
inhibition mediated by dopamine autoreceptors in the midbrain, noradrenaline
autoreceptors in the LC, and serotonin autoreceptors in the DRN. The experiments
tested the contributions of spillover, extrasynaptic pooling, and transporter-
mediated clearance in shaping autoreceptor activity and the resulting inhibition.
Perhaps the most significant finding of this thesis work is that, despite similar
ultrastructure and electrochemical profiles, the mechanisms underlying
autoreceptor activation are substantially different between central monoamines.
Dopamine transmission occurred in a point-to-point fashion in the absence of
transmitter spillover or pooling resulting in a highly reproducible time course of
inhibition. While serotonin spillover activated some extrasynaptic autoreceptors,
the lack of pooling and synaptic crosstalk still resulted in point-to-point serotonin
transmission. Finally, noradrenaline spillover resulted in extrasynaptic pooling to
extend autoreceptor activity and prolong the time course of inhibition. Knowing
that monoamine autoreceptor activation is governed by different transmission
mechanisms demonstrates that GPCR-mediated transmission, in general, is not
confined to loosely regulated paracrine mechanisms and that point-to-point transmission similar to classical synapses is possible. Thus, each instance of GPCR- mediated transmission must be individually evaluated for its properties and synaptic mechanisms and it cannot be assuming that canonical paracrine mechanisms govern all metabotropic transmission.
This chapter will review the results and implications of the previous three chapters to address general issues and remaining questions.
163 The Importance of Inhibitory Timing and the Activity of Monoamine Neurons
The primary meaningful output of any neuron is neurotransmitter release
that successfully influences a neuron or other effector through the activation of
receptors. From this perspective, the ultimate consequence of excitatory inputs
onto neurons can be evaluated by asking whether transmitter release, action
potential driven or otherwise, resulted because of those excitatory inputs.
Determining whether inhibitory inputs had meaningfully consequences, however,
requires a more nuanced evaluation that instead asks whether an action potential or
release event that otherwise would have occurred was prevented by the inhibitory
transmission. Because of this, a thorough understanding of the mechanisms that
shape the duration and variability of inhibitory transmission is essential to
elucidating its role in the context of neuronal networks. However, while this thesis
largely focused on the implications of paracrine vs. point-to-point transmission on
the timing of monoamine autoreceptor-mediated transmission, spatial selectivity
and whether information can be transmitted exclusively between paired pre- and
post-synaptic neurons is also important. This could be especially important in
respect to monoamine heteroreceptors or other metabotropic receptors that do not
couple to GIRK channels and instead modulate neurons by slower signaling cascades. Regulating the duration of receptor activity on the order of seconds may or may not have large net effects on the timing of effector signaling when considering the slow, GPCR-mediated signaling cascades that can affect targets for minutes or longer. In contrast, limiting which neurons receive that signaling and
164 which neurons do not could easily have profound consequences for network
functions.
The absence of spillover and pooling in D2-autoreceptor activation results in a time course of inhibition that is both consistent cell-to-cell and unaffected by the strength of stimulation eliciting dopamine release. In vivo, burst firing of dopamine neurons (Bunney et al., 1973; Grace and Bunney, 1983, 1984) driven by excitatory inputs from the subthalamic or pedunculopontine nucleus (Overton and Clark,
1997; Smith and Grace, 1992) facilitates dopamine release locally in the midbrain and globally in projection regions. These bursts are thought to be particularly important in reward-dependent learning (Schultz et al., 1997). Bursts are comprised of 2 - 10 action potentials fired at ~10 Hz (Grace and Bunney, 1984) and induce a pause in the tonic firing of midbrain dopamine neurons (Paladini and
Roeper, 2014). Although additional action potentials in a burst delay the peak time of GIRK-mediated currents, the lack of spillover and pooling in midbrain dopamine transmission results in a post-burst pause with a consistent time course of receptor activation that is independent of the number of action potentials in the burst. This suggests that bursts of dopamine neuron activity with greater numbers of action potentials could potentially serve to increase dopamine release in terminal regions without substantially altering the timing of autoreceptor-mediated inhibition in midbrain dopamine neurons.
Similar to dopamine neurons, noradrenaline neurons of the locus coeruleus can also discharge bursts of activity (Aston-Jones and Bloom, 1981) that enhance noradrenaline release both locally and in terminal regions (Florin-Lechner et al.,
165 1996). In vivo, these bursts of activity are typically evoked by an arousing stimulus
(Florin-Lechner et al., 1996), generate a pause in the firing of locus coeruleus
neurons, and are thought to be important in resetting the activity of LC neurons to
lower baseline firing rates (Aston-Jones and Cohen, 2005b). Unlike burst firing in dopamine neurons, our results demonstrate that bursts prolong the duration of auto-receptor mediated inhibition in the locus coeruleus due to transmitter spillover and resulting pooling. Thus, longer bursts enhance noradrenaline release in terminal regions and generate longer windows of inhibition in locus coeruleus activity. The extended duration of local inhibition in response to bursts of activity may prove important mechanistically in how bursts reset the activity of LC neurons.
Despite a similar lack of pooling and crosstalk, the duration of 5-HT1A-
autoreceptor mediated IPSCs was significantly more variable from cell-to-cell when
compared to dopamine signaling. To account for this variability, one could
hypothesize that the duration of inhibitory serotonergic signaling may be tuned for
individual neurons. This could be accomplished either by altering the extrasynaptic
expression of 5-HT1A receptors and thus the contribution of spillover in extending
signaling duration, or potentially by modulating intracellular mechanisms such as
RGS expression. While both of these speculations are yet untested, individual tuning
of the duration of signaling would be consistent with the developing theme of
heterogeneity in DRN serotonin neurons (Abrams et al., 2004; Andrade and Haj-
Dahmane, 2013). Burst firing of DRN serotonin neurons has only recently been
described (Hajós et al., 2007) and little is known about the physiological drivers or
relevance of bursting. Our results suggest that, like dopamine transmission,
166 increased serotonin release driven by bursts would not have a substantial impact on
the time course of 5-HT1A-autoreceptor mediated inhibition in DRN serotonin
neurons due to the lack of transmitter pooling during transmission.
Synchronous verse Asynchronous Transmitter Release.
During the vast majority of the experiments presented in chapters 2 through
4, electrical stimulation was used to drive transmitter release. This stimulation
protocol synchronously activates release sites in a fashion that does not always
reflect normal physiology.
Dopamine neurons in the midbrain are intrinsically active and tonically fire
at a rate of 1 – 2 Hz. During resting conditions, only a small minority of neurons
display coordinated firing activity (Li et al., 2011) suggesting that the pacemaker
firing of dopamine neurons is primarily asynchronous at baseline. Several
conditions are known to increase the synchrony of dopamine neuron firing,
including nicotine administration (Li et al., 2011) and excitatory input-driven burst firing (Canavier and Landry, 2006; Komendantov and Canavier, 2002; Lodge and
Grace, 2005). In our recordings, synchronous dopamine release was evoked on top of a background of intrinsic, likely asynchronous dopamine neuron pacemaker firing. Application of the D2-receptor antagonist sulpiride did not significantly change the resting baseline current, suggesting that the pacemaker activity of DA neurons was insufficient to generate tonic post-synaptic receptor activity. Whether pacemaker activity has any effect on dopamine neuron excitability remains
167 unknown. Nonetheless, the modulation of firing patterns by evoked activity, as
shown in chapters 2 and 3, may be reflective of signaling that occurs when
synchrony is behaviorally or pharmacologically induced. Even in the absence of
synchronous release from multiple sites, the point-to-point transmission displayed
by the dopamine system suggests that it should be possible to make paired
recording of dopamine neurons whereby generating an action potential in one
neuron causes D2-receptor activation in the paired neuron. While this is potentially
supported by the observation of spontaneous D2-receptor mediated currents (Gantz et al., 2013), paired dopamine neurons have not be observed at the time of this writing.
Similar to dopamine neurons, noradrenergic neurons of the locus coeruleus are also intrinsically active and fire between 0.5 - 2 Hz. Synchrony between the firing of multiple noradrenergic neurons is induced in vivo by various stimuli,
especially during bursts of activity presumably driven by common excitatory inputs
(Finlayson and Marshall, 1988). Furthermore, the application of cocaine or
morphine decreases overall firing rates while inducing synchronous firing in
noradrenergic LC neurons (Harris et al., 1992; Zhu and Zhou, 2001), with cocaine-
induced synchrony being dependent on α2-receptor activity (Harris et al., 1992).
Finally, slower tonic firing rates in general increase the synchrony of firing due
partially to electrotonic coupling (Alvarez et al., 2002). The experiments presented
in chapter 3 resulted from the synchronous activation of noradrenaline release sites
and they are likely reflective of noradrenaline signaling during input-induced
bursting or in pharmacologically induced synchronous firing. However, it is unclear
168 how the mechanisms demonstrated to underlie LC noradrenaline transmission
might function during the more asynchronous resting state. LC neurons were
unique among the studied monoamine neurons in that they did not display
spontaneous autoreceptor currents. Thus, it is possible that noradrenaline release
from a single neuron is insufficient to generate a resolvable post-synaptic
consequence. This suggests that some degree of synchronization of release or obligate pooling of noradrenaline might be required to generate phasic pauses during noradrenaline feedback inhibition.
Synchrony has been less studied in the serotonin neurons of the dorsal raphe both during resting conditions and in response to stimuli. However, the point-to-
point nature of transmission identified in chapter 4 of this thesis results in a time
course of inhibitory transmission that is independent of the number of
synchronously activated release sites. During some experiments in chapter 4,
serotonin release was successfully evoked on top of a background of tonic firing
induced by α1-receptor activation similar to that which occurs in vivo (Baraban and
Aghajanian, 1981; Haddjeri et al., 2004). Thus, evoked synchronous serotonin
discharge can modulate firing patterns and excitability on top of background firing
which is presumably asynchronous. Whether these two modes of release both
shape DRN activity and when, physiologically, one type would become more
influential remains to be examined. Chapter 4 demonstrated spontaneous 5-HT1A
receptor-mediated currents similar to what has been observed in dopamine
neurons (Gantz et al., 2013). While the pre-synaptic source of these events remains
unclear, their existence suggests that it might be possible to find pairs of connected
169 serotonin neurons. If this were to prove feasible, performing paired recordings could illustrate the interconnectedness of serotonin regions that may or may not reflect regional heterogeneity.
Reuptake Transporters Actively Shape Monoamine Transmission
The role of monoaminergic reuptake transporters in shaping monoaminergic transmission is debated. In the case of midbrain dopamine transmission, electrochemical and modeling studies have concluded that reuptake transporters do not meaningfully impact extracellular dopamine transients until dopamine has traveled nearly 10 µm away from release sites (Cragg and Rice, 2004; Rice and
Cragg, 2008). By modeling receptor activation based on EC50 values, these studies conclude that transporter-mediated reuptake has little to no bearing on transmission (Cragg and Rice, 2004; Rice and Cragg, 2008). This conclusion is thought to be supported by the relatively slow turnover rate of dopamine transporters (Povlock and Schenk, 1997), which is interpreted to mean that dopamine transporters buffer dopamine during the time scale of transmission and primarily function to slowly clear it from the bulk extracellular space (Cragg and
Greenfield, 1997; Cragg et al., 2001; Rice, 2000). This differs from dopamine release in the striatum, where greater DAT expression can strongly regulate extracellular dopamine levels and thus presumably dopamine transmission (Floresco et al., 2003;
Giros et al., 1996; Stamford et al., 1988).
170 Previous studies of evoked dopamine-mediated GIRK currents have
contested this passive role of dopamine reuptake transporters in shaping midbrain
dopamine transmission. Inhibiting reuptake transporters while evoking dopamine
release results in GIRK-mediated currents that are significantly larger in amplitude
and longer in duration (Beckstead et al., 2004; Ford et al., 2009) and lower doses of
DAT antagonists preferentially increase the amplitude of evoked transmission
without significantly altering the time course (Ford et al., 2009). Thus while some
mechanistic ambiguities remain, reuptake transporters likely gate or restrict the
activation D2-autoreceptors in a manner not suggested by electrochemical studies.
In chapter 3 of this thesis, the role of dopamine reuptake transporters was
further examined. Because evidence for transmitter spillover, pooling, and synaptic
crosstalk was present only when reuptake transporters were inhibited, dopamine
reuptake transporters prevent the spillover activation of extrasynaptic D2-
receptors and maintain the point-to-point fidelity of transmission sites.
Understanding this role of reuptake transporters may have significant consequences for a mechanistic understanding of the effects of psychostimulants. While the application of psychostimulants such as cocaine and amphetamine undoubtedly increases the amount of dopamine present in the extracellular space, these drugs may also disrupt the point-to-point nature of communication and the temporal precision of midbrain dopamine signaling by allowing for transmitter pooling.
Whether or not this significantly alters the acute actions of psychostimulants or their longer-term effects leading to addiction remains unknown.
171 In chapter 4, serotonin reuptake transporters were found to fulfill a role
somewhat similar to that of dopamine reuptake transporters. Though serotonin
reuptake transporters did not entirely prevent the spillover activation of
extrasynaptic receptors under control conditions, they isolated synaptic
transmission sites by preventing synaptic crosstalk. Inhibition of serotonin
reuptake by selective serotonin reuptake transporter inhibitors (SSRIs) allowed for
pooling and resulted in the duration of evoked inhibition being dependent on the
strength of stimulation. Serotonin reuptake transporters are thought to be the
primary effector behind the actions of many anti-depressant drug categories. As
such, it is unclear whether or how induced serotonin pooling and resulting synaptic
crosstalk may contribute to their effectiveness.
Implications for Monoamine Transmission in Terminal Regions
In this thesis, the mechanisms governing how locally released monoamine
neurotransmitters activated somatodendritic autoreceptors could be examined in
part due to the relatively fast coupling between autoreceptors and GIRK channels
present in nuclei regions. However, a lack of GIRK channel expression in post-
synaptic neurons in projection regions renders it more difficult to mechanistically
study how terminally released monoamines activate heteroreceptors located on
non-monoaminergic post-synaptic neurons. While it cannot be merely assumed that
the mechanisms underlying local monoamine transmission are fully conserved in
terminal regions, this thesis works at least provides a basis to hypothesizes that 1)
172 terminal release of different monoamines may occur by distinct mechanisms and 2)
terminal release of some monoamines, such as dopamine, may be highly regulated
and more analogous to point-to-point synaptic transmission.
These hypotheses seem to be supported by the very distinct differences in
the anatomy of noradrenaline and dopamine terminals. Noradrenaline axons are
known to form varicosities (Beaudet and Descarries, 1978; Farb et al., 2010;
Marotte and Raisman, 1974) that contain fully functional release machinery while
lacking any clearly apposed post-synaptic contacts. If point-to-point contacts and synapses are unnecessary for the function of terminally released noradrenaline, these varicosities may be an adaptation that is well suited for long ranging, loosely regulated transmission similar the noradrenaline transmission observed within the locus coeruleus. Furthermore, by centralizing noradrenaline release to these varicosities, plasticity or regulation by other neurotransmitters might be enabled to efficiently affect noradrenaline release region-wide by acting only at these varicosities instead of having to individually modulate many separate release sites.
In contrast to the varicosities observed in noradrenaline axons, many of the dopamine terminals in the striatum form contacts onto the necks of dendritic spines
(Descarries et al., 1996; Freund et al., 1984). This arrangement is commonplace for inhibitory neurotransmission, as it enables inhibitory inputs on spine necks to selectively modulate and dampen only the excitatory inputs that synapse onto the head of that same spine. Having point-to-point transmission for terminal dopamine release would then similarly allow dopaminergic inputs to selectively modulate excitatory inputs in a spine-specific manner. In addition, dopamine can be released
173 from selective terminals by nicotinic acetylcholine receptors located on axons
independent of impulse activity from the midbrain (Giorguieff-Chesselet et al.,
1979). Together, this evidence leads to a speculative system of striatal dopamine transmission in which synaptic dopamine transmission onto spine necks may
selectively attenuate paired inputs in a spine-specific manner. Propagating action potentials from the midbrain may allow for dopaminergic modulation of many dendritic spines and their paired inputs due to the highly branching nature of dopamine axons. Release of dopamine driven by terminal nicotinic receptors, in contrast, could allow for the modulation of a different population of striatal spines
depending on the mechanisms of acetylcholine transmission and the arborization of
cholinergic interneurons. From a circuit perspective, this speculative system of
dopamine transmission in the striatum, and perhaps other terminal regions, is far
more complex than the previously hypothesized mechanisms of paracrine
transmission, capable of encoding far more nuanced information, and if supported
by other studies, may be consequential for understanding how dopamine encodes the prediction of rewards and other behaviors. In support of this hypothesis, a study published during the course of my thesis work, in which GIRK channels were virally expressed in striatal neurons to assess dopamine receptor activity in response to evoked dopamine release from axon terminals, concluded that D2- receptors on striatal neurons could encode dopamine release on a phasic, second to sub-second timescale (Marcott et al., 2014). These D2-heteroreceptors were also likely exposed to >10 µM dopamine during evoked transmission (Marcott et al.,
2014). While this study did not test whether this transmission occurs by
174 independent, point-to-point synapses, it at least suggests that dopamine transmission in the striatum, similar to transmission in the midbrain, can occur with
a tightly regulated time course. Furthermore, the high concentrations of dopamine
found to activate receptors suggest that the synaptic distances between dopamine axon terminals and D2-heteroreceptors may be shorter than previously believed.
While further experiments testing the contributions of spillover and pooling to
striatal D2-receptor activation are necessary to assess the independence of
dopamine inputs, this study nonetheless supports that dopamine modulation of
striatal neurons may be far more complex than simple paracrine transmission and
capable of modulating individual dendritic spines and their associated inputs.
Future Directions
Many questions were raised during the course of this thesis work that remain unanswered. First, what is the anatomical origin of the monoamine release evoked during the course of these experiments? While dopamine release in the SNc is thought to exclusively occur at somatodendritic sites, release of monoamines in the LC, VTA, and DRN contains elements of both somatodendritic and axonal release.
Defining the relative contributions of release from somatodendritic sites, axon collaterals, and projection axons from other monoamine nuclei would contribute to an understanding of the overall circuitry of central monoamines. Axonal release in
these three areas may originate either locally (axon collaterals) or as projections
from other monoaminergic nuclei. The ratio of locally originating innervation
175 compared to innervation by projection axons from other monoamine nuclei regions
could be tested using optogenetics. Expressing channel rhodopsin in select nuclei
regions, targeted by viral injection, would isolate potential input sources and allow
for the relative contributions of each source, in relation to autoreceptor activation, to be assessed. Furthermore, in the case of point-to-point transmission in the DRN and VTA, it would be interesting to examine whether all neurons received equal inputs from potential innervation sources or whether there was organization of inputs such that certain neurons preferentially encoded monoamine release only from certain sources. This is quite possibly the case at least for serotonin neurons, where studies have demonstrated heterogeneity in serotonin neurons and anatomical evidence for serotonergic microcircuits (Altieri et al., 2013; Bang et al.,
2012; Gaspar and Lillesaar, 2012). Knowing these details would help expand the metaphorical circuit diagram of monoamine signaling and could provide insights into how overall central monoamine systems function.
Another potential future direction is to develop a more sophisticated model of transmission for dopamine, noradrenaline, and serotonin. A few of the shortcomings of the model presented in chapter 3 are: it is unable to handle the removal of transmitter by transporters in an elegant fashion; it cannot account for multiple release events; it lacks the ability to distribute receptors and transporters unevenly in space; and it does not account for receptor activation beyond what occurs at the transmitter concentrations necessary to mimic the initial rise kinetics of the recorded IPSCs. To overcome these limitations, I expect that a new modeling approach based on computationally solving a system of differential equations and
176 simulated diffusion, reuptake, and receptor binding would be required. A more
elegant model could predict the differences in configuration of reuptake
transporters that allow for pooling in noradrenaline transmission and prevent it in
dopamine transmission. Furthermore, the spread of noradrenaline and how far it is
able to interact with receptors beyond those encoding the initial rise of IPSCs may
be answerable by this model. Finally, this model may even predict the minimal
spatial separation between sites of dopamine transmission required for their
synaptic independence.
The point-to-point nature of transmission in the dopamine and serotonin
system suggests that there must be some overarching architecture or organization
of transmission sites. How these transmission sites are shaped and maintained at
the ultrastructural level would be interesting to pursue. For example, why do some
D2-receptors respond to synaptic dopamine while others are likely located extrasynaptically? At ionotropic synapses, glutamate receptors are often held in synaptic locations by scaffolding proteins with membrane circulating, extrasynaptic
receptors providing a reserve pool for plasticity-induced changes to synaptic strength. Thus it is possible, and perhaps even likely, that dopamine and serotonin receptors could have intracellular scaffolding proteins that organize receptors to synaptic locations. Identifying what these scaffolding proteins are could have implications for synaptic organization, plasticity, or even possible dysfunctions in
dopamine signaling.
Finally, the highly branching nature of dopamine axons in the striatum and
the point-to-point nature of dopamine transmission, at least in the midbrain, form
177 an interesting dichotomy of purposes. If dopamine transmission in the striatum occurs by similar mechanisms to midbrain transmission, point-to-point synapses are meant to confine information flow to select linear pathways whereas highly branching axons leading to widespread release sites are oppositely intended to spread similar information over broad areas. This dichotomy may indicate that midbrain dopamine neurons modulate widespread, yet still selective, post-synaptic targets to accomplish their functions. This architecture could suggest that the widespread post-synaptic spines or neurons receiving common innervation from a
single midbrain dopamine neuron may have a common feature or purpose in relation to their circuit function or behavioral role. This common purpose may be that these post-synaptic neurons have similar projection targets of their own or that the inputs onto individual, dopamine-modulated spines many originate from common sources. While this idea would without doubt be very difficult to study, it could significantly advance our understanding of the role of individual dopamine neurons and the overall circuitry of the dopamine system in behaviors such as reward prediction.
178 REFERENCES
Abrams, J.K., Johnson, P.L., Hollis, J.H., and Lowry, C.A. (2004). Anatomic and
functional topography of the dorsal raphe nucleus. Ann. N. Y. Acad. Sci. 1018, 46–57.
Adell, A., Sarna, G.S., Hutson, P.H., and Curzon, G. (1989). An in vivo dialysis and behavioural study of the release of 5-HT by p-chloroamphetamine in reserpine- treated rats. Br. J. Pharmacol. 97, 206–212.
Adell, A., Celada, P., Abellán, M.T., and Artigas, F. (2002). Origin and functional role of the extracellular serotonin in the midbrain raphe nuclei. Brain Res. Rev. 39, 154–
180.
Aghajanian, G.K., and Bunney, B.S. (1977). Dopamine“autoreceptors”: pharmacological characterization by microiontophoretic single cell recording studies. Naunyn. Schmiedebergs Arch. Pharmacol. 297, 1–7.
Aghajanian, G.K., and Lakoski, J.M. (1984). Hyperpolarization of serotonergic neurons by serotonin and LSD: Studies in brain slices showing increased K+ conductance. Brain Res. 305, 181–185.
Aghajanian, G.K., and Vandermaelen, C.P. (1982). Intracellular identification of central noradrenergic and serotonergic neurons by a new double labeling procedure. J. Neurosci. 2, 1786–1792.
179 Aghajanian, G.K., Cedarbaum, J.M., and Wang, R.Y. (1977). Evidence for
norepinephrine-mediated collateral inhibition of locus coeruleus neurons. Brain
Res. 136, 570–577.
Agnati, L.F., Fuxe, K., Zini, I., Benfenati, F., Hökfelt, T., and de Mey, J. (1982).
Principles for the morphological characterization of transmitter-identified nerve cell
groups. J. Neurosci. Methods 6, 157–167.
Agnati, L.F., Guidolin, D., Guescini, M., Genedani, S., and Fuxe, K. (2010).
Understanding wiring and volume transmission. Brain Res. Rev. 64, 137–159.
Ahern, T.H., Javors, M.A., Eagles, D.A., Martillotti, J., Mitchell, H.A., Liles, L.C., and
Weinshenker, D. (2005). The Effects of Chronic Norepinephrine Transporter
Inactivation on Seizure Susceptibility in Mice. Neuropsychopharmacology 31, 730–
738.
Altieri, S.C., Garcia-Garcia, A.L., Leonardo, E.D., and Andrews, A.M. (2013).
Rethinking 5-HT 1A Receptors: Emerging Modes of Inhibitory Feedback of Relevance
to Emotion-Related Behavior. ACS Chem. Neurosci. 4, 72–83.
Alvarez, V.A., Chow, C.C., Van Bockstaele, E.J., and Williams, J.T. (2002). Frequency- dependent synchrony in locus ceruleus: Role of electrotonic coupling. Proc. Natl.
Acad. Sci. U. S. A. 99, 4032–4036.
Andrade, R., and Haj-Dahmane, S. (2013). Serotonin Neuron Diversity in the Dorsal
Raphe. ACS Chem. Neurosci. 4, 22–25.
180 Andrade, R., Huereca, D., Lyons, J.G., Andrade, E.M., and McGregor, K.M. (2015). 5-
HT1A Receptor-Mediated Autoinhibition and the Control of Serotonergic Cell Firing.
ACS Chem. Neurosci.
Anzalone, A., Lizardi-Ortiz, J.E., Ramos, M., Mei, C.D., Hopf, F.W., Iaccarino, C.,
Halbout, B., Jacobsen, J., Kinoshita, C., Welter, M., et al. (2012). Dual Control of
Dopamine Synthesis and Release by Presynaptic and Postsynaptic Dopamine D2
Receptors. J. Neurosci. 32, 9023–9034.
Arai, R., Horiike, K., and Hasegawa, Y. (1998). Dopamine-degrading activity of monoamine oxidase is not detected by histochemistry in neurons of the substantia nigra pars compacta of the rat. Brain Res. 812, 275–278.
Arbuthnott, G.W., and Wickens, J. (2007). Space, time and dopamine. Trends
Neurosci. 30, 62–69.
Arnth-Jensen, N., Jabaudon, D., and Scanziani, M. (2002). Cooperation between independent hippocampal synapses is controlled by glutamate uptake. Nat.
Neurosci. 5, 325–331.
Aston-Jones, G., and Bloom, F.E. (1981). Activity of norepinephrine-containing locus coeruleus neurons in behaving rats anticipates fluctuations in the sleep-waking cycle. J. Neurosci. 1, 876–886.
181 Aston-Jones, G., and Cohen, J.D. (2005a). Adaptive gain and the role of the locus
coeruleus–norepinephrine system in optimal performance. J. Comp. Neurol. 493, 99–
110.
Aston-Jones, G., and Cohen, J.D. (2005b). An Integrative Theory of Locus Coeruleus-
Norepinephrine Function: Adaptive Gain and Optimal Performance. Annu. Rev.
Neurosci. 28, 403–450.
Aston-Jones, G., Rajkowski, J., and Cohen, J. (1999). Role of locus coeruleus in
attention and behavioral flexibility. Biol. Psychiatry 46, 1309–1320.
Asztely, F., Erdemli, G., and Kullmann, D.M. (1997). Extrasynaptic Glutamate
Spillover in the Hippocampus: Dependence on Temperature and the Role of Active
Glutamate Uptake. Neuron 18, 281–293.
Balakrishnan, V., Kuo, S.P., Roberts, P.D., and Trussell, L.O. (2009). Slow glycinergic
transmission mediated by transmitter pooling. Nat. Neurosci. 12, 286–294.
Bang, S.J., Jensen, P., Dymecki, S.M., and Commons, K.G. (2012). Projections and interconnections of genetically defined serotonin neurons in mice: Networks of rhombomere-specific serotonin neurons. Eur. J. Neurosci. 35, 85–96.
Baraban, J., and Aghajanian, G.K. (1981). Noradrenergic innervation of serotonergic neuronis in the dorsal raphe: Demonstration by electron microscopic autoradiography. Brain Res. 204, 1–11.
182 Barbour, B., and Hausser, M. (1997). Intersynaptic diffusion of neurotransmitter.
Trends Neurosci. 20, 377–384.
Barbour, B., Keller, B.U., Llano, I., and Marty, A. (1994). Prolonged presence of glutamate during excitatory synaptic transmission to cerebellar Purkinje cells.
Neuron 12, 1331–1343.
Bayer, V.E., and Pickel, V.M. (1990). Ultrastructural localization of tyrosine hydroxylase in the rat ventral tegmental area: relationship between immunolabeling density and neuronal associations. J. Neurosci. 10, 2996–3013.
Bayliss, D.A., Li, Y., and Talley, E.M. (1997). Effects of Serotonin on Caudal Raphe
Neurons: Activation of an Inwardly Rectifying Potassium Conductance. J.
Neurophysiol. 77, 1349–1361.
Beato, M. (2008). The time course of transmitter at glycinergic synapses onto motoneurons. J. Neurosci. Off. J. Soc. Neurosci. 28, 7412–7425.
Beaudet, A., and Descarries, L. (1978). The monoamine innervation of rat cerebral cortex: Synaptic and nonsynaptic axon terminals. Neuroscience 3, 851–860.
Beaulieu, J.-M., and Gainetdinov, R.R. (2011). The Physiology, Signaling, and
Pharmacology of Dopamine Receptors. Pharmacol. Rev. 63, 182–217.
Beckstead, M.J., and Williams, J.T. (2007). Long-Term Depression of a Dopamine
IPSC. J. Neurosci. 27, 2074–2080.
183 Beckstead, M.J., Grandy, D.K., Wickman, K., and Williams, J.T. (2004). Vesicular
Dopamine Release Elicits an Inhibitory Postsynaptic Current in Midbrain Dopamine
Neurons. Neuron 42, 939–946.
Beenhakker, M.P., and Huguenard, J.R. (2010). Astrocytes as gatekeepers of GABAB receptor function. J. Neurosci. Off. J. Soc. Neurosci. 30, 15262–15276.
Bello, E.P., Mateo, Y., Gelman, D.M., Noaín, D., Shin, J.H., Low, M.J., Alvarez, V.A.,
Lovinger, D.M., and Rubinstein, M. (2011). Cocaine supersensitivity and enhanced motivation for reward in mice lacking dopamine D2 autoreceptors. Nat. Neurosci.
14, 1033–1038.
Bengel, D., Murphy, D.L., Andrews, A.M., Wichems, C.H., Feltner, D., Heils, A., Mössner,
R., Westphal, H., and Lesch, K.-P. (1998). Altered Brain Serotonin Homeostasis and
Locomotor Insensitivity to 3,4-Methylenedioxymethamphetamine (“Ecstasy”) in
Serotonin Transporter-Deficient Mice. Mol. Pharmacol. 53, 649–655.
Bennett, C., Arroyo, S., Berns, D., and Hestrin, S. (2012). Mechanisms generating dual-component nicotinic EPSCs in cortical interneurons. J. Neurosci. Off. J. Soc.
Neurosci. 32, 17287–17296.
Bennett, M.R., Farnell, L., Gibson, W.G., and Blair, D. (2004). A quantitative description of the diffusion of noradrenaline in the media of blood vessels following its release from sympathetic varicosities. J. Theor. Biol. 226, 359–372.
184 Benoit-Marand, M., Borrelli, E., and Gonon, F. (2001). Inhibition of dopamine release
via presynaptic D2 receptors: time course and functional characteristics in vivo. J.
Neurosci. Off. J. Soc. Neurosci. 21, 9134–9141.
Bergles, D.E., Diamond, J.S., and Jahr, C.E. (1999). Clearance of glutamate inside the
synapse and beyond. Curr. Opin. Neurobiol. 9, 293–298.
Bergquist, F., Niazi, H.S., and Nissbrandt, H. (2002). Evidence for different exocytosis pathways in dendritic and terminal dopamine release in vivo. Brain Res. 950, 245–
253.
Bérubé-Carrière, N., Guay, G., Fortin, G.M., Kullander, K., Olson, L., Wallén-Mackenzie,
Å., Trudeau, L.-E., and Descarries, L. (2012). Ultrastructural characterization of the mesostriatal dopamine innervation in mice, including two mouse lines of conditional VGLUT2 knockout in dopamine neurons. Eur. J. Neurosci. 35, 527–538.
Birnbaumer, L. (1992). Receptor-to-effector signaling through G proteins: Roles for
dimers as well as subunits. Cell 71, 1069–1072.
βγ α Bogerts, B. (1981). A brainstem atlas of catecholaminergic neurons in man, using melanin as a natural marker. J. Comp. Neurol. 197, 63–80.
Bohn, L.M., Xu, F., Gainetdinov, R.R., and Caron, M.G. (2000). Potentiated Opioid
Analgesia in Norepinephrine Transporter Knock-Out Mice. J. Neurosci. 20, 9040–
9045.
185 Bonci, A., and Williams, J.T. (1996). A common mechanism mediates long-term changes in synaptic transmission after chronic cocaine and morphine. Neuron 16,
631–639.
Boothman, L.J., Allers, K.A., Rasmussen, K., and Sharp, T. (2003). Evidence that central 5-HT 2A and 5-HT 2B/C receptors regulate 5-HT cell firing in the dorsal raphe
nucleus of the anaesthetised rat. Br. J. Pharmacol. 139, 998–1004.
Brasnjo, G., and Otis, T.S. (2001). Neuronal glutamate transporters control activation
of postsynaptic metabotropic glutamate receptors and influence cerebellar long-
term depression. Neuron 31, 607–616.
Bromberg-Martin, E.S., and Hikosaka, O. (2009). Midbrain dopamine neurons signal preference for advance information about upcoming rewards. Neuron 63, 119–126.
Bruns, D., Riedel, D., Klingauf, J., and Jahn, R. (2000). Quantal release of serotonin.
Neuron 28, 205–220.
Buckholtz, J.W., Treadway, M.T., Cowan, R.L., Woodward, N.D., Li, R., Ansari, M.S.,
Baldwin, R.M., Schwartzman, A.N., Shelby, E.S., Smith, C.E., et al. (2010).
Dopaminergic network differences in human impulsivity. Science 329, 532.
Buffalari, D.M., and Grace, A.A. (2007). Noradrenergic Modulation of Basolateral
Amygdala Neuronal Activity: Opposing Influences of -
J. Neurosci. 27, 12358–12366. α 2 and β Receptor Activation.
186 Bünemann, M., Frank, M., and Lohse, M.J. (2003). Gi protein activation in intact cells
involves subunit rearrangement rather than dissociation. Proc. Natl. Acad. Sci. U. S.
A. 100, 16077–16082.
Bunin, M.A., and Wightman, R.M. (1998). Quantitative evaluation of 5-
hydroxytryptamine (serotonin) neuronal release and uptake: an investigation of
extrasynaptic transmission. J. Neurosci. 18, 4854–4860.
Bunin, M.A., and Wightman, R.M. (1999). Paracrine neurotransmission in the CNS: involvement of 5-HT. Trends Neurosci. 22, 377–382.
Bunin, M.A., Prioleau, C., Mailman, R.B., and Wightman, R.M. (1998). Release and
uptake rates of 5-hydroxytryptamine in the dorsal raphe and substantia nigra
reticulata of the rat brain. J. Neurochem. 70, 1077–1087.
Bunney, B.S., Walters, J.R., Roth, R.H., and Aghajanian, G.K. (1973). Dopaminergic
neurons: effect of antipsychotic drugs and amphetamine on single cell activity. J.
Pharmacol. Exp. Ther. 185, 560–571.
Burchett, S.A., and Bannon, M.J. (1997). Serotonin, dopamine and norepinephrine
transporter mRNAs: heterogeneity of distribution and response to “binge” cocaine
administration. Brain Res. Mol. Brain Res. 49, 95–102.
del Burgo, L.S., Cortes, R., Mengod, G., Zarate, J., Echevarria, E., and Salles, J. (2008).
Distribution and neurochemical characterization of neurons expressing GIRK
channels in the rat brain. J. Comp. Neurol. 510, 581–606.
187 Burns, R.S., Chiueh, C.C., Markey, S.P., Ebert, M.H., Jacobowitz, D.M., and Kopin, I.J.
(1983). A primate model of parkinsonism: selective destruction of dopaminergic
neurons in the pars compacta of the substantia nigra by N-methyl-4-phenyl-1,2,3,6-
tetrahydropyridine. Proc. Natl. Acad. Sci. 80, 4546–4550.
Calebiro, D., and Maiellaro, I. (2014). cAMP signaling microdomains and their
observation by optical methods. Front. Cell. Neurosci. 8.
Calizo, L.H., Akanwa, A., Ma, X., Pan, Y., Lemos, J.C., Craige, C., Heemstra, L.A., and
Beck, S.G. (2011). Raphe serotonin neurons are not homogenous:
Electrophysiological, morphological and neurochemical evidence.
Neuropharmacology 61, 524–543.
Cameron, D.L., and Williams, J.T. (1993). Dopamine D1 receptors facilitate transmitter release. Nature 366, 344–347.
Camps, M., Kelly, P.H., and Palacios, J.M. (1990). Autoradiographic localization of dopamine D 1 and D 2 receptors in the brain of several mammalian species. J. Neural
Transm. Gen. Sect. 80, 105–127.
Canavier, C.C., and Landry, R.S. (2006). An increase in AMPA and a decrease in SK conductance increase burst firing by different mechanisms in a model of a dopamine neuron in vivo. J. Neurophysiol. 96, 2549–2563.
Carlsson, A., Falck, B., and Hillarp, N.A. (1962). Cellular localization of brain monoamines. Acta Physiol. Scand. Suppl. 56, 1–28.
188 Carter, A.G., and Regehr, W.G. (2000). Prolonged Synaptic Currents and Glutamate
Spillover at the Parallel Fiber to Stellate Cell Synapse. J. Neurosci. 20, 4423–4434.
Castro, S.W., and Strange, P.G. (1993). Differences in the Ligand Binding Properties of the Short and Long Versions of the D2 Dopamine Receptor. J. Neurochem. 60,
372–375.
Cathala, L., Holderith, N.B., Nusser, Z., DiGregorio, D.A., and Cull-Candy, S.G. (2005).
Changes in synaptic structure underlie the developmental speeding of AMPA receptor-mediated EPSCs. Nat. Neurosci. 8, 1310–1318.
Cedarbaum, J.M., and Aghajanian, G.K. (1978). Afferent projections to the rat locus coeruleus as determined by a retrograde tracing technique. J. Comp. Neurol. 178, 1–
16.
Charara, A., Pare, J.-F., Levey, A.I., and Smith, Y. (2005). Synaptic and extrasynaptic
GABA-A and GABA-B receptors in the globus pallidus: an electron microscopic immunogold analysis in monkeys. Neuroscience 131, 917–933.
Chaudhry, F.A., Edwards, R.H., and Fonnum, F. (2008). Vesicular Neurotransmitter
Transporters as Targets for Endogenous and Exogenous Toxic Substances. Annu.
Rev. Pharmacol. Toxicol. 48, 277–301.
Chen, B.T., and Rice, M.E. (2001). Novel Ca2+ dependence and time course of somatodendritic dopamine release: substantia nigra versus striatum. J. Neurosci.
Off. J. Soc. Neurosci. 21, 7841–7847.
189 Chen, Z., and Minneman, K.P. (2005). Recent progress in 1-adrenergic receptor research. Acta Pharmacol. Sin. 26, 1281–1287. α
Chen, B.T., Patel, J.C., Moran, K.A., and Rice, M.E. (2011). Differential Calcium
Dependence of Axonal Versus Somatodendritic Dopamine Release, with
Characteristics of Both in the Ventral Tegmental Area. Front. Syst. Neurosci. 5.
Cheramy, A., Leviel, V., and Glowinski, J. (1981). Dendritic release of dopamine in the
substantia nigra. Nature 289, 537–542.
Chieng, B., Azriel, Y., Mohammadi, S., and Christie, M.J. (2011). Distinct cellular
properties of identified dopaminergic and GABAergic neurons in the mouse ventral
tegmental area. J. Physiol. 589, 3775–3787.
Chiodo, L.A., Bannon, M.J., Grace, A.A., Roth, R.H., and Bunney, B.S. (1984). Evidence
for the absence of impulse-regulating somatodendritic and synthesis-modulating nerve terminal autoreceptors on subpopulations of mesocortical dopamine neurons.
Neuroscience 12, 1–16.
Chiti, Z., and Teschemacher, A.G. (2007). Exocytosis of norepinephrine at axon varicosities and neuronal cell bodies in the rat brain. FASEB J. 21, 2540–2550.
Christie, J.M., and Jahr, C.E. (2006). Multivesicular release at Schaffer collateral-CA1 hippocampal synapses. J. Neurosci. Off. J. Soc. Neurosci. 26, 210–216.
190 Chung, H.J., Qian, X., Ehlers, M., Jan, Y.N., and Jan, L.Y. (2009). Neuronal activity regulates phosphorylation-dependent surface delivery of G protein-activated inwardly rectifying potassium channels. Proc. Natl. Acad. Sci. U. S. A. 106, 629–634.
Clancy, S.M., Fowler, C.E., Finley, M., Suen, K.F., Arrabit, C., Berton, F., Kosaza, T.,
Casey, P.J., and Slesinger, P.A. (2005). Pertussis-toxin-sensitive G subunits selectively bind to C-terminal domain of neuronal GIRK channels:α evidence for a heterotrimeric G-protein-channel complex. Mol. Cell. Neurosci. 28, 375–389.
Clancy, S.M., Boyer, S.B., and Slesinger, P.A. (2007). Coregulation of Natively
Expressed Pertussis Toxin-Sensitive Muscarinic Receptors with G-Protein-Activated
Potassium Channels. J. Neurosci. 27, 6388–6399.
Clements, J.D., Lester, R.A., Tong, G., Jahr, C.E., and Westbrook, G.L. (1992). The time course of glutamate in the synaptic cleft. Science 258, 1498–1501.
Cohen, J.Y., Amoroso, M.W., and Uchida, N. (2015). Serotonergic neurons signal reward and punishment on multiple timescales. eLife 4.
Colgan, L.A., Putzier, I., and Levitan, E.S. (2009). Activity-Dependent Vesicular
Monoamine Transporter-Mediated Depletion of the Nucleus Supports Somatic
Release by Serotonin Neurons. J. Neurosci. 29, 15878–15887.
Colgan, L.A., Cavolo, S.L., Commons, K.G., and Levitan, E.S. (2012). Action Potential-
Independent and Pharmacologically Unique Vesicular Serotonin Release from
Dendrites. J. Neurosci. 32, 15737–15746.
191 Commons, K.G. (2009). Locally Collateralizing Glutamate Neurons in the Dorsal
Raphe Nucleus Responsive to Substance P Contain Vesicular Glutamate Transporter
3 (VGLUT3). J. Chem. Neuroanat. 38, 273–281.
Courtney, N.A., and Ford, C.P. (2014). The Timing of Dopamine- and Noradrenaline-
Mediated Transmission Reflects Underlying Differences in the Extent of Spillover
and Pooling. J. Neurosci. 34, 7645–7656.
Courtney, N.A., Mamaligas, A.A., and Ford, C.P. (2012). Species Differences in
Somatodendritic Dopamine Transmission Determine D2-Autoreceptor-Mediated
Inhibition of Ventral Tegmental Area Neuron Firing. J. Neurosci. 32, 13520–13528.
Cragg, S.J., and Greenfield, S.A. (1997). Differential Autoreceptor Control of
Somatodendritic and Axon Terminal Dopamine Release in Substantia Nigra, Ventral
Tegmental Area, and Striatum. J. Neurosci. 17, 5738–5746.
Cragg, S.J., and Rice, M.E. (2004). DAncing past the DAT at a DA synapse. Trends
Neurosci. 27, 270–277.
Cragg, S.J., Nicholson, C., Kume-Kick, J., Tao, L., and Rice, M.E. (2001). Dopamine-
Mediated Volume Transmission in Midbrain Is Regulated by Distinct Extracellular
Geometry and Uptake. J. Neurophysiol. 85, 1761–1771.
Dahlström, A., and Fuxe, K. (1964). Localization of monoamines in the lower brain stem. Experientia 20, 398–399.
192 Dal Toso, R., Sommer, B., Ewert, M., Herb, A., Pritchett, D.B., Bach, A., Shivers, B.D.,
and Seeburg, P.H. (1989). The dopamine D2 receptor: two molecular forms
generated by alternative splicing. EMBO J. 8, 4025–4034.
Daubner, S.C., Le, T., and Wang, S. (2011). Tyrosine hydroxylase and regulation of
dopamine synthesis. Arch. Biochem. Biophys. 508, 1–12.
David, M., Richer, M., Mamarbachi, A.M., Villeneuve, L.R., Dupré, D.J., and Hebert, T.E.
(2006). Interactions between GABA-B1 receptors and Kir 3 inwardly rectifying potassium channels. Cell. Signal. 18, 2172–2181.
Davila, V., Yan, Z., Craciun, L.C., Logothetis, D., and Sulzer, D. (2003). D3 Dopamine
Autoreceptors Do Not Activate G-Protein-Gated Inwardly Rectifying Potassium
Channel Currents in Substantia Nigra Dopamine Neurons. J. Neurosci. 23, 5693–
5697.
Day, H.E.W., Greenwood, B.N., Hammack, S.E., Watkins, L.R., Fleshner, M., Maier, S.F.,
and Campeau, S. (2004). Differential Expression of 5HT-1A, 1b Adrenergic, CRF-R1,
and CRF- -Aminobutyricα Acidergic, and
CatecholaminerR2 Receptorgic Cells mRNA of the in Rat Serotonergic, Dorsal Raphe γ Nucleus. J. Comp. Neurol. 474, 364–
378.
Dearry, A., Gingrich, J.A., Falardeau, P., Fremeau, R.T., Bates, M.D., and Caron, M.G.
(1990). Molecular cloning and expression of the gene for a human D1 dopamine
receptor. Nature 347, 72–76.
193 Demarest, K.T., and Moore, K.E. (1979). Comparison of dopamine synthesis
regulation in the terminals of nigrostriatal, mesolimbic, tuberoinfundibular and
tuberohypophyseal neurons. J. Neural Transm. 46, 263–277.
Denney, R.M., and Denney, C.B. (1985). An update on the identity crisis of monoamine oxidase: new and old evidence for the independence of MAO A and B.
Pharmacol. Ther. 30, 227–258.
Descarries, L., Berthelet, F., Garcia, S., and Beaudet, A. (1986). Dopaminergic projection from nucleus raphe dorsalis to neostriatum in the rat. J. Comp. Neurol.
249, 511–520, 484–485.
Descarries, L., Watkins, K.C., Garcia, S., Bosler, O., and Doucet, G. (1996). Dual character, asynaptic and synaptic, of the dopamine innervation in adult rat neostriatum: a quantitative autoradiographic and immunocytochemical analysis. J.
Comp. Neurol. 375, 167–186.
Descarries, L., Bérubé-Carrière, N., Riad, M., Bo, G.D., Mendez, J.A., and Trudeau, L.-É.
(2008). Glutamate in dopamine neurons: Synaptic versus diffuse transmission. Brain
Res. Rev. 58, 290–302.
Deutch, A.Y., Goldstein, M., Baldino, F., and Roth, R.H. (1988). Telencephalic projections of the A8 dopamine cell group. Ann. N. Y. Acad. Sci. 537, 27–50.
194 Diamond, J.S. (2001). Neuronal Glutamate Transporters Limit Activation of NMDA
Receptors by Neurotransmitter Spillover on CA1 Pyramidal Cells. J. Neurosci. 21,
8328–8338.
Diaz, J., Pilon, C., Foll, B.L., Gros, C., Triller, A., Schwartz, J.-C., and Sokoloff, P. (2000).
Dopamine D3 Receptors Expressed by All Mesencephalic Dopamine Neurons. J.
Neurosci. 20, 8677–8684.
DiGregorio, D.A., Nusser, Z., and Silver, R.A. (2002). Spillover of glutamate onto
synaptic AMPA receptors enhances fast transmission at a cerebellar synapse.
Neuron 35, 521–533.
Dougalis, A.G., Matthews, G.A.C., Bishop, M.W., Brischoux, F., Kobayashi, K., and
Ungless, M.A. (2012). Functional properties of dopamine neurons and co-expression
of vasoactive intestinal polypeptide in the dorsal raphe nucleus and ventro-lateral periaqueductal grey: DRN/vlPAG dopamine neurons. Eur. J. Neurosci. 36, 3322–
3332.
Dreyer, J.K., Herrik, K.F., Berg, R.W., and Hounsgaard, J.D. (2010). Influence of phasic and tonic dopamine release on receptor activation. J. Neurosci. Off. J. Soc. Neurosci.
30, 14273–14283.
Dunkley, P.R., Bobrovskaya, L., Graham, M.E., Von Nagy-Felsobuki, E.I., and Dickson,
P.W. (2004). Tyrosine hydroxylase phosphorylation: regulation and consequences. J.
Neurochem. 91, 1025–1043.
195 Edmondson, D.E., Binda, C., Wang, J., Upadhyay, A.K., and Mattevi, A. (2009).
Molecular and mechanistic properties of the membrane-bound mitochondrial monoamine oxidases. Biochemistry (Mosc.) 48, 4220–4230.
Egan, T.M., Henderson, G., North, R.A., and Williams, J.T. (1983). Noradrenaline- mediated synaptic inhibition in rat locus coeruleus neurones. J. Physiol. 345, 477–
488.
Erickson, J.D., Schafer, M.K., Bonner, T.I., Eiden, L.E., and Weihe, E. (1996). Distinct pharmacological properties and distribution in neurons and endocrine cells of two isoforms of the human vesicular monoamine transporter. Proc. Natl. Acad. Sci. 93,
5166–5171.
Ewing, A.G., Wightman, R.M., and Dayton, M.A. (1982). In vivo voltammetry with electrodes that discriminate between dopamine and ascorbate. Brain Res. 249, 361–
370.
Fagervall, I., and Ross, S.B. (1986). A and B Forms of Monoamine Oxidase Within the
Monoaminergic Neurons of the Rat Brain. J. Neurochem. 47, 569–576.
Farb, C.R., Chang, W., and LeDoux, J.E. (2010). Ultrastructural Characterization of
Noradrenergic Axons and Beta-Adrenergic Receptors in the Lateral Nucleus of the
Amygdala. Front. Behav. Neurosci. 4.
196 Felten, D.L., and Sladek, J.R. (1983). Monoamine distribution in primate brain V.
Monoaminergic nuclei: anatomy, pathways and local organization. Brain Res. Bull.
10, 171–284.
Ferguson, S.S.G. (2001). Evolving Concepts in G Protein-Coupled Receptor
Endocytosis: The Role in Receptor Desensitization and Signaling. Pharmacol. Rev.
53, 1–24.
Finberg, J.P.M. (2014). Update on the pharmacology of selective inhibitors of MAO-A
and MAO-B: Focus on modulation of CNS monoamine neurotransmitter release.
Pharmacol. Ther. 143, 133–152.
Finlayson, P.G., and Marshall, K.C. (1988). Synchronous bursting of locus coeruleus
neurons in tissue culture. Neuroscience 24, 217–225.
Fiorillo, C.D., and Williams, J.T. (1998). Glutamate mediates an inhibitory
postsynaptic potential in dopamine neurons. Nature 394, 78–82.
Flores, J.A., Banoua, F. El, Galán-Rodríguez, B., and Fernandez-Espejo, E. (2004).
Opiate anti-nociception is attenuated following lesion of large dopamine neurons of
the periaqueductal grey: critical role for D1 (not D2) dopamine receptors. Pain 110,
205–214.
Floresco, S.B., West, A.R., Ash, B., Moore, H., and Grace, A.A. (2003). Afferent
modulation of dopamine neuron firing differentially regulates tonic and phasic
dopamine transmission. Nat. Neurosci. 6, 968–973.
197 Florin-Lechner, S.M., Druhan, J.P., Aston-Jones, G., and Valentino, R.J. (1996).
Enhanced norepinephrine release in prefrontal cortex with burst stimulation of the
locus coeruleus. Brain Res. 742, 89–97.
Foote, S.L., Bloom, F.E., and Aston-Jones, G. (1983). Nucleus locus ceruleus: new
evidence of anatomical and physiological specificity. Physiol. Rev. 63, 844–914.
Ford, C.P. (2014). The role of D2-autoreceptors in regulating dopamine neuron
activity and transmission. Neuroscience 282, 13–22.
Ford, C.P., Mark, G.P., and Williams, J.T. (2006). Properties and opioid inhibition of mesolimbic dopamine neurons vary according to target location. J. Neurosci. Off. J.
Soc. Neurosci. 26, 2788–2797.
Ford, C.P., Beckstead, M.J., and Williams, J.T. (2007). Kappa opioid inhibition of somatodendritic dopamine inhibitory postsynaptic currents. J. Neurophysiol. 97,
883–891.
Ford, C.P., Phillips, P.E.M., and Williams, J.T. (2009). The Time Course of Dopamine
Transmission in the Ventral Tegmental Area. J. Neurosci. 29, 13344–13352.
Ford, C.P., Gantz, S.C., Phillips, P.E.M., and Williams, J.T. (2010). Control of
Extracellular Dopamine at Dendrite and Axon Terminals. J. Neurosci. 30, 6975–6983.
Fornal, C.A., Metzler, C.W., Gallegos, R.A., Veasey, S.C., McCreary, A.C., and Jacobs, B.L.
(1996). WAY-100635, a potent and selective 5-hydroxytryptamine1A antagonist,
198 increases serotonergic neuronal activity in behaving cats: comparison with (S)-
WAY-100135. J. Pharmacol. Exp. Ther. 278, 752–762.
Fortin, G.D., Desrosiers, C.C., Yamaguchi, N., and Trudeau, L.-E. (2006). Basal somatodendritic dopamine release requires snare proteins. J. Neurochem. 96, 1740–
1749.
Freund, T.F., Powell, J.F., and Smith, A.D. (1984). Tyrosine hydroxylase- immunoreactive boutons in synaptic contact with identified striatonigral neurons, with particular reference to dendritic spines. Neuroscience 13, 1189–1215.
Gainetdinov, R.R., and Caron, M.G. (2003). Monoamine Transporters: From Genes to
Behavior. Annu. Rev. Pharmacol. Toxicol. 43, 261–284.
Gainetdinov, R.R., Premont, R.T., Bohn, L.M., Lefkowitz, R.J., and Caron, M.G. (2004).
Desensitization of G Protein–Coupled Receptors and Neuronal Functions. Annu. Rev.
Neurosci. 27, 107–144.
Galvez, R., Mesches, M.H., and Mcgaugh, J.L. (1996). Norepinephrine Release in the
Amygdala in Response to Footshock Stimulation. Neurobiol. Learn. Mem. 66, 253–
257.
Gannon, M., Che, P., Chen, Y., Jiao, K., Roberson, E.D., and Wang, Q. (2015).
Noradrenergic dysfunction in Alzheimer’s disease. Front. Neurosci. 9.
Gantz, S.C., Bunzow, J.R., and Williams, J.T. (2013). Spontaneous Inhibitory Synaptic
Currents Mediated by a G Protein-Coupled Receptor. Neuron 78, 807–812.
199 Gantz, S.C., Levitt, E.S., Llamosas, N., Neve, K.A., and Williams, J.T. (2015). Depression
of Serotonin Synaptic Transmission by the Dopamine Precursor L-DOPA. Cell Rep.
12, 944–954.
Garris, P.A., Ciolkowski, E.L., Pastore, P., and Wightman, R.M. (1994). Efflux of
dopamine from the synaptic cleft in the nucleus accumbens of the rat brain. J.
Neurosci. Off. J. Soc. Neurosci. 14, 6084–6093.
Gaspar, P., and Lillesaar, C. (2012). Probing the diversity of serotonin neurons.
Philos. Trans. R. Soc. B Biol. Sci. 367, 2382–2394.
Geffen, L.B., Jessell, T.M., Cuello, A.C., and Iversen, L.L. (1976). Release of dopamine
from dendrites in rat substantia nigra. Nature 260, 258–260.
German, D.C., Schlusselberg, D.S., and Woodward, D.J. (1983). Three-dimensional
computer reconstruction of midbrain dopaminergic neuronal populations: from
mouse to man. J. Neural Transm. 57, 243–254.
Gingrich, J.A., and Caron, M.G. (1993). Recent Advances in the Molecular Biology of
Dopamine Receptors. Annu. Rev. Neurosci. 16, 299–321.
Giorguieff-Chesselet, M.F., Kemel, M.L., Wandscheer, D., and Glowinski, J. (1979).
Regulation of dopamine release by presynaptic nicotinic receptors in rat striatal slices: Effect of nicotine in a low concentration. Life Sci. 25, 1257–1261.
200 Giros, B., Jaber, M., Jones, S.R., Wightman, R.M., and Caron, M.G. (1996).
Hyperlocomotion and indifference to cocaine and amphetamine in mice lacking the dopamine transporter. Nature 379, 606–612.
Grace, A.A., and Bunney, B.S. (1983). Intracellular and extracellular electrophysiology of nigral dopaminergic neurons--2. Action potential generating mechanisms and morphological correlates. Neuroscience 10, 317–331.
Grace, A.A., and Bunney, B.S. (1984). The control of firing pattern in nigral dopamine neurons: burst firing. J. Neurosci. 4, 2877–2890.
Grandy, D.K., Zhang, Y.A., Bouvier, C., Zhou, Q.Y., Johnson, R.A., Allen, L., Buck, K.,
Bunzow, J.R., Salon, J., and Civelli, O. (1991). Multiple human D5 dopamine receptor genes: a functional receptor and two pseudogenes. Proc. Natl. Acad. Sci. U. S. A. 88,
9175–9179.
Groves, P.M., and Linder, J.C. (1983). Dendro-dendritic synapses in substantia nigra:
Descriptions based on analysis of serial sections. Exp. Brain Res. 49, 209–217.
Groves, P.M., and Wilson, C.J. (1980). Monoaminergic presynaptic axons and dendrites in rat locus coeruleus seen in reconstructions of serial sections. J. Comp.
Neurol. 193, 853–862.
Haddjeri, N., Lavoie, N., and Blier, P. (2004). Electrophysiological Evidence for the
Tonic Activation of 5-HT1A Autoreceptors in the Rat Dorsal Raphe Nucleus.
Neuropsychopharmacology 29, 1800–1806.
201 Hajos, M., Hoffmann, W.E., Tetko, I.V., Hyland, B., Sharp, T., and Villa, A.E.P. (2001).
Different tonic regulation of neuronal activity in the rat dorsal raphe and medial prefrontal cortex via 5-HT1A receptors. Neurosci. Lett. 304, 129–132.
Hajós, M., Allers, K.A., Jennings, K., Sharp, T., Charette, G., Sík, A., and Kocsis, B.
(2007). Neurochemical identification of stereotypic burst-firing neurons in the rat dorsal raphe nucleus using juxtacellular labelling methods. Eur. J. Neurosci. 25, 119–
126.
Hall, R.A., Premont, R.T., and Lefkowitz, R.J. (1999). Heptahelical Receptor Signaling:
Beyond the G Protein Paradigm. J. Cell Biol. 145, 927–932.
Haller, J., Bakos, N., Rodriguiz, R.M., Caron, M.G., Wetsel, W.C., and Liposits, Z. (2002).
Behavioral responses to social stress in noradrenaline transporter knockout mice:
Effects on social behavior and depression. Brain Res. Bull. 58, 279–284.
Han, D.D., and Gu, H.H. (2006). Comparison of the monoamine transporters from human and mouse in their sensitivities to psychostimulant drugs. BMC Pharmacol. 6,
6.
Hare, M.L. (1928). Tyramine oxidase: A new enzyme system in liver. Biochem. J. 22,
968–979.
Harris, G.C., Hausken, Z.E., and Williams, J.T. (1992). Cocaine induced synchronous oscillations in central noradrenergic neurons In vitro. Neuroscience 50, 253–257.
202 Harrison, J.K., Pearson, W.R., and Lynch, K.R. (1991). Molecular characterization of alpha 1- and alpha 2-adrenoceptors. Trends Pharmacol. Sci. 12, 62–67.
Hartzell, H.C., Kuffler, S.W., and Yoshikami, D. (1975). Post-synaptic potentiation: interaction between quanta of acetylcholine at the skeletal neuromuscular synapse.
J. Physiol. 251, 427–463.
Hatfield, T., Spanis, C., and McGaugh, J.L. (1999). Response of amygdalar norepinephrine to footshock and GABAergic drugs using in vivo microdialysis and
HPLC. Brain Res. 835, 340–345.
Häusser, M., and Roth, A. (1997). Estimating the time course of the excitatory synaptic conductance in neocortical pyramidal cells using a novel voltage jump method. J. Neurosci. Off. J. Soc. Neurosci. 17, 7606–7625.
Heien, M.L.A.V., Phillips, P.E.M., Stuber, G.D., Seipel, A.T., and Wightman, R.M. (2003).
Overoxidation of carbon-fiber microelectrodes enhances dopamine adsorption and increases sensitivity. The Analyst 128, 1413–1419.
Herdon, H., and Nahorski, S.R. (1989). Investigations of the roles of dihydropyridine and omega-conotoxin-sensitive calcium channels in mediating depolarisation- evoked endogenous dopamine release from striatal slices. Naunyn. Schmiedebergs
Arch. Pharmacol. 340, 36–40.
203 Hersch, S.M., Yi, H., Heilman, C.J., Edwards, R.H., and Levey, A.I. (1997). Subcellular
localization and molecular topology of the dopamine transporter in the striatum and
substantia nigra. J. Comp. Neurol. 388, 211–227.
Hestrin, S., Sah, P., and Nicoll, R.A. (1990). Mechanisms generating the time course of
dual component excitatory synaptic currents recorded in hippocampal slices.
Neuron 5, 247–253.
Hioki, H., Nakamura, H., Ma, Y.-F., Konno, M., Hayakawa, T., Nakamura, K.C.,
Fujiyama, F., and Kaneko, T. (2010). Vesicular glutamate transporter 3-expressing
nonserotonergic projection neurons constitute a subregion in the rat midbrain
raphe nuclei. J. Comp. Neurol. 518, 668–686.
Hjorth, S., and Sharp, T. (1991). Effect of the 5-HT1A receptor agonist 8-OH-DPAT on the release of 5-HT in dorsal and median raphe-innervated rat brain regions as measured by in vivo microdialysis. Life Sci. 48, 1779–1786.
Hoffman, A.F., and Gerhardt, G.A. (1999). Differences in pharmacological properties of dopamine release between the substantia nigra and striatum: an in vivo electrochemical study. J. Pharmacol. Exp. Ther. 289, 455–463.
Hollinger, S., and Hepler, J.R. (2002). Cellular Regulation of RGS Proteins:
Modulators and Integrators of G Protein Signaling. Pharmacol. Rev. 54, 527–559.
204 Holmes, A., Murphy, D.L., and Crawley, J.N. (2003). Abnormal behavioral phenotypes
of serotonin transporter knockout mice: parallels with human anxiety and
depression. Biol. Psychiatry 54, 953–959.
Hoyer, D., Clarke, D.E., Fozard, J.R., Hartig, P.R., Martin, G.R., Mylecharane, E.J.,
Saxena, P.R., and Humphrey, P.P. (1994). International Union of Pharmacology
classification of receptors for 5-hydroxytryptamine (Serotonin). Pharmacol. Rev. 46,
157–203.
Huang, E.P. (1998). Synaptic transmission: Spillover at central synapses. Curr. Biol.
8, R613–R615.
Huang, C.-L., Slesinger, P.A., Casey, P.J., Jan, Y.N., and Jan, L.Y. (1995). Evidence that
direct binding of G to the GIRK1 G protein-gated inwardly rectifying K+ channel is
important for channelβγ activation. Neuron 15, 1133–1143.
Huang, C.S., Shi, S.-H., Ule, J., Ruggiu, M., Barker, L.A., Darnell, R.B., Jan, Y.N., and Jan,
L.Y. (2005). Common Molecular Pathways Mediate Long-Term Potentiation of
Synaptic Excitation and Slow Synaptic Inhibition. Cell 123, 105–118.
Huang, H.-P., Wang, S.-R., Yao, W., Zhang, C., Zhou, Y., Chen, X.-W., Zhang, B., Xiong,
W., Wang, L.-Y., Zheng, L.-H., et al. (2007). Long latency of evoked quantal transmitter release from somata of locus coeruleus neurons in rat pontine slices.
Proc. Natl. Acad. Sci. U. S. A. 104, 1401–1406.
205 Huang, H.-P., Zhu, F.-P., Chen, X.-W., Xu, Z.-Q.D., Zhang, C.X., and Zhou, Z. (2012).
Physiology of quantal norepinephrine release from somatodendritic sites of neurons
in locus coeruleus. Front. Mol. Neurosci. 5.
Imai, H., Steindler, D.A., and Kitai, S.T. (1986). The organization of divergent axonal
projections from the midbrain raphe nuclei in the rat. J. Comp. Neurol. 243, 363–
380.
Inaba, K., Mizuhiki, T., Setogawa, T., Toda, K., Richmond, B.J., and Shidara, M. (2013).
Neurons in Monkey Dorsal Raphe Nucleus Code Beginning and Progress of Step-by-
Step Schedule, Reward Expectation, and Amount of Reward Outcome in the Reward
Schedule Task. J. Neurosci. 33, 3477–3491.
Ingram, S., Wilding, T.J., McCleskey, E.W., and Williams, J.T. (1997). Efficacy and
kinetics of opioid action on acutely dissociated neurons. Mol. Pharmacol. 52, 136–
143.
Invernizzi, R.W. (2007). Role of TPH-2 in brain function: News from behavioral and pharmacologic studies. J. Neurosci. Res. 85, 3030–3035.
Isaacson, J.S. (1999). Glutamate Spillover Mediates Excitatory Transmission in the
Rat Olfactory Bulb. Neuron 23, 377–384.
Isaacson, J.S., and Nicoll, R.A. (1993). The uptake inhibitor L-trans-PDC enhances responses to glutamate but fails to alter the kinetics of excitatory synaptic currents in the hippocampus. J. Neurophysiol. 70, 2187–2191.
206 Isaacson, J.S., Solis, J.M., and Nicoll, R.A. (1993). Local and diffuse synaptic actions of
GABA in the hippocampus. Neuron 10, 165–175.
Ishimatsu, M., and Williams, J.T. (1996). Synchronous Activity in Locus Coeruleus
Results from Dendritic Interactions in Pericoerulear Regions. J. Neurosci. 16, 5196–
5204.
Itoi, K. (2008). Ablation of the Central Noradrenergic Neurons for Unraveling Their
Roles in Stress and Anxiety. Ann. N. Y. Acad. Sci. 1129, 47–54.
Itoi, K., and Sugimoto, N. (2010). The Brainstem Noradrenergic Systems in Stress,
Anxiety and Depression. J. Neuroendocrinol. 22, 355–361.
Jackson, B.P., Dietz, S.M., and Wightman, R.M. (1995). Fast-scan cyclic voltammetry of 5-hydroxytryptamine. Anal. Chem. 67, 1115–1120.
Jacobowitz, D.M., and MacLean, P.D. (1978). A brainstem atlas of catecholaminergic neurons and serotonergic perikarya in a pygmy primate (Cebuella pygmaea). J.
Comp. Neurol. 177, 397–416.
Jacobs, B.L., and Azmitia, E.C. (1992). Structure and function of the brain serotonin system. Physiol Rev 72, 165–229.
Jaffe, E.H., Marty, A., Schulte, A., and Chow, R.H. (1998). Extrasynaptic Vesicular
Transmitter Release from the Somata of Substantia Nigra Neurons in Rat Midbrain
Slices. J. Neurosci. 18, 3548–3553.
207 Jahr, C.E., and Lester, R.A.J. (1992). Synaptic excitation mediated by glutamate-gated ion channels. Curr. Opin. Neurobiol. 2, 270–274.
Jelacic, T.M., Sims, S.M., and Clapham, D.E. (1999). Functional Expression and
Characterization of G-protein-gated Inwardly Rectifying K+ Channels Containing
GIRK3. J. Membr. Biol. 169, 123–129.
Jelacic, T.M., Kennedy, M.E., Wickman, K., and Clapham, D.E. (2000). Functional and
Biochemical Evidence for G-protein-gated Inwardly Rectifying K+ (GIRK) Channels
Composed of GIRK2 and GIRK3. J. Biol. Chem. 275, 36211–36216.
Jennings, K.A. (2013). A Comparison of the Subsecond Dynamics of
Neurotransmission of Dopamine and Serotonin. ACS Chem. Neurosci. 4, 704–714.
Jiang, Y., Ma, W., Wan, Y., Kozasa, T., Hattori, S., and Huang, X.-Y. (1998). The G protein G 12 stimulates Bruton’s tyrosine kinase and a rasGAP through a conserved
PH/BM domain.α Nature 395, 808–813.
John, C.E., Budygin, E.A., Mateo, Y., and Jones, S.R. (2006). Neurochemical characterization of the release and uptake of dopamine in ventral tegmental area and serotonin in substantia nigra of the mouse. J. Neurochem. 96, 267–282.
Jones, M.V., and Westbrook, G.L. (1996). The impact of receptor desensitization on fast synaptic transmission. Trends Neurosci. 19, 96–101.
208 Judge, S.J., and Gartside, S.E. (2006). Firing of 5-HT neurones in the dorsal and median raphe nucleus in vitro shows differential 1-adrenoceptor and 5-HT1A receptor modulation. Neurochem. Int. 48, 100–107.α
Juraska, J.M., Wilson, C.J., and Groves, P.M. (1977). The substantia nigra of the rat: a
Golgi study. J. Comp. Neurol. 172, 585–600.
Kach, J., Sethakorn, N., and Dulin, N.O. (2012). A finer tuning of G-protein signaling through regulated control of RGS proteins. Am. J. Physiol. - Heart Circ. Physiol. 303,
H19–H35.
Kalgutkar, A.S., Dalvie, D.K., Castagnoli, N., and Taylor, T.J. (2001). Interactions of nitrogen-containing xenobiotics with monoamine oxidase (MAO) isozymes A and B:
SAR studies on MAO substrates and inhibitors. Chem. Res. Toxicol. 14, 1139–1162.
Kasparov, S., and Teschemacher, A.G. (2008). Altered central catecholaminergic transmission and cardiovascular disease. Exp. Physiol. 93, 725–740.
Katayama, J., Yakushiji, T., and Akaike, N. (1997). Characterization of the K+ current mediated by 5-HT1A receptor in the acutely dissociated rat dorsal raphe neurons.
Brain Res. 745, 283–292.
Katz, B., and Miledi, R. (1973). The binding of acetylcholine to receptors and its removal from the synaptic cleft. J. Physiol. 231, 549–574.
Kaushalya, S. k., Desai, R., Arumugam, S., Ghosh, H., Balaji, J., and Maiti, S. (2008).
Three-photon microscopy shows that somatic release can be a quantitatively
209 significant component of serotonergic neurotransmission in the mammalian brain. J.
Neurosci. Res. 86, 3469–3480.
Kavalali, E.T. (2015). The mechanisms and functions of spontaneous neurotransmitter release. Nat. Rev. Neurosci. 16, 5–16.
Kawagoe, K.T., Garris, P.A., Wiedemann, D.J., and Wightman, R.M. (1992). Regulation of transient dopamine concentration gradients in the microenvironment surrounding nerve terminals in the rat striatum. Neuroscience 51, 55–64.
Kawano, T., Zhao, P., Nakajima, S., and Nakajima, Y. (2004). Single-cell RT-PCR analysis of GIRK channels expressed in rat locus coeruleus and nucleus basalis neurons. Neurosci. Lett. 358, 63–67.
Kelly, R.S., and Wightman, R.M. (1987). Detection of dopamine overflow and diffusion with voltammetry in slices of rat brain. Brain Res. 423, 79–87.
Kia, H.K., Brisorgueil, M.-J., Hamon, M., Calas, A., and Vergé, D. (1996). Ultrastructural localization of 5-hydroxytryptamine1A receptors in the rat brain. J. Neurosci. Res.
46, 697–708.
Kinney, G.A., Overstreet, L.S., and Slater, N.T. (1997). Prolonged Physiological
Entrapment of Glutamate in the Synaptic Cleft of Cerebellar Unipolar Brush Cells. J.
Neurophysiol. 78, 1320–1333.
210 Kitayama, S., Morita, K., and Dohi, T. (2001). Functional characterization of the
splicing variants of human norepinephrine transporter. Neurosci. Lett. 312, 108–
112.
De Kock, C.P.J., Cornelisse, L.N., Burnashev, N., Lodder, J.C., Timmerman, A.J., Couey,
J.J., Mansvelder, H.D., and Brussaard, A.B. (2006). NMDA receptors trigger
neurosecretion of 5-HT within dorsal raphé nucleus of the rat in the absence of
action potential firing: NMDA receptors trigger secretion of 5-HT. J. Physiol. 577,
891–905.
Kofuji, P., Davidson, N., and Lester, H.A. (1995). Evidence that neuronal G-protein- gated inwardly rectifying K+ channels are activated by G beta gamma subunits and function as heteromultimers. Proc. Natl. Acad. Sci. U. S. A. 92, 6542–6546.
Köhler, C., and Steinbusch, H. (1982). Identification of serotonin and non-serotonin- containing neurons of the mid-brain raphe projecting to the entorhinal area and the hippocampal formation. A combined immunohistochemical and fluorescent retrograde tracing study in the rat brain. Neuroscience 7, 951–975.
Komendantov, A.O., and Canavier, C.C. (2002). Electrical Coupling Between Model
Midbrain Dopamine Neurons: Effects on Firing Pattern and Synchrony. J.
Neurophysiol. 87, 1526–1541.
Koob, G.F. (1992). Drugs of abuse: anatomy, pharmacology and function of reward pathways. Trends Pharmacol. Sci. 13, 177–184.
211 Kuffler, S.W., and Yoshikami, D. (1975). The distribution of acetylcholine sensitivity
at the post-synaptic membrane of vertebrate skeletal twitch muscles: iontophoretic mapping in the micron range. J. Physiol. 244, 703–730.
Kuhn, D.M., Wolf, W.A., and Youdim, M.B. (1985). 5-Hydroxytryptamine release in vivo from a cytoplasmic pool: studies on the 5-HT behavioural syndrome in reserpinized rats. Br. J. Pharmacol. 84, 121–129.
Kuhr, W.G., and Wightman, R.M. (1986). Real-time measurement of dopamine release in rat brain. Brain Res. 381, 168–171.
Kulik, A., Nakadate, K., Nyíri, G., Notomi, T., Malitschek, B., Bettler, B., and Shigemoto,
R. (2002). Distinct localization of GABA(B) receptors relative to synaptic sites in the rat cerebellum and ventrobasal thalamus. Eur. J. Neurosci. 15, 291–307.
Kullmann, D.M., Erdemli, G., and Asztély, F. (1996). LTP of AMPA and NMDA
Receptor–Mediated Signals: Evidence for Presynaptic Expression and Extrasynaptic
Glutamate Spill-Over. Neuron 17, 461–474.
Lacey, M.G., Mercuri, N.B., and North, R.A. (1987). Dopamine acts on D2 receptors to increase potassium conductance in neurones of the rat substantia nigra zona compacta. J. Physiol. 392, 397–416.
Lalive, A.L., Munoz, M.B., Bellone, C., Slesinger, P.A., Lüscher, C., and Tan, K.R. (2014).
Firing Modes of Dopamine Neurons Drive Bidirectional GIRK Channel Plasticity. J.
Neurosci. 34, 5107–5114.
212 Lammel, S., Hetzel, A., Häckel, O., Jones, I., Liss, B., and Roeper, J. (2008). Unique
Properties of Mesoprefrontal Neurons within a Dual Mesocorticolimbic Dopamine
System. Neuron 57, 760–773.
Lammel, S., Lim, B.K., and Malenka, R.C. (2014). Reward and aversion in a heterogeneous midbrain dopamine system. Neuropharmacology 76.
Laporte, S.A., Miller, W.E., Kim, K.-M., and Caron, M.G. (200 -Arrestin/AP-2
Interaction in G Protein- 2). β -
coupled-Adaptin. Receptor J. Biol. Internalization Chem. 277, 9247 Identification–9254. of a β
Arrestin Binding Site in β2 Lautenschlager, M., Höltje, M., von Jagow, B., Veh, R.W., Harms, C., Bergk, A., Dirnagl,
U., Ahnert-Hilger, G., and Hörtnagl, H. (2000). Serotonin uptake and release mechanisms in developing cultures of rat embryonic raphe neurons: age- and
region-specific differences. Neuroscience 99, 519–527.
Lavine, N., Ethier, N., Oak, J.N., Pei, L., Liu, F., Trieu, P., Rebois, R.V., Bouvier, M.,
Hébert, T.E., and Tol, H.H.M.V. (2002). G Protein-coupled Receptors Form Stable
Complexes with Inwardly Rectifying Potassium Channels and Adenylyl Cyclase. J.
Biol. Chem. 277, 46010–46019.
Lee, A., Rosin, D.L., and Van Bockstaele, E.J. (1998a). 2A-adrenergic receptors in the
rat nucleus locus coeruleus: subcellular localization inα catecholaminergic dendrites,
astrocytes, and presynaptic axon terminals. Brain Res. 795, 157–169.
213 Lee, A., Wissekerke, A.E., Rosin, D.L., and Lynch, K.R. (1998b). Localization of 2c- adrenergic receptor immunoreactivity in catecholaminergic neurons in the ratα central nervous system. Neuroscience 84, 1085–1096.
Lee, A., Rosin, D.L., and Van Bockstaele, E.J. (1998c). 2A-adrenergic receptors in the rat nucleus locus coeruleus: subcellular localization αin catecholaminergic dendrites, astrocytes, and presynaptic axon terminals. Brain Res. 795, 157–169.
Lee, A., Rosin, D.L., and Van Bockstaele, E.J. (1998d). Ultrastructural evidence for prominent postsynaptic localization of 2C-adrenergic receptors in catecholaminergic dendrites in the rat nucleusα locus coeruleus. J. Comp. Neurol. 394,
218–229.
Lejeune, F., and Millan, M.J. (1995). Activation of dopamine D3 autoreceptors inhibits firing of ventral tegmental dopaminergic neurones in vivo. Eur. J.
Pharmacol. 275, R7–R9.
Lesage, F., Guillemare, E., Fink, M., Duprat, F., Heurteaux, C., Fosset, M., Romey, G.,
Barhanin, J., and Lazdunski, M. (1995). Molecular Properties of Neuronal G-protein- activated Inwardly Rectifying K+ Channels. J. Biol. Chem. 270, 28660–28667.
Lester, R.A.J., Clements, J.D., Westbrook, G.L., and Jahr, C.E. (1990). Channel kinetics determine the time course of NMDA receptor-mediated synaptic currents. Nature
346, 565–567.
214 Levi, G., and Raiteri, M. (1993). Carrier-mediated release of neurotransmitters.
Trends Neurosci. 16, 415–419.
Levitt, E.S., Hunnicutt, B.J., Knopp, S.J., Williams, J.T., and Bissonnette, J.M. (2013). A
selective 5-HT1a receptor agonist improves respiration in a mouse model of Rett
syndrome. J. Appl. Physiol. 115, 1626–1633.
Li, Q., Wichems, C., Heils, A., Kar, L.D.V. de, Lesch, K.-P., and Murphy, D.L. (1999).
Reduction of 5-Hydroxytryptamine (5-HT)1A-Mediated Temperature and
Neuroendocrine Responses and 5-HT1A Binding Sites in 5-HT Transporter
Knockout Mice. J. Pharmacol. Exp. Ther. 291, 999–1007.
Li, W., Doyon, W.M., and Dani, J.A. (2011). Acute In Vivo Nicotine Administration
Enhances Synchrony Among Dopamine Neurons. Biochem. Pharmacol. 82, 977–983.
Liao, Y.J., Jan, Y.N., and Jan, L.Y. (1996). Heteromultimerization of G-Protein-Gated
Inwardly Rectifying K+ Channel Proteins GIRK1 and GIRK2 and Their Altered
Expression in weaver Brain. J. Neurosci. 16, 7137–7150.
Llamosas, N., Bruzos-Cidón, C., Rodríguez, J.J., Ugedo, L., and Torrecilla, M. (2015).
Deletion of GIRK2 Subunit of GIRK Channels Alters the 5-HT1A Receptor-Mediated
Signaling and Results in a Depression-Resistant Behavior. Int. J.
Neuropsychopharmacol. 1–12.
215 L. Mohan, M., T. Vasudevan, N., K. Gupta, M., E. Martelli, E., and Naga Prasad, S. V.
(2012). G-Protein Coupled Receptor Resensitization - Appreciating the Balancing
Act of Receptor Function. Curr. Mol. Pharmacol. 5, 350–361.
Lober, R.M., Pereira, M.A., and Lambert, N.A. (2006). Rapid Activation of Inwardly
Rectifying Potassium Channels by Immobile G-Protein-Coupled Receptors. J.
Neurosci. 26, 12602–12608.
Lodge, D.J., and Grace, A.A. (2005). The Hippocampus Modulates Dopamine Neuron
Responsivity by Regulating the Intensity of Phasic Neuron Activation.
Neuropsychopharmacology 31, 1356–1361.
Logothetis, D.E., Kurachi, Y., Galper, J., Neer, E.J., and Clapham, D.E. (1987). The subunits of GTP-binding proteins activate the muscarinic K+ channel in heart. βγ
Nature 325, 321–326.
Lohse, M.J., Benovic, J.L., Codina, J., Caron, M.G., and Lefkowitz, R.J. (1990). beta-
Arrestin: a protein that regulates beta-adrenergic receptor function. Science 248,
1547–1550.
Lovenberg, T.W., Baron, B.M., de Lecea, L., Miller, J.D., Prosser, R.A., Rea, M.A., Foye,
P.E., Racke, M., Slone, A.L., Siegel, B.W., et al. (1993). A novel adenylyl cyclase- activating serotonin receptor (5-HT7) implicated in the regulation of mammalian circadian rhythms. Neuron 11, 449–458.
216 Lowry, C.A. (2002). Functional Subsets of Serotonergic Neurones: Implications for
Control of the Hypothalamic-Pituitary-Adrenal Axis. J. Neuroendocrinol. 14, 911–
923.
Lucki, I. (1998). The spectrum of behaviors influenced by serotonin. Biol. Psychiatry
44, 151–162.
Luján, R., de Velasco, E.M.F., Aguado, C., and Wickman, K. (2014). New insights into the therapeutic potential of Girk channels. Trends Neurosci. 37.
Luppi, P.-H., Aston-Jones, G., Akaoka, H., Chouvet, G., and Jouvet, M. (1995). Afferent projections to the rat locus coeruleus demonstrated by retrograde and anterograde tracing with cholera-toxin B subunit and Phaseolus vulgaris leucoagglutinin.
Neuroscience 65, 119–160.
Lüscher, C., and Slesinger, P.A. (2010). Emerging roles for G protein-gated inwardly rectifying potassium (GIRK) channels in health and disease. Nat. Rev. Neurosci. 11,
301–315.
Luttrell, L.M., Daaka, Y., and Lefkowitz, R.J. (1999). Regulation of tyrosine kinase cascades by G-protein-coupled receptors. Curr. Opin. Cell Biol. 11, 177–183.
Madras, B.K., Miller, G.M., and Fischman, A.J. (2005). The Dopamine Transporter and
Attention-Deficit/Hyperactivity Disorder. Biol. Psychiatry 57, 1397–1409.
217 Maier, S.F., Amat, J., Baratta, M.V., Paul, E., and Watkins, L.R. (2006). Behavioral
control, the medial prefrontal cortex, and resilience. Dialogues Clin. Neurosci. 8,
397–406.
Marcott, P.F., Mamaligas, A.A., and Ford, C.P. (2014). Phasic Dopamine Release
Drives Rapid Activation of Striatal D2-Receptors. Neuron 84, 164–176.
Marinelli, S., Schnell, S.A., Hack, S.P., Christie, M.J., Wessendorf, M., and Vaughan, C.W.
(2004). Serotonergic and Nonserotonergic Dorsal Raphe Neurons Are
Pharmacologically and Electrophysiologically Heterogeneous. J. Neurophysiol. 92,
3532–3537.
Markwardt, S.J., Wadiche, J.I., and Overstreet-Wadiche, L.S. (2009). Input-Specific
GABAergic Signaling to Newborn Neurons in Adult Dentate Gyrus. J. Neurosci. 29,
15063–15072.
Marotte, L.R., and Raisman, G. (1974). An experimental investigation of the relationship between granular vesicles and noradrenergic axons in the preoptic area of the rat. Brain Res. 70, 335–339.
Matsuda, W., Furuta, T., Nakamura, K.C., Hioki, H., Fujiyama, F., Arai, R., and Kaneko,
T. (2009). Single Nigrostriatal Dopaminergic Neurons Form Widely Spread and
Highly Dense Axonal Arborizations in the Neostriatum. J. Neurosci. 29, 444–453.
Matthes, H., Boschert, U., Amlaiky, N., Grailhe, R., Plassat, J.L., Muscatelli, F., Mattei,
M.G., and Hen, R. (1993). Mouse 5-hydroxytryptamine5A and 5-
218 hydroxytryptamine5B receptors define a new family of serotonin receptors: cloning,
functional expression, and chromosomal localization. Mol. Pharmacol. 43, 313–319.
McCorvy, J.D., and Roth, B.L. (2015). Structure and function of serotonin G protein-
coupled receptors. Pharmacol. Ther. 150, 129–142.
McDevitt, R.A., Tiran-Cappello, A., Shen, H., Balderas, I., Britt, J.P., Marino, R.A.M.,
Chung, S.L., Richie, C.T., Harvey, B.K., and Bonci, A. (2014). Serotonergic versus
Nonserotonergic Dorsal Raphe Projection Neurons: Differential Participation in
Reward Circuitry. Cell Rep. 8, 1857–1869.
Meisenzahl, E.M., Schmitt, G.J., Scheuerecker, J., and Möller, H.-J. (2007). The role of dopamine for the pathophysiology of schizophrenia. Int. Rev. Psychiatry Abingdon
Engl. 19, 337–345.
Meyer, P.J., Morgan, M.M., Kozell, L.B., and Ingram, S.L. (2009). Contribution of dopamine receptors to periaqueductal gray-mediated antinociception.
Psychopharmacology (Berl.) 204, 531–540.
Michelsen, K., Prickaerts, J., and Steinbusch, H. (2008). The dorsal raphe nucleus and serotonin: implications for neuroplasticity linked to major depression and
Alzheimer’s disease. Prog. Brain Res. 172, 233–264.
Millar, J., Stamford, J.A., Kruk, Z.L., and Wightman, R.M. (1985). Electrochemical, pharmacological and electrophysiological evidence of rapid dopamine release and
219 removal in the rat caudate nucleus following electrical stimulation of the median
forebrain bundle. Eur. J. Pharmacol. 109, 341–348.
Min, M.-Y., Asztely, F., Kokaia, M., and Kullmann, D.M. (1998a). Long-term potentiation and dual-component quantal signaling in the dentate gyrus. Proc. Natl.
Acad. Sci. U. S. A. 95, 4702–4707.
Min, M.-Y., Rusakov, D.A., and Kullmann, D.M. (1998b). Activation of AMPA, Kainate, and Metabotropic Receptors at Hippocampal Mossy Fiber Synapses: Role of
Glutamate Diffusion. Neuron 21, 561–570.
Moghaddam, B., and Bunney, B.S. (1989). Ionic composition of microdialysis perfusing solution alters the pharmacological responsiveness and basal outflow of striatal dopamine. J. Neurochem. 53, 652–654.
Monsma, F.J., Mahan, L.C., McVittie, L.D., Gerfen, C.R., and Sibley, D.R. (1990).
Molecular cloning and expression of a D1 dopamine receptor linked to adenylyl cyclase activation. Proc. Natl. Acad. Sci. U. S. A. 87, 6723–6727.
Do Monte, F.H., Souza, R.R., Wong, T.T., and Carobrez, A. de P. (2013). Systemic or intra-prelimbic cortex infusion of prazosin impairs fear memory reconsolidation.
Behav. Brain Res. 244, 137–141.
Morikawa, H., and Paladini, C.A. (2011). Dynamic regulation of midbrain dopamine neuron activity: intrinsic, synaptic, and plasticity mechanisms. Neuroscience 198,
95–111.
220 Morikawa, H., Manzoni, O.J., Crabbe, J.C., and Williams, J.T. (2000). Regulation of
Central Synaptic Transmission by 5-HT1BAuto- and Heteroreceptors. Mol.
Pharmacol. 58, 1271–1278.
Morikawa, H., Khodakhah, K., and Williams, J.T. (2003). Two intracellular pathways
mediate metabotropic glutamate receptor-induced Ca2+ mobilization in dopamine neurons. J. Neurosci. Off. J. Soc. Neurosci. 23, 149–157.
Morón, J.A., Brockington, A., Wise, R.A., Rocha, B.A., and Hope, B.T. (2002). Dopamine
Uptake through the Norepinephrine Transporter in Brain Regions with Low Levels of the Dopamine Transporter: Evidence from Knock-Out Mouse Lines. J. Neurosci.
22, 389–395.
Mouton, P.R., Pakkenberg, B., Gundersen, H.J., and Price, D.L. (1994). Absolute number and size of pigmented locus coeruleus neurons in young and aged individuals. J. Chem. Neuroanat. 7, 185–190.
Muly, E.C., Maddox, M., and Khan, Z.U. (2010). Distribution of D1 and D5 dopamine receptors in the primate nucleus accumbens. Neuroscience 169, 1557–1566.
Mundey, M.K., Fletcher, A., and Marsden, C.A. (1996). Effects of 8‐OHDPAT and 5‐
HT1A antagonists WAY100135 and WAY100635, on guinea‐pig behaviour and dorsal raphe 5‐HT neurone firing. Br. J. Pharmacol. 117.
Nagatsu, T. (1995). Tyrosine hydroxylase: human isoforms, structure and regulation in physiology and pathology. Essays Biochem. 30, 15–35.
221 Nair-Roberts, R.G., Chatelain-Badie, S.D., Benson, E., White-Cooper, H., Bolam, J.P.,
and Ungless, M.A. (2008). Stereological estimates of dopaminergic, GABAergic and
glutamatergic neurons in the ventral tegmental area, substantia nigra and
retrorubral field in the rat. Neuroscience 152, 1024–1031.
Nassirpour, R., Bahima, L., Lalive, A.L., Lüscher, C., Luján, R., and Slesinger, P.A.
(2010). Morphine- and CaMKII dependent enhancement of GIRK channel signaling
in hippocampal neurons. J. Neurosci. Off. J. Soc. Neurosci. 30, 13419–13430.
Neer, E.J. (1995). Heterotrimeric C proteins: Organizers of transmembrane signals.
Cell 80, 249–257.
Neves, S.R., Ram, P.T., and Iyengar, R. (2002). G Protein Pathways. Science 296,
1636–1639.
Nielsen, T.A., DiGregorio, D.A., and Silver, R.A. (2004). Modulation of glutamate
mobility reveals the mechanism underlying slow-rising AMPAR EPSCs and the diffusion coefficient in the synaptic cleft. Neuron 42, 757–771.
Nirenberg, M.J., Liu, Y., Peter, D., Edwards, R.H., and Pickel, V.M. (1995). The vesicular monoamine transporter 2 is present in small synaptic vesicles and preferentially localizes to large dense core vesicles in rat solitary tract nuclei. Proc.
Natl. Acad. Sci. 92, 8773–8777.
Nirenberg, M.J., Chan, J., Liu, Y., Edwards, R.H., and Pickel, V.M. (1996).
Ultrastructural Localization of the Vesicular Monoamine Transporter-2 in Midbrain
222 Dopaminergic Neurons: Potential Sites for Somatodendritic Storage and Release of
Dopamine. J. Neurosci. 16, 4135–4145.
Nobles, M., Benians, A., and Tinker, A. (2005). Heterotrimeric G proteins precouple
with G protein-coupled receptors in living cells. Proc. Natl. Acad. Sci. U. S. A. 102,
18706–18711.
O’Connor, J.J., and Kruk, Z.L. (1991). Fast cyclic voltammetry can be used to measure stimulated endogenous 5-hydroxytryptamine release in untreated rat brain slices. J.
Neurosci. Methods 38, 25–33.
Oláh, S., Füle, M., Komlósi, G., Varga, C., Báldi, R., Barzó, P., and Tamás, G. (2009).
Regulation of cortical microcircuits by unitary GABA-mediated volume transmission. Nature 461, 1278–1281.
Otis, T.S., and Mody, I. (1992). Differential activation of GABAA and GABAB receptors by spontaneously released transmitter. J. Neurophysiol. 67, 227–235.
Otis, T.S., Wu, Y.C., and Trussell, L.O. (1996). Delayed clearance of transmitter and the role of glutamate transporters at synapses with multiple release sites. J.
Neurosci. 16, 1634–1644.
Overstreet, L.S., and Westbrook, G.L. (2003). Synapse density regulates independence at unitary inhibitory synapses. J. Neurosci. Off. J. Soc. Neurosci. 23,
2618–2626.
223 Overton, P.G., and Clark, D. (1997). Burst firing in midbrain dopaminergic neurons.
Brain Res. Rev. 25, 312–334.
Paladini, C.A., and Roeper, J. (2014). Generating bursts (and pauses) in the dopamine
midbrain neurons. Neuroscience 282, 109–121.
Paladini, C.A., and Tepper, J.M. (1999). GABA(A) and GABA(B) antagonists differentially affect the firing pattern of substantia nigra dopaminergic neurons in vivo. Synap. N. Y. N 32, 165–176.
Paladini, C.A., and Williams, J.T. (2004). Noradrenergic Inhibition of Midbrain
Dopamine Neurons. J. Neurosci. 24, 4568–4575.
Palij, P., and Stamford, J.A. (1994). Real-time monitoring of endogenous
noradrenaline release in rat brain slices using fast cyclic voltammetry: 3. Selective
detection of noradrenaline efflux in the locus coeruleus. Brain Res. 634, 275–282.
Palkovits, M., Brownstein, M., and Saavedra, J.M. (1974). Serotonin content of the
brain stem nuclei in the rat. Brain Res. 80, 237–249.
Pan, Z.Z., and Williams, J.T. (1989). GABA-and glutamate-mediated synaptic
potentials in rat dorsal raphe neurons in vitro. J. Neurophysiol. 61, 719–726.
Pan, Z.Z., Colmers, W.F., and Williams, J.T. (1989). 5-HT-mediated synaptic potentials in the dorsal raphe nucleus: interactions with excitatory amino acid and GABA neurotransmission. J. Neurophysiol. 62, 481–486.
224 Patel, J.C., Witkovsky, P., Avshalumov, M.V., and Rice, M.E. (2009). Mobilization of
Calcium from Intracellular Stores Facilitates Somatodendritic Dopamine Release. J.
Neurosci. 29, 6568–6579.
Pearce, R.A. (1993). Physiological evidence for two distinct GABAA responses in rat
hippocampus. Neuron 10, 189–200.
Perez, D.M. (2007). Structure-Function of 1-Adrenergic Receptors. Biochem.
Pharmacol. 73, 1051–1062. α
Perez, R.G., Waymire, J.C., Lin, E., Liu, J.J., Guo, F., and Zigmond, M.J. (2002). A Role for
-Synuclein in the Regulation of Dopamine Biosynthesis. J. Neurosci. 22, 3090–3099.
α Peter, D., Jimenez, J., Liu, Y., Kim, J., and Edwards, R.H. (1994). The chromaffin
granule and synaptic vesicle amine transporters differ in substrate recognition and
sensitivity to inhibitors. J. Biol. Chem. 269, 7231–7237.
Peyron, C., Petit, J.-M., Rampon, C., Jouvet, M., and Luppi, P.-H. (1998). Forebrain
afferents to the rat dorsal raphe nucleus demonstrated by retrograde and
anterograde tracing methods. Neuroscience 82, 443–468.
Pfaffinger, P.J., Martin, J.M., Hunter, D.D., Nathanson, N.M., and Hille, B. (1985). GTP-
binding proteins couple cardiac muscarinic receptors to a K channel. Nature 317,
536–538.
225 Phillips, P.E.M., Hancock, P.J., and Stamford, J.A. (2002). Time window of autoreceptor-mediated inhibition of limbic and striatal dopamine release. Synap. N.
Y. N 44, 15–22.
Phillips, P.E.M., Stuber, G.D., Heien, M.L.A.V., Wightman, R.M., and Carelli, R.M.
(2003). Subsecond dopamine release promotes cocaine seeking. Nature 422, 614–
618.
Pickel, V.M., Joh, T.H., and Reis, D.J. (1976). Monoamine-synthesizing enzymes in central dopaminergic, noradrenergic and serotonergic neurons.
Immunocytochemical localization by light and electron microscopy. J. Histochem.
Cytochem. Off. J. Histochem. Soc. 24, 792–306.
Pickel, V.M., Chan, J., and Nirenberg, M.J. (2002). Region-specific targeting of dopamine D2-receptors and somatodendritic vesicular monoamine transporter 2
(VMAT2) within ventral tegmental area subdivisions. Synap. N. Y. N 45, 113–124.
Piñeyro, G., and Blier, P. (1999). Autoregulation of serotonin neurons: role in antidepressant drug action. Pharmacol. Rev. 51, 533–591.
Pitcher, J.A., Hall, R.A., Daaka, Y., Zhang, J., Ferguson, S.S.G., Hester, S., Miller, S.,
Caron, M.G., Lefkowitz, R.J., and Barak, L.S. (1998). The G Protein-coupled Receptor
Kinase 2 Is a Microtubule-associated Protein Kinase That Phosphorylates Tubulin. J.
Biol. Chem. 273, 12316–12324.
226 Portas, C.M., Thakkar, M., Rainnie, D., and McCarley, R.W. (1996). Microdialysis
perfusion of 8-hydroxy-2-(di-n-propylamino)tetralin (8-OH-DPAT) in the dorsal raphe nucleus decreases serotonin release and increases rapid eye movement sleep in the freely moving cat. J. Neurosci. Off. J. Soc. Neurosci. 16, 2820–2828.
Pothos, E., Desmond, M., and Sulzer, D. (1996). l-3,4-Dihydroxyphenylalanine
Increases the Quantal Size of Exocytotic Dopamine Release In Vitro. J. Neurochem.
66, 629–636.
De Potter, W.P., Smith, A.D., and De Schaepdryver, A.F. (1970). Subcellular
-hydroxylase in noradrenergicfractionation of vesicles. splenic nerve:Tissue ATP,Cell 2 chromogranin, 529–546. A and dopamine β
Povlock, S.L., and Schenk, J.O. (1997). A Multisubstrate Kinetic Mechanism of
Dopamine Transport in the Nucleus Accumbens and Its Inhibition by Cocaine. J.
Neurochem. 69, 1093–1105.
Puumala, T., Greijus, S., Narinen, K., Haapalinna, A., Riekkinen Sr, P., and Sirviö, J.
(1998). Stimulation of alpha-1 adrenergic receptors facilitates spatial learning in rats. Eur. Neuropsychopharmacol. 8, 17–26.
Qi, J., Zhang, S., Wang, H.-L., Wang, H., de Jesus Aceves Buendia, J., Hoffman, A.F.,
Lupica, C.R., Seal, R.P., and Morales, M. (2014). A glutamatergic reward input from the dorsal raphe to ventral tegmental area dopamine neurons. Nat. Commun. 5,
5390.
227 Rall, T.W., and Sutherland, E.W. (1962). Adenyl Cyclase II. The Enzymatically
Catalyzed Formation of Adenosine 3’,5’-Phosphate and Inorganic Pyrophosphate from Adenosine Triphosphate. J. Biol. Chem. 237, 1228–1232.
Ramamoorthy, S., Bauman, A.L., Moore, K.R., Han, H., Yang-Feng, T., Chang, A.S.,
Ganapathy, V., and Blakely, R.D. (1993). Antidepressant- and cocaine-sensitive human serotonin transporter: molecular cloning, expression, and chromosomal localization. Proc. Natl. Acad. Sci. U. S. A. 90, 2542–2546.
Ramamoorthy, S., Shippenberg, T.S., and Jayanthi, L.D. (2011). Regulation of
Monoamine Transporters: Role of Transporter Phosphorylation. Pharmacol. Ther.
129, 220–238.
Ramirez, D.M., and Kavalali, E.T. (2011). Differential regulation of spontaneous and evoked neurotransmitter release at central synapses. Curr. Opin. Neurobiol. 21,
275–282.
Ranade, S.P., and Mainen, Z.F. (2009). Transient Firing of Dorsal Raphe Neurons
Encodes Diverse and Specific Sensory, Motor, and Reward Events. J. Neurophysiol.
102, 3026–3037.
Rasmussen, S.G.F., Choi, H.-J., Fung, J.J., Pardon, E., Casarosa, P., Chae, P.S., DeVree,
B.T., Rosenbaum, D.M., Thian, F.S., Kobilka, T.S., et al. (2011). Structure of a nanobody-stabilized active state of 469, 175–180.
the β2 adrenoceptor. Nature
228 Reyes, S., Fu, Y., Double, K., Thompson, L., Kirik, D., Paxinos, G., and Halliday, G.M.
(2012). GIRK2 expression in dopamine neurons of the substantia nigra and ventral
tegmental area. J. Comp. Neurol. 520, 2591–2607.
Riad, M., Garcia, S., Watkins, K.C., Jodoin, N., Doucet, É., Langlois, X., Mestikawy, S. El,
Hamon, M., and Descarries, L. (2000). Somatodendritic localization of 5-HT1A and preterminal axonal localization of 5-HT1B serotonin receptors in adult rat brain. J.
Comp. Neurol. 417, 181–194.
Rice, M.E. (2000). Distinct regional differences in dopamine-mediated volume transmission. B.-P. in B. Research, ed. (Elsevier), pp. 277–290.
Rice, M.E., and Cragg, S.J. (2008). Dopamine Spillover after Quantal Release:
Rethinking Dopamine Transmission in the Nigrostriatal Pathway. Brain Res. Rev. 58,
303–313.
Rice, M.E., Richards, C.D., Nedergaard, S., Hounsgaard, J., Nicholson, C., and
Greenfield, S.A. (1994). Direct monitoring of dopamine and 5-HT release in substantia nigra and ventral tegmental area in vitro. Exp. Brain Res. 100, 395–406.
Rice, M.E., Cragg, S.J., and Greenfield, S.A. (1997). Characteristics of Electrically
Evoked Somatodendritic Dopamine Release in Substantia Nigra and Ventral
Tegmental Area In Vitro. J. Neurophysiol. 77, 853–862.
Richardson-Jones, J.W., Craige, C.P., Guiard, B.P., Stephen, A., Metzger, K.L., Kung,
H.F., Gardier, A.M., Dranovsky, A., David, D.J., Beck, S.G., et al. (2010). 5-HT1A
229 Autoreceptor Levels Determine Vulnerability to Stress and Response to
Antidepressants. Neuron 65, 40–52.
Rosenbaum, D.M., Rasmussen, S.G.F., and Kobilka, B.K. (2009). The structure and function of G-protein-coupled receptors. Nature 459, 356–363.
Rossi, D.J., and Hamann, M. (1998). Spillover-Mediated Transmission at Inhibitory
Synapses Promoted by High Affinity 6 Subunit GABAA Receptors and Glomerular
Geometry. Neuron 20, 783–795. α
Rusakov, D.A., and Kullmann, D.M. (1998a). Extrasynaptic Glutamate Diffusion in the
Hippocampus: Ultrastructural Constraints, Uptake, and Receptor Activation. J.
Neurosci. 18, 3158–3170.
Rusakov, D.A., and Kullmann, D.M. (1998b). Geometric and viscous components of the tortuosity of the extracellular space in the brain. Proc. Natl. Acad. Sci. U. S. A. 95,
8975–8980.
Sánchez-Fernández, G., Cabezudo, S., García-Hoz, C., Benincá, C., Aragay, A.M., Mayor
Jr., F., and Ribas, C. (2014). G q signalling: The new and the old. Cell. Signal. 26, 833–
848. α
Sara, S.J. (2009). The locus coeruleus and noradrenergic modulation of cognition.
Nat. Rev. Neurosci. 10, 211–223.
230 Sarantis, M., Ballerini, L., Miller, B., Silver, R.A., Edwards, M., and Attwell, D. (1993).
Glutamate uptake from the synaptic cleft does not shape the decay of the non-NMDA
component of the synaptic current. Neuron 11, 541–549.
Sari, Y. (2004). Serotonin receptors: from protein to physiological function and
behavior. Neurosci. Biobehav. Rev. 28, 565–582.
Sari, Y., Miquel, M.C., Brisorgueil, M.J., Ruiz, G., Doucet, E., Hamon, M., and Verge, D.
(1999). Cellular and subcellular localization of 5-hydroxytryptamine1B receptors in the rat central nervous system: immunocytochemical autoradiographic and lesion studies. Neuroscience 88, 899–915.
Scanziani, M. (2000). GABA Spillover Activates Postsynaptic GABAB Receptors to
Control Rhythmic Hippocampal Activity. Neuron 25, 673–681.
Schechter, L.E., Lin, Q., Smith, D.L., Zhang, G., Shan, Q., Platt, B., Brandt, M.R., Dawson,
L.A., Cole, D., Bernotas, R., et al. (2007). Neuropharmacological Profile of Novel and
Selective 5-HT6 Receptor Agonists: WAY-181187 and WAY-208466.
Neuropsychopharmacology 33, 1323–1335.
Scheinin, M., Lomasney, J.W., Hayden-Hixson, D.M., Schambra, U.B., Caron, M.G.,
Lefkowitz, R.J., and Fremeau Jr., R.T. (1994). Distribution of 2-adrenergic receptor subtype gene expression in rat brain. Mol. Brain Res. 21, 133α–149.
231 Schiöth, H.B., and Fredriksson, R. (2005). The GRAFS classification system of G-
protein coupled receptors in comparative perspective. Gen. Comp. Endocrinol. 142,
94–101.
Schmitz, Y., Schmauss, C., and Sulzer, D. (2002). Altered dopamine release and uptake kinetics in mice lacking D2 receptors. J. Neurosci. Off. J. Soc. Neurosci. 22,
8002–8009.
Schmitz, Y., Benoit-Marand, M., Gonon, F., and Sulzer, D. (2003). Presynaptic regulation of dopaminergic neurotransmission. J. Neurochem. 87, 273–289.
Schoffelmeer, A.N., Hogenboom, F., Mulder, A.H., Ronken, E., Stoof, J.C., and Drukarch,
B. (1994). Dopamine displays an identical apparent affinity towards functional
dopamine D1 and D2 receptors in rat striatal slices: possible implications for the
regulatory role of D2 receptors. Synap. N. Y. N 17, 190–195.
Schott, B.H., Frischknecht, R., Debska-Vielhaber, G., John, N., Behnisch, G., Düzel, E.,
Gundelfinger, E.D., and Seidenbecher, C.I. (2010). Membrane-bound catechol-O- methyl transferase in cortical neurons and glial cells is intracellularly oriented. Mol.
Psychiatry 1, 142.
Schultz, W. (1997). Dopamine neurons and their role in reward mechanisms. Curr.
Opin. Neurobiol. 7, 191–197.
Schultz, W. (2007). Behavioral dopamine signals. Trends Neurosci. 30, 203–210.
232 Schultz, D.W., Ruffieux, A., and Aebischer, P. (1983). The activity of pars compacta
neurons of the monkey substantia nigra in relation to motor activation. Exp. Brain
Res. 51, 377–387.
Schultz, W., Studer, A., Romo, R., Sundstrom, E., Jonsson, G., and Scarnati, E. (1989).
Deficits in reaction times and movement times as correlates of hypokinesia in monkeys with MPTP-induced striatal dopamine depletion. J. Neurophysiol. 61, 651–
668.
Schultz, W., Dayan, P., and Montague, P.R. (1997). A Neural Substrate of Prediction
and Reward. Science 275, 1593–1599.
Scott, M.M., Wylie, C.J., Lerch, J.K., Murphy, R., Lobur, K., Herlitze, S., Jiang, W., Conlon,
R.A., Strowbridge, B.W., and Deneris, E.S. (2005). A genetic approach to access
serotonin neurons for in vivo and in vitro studies. Proc. Natl. Acad. Sci. U. S. A. 102,
16472–16477.
Sesack, S.R., Aoki, C., and Pickel, V.M. (1994). Ultrastructural localization of D2
receptor-like immunoreactivity in midbrain dopamine neurons and their striatal
targets. J. Neurosci. 14, 88–106.
Shipley, M.T., Fu, L., Ennis, M., Liu, W.-L., and Aston-Jones, G. (1996). Dendrites of
locus coeruleus neurons extend preferentially into two pericoerulear zones. J. Comp.
Neurol. 365, 56–68.
233 Sibley, D.R., Mahan, L.C., and Creese, I. (1983). Dopamine receptor binding on intact cells. Absence of a high-affinity agonist-receptor binding state. Mol. Pharmacol. 23,
295–302.
Siehler, S. (2009). Regulation of RhoGEF proteins by G12/13-coupled receptors. Br.
J. Pharmacol. 158, 41–49.
Silver, R.A., Cull-Candy, S.G., and Takahashi, T. (1996). Non-NMDA glutamate receptor occupancy and open probability at a rat cerebellar synapse with single and multiple release sites. J. Physiol. 494 ( Pt 1), 231–250.
Singewald, N., and Philippu, A. (1998). Release of neurotransmitters in the locus coeruleus. Prog. Neurobiol. 56, 237–267.
Slesinger, P.A., Patil, N., Liao, Y.J., Jan, Y.N., Jan, L.Y., and Cox, D.R. (1996). Functional
Effects of the Mouse weaver Mutation on G Protein–Gated Inwardly Rectifying K+
Channels. Neuron 16, 321–331.
Smith, I.D., and Grace, A.A. (1992). Role of the subthalamic nucleus in the regulation of nigral dopamine neuron activity. Synap. N. Y. N 12, 287–303.
Sodickson, D.L., and Bean, B.P. (1996). GABAB Receptor-Activated Inwardly
Rectifying Potassium Current in Dissociated Hippocampal CA3 Neurons. J. Neurosci.
16, 6374–6385.
234 Sokoloff, P., Giros, B., Martres, M.-P., Bouthenet, M.-L., and Schwartz, J.-C. (1990).
Molecular cloning and characterization of a novel dopamine receptor (D3) as a target for neuroleptics. Nature 347, 146–151.
Sora, I., Wichems, C., Takahashi, N., Li, X.-F., Zeng, Z., Revay, R., Lesch, K.-P., Murphy,
D.L., and Uhl, G.R. (1998). Cocaine reward models: Conditioned place preference can be established in dopamine- and in serotonin-transporter knockout mice. Proc. Natl.
Acad. Sci. U. S. A. 95, 7699–7704.
Spano, P.F., Govoni, S., and Trabucchi, M. (1978). Studies on the pharmacological properties of dopamine receptors in various areas of the central nervous system.
Adv. Biochem. Psychopharmacol. 19, 155–165.
Stamford, J.A., Kruk, Z.L., Palij, P., and Millar, J. (1988). Diffusion and uptake of dopamine in rat caudate and nucleus accumbens compared using fast cyclic voltammetry. Brain Res. 448, 381–385.
Stamp, J.A., and Semba, K. (1995). Extent of colocalization of serotonin and GABA in the neurons of the rat raphe nuclei. Brain Res. 677, 39–49.
Steinbusch, H.W., Nieuwenhuys, R., Verhofstad, A.A., and Van der Kooy, D. (1981).
The nucleus raphe dorsalis of the rat and its projection upon the caudatoputamen. A combined cytoarchitectonic, immunohistochemical and retrograde transport study.
J. Physiol. (Paris) 77, 157–174.
235 Stratford, T.R., and Wirtshafter, D. (1990). Ascending dopaminergic projections from
the dorsal raphe nucleus in the rat. Brain Res. 511, 173–176.
Stuber, G.D., Hnasko, T.S., Britt, J.P., Edwards, R.H., and Bonci, A. (2010).
Dopaminergic terminals in the nucleus accumbens but not the dorsal striatum co- release glutamate. J. Neurosci. Off. J. Soc. Neurosci. 30, 8229–8233.
Sunahara, R.K., Guan, H.-C., O’Dowd, B.F., Seeman, P., Laurier, L.G., Ng, G., George, S.R.,
Torchia, J., Van Tol, H.H.M., and Niznik, H.B. (1991). Cloning of the gene for a human dopamine D5 receptor with higher affinity for dopamine than D1. Nature 350, 614–
619.
Szabadi, E. (2013). Functional neuroanatomy of the central noradrenergic system. J.
Psychopharmacol. (Oxf.) 27, 659–693.
Szabadics, J., Tamás, G., and Soltesz, I. (2007). Different transmitter transients underlie presynaptic cell type specificity of GABAA,slow and GABAA,fast. Proc. Natl.
Acad. Sci. U. S. A. 104, 14831–14836.
Szapiro, G., and Barbour, B. (2007). Multiple climbing fibers signal to molecular layer interneurons exclusively via glutamate spillover. Nat. Neurosci. 10, 735–742.
Szot, P., Reigel, C.E., White, S.S., and Veith, R.C. (1996). Alterations in mRNA expression of systems that regulate neurotransmitter synaptic content in seizure- naive genetically epilepsy-prone rat (GEPR): transporter proteins and rate-limiting
236 synthesizing enzymes for norepinephrine, dopamine and serotonin. Brain Res. Mol.
Brain Res. 43, 233–245.
Tiberi, M., Jarvie, K.R., Silvia, C., Falardeau, P., Gingrich, J.A., Godinot, N., Bertrand, L.,
Yang-Feng, T.L., Fremeau, R.T., and Caron, M.G. (1991). Cloning, molecular
characterization, and chromosomal assignment of a gene encoding a second D1
dopamine receptor subtype: differential expression pattern in rat brain compared
with the D1A receptor. Proc. Natl. Acad. Sci. U. S. A. 88, 7491–7495.
Tobler, P.N., Fiorillo, C.D., and Schultz, W. (2005). Adaptive coding of reward value
by dopamine neurons. Science 307, 1642–1645.
Van Tol, H.H.M., Bunzow, J.R., Guan, H.-C., Sunahara, R.K., Seeman, P., Niznik, H.B.,
and Civelli, O. (1991). Cloning of the gene for a human dopamine D4 receptor with
high affinity for the antipsychotic clozapine. Nature 350, 610–614.
Torkaman-Boutorabi, A., Danyali, F., Oryan, S., Ebrahimi-Ghiri, M., and Zarrindast,
M.-R. (2014). Hippocampal -adrenoceptors involve in the effect of histamine on
spatial learning. Physiol. Behav.α 129, 17–24.
Torrecilla, M., Marker, C.L., Cintora, S.C., Stoffel, M., Williams, J.T., and Wickman, K.
(2002). G-Protein-Gated Potassium Channels Containing Kir3.2 and Kir3.3 Subunits
Mediate the Acute Inhibitory Effects of Opioids on Locus Ceruleus Neurons. J.
Neurosci. 22, 4328–4334.
237 Tritsch, N.X., Oh, W.-J., Gu, C., and Sabatini, B.L. (2014). Midbrain dopamine neurons sustain inhibitory transmission using plasma membrane uptake of GABA, not synthesis. eLife 3.
Trussell, L.O., Zhang, S., and Ramant, I.M. (1993). Desensitization of AMPA receptors upon multiquantal neurotransmitter release. Neuron 10, 1185–1196.
Trzaskowski, B., Latek, D., Yuan, S., Ghoshdastider, U., Debinski, A., and Filipek, S.
(2012). Action of Molecular Switches in GPCRs - Theoretical and Experimental
Studies. Curr. Med. Chem. 19, 1090–1109.
Tucker, S.J., Pessia, M., and Adelman, J.P. (1996). Muscarine-gated K+ channel: subunit stoichiometry and structural domains essential for G protein stimulation.
Am. J. Physiol. - Heart Circ. Physiol. 271, H379–H385.
Ulmanen, I., Peränen, J., Tenhunen, J., Tilgmann, C., Karhunen, T., Panula, P.,
Bernasconi, L., Aubry, J.-P., and Lundström, K. (1997). Expression and Intracellular
Localization of Catechol O-methyltransferase in Transfected Mammalian Cells. Eur. J.
Biochem. 243, 452–459.
Ungless, M.A. (2004). Dopamine: the salient issue. Trends Neurosci. 27, 702–706.
Ungless, M.A., Argilli, E., and Bonci, A. (2010). Effects of stress and aversion on dopamine neurons: Implications for addiction. Neurosci. Biobehav. Rev. 35, 151–
156.
238 Vandermaelen, C.P., and Aghajanian, G.K. (1983). Electrophysiological and
pharmacological characterization of serotonergic dorsal raphe neurons recorded
extracellularly and intracellularly in rat brain slices. Brain Res. 289, 109–119.
Vasudeva, R.K., Lin, R.C.S., Simpson, K.L., and Waterhouse, B.D. (2011). Functional organization of the dorsal raphe efferent system with special consideration of nitrergic cell groups. J. Chem. Neuroanat. 41, 281–293.
Vertes, R.P. (1991). A PHA-L analysis of ascending projections of the dorsal raphe nucleus in the rat. J. Comp. Neurol. 313, 643–668.
Vivo, M.D., and Maayani, S. (1986). Characterization of the 5-hydroxytryptamine1a receptor-mediated inhibition of forskolin-stimulated adenylate cyclase activity in guinea pig and rat hippocampal membranes. J. Pharmacol. Exp. Ther. 238, 248–253.
Volkow, N.D., Fowler, J.S., Wang, G.J., Baler, R., and Telang, F. (2009). Imaging
dopamine’s role in drug abuse and addiction. Neuropharmacology 56 Suppl 1, 3–8.
Wachtel, S.R., and Abercrombie, E.D. (1994). L-3,4-dihydroxyphenylalanine-induced dopamine release in the striatum of intact and 6-hydroxydopamine-treated rats: differential effects of monoamine oxidase A and B inhibitors. J. Neurochem. 63, 108–
117.
Wadiche, J.I., and Jahr, C.E. (2005). Patterned expression of Purkinje cell glutamate transporters controls synaptic plasticity. Nat. Neurosci. 8, 1329–1334.
239 Wassef, M., Berod, A., and Sotelo, C. (1981). Dopaminergic dendrites in the pars reticulata of the rat substantia nigra and their striatal input. Combined immunocytochemical localization of tyrosine hydroxylase and anterograde degeneration. Neuroscience 6, 2125–2139.
Watabe-Uchida, M., Zhu, L., Ogawa, S.K., Vamanrao, A., and Uchida, N. (2012). Whole-
Brain Mapping of Direct Inputs to Midbrain Dopamine Neurons. Neuron 74, 858–
873.
Werkman, T.R., McCreary, A.C., Kruse, C.G., and Wadman, W.J. (2011). NK3 receptors mediate an increase in firing rate of midbrain dopamine neurons of the rat and the guinea pig. Synap. N. Y. N 65, 814–826.
Westlund, K.N., Krakower, T.J., Kwan, S.W., and Abell, C.W. (1993). Intracellular distribution of monoamine oxidase A in selected regions of rat and monkey brain and spinal cord. Brain Res. 612, 221–230.
Wickman, K., Karschin, C., Karschin, A., Picciotto, M.R., and Clapham, D.E. (2000).
Brain Localization and Behavioral Impact of the G-Protein-Gated K+ Channel
Subunit GIRK4. J. Neurosci. 20, 5608–5615.
Wickman, K.D., Iñiguez-Lluhi, J.A., Davenport, P.A., Taussig, R., Krapivinsky, G.B.,
Linder, M.E., Gilman, A.G., and Clapham, D.E. (1994). Recombinant G-protein - subunits activate the muscarinic-gated atrial potassium channel. Nature 368,βγ 255–
257.
240 Williams, J.T., North, R.A., Shefner, S.A., Nishi, S., and Egan, T.M. (1984). Membrane
properties of rat locus coeruleus neurones. Neuroscience 13, 137–156.
Williams, J.T., Colmers, W.F., and Pan, Z.Z. (1988). Voltage- and ligand-activated inwardly rectifying currents in dorsal raphe neurons in vitro. J. Neurosci. 8, 3499–
3506.
Wilson, C.J., Groves, P.M., and Fifková, E. (1977). Monoaminergic synapses, including dendro-dendritic synapses in the rat substantia nigra. Exp. Brain Res. 30, 161–174.
Wimalasena, K. (2011). Vesicular monoamine transporters: Structure-function, pharmacology, and medicinal chemistry. Med. Res. Rev. 31, 483–519.
Wise, R.A. (2004). Dopamine, learning and motivation. Nat. Rev. Neurosci. 5, 483–
494.
Witkovsky, P., Patel, J.C., Lee, C.R., and Rice, M.E. (2009). Immunocytochemical
identification of proteins involved in dopamine release from the somatodendritic
compartment of nigral dopaminergic neurons. Neuroscience 164, 488–496.
Yang, J., Jan, Y.N., and Jan, L.Y. (1995). Determination of the subunit stoichiometry of
an inwardly rectifying potassium channel. Neuron 15, 1441–1447.
Yeagle, P.L., and Albert, A.D. (2003). A Conformational Trigger for Activation of a G
Protein by a G Protein-Coupled Receptor†. Biochemistry (Mosc.) 42, 1365–1368.
Zhou, Q.-Y., and Palmiter, R.D. (1995). Dopamine-deficient mice are severely
hypoactive, adipsic, and aphagic. Cell 83, 1197–1209.
241 Zhou, Q.-Y., Grandy, D.K., Thambi, L., Kushner, J.A., Tol, H.H.M.V., Cone, R., Pribnow,
D., Salon, J., Bunzow, J.R., and Civelli, O. (1990). Cloning and expression of human and rat Dt dopamine receptors. Nature 347, 76–80.
Zhu, H., and Zhou, W. (2001). Morphine Induces Synchronous Oscillatory Discharges
in the Rat Locus Coeruleus. J. Neurosci. 21, RC179–RC179.
Zoli, M., Torri, C., Ferrari, R., Jansson, A., Zini, I., Fuxe, K., and Agnati, L.F. (1998). The emergence of the volume transmission concept. Brain Res. Brain Res. Rev. 26, 136–
147.
242