A Dissertation
entitled
Serotonin Modulates a Calcium-Driven Negative Feedback Loop in a C. elegans
Nociceptor
by
Jeffrey A Zahratka
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in
Biology
_________________________________________
Dr. Bruce Bamber, Committee Chair
_________________________________________
Dr. Patricia R. Komuniecki, Dean
College of Graduate Studies
The University of Toledo
December 2015
Copyright 2015, Jeffrey Allen Zahratka
This document is copyrighted material. Under copyright law, no parts of this document
may be reproduced without the expressed permission of the author.
An Abstract of
Serotonin Modulates a Calcium-Driven Negative Feedback Loop in a C. elegans
Nociceptor
by
Jeffrey A Zahratka
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in
Biology
The University of Toledo
December 2015
Neuromodulation in sensory circuits is critical because it allows an organism to
respond appropriately to a given stimulus. Sensory systems are modulated by monoamine
neurotransmitters as well as neuropeptides, which act in concert to regulate sensory
circuits to give rise to complex behavioral states. One common technique for studying
sensory circuits is brain activity mapping, where circuits are probed with fluorescent
indicators whose readouts are related directly to neuronal activity. In the present work,
we focus on the modulation of a pair of sensory neurons, the ASHs, in the nematode
Caenorhabditis elegans. ASHs are polymodal, nociceptive neurons that are extensively
modulated by monoamines and neuropeptides. Using a combination of genetics, Ca2+
imaging, electrophysiology, and behavioral assays, we have identified a complex instance
where the monoamine serotonin (5-HT) stimulates aversive behaviors and neuronal
depolarization, but decreases sensory-evoked Ca2+ signals, indicating that the recorded
Ca2+ levels do not positively correlate with neuronal activity. Mechanistically, 5-HT is
likely acting through the SER-5 receptor and Gαq signaling in ASHs to downregulate
Ca2+ directly by initiating a Ca2+-driven negative feedback loop targeting the L-type Ca2+
iii
channel EGL-19. Together, these studies reveal a complex inhibitory feedback
mechanism for sensory modulation, and have broad implications for activity mapping of
complex neural circuits.
iv
My dissertation is dedicated to my friends, family, and fiancée Abby, for all of their
outstanding love and support throughout the years. It is also dedicated to Bill Molina,
Cindy Llewellyn, the late Bill Kipp, Dr. Katie Alex, Bruce, and Rick for making me into
the scientist I am today.
I graciously thank you all.
Acknowledgements
I would like to thank Drs. Bruce Bamber and Richard Komuniecki for their
immense help and guidance that was necessary to complete this project, Bruce for
allowing me the freedom to see where the project would take me, and Rick for always
challenging my line of thinking to make sure I was staying on track. I would also like to
thank Dr. Robert Steven and Amanda Korchnak for their continued help through the
duration of my PhD training, as without their help quite a bit of this work would not have
been possible. I also would like to acknowledge members of the Bamber and Komuniecki
labs: Paul Williams, Robert Layne, Hilary Linzie, Dan Morrissey, Matt Rodenbeck,
Emily Scott, Lama Karam, Dr. Gareth Harris, Dr. Holly Mills, and Philip Summers for
their input and help across all aspects of these projects. Finally, I would also like to
acknowledge my committee, Drs. Bruce Bamber, Richard Komuniecki, Robert Steven,
Guofa Liu, Scott Molitor, and Ajith Karunarathne.
v
Table of Contents
Abstract .............................................................................................................................. iii
Acknowledgements ..............................................................................................................v
Table of Contents ............................................................................................................... vi
List of Figures ......................................................................................................................x
List of Abbreviations ......................................................................................................... xi
List of Symbols ................................................................................................................ xiv
1
Introduction…………. .............................................................................................1
1.1 Voltage-gated Calcium Channels ......................................................................2
1.1.1 CaV1 (L-type) Channels ......................................................................7
1.1.1.1 Modulation of CaV1 Channels by Monoamines ................10
1.1.2 CaV2 (N/P/Q/R-type) Channels ........................................................12
1.1.3 CaV3 (T-type) Channels ....................................................................13
1.2 Intracellular Ca2+ Receptors: IP3Rs and RyRs .................................................16
1.2.1 IP3 Receptors.....................................................................................16
1.2.2 Ryanodine Receptors ........................................................................18
1.3 C.elegans: A model organism for studying sensory neuron modulation .........20
1.3.1 The C. elegans Nervous System: Small but Complex ......................21
1.3.2 The ASH nociceptors as a model for sensory modulation ................22
1.3.3 In vivo recordings from C. elegans neurons .....................................24
vi
1.3.4 Calcium channels in C. elegans ........................................................27
1.3.4.1 EGL-19 ..............................................................................28
1.3.4.2 UNC-2 ................................................................................29
1.3.4.3 CCA-1 ................................................................................30
1.3.4.4 ITR-1 ..................................................................................31
1.3.4.5 UNC-68 ..............................................................................32
1.3.5 Monoamines and neuropeptides extensively modulate ASHs ..................................33
1.3.5.1 NT modulation of ASHs is complex and requires multiple
layers of neurons…….. ......................................................................................................33
1.3.5.2 Calcium Channels as Sensory Modulators ........................36
2
Materials and Methods...........................................................................................37
2.1 Worm strains and maintenance ........................................................................37
2.2 RNA interference .............................................................................................37
2.3 Behavioral assays .............................................................................................38
2.4 Calcium imaging ..............................................................................................39
2.5 Electrophysiology ............................................................................................41
2.6 Statistical analysis ............................................................................................42
3
Monoamines and neuropeptides cell-specifically modulate ASH-mediated
aversive behavior in Caenorhabditis elegans ....................................................................45
3.1 Results…… ......................................................................................................45
3.1.1 ASH neurons, but not ADL or AWB neurons, are required for
saturating 1-octanol sensation. ...........................................................................................45
3.1.2 Basic characterization of ASH 1-octanol responses .........................48
vii
3.1.3 ASH-mediated 1-octanol responses are modulated by monoamines
and neuropeptides…... .......................................................................................................51
3.1.4 5-HT modulates responses to multiple ASH-sensed stimuli ............54
3.2 Discussion…… ................................................................................................56
3.2.1 ASHs are the primary sensors of saturating 1-octanol......................56
3.2.2 A prominent role for 5-HT in regulation of ASH stimuli .................58
3.2.3 Neuropeptides interact with monoamines to modulate ASH-mediated
aversive behaviors..............................................................................................................61
3.2.4 5-HT inhibits sensory-evoked Ca2+ from multiple chemorepellents 64
4
Serotonin differentially modulates Ca2+ transients and depolarization in a C.
elegans nociceptor….. .......................................................................................................67
4.1 Results…… ......................................................................................................67
4.1.1 Distinct Ca2+ pools mediate ASH aversive responses ......................67
4.1.2 Intracellular stores receptors shape ASH Ca2+ kinetics ....................70
4.1.3 RNAi-mediated knockdown of EGL-19 has no behavioral effect....72
4.1.4 5-HT likely inhibits ASH Ca2+ via SER-5 and EGL-30 ...................74
4.1.5 5-HT inversely modulates somal Ca2+ and depolarization ...............77
4.2 Discussion…… ................................................................................................79
4.2.1 The ASH soma & axon have distinct Ca2+ channel configurations ..79
4.2.2 Sensory-evoked ASH Ca2+ levels are not predictive of behavior .....81
4.2.3 5-HT decreases sensory-evoked Ca2+ influx, but decreases sensoryevoked depolarization via a Gαq signaling pathway in the soma ......................................82
viii
5
Serotonin negatively regulates C. elegans L-type VGCCs via a Ca2+-dependent
negative feedback loop ......................................................................................................87
5.1 Results…… ......................................................................................................87
5.1.1 5-HT modulates ASH Ca2+ downstream of sensory transduction ....87
5.1.2 5-HT inhibits ASH Ca2+ transients through SER-5 and EGL-30 .....89
5.1.3 CDI may act in the ASH circuit to modulate sensory responses ......91
5.1.4 The canonical CDI pathway is not present in ASH ..........................95
5.1.5 5-HT activates TAX-6 via SER-5 and EGL-30 to inhibit EGL-19 .97
5.2 Discussion…… ................................................................................................99
5.2.1 High concentrations of extracellular K+ can depolarize ASHs
bypassing sensory transduction..........................................................................................99
5.2.2 5-HT acts through Gαq directly to modulate Ca2+ signaling ...........100
5.2.3 The classical CDI pathway is not acting in ASHs, but each of the
proteins are still important for basal evoked Ca2+ signals and behavior..........................101
5.2.4 A model for serotonergic modulation of EGL-19 ..........................104
References ........................................................................................................................107
ix
List of Figures
1-1
The typical structure of a VGCC .............................................................................6
3-1
1-octanol responses of ASH/ADL/AWB sensory neurons ....................................47
3-2
Basic characterization of ASH 1-octanol response ................................................50
3-3
Monoamines and neuropeptides modulate 1-octanol responses ............................53
3-4
Serotonin modulates ASH activity in response to multiple odorants ....................55
4-1
VGCCs pass Ca2+ in ASH somas and axons .........................................................69
4-2
Internal Ca2+ stores amplify and shape ASH Ca2+ responses ................................71
4-3
Cell-specific RNAi knockdown of EGL-19 does not affect behavior ...................73
4-4
Serotonergic modulation of ASH 1-octanol-evoked Ca2+ transients .....................76
4-5
Serotonin potentiates 1-octanol-evoked ASH depolarization ................................78
5-1
Serotonin mediates ASH Ca2+ downstream of sensory transduction.....................87
5-2
Serotonin inhibits Ca2+ downstream of sensory transduction via Gαq...................90
5-3
CDI may act in the aversive circuit to modulate behavior.....................................93
5-4
A canonical CDI pathway is not present in ASHs .................................................96
5-5
TAX-6 acts downstream of 5-HT in ASHs............................................................98
5-6
Model of a 5-HT activated, Ca2+-driven negative feedback loop in ASHs .........106
x
List of Abbreviations
5-HT ...........................Serotonin
ACh ............................Acetylcholine
aka ..............................A Kinase Anchor Protein (C. elegans gene class)
AKAP .........................A Kinase Anchor Protein
Akt/PKB.....................Protein kinase B
anoh ............................Anoctamin (calcium-activated chloride channel) homolog
best .............................Bestrophin (chloride channel) homolog
BK ..............................Big K potassium channel
Ca2+ ............................Calcium ion
calf..............................Calcium channel localization factor
CaM............................Calmodulin
CaMKII ......................Ca2+/calmodulin-dependent protein kinase II
CaN ............................Calcineurin (PP2B), a Ca2+-dependent phosphatase
Cas9............................CRISPR-associated protein 9
CaV .............................Voltage-gated Ca2+ channel (number indicates family)
cca ..............................Calcium channel, alpha subunit
ccb ..............................Calcium channel, beta subunit
CDI .............................Ca2+-dependent inactivation
CDK1 .........................Cyclin-dependent kinase 1
CICR ..........................Ca2+-induced Ca2+ release
Cl- ...............................Chloride ion
CREB .........................Cyclic AMP response element binding protein
crh ..............................CREB homolog
CRISPR ......................Clustered regularly interspaced short palindromic repeats
crtc..............................CREB-regulated transcriptional coactivator
Cu2+ ............................Copper ion
D2 ...............................Dopamine receptor, type 2
DA ..............................Dopamine
DAG ...........................Diacylglycerol
DHCA ........................Dihydrocaffeic acid
eat ...............................Eating: abnormal pharyngeal pumping
xi
egl...............................Egg laying defective
ER ..............................Endoplasmic reticulum
ERK............................Extracellular signal-regulated kinase
FKBP..........................FK-506 binding protein
flp ...............................FMRFamide-like Peptide
Fyn .............................Proto-oncogene tyrosine-protein kinase Fyn
GABA ........................γ-aminobutyric acid
gpa ..............................G Protein, alpha subunit
gpc ..............................G Protein, gamma subunit
GPCR .........................G protein-coupled receptor
IAA ............................Isoamyl alcohol
ins ...............................Insulin related
IP3...............................Inositol 1, 4, 5-trisphosphate
IP3R ............................Inositol 1, 4, 5-trisphosphate receptor
itr ................................Inositol Trisphosphate Receptor
K+ ...............................Potassium ion
L4 ...............................Larval stage 4
LVA ...........................Low-voltage activated
M1...............................Muscarinic acetylcholine receptor, type 1
Na+ .............................Sodium ion
NemA .........................Nemadipine-A (L-type channel antagonist)
NGM ..........................Nematode growth medium
nlp ..............................Neuropeptide-like Protein
npr ..............................Neuropeptide Receptor
ntc...............................Nematocin (vasopressin-like peptide)
ntr ...............................Nematocin receptor
OA ..............................Octopamine
ocr ..............................OSM-9 and Capsaicin Receptor Related
octr .............................Octopamine receptor
odr ..............................Odorant response abnormal
osm .............................Osmotic avoidance abnormal
PCR ............................Polymerase chain reaction
PDZ ............................Postsynaptic density-95/Drosophila disc large tumor suppressor1/zonula occludens-1 domain
PIP2 ............................Phosphatidylinositol 4, 5-bisphosphate
PKA............................Protein kinase A
PKC ............................Protein kinase C
xii
PLC ............................Phospholipase C
PP ...............................Protein phosphatase
qui ..............................Quinine non-avoider
RACK ........................Receptor for Activated C Kinase
RNAi ..........................RNA interference
RyR ............................Ryanodine receptor
SDS ............................Sodium dodecyl sulfate
SEM ...........................Standard error of the mean
ser ...............................Serotonin/octopamine receptor family
SH3 ............................Src homology 3 domain
shn ..............................Shank (SH3/ankyrin domain scaffold protein) related
slo ...............................Slowpoke potassium channel family
SR101 .........................Sulforhodamine 101
sra ...............................Serpentine receptor, class A (alpha)
TA ..............................Tyramine
tag...............................Temporarily Assigned Gene Name
TALEN ......................Transcription activator-like effector nuclease
tax...............................Abnormal chemotaxis
TRPV .........................Transient receptor potential cation channel subfamily V
unc ..............................Uncoordinated
VDI ............................Voltage-dependent inactivation
VGCC ........................Voltage-gated calcium channel
xiii
List of Symbols
α .................................Alpha subunit
β .................................Beta subunit
γ ..................................Gamma subunit
Δ .................................Change in
δ..................................Delta subunit
µ .................................micro- (10-6)
Ω.................................ohms
ω .................................omega
A.................................Amperes
ºC................................degrees Celsius
F .................................Fluorescence
F0 ................................Baseline fluorescence
M ................................Molar (mol/L)
m ................................Meters
S .................................Siemens
V.................................Volts
xiv
Chapter 1
Introduction
Neuromodulation in sensory circuitry is critical for organisms integrating
environmental factors and nutritional state to exhibit appropriate behaviors. Smallmolecule neurotransmitters operate by directing information in disparate pathways
through the same physical hard-wired circuits, generating multiple behaviors dependent
on which pathways are activated. A common approach for dissecting behavioral
modulation is to compare information flow through physical circuits under different
circumstances, often with drug treatments or genetic mutations in order to probe neuronal
function (Chao, Komatsu, Fukuto, Dionne, & Hart, 2004; G. P. Harris et al., 2009). One
set of techniques that has emerged is brain activity mapping, where circuits are probed
using fluorescent imaging, with calcium (Ca2+) signals being used as a readout for
neuronal activity and depolarization (Grienberger & Konnerth, 2012; Mao, HamzeiSichani, Aronov, Froemke, & Yuste, 2001; Stosiek, Garaschuk, Holthoff, & Konnerth,
2003; Zimmer et al., 2009). While Ca2+ imaging has allowed neuroscientists to make
great strides in probing nervous system function, the functional significance of Ca2+
transients have not been rigorously examined. Therefore, exploring the interplay between
1
intracellular Ca2+ transients, neuronal excitability, and behavioral outputs is an important
priority.
However, performing precise optical imaging experiments in the mammalian
brain is incredibly difficult in vivo due to the sheer number of neurons and complexity of
the mammalian brain (Azevedo et al., 2009). To examine single-neuron Ca2+ dynamics
and behavioral circuits, we use the small nervous system of the model organism
Caenorhabditis elegans. Despite having a small nervous system consisting of only 302
neurons, C. elegans can sense multiple stimuli while modulating its behavioral outputs
based on nutritional status, environmental factors, and past experience. Additionally, C.
elegans has the first fully-mapped connectome and has conservation of many of the same
proteins and neurotransmitters with mammals (C I Bargmann & Kaplan, 1998; C. I.
Bargmann, 1998; White, Southgate, Thomson, & Brenner, 1986). Here, we provide a
novel pathway of how monoamine neurotransmitters, like serotonin (5-HT), can directly
modulate voltage-gated Ca2+ channels (VGCCs) through a Ca2+-dependent negative
feedback loop to influence Ca2+ transients and behavior in a polymodal sensory neuron.
In the following sections of this chapter, I will discuss common proteins responsible for
neuronal Ca2+ influx in mammals and C. elegans, focusing on structure, function, and
regulation.
1.1 Voltage-gated Ca2+ Channels
VGCCs are the primary method for Ca2+ entry into neurons, and drive neuronal
processes such as gene expression (through excitation-transcription coupling),
2
neurotransmitter release, and burst firing (Dai, Hall, & Hell, 2009; Welling, 2009).
Mammalian VGCCs typically contain up to five subunits: the α1-subunit, the poreforming subunit that confers most channel properties, and the β, α2, δ, and γ auxiliary
subunits that act as regulators of channel function (Dai et al., 2009; Dolphin, 2009; Laine,
Frokjaer-Jensen, Couchoux, & Jospin, 2011; Saheki & Bargmann, 2009; Welling, 2009).
α1-subunits contain four homologous domains, I – IV, each consisting of six
transmembrane segments with a voltage-sensing glutamate residue in the fourth segment
and a pore-forming P-loop segment in between the fifth and sixth transmembrane
segments of each domain (see Figure 1-1) (Dai et al., 2009). β-subunits aid in channel
expression, likely playing a role in trafficking channels to the plasma membrane, as well
as increasing peak currents, accelerating channel activation and inactivation, and shift
steady-state inactivation levels to hyperpolarizing membrane voltages when co-expressed
with their α1-subunits (Dolphin, 2009; L’ Lacinová & Hofmann, 2005; L Lacinová,
2005). Knockout of β-subunits has produced some unexpected results, as CaVβ3
knockouts demonstrate no obvious phenotype in mice despite its wide expression in the
brain, suggesting the possibility of some redundancy or developmental compensation
amongst subunits (Dolphin, 2009). The α2δ-subunits are also involved with steady-state
inactivation and voltage-dependent activation, but are not as well-studied compared to
their β counterparts. The α2 and δ-subunits are encoded by the same gene, but are posttranslationally modified to yield two distinct subunits linked by disulfide bonds (Dolphin,
2009; L Lacinová, 2005; Saheki & Bargmann, 2009; Striessnig, Pinggera, Kaur, Bock, &
Tuluc, 2014). The α2 protein is peculiar in that its localization is completely extracellular,
and is only bound to the rest of the channel (and membrane) via the δ subunit, which has
3
a single transmembrane domain (Dolphin, 2009; L Lacinová, 2005; Striessnig et al.,
2014). The α2-subunit seems to provide the structural elements required for channel
activation, where the δ-subunit modifies activation and inactivation kinetics (Dolphin,
2009; L Lacinová, 2005). Knockouts of α2δ subunits produce a variety of phenotypes
based on where they are expressed, with mild α2δ1 mutants demonstrating cardiac
phenotypes while stronger mutations are embryonic lethal (Dolphin, 2009). In contrast,
α2δ2 “Ducky” mutants exhibit decreased channel function due to decreased trafficking of
the channels to the membrane and cell activity in Purkinje cells, the cell type where they
are most often expressed (Dolphin, 2009; L Lacinová, 2005). Finally, the γ-subunits are
the least characterized of all VGCC subunits, and are not always present in channel
heteromers (Catterall, 2010, 2011; Laine et al., 2011; Saheki & Bargmann, 2009). The γsubunit is an integral membrane protein with four transmembrane domains, and is also
implicated in affecting the channel’s steady-state inactivation, but much of this subunit’s
function is likely involved in interactions with other membrane proteins via the protein’s
PDZ domain (Dai et al., 2009; L’ Lacinová & Hofmann, 2005; L Lacinová, 2005).
Interestingly, γ-subunits have only been co-immunoprecipitated with neuronal CaV2
channels, suggesting that these subunits are dispensable for function of CaV1 and CaV3
channels in the brain and likely play a special role in CaV2 channel signaling (Dai et al.,
2009). Therefore, by expressing channels with different configurations of subunits, Ca2+
entry can be regulated for a given cell type’s specific needs.
There are three major classes of VGCCs, classified based on the α1-subunit,
consisting of CaV1 (L-type), CaV2 (N/P/Q/R-type), and CaV3 (T-type) channels
4
(Catterall, 2010, 2011; Dolphin, 2009; Welling, 2009). Each of these channel subtypes
will be described in greater detail below.
5
Figure 1-1 The typical structure of a VGCC. The pore-forming α1-subunit (colored blue)
contains multiple repeating subunits and confers most channel properties. Channel
complexes also contain α2- (green), β- (red), and δ-subunits (green) which modulate
channel kinetics and activity. Some channel complexes also contain γ-subunits, which are
not pictured here. Figure adapted from Budde et al. 2002.
6
1.1.1 CaV1 (L-type) Channels
CaV1 channels were the first type of VGCCs to be discovered, and are
characterized by high voltage activation (around -30 mV), large single-channel
conductances, exhibit voltage-dependent inactivation (VDI), and antagonism by a class of
alkaloid molecules known as dihydropyridines (Catterall, 2011; Kwok et al., 2006, 2008;
Welling, 2009). The channels are composed of each of the VGCC subunits described
above, and are referred to as “L” type because they have a strong, long-lasting current
and were first discovered in skeletal muscle (Catterall, 2010, 2011; Dolphin, 2009; L
Lacinová, 2005). There are four known types of L-type channels: CaV1.1, which is
localized to skeletal muscle and is involved in excitation-contraction coupling as well as
transcriptional regulation; CaV1.2, which are localized to cardiac and smooth muscle for
excitation-contraction coupling, neuronal cell bodies and dendrites for activating Ca2+
transients, as well as regulation of enzymatic activity and transcription; CaV1.3, with
roles in cardiac pacemaking, Ca2+ transients in neurons, endocrine secretion, and auditory
transduction; and CaV1.4, a poorly-characterized channel involved in visual transduction
in the retina that completely lacks Ca2+ dependent inactivation (CDI) due to a novel CDIinhibiting domain (Catterall, 2011; Dai et al., 2009; Griessmeier et al., 2009; Striessnig et
al., 2014; Wahl-Schott et al., 2006; Welling, 2009). CaV1 channels are also capable of
forming large supramolecular signaling complexes, often acting in association with Gprotein coupled receptors, inositol trisphosphate receptors (IP3Rs), ryanodine receptors
(RyRs), and large-current Ca2+-dependent K+ channels (BK channels, “big K”) to tightly
control channel activity and regulate Ca2+ entry into the cell (Dai et al., 2009; Emrick,
7
Sadilek, Konoki, & Catterall, 2010; Hongkyun Kim et al., 2009; Parys & Smedt, 2012).
These signaling complexes perform a variety of functions, but often act upstream of
phospho-CREB –mediated transcriptional regulation, effectively allowing the channels to
participate in “excitation-transcription coupling” and regulation of gene expression
(Marshall et al., 2011; Simms, Souza, & Zamponi, 2013; Tuckwell, 2012; Wheeler,
Barrett, Groth, Safa, & Tsien, 2008). Interestingly, a conformational change of the CaV1
C-terminal tail may be necessary to invoke CREB-mediated signaling, with the channel’s
PDZ domain playing a prominent role (Morad & Soldatov, 2005; Weick, Groth, Isaksen,
& Mermelstein, 2003). CaV1-mediated Ca2+ release can act upstream of multiple
transcription factors, such as c-Jun and c-Fos, to contribute to gene expression in neurons,
skeletal muscle, and cardiac muscle through a mechanism requiring the activities of
serine-113-phosphorylated CREB, Ca2+/calmodulin-dependent protein kinase II
(CaMKII), and protein kinase A (PKA) (Brittain, Wang, Wilson, & Khanna, 2012;
Cruzalegui, Hardingham, & Bading, 1999; Morad & Soldatov, 2005; Thompson, Ginty,
Bonni, & Greenberg, 1995). However, despite extensive work on CaV1’s role in gene
expression, more studies will be required in the future to elucidate the complex interplay
between second messenger systems and gene expression in neurons.
Ca2+ signaling is a tightly-controlled process, as the ion acts as both a driver of
intracellular signaling as well as sometimes acting as a charge carrier for cellular
depolarization. VGCCs are only activated by transient increases in depolarization at the
plasma membrane, but have multiple mechanisms for negative feedback and gain control
of channel function. CaV1 channels have two primary methods of inactivation, VDI and
CDI, which are necessary to prevent overstimulation. VDI is a slow (over a time course
8
of seconds), two-part inactivation that consists of a conformational change in the channel
pore region, preventing the passage of Ca2+ while the channel would still be at an active
voltage (Kwok et al., 2008; L’ Lacinová & Hofmann, 2005; L Lacinová, 2005). VDI is
present in all CaV1 channels; however, the relatively slow kinetics and masking by other
inactivation mechanisms lead to difficulty in studying the VDI process. In contrast, CDI
is the faster of the two inactivation mechanisms, and is well-described, especially in the
context of CaV1 channels. CDI provides a mechanism for shutting a channel off that can
be modulated by intracellular signaling. The canonical CDI mechanism consists of six
core proteins: the CaV1 channel, the IP3R, the Ca2+-dependent serine/threonine
phosphatase calcineurin (CaN), a kinase (either CaMKII or PKA), and the scaffold
proteins Shank and A-kinase anchor protein (AKAP5) (Budde, Meuth, & Pape, 2002; Jee
et al., 2004; Morad & Soldatov, 2005; Oliveria, Dittmer, Youn, Dell’Acqua, & Sather,
2012; Olson et al., 2005). In this complex, the CaV1 channel is tethered to the IP3R via
the Shank scaffolding protein via its SH3 and PDZ domains, is also tethered to PKA and
CaN by AKAP5, and disruption of the complex impedes CDI activity (Marshall et al.,
2011; Oliveria et al., 2012; Olson et al., 2005; Rankovic et al., 2011; Zhang et al., 2005).
Upon L-type VGCC activation and Ca2+ influx, IP3Rs are activated, leading to Ca2+induced Ca2+ release (CICR) from the endoplasmic reticulum. Together, the Ca2+ influx
from both the CaV1 channel and IP3R is sufficient to activate CaN, which
dephosphorylates a conserved serine residue on the L-type channel’s C-terminus,
resulting in inactivation (Budde et al., 2002; Oliveria et al., 2012). CaN activity is reliant
on Ca2+/CaM binding the CaV1 channel’s C-terminal IQ domain for proper CDI
activation, and mutation of the conserved isoleucine to glutamate (IQ  EQ) is sufficient
9
to abolish regulation of channels by CDI (Blaich, Pahlavan, & Tian, 2012; Lian, Myatt,
& Kitmitto, 2007; Poomvanicha et al., 2011). Further supporting these data, the hearts of
EQ-mutated mice exhibited dilated cardiomyopathy and death indicative of aberrant
channel activity (Blaich et al., 2012; Poomvanicha et al., 2011). Similarly, CaM binding
has been implicated for two other important domains on the CaV1 channel as well: the Nterminal “NSCaTE” domain as well as a highly-conserved region directly upstream of the
IQ domain, termed the “pre-IQ domain” (Bers & Morotti, 2014; Christel & Lee, 2012;
Dolphin, 2009; Griessmeier et al., 2009; Morad & Soldatov, 2005; Simms et al., 2013;
Striessnig et al., 2014; Taiakina et al., 2013; Wahl-Schott et al., 2006). These domains
have differing functions than the IQ domain, but also likely play a role in Ca2+ regulation
(Lian et al., 2007; Simms et al., 2013; Taiakina et al., 2013). Taken together, these results
suggest CaM interactions with CaV1 channels are a multifaceted process that allows
multiple levels of tight regulation for Ca2+ entry in excitable cells.
1.1.1.1 Modulation of CaV1 Channels by Monoamine Neurotransmitters
The most well-studied model of CaV1 channels by monoamines is the βadrenergic modulation of CaV1.2 channels in cardiac muscle, but similar signaling
complexes have been described in the brain with regards to CaV1.2 and CaV1.3 channels,
also involving dopamine (DA), acetylcholine (ACh), and 5-HT (Day, Olson, Platzer,
Striessnig, & Surmeier, 2002; Olson et al., 2005; Rankovic et al., 2011; Zhang et al.,
2005). In the case of CaV1.2 channels, 5-HT2A and 5-HT2C receptors act through Gαq
signaling to inhibit channel function in prefrontal pyramidal neurons (Day et al., 2002).
10
Upon 5-HT binding the GPCR, Gαq is activated, stimulating the activation of
phospholipase Cβ (PLCβ), which cleaves phosphotidylinositol 4, 5-bisphosphate (PIP2)
into inositol 1, 4, 5-trisphosphate (IP3) and diacylglycerol (DAG). IP3 then activates its
IP3R, resulting in CICR from the endoplasmic reticulum and CaN activation as described
above. Furthermore, blockage of CaN by inhibitory peptides and FK-506 abolished Gαqmediated downregulation of CaV1.2 channels, suggesting that 5-HT was directly leading
to decreased channel function via CDI (Day et al., 2002). Similarly, CaV1.3 channels in
striatal medium spiny neurons can be bi-directionally modulated by dopaminergic D2
receptors and cholinergic M1 receptors (Olson et al., 2005). While the M1 receptors work
through a Gαq pathway as previously described, the D2 receptors act through Gαi, which
antagonizes the Gαq pathway. Interestingly, disrupting channel interactions with the
Shank scaffolding protein with an inhibitory PDZ-domain-targeting peptide disrupted DA
and ACh modulation, suggesting that Shank scaffolding is critical for proper regulation
of this signaling complex (Olson et al., 2005). Disruption of a secondary scaffolding
protein called Homer, which tethers Shank to the IP3R, also disrupted DA and ACh
modulation (Olson et al., 2005). Finally, in cultured rat hippocampal neurons, Shank
directly interacts with CaV1.3 channels via its SH3 and PDZ domains, with Homer via a
proline-rich region, and indirectly with IP3Rs and metabotropic glutamate receptors via
Homer (Zhang et al., 2005). This macromolecular signaling complex is thought to
properly localize the channel apparatus to postsynaptic locations, as well as acting a
modulator of pCREB signaling (Zhang et al., 2005). Together these results indicate that
CaV1 channels can be modulated by a variety of small molecule neurotransmitters
11
throughout the brain, with wide-reaching effects on gene expression, cell excitability, and
proper synaptic function.
1.1.2 CaV2 (N/P/Q/R-type) Channels
CaV2 channels are another family of VGCCs, consisting of α1, α2δ, and β, but not
γ-subunits. Like CaV1 channels, CaV2 channels are also activated at high voltages, have
large single-channel conductance, and most subtypes exhibit VDI and CDI. However,
unlike CaV1 channels, which are broadly antagonized by dihydropyridines and located in
many cell types, CaV2 channel classification is broken down further based on channel
subtype-specific peptide antagonists and are primarily expressed in neurons. Specifically,
CaV2.1 (P/Q-type) channels are antagonized by the web spider toxin ω-agatoxin IVA,
CaV2.2 channels (N-type) channels are antagonized by the cone snail toxin ω-conotoxin
GVIA, and CaV2.3 (R-type) channels are antagonized by SNX-482 (Catterall, 2011; L
Lacinová, 2005; Welling, 2009). CaV2.1 channels are referred to as “P-type” due to their
first characterization in Purkinje cells, but later other isoforms of the gene were found in
cerebellar granule cells and other regions of the brain, which are referred to as “Q-type”
(Welling, 2009). In contrast, CaV2.2 channels are “N-type” (“non-L”, “neuronal”), were
first characterized in the brain, and are highly concentrated at presynaptic terminals and
dendrites, where they play multiple roles in neuronal functions including neurotransmitter
release (L Lacinová, 2005; Saheki & Bargmann, 2009; Welling, 2009). Finally, CaV2.3
channels (R-type) are divergent from the other channel family members, being activated
at lower voltages than their CaV2.1 and CaV2.2 counterparts but still retaining high12
voltage-activated characteristics (Christel & Lee, 2012; Tuckwell, 2012; Welling, 2009).
Additionally, these R-type channels do not undergo CDI with the same mechanism of
other CaV1 and CaV2 channels, instead inactivating in a way that is independent of
divalent cations (L Lacinová, 2005).
As noted above, CaV2 channels are notable for their importance in driving fast
neurotransmission of small molecule neurotransmitters such as glutamate, GABA, 5-HT,
and DA, and as such require considerable regulation (Catterall, 2011). Like their CaV1
counterparts, CaV2 channels are extensively modulated by GPCRs, although these
channels are primarily regulated by the Gβγ subunits (Catterall, 2011; Dai et al., 2009;
Jansen, Weinkove, & Plasterk, 2002; Yamada, Hirotsu, Matsuki, Kunitomo, & Iino,
2009). Upon GPCR stimulation, Gβγ separates from Gα, and interacts with the first
intracellular loop of the channel to slow channel activation (Dai et al., 2009).
Interestingly, The Gβγ-CaV2 channel interaction is constrained to a short physical
distance, and is likely tonically active, with strong depolarization displacing the Gβγ
subunits from the loop (Dai et al., 2009). Similarly, activation of protein kinase C (PKC)
and CaMKII can phosphorylate key subunits in the first intracellular loop, blocking Gβγ
binding and facilitating channel activation (Dai et al., 2009; Mathews et al., 2003). In
summary, much like CaV1 subunits, CaV2 channels can be actively regulated by second
messenger systems, allowing for tight control of Ca2+ entry and downstream
neurotransmitter release.
1.1.3 CaV3 (T-type) Channels
13
The third major class of VGCCs is CaV3, the “T-type” channels. CaV3 channels
are distinct from their CaV1 and CaV2 counterparts because they activate at comparatively
lower (more hyperpolarized) voltages (around -60mV), and are referred to as low-voltage
activated (LVA) channels (Burgess, Crawford, Delisle, & Satin, 2002; L Lacinová, 2005;
Welling, 2009; Zamponi, Lewis, Todorovic, Arneric, & Snutch, 2009). T-type channels
also inactivate quickly after depolarization and demonstrate very low single channel
conductance between 5 – 9 pS (Catterall, 2011; L Lacinová, 2005). CaV3 channels have a
similar protein structure to their CaV1 and CaV2 counterparts, but only share about 25%
sequence identity in their α1-subunits, with no direct evidence that these channels interact
with other auxiliary subunits (Catterall, 2011; L Lacinová, 2005). The replacement of key
glutamate residues in the ion selectivity filter with aspartate (EEEE in CaV1 and CaV2;
EEDD in CaV3) is the likely reason these channels have low conductance, and may allow
the channels to pass Na+ in species lacking traditional voltage-gated Na+ channels, such
as Caenorhabditis elegans (Cribbs et al., 1998; L Lacinová, 2005; Senatore, Guan, &
Spafford, 2014). There are three types of CaV3 channels: CaV3.1, CaV3.2, and CaV3.3,
which are blocked by mibefradil, the scorpion toxin α-kurtoxin, and Ni2+ ions, although
none of these are specific channel antagonists (Cribbs et al., 1998; Dolphin, 2009;
Welling, 2009). Like CaV1, CaV3 channels are present in multiple cell types including the
heart, brain, and adrenal cortex, where they provide multiple functions (Burgess et al.,
2002; Catterall, 2011; Cribbs et al., 1998; L Lacinová, 2005). Notably, each CaV3
channel subtype localizes to the soma and dendrites of neurons, not being present in the
axon (Cheong & Shin, 2013). The most prominent role for CaV3 channels is their role in
burst firing, where activation of K+ channels or transient inhibitory postsynaptic
14
potentials deinactivate T-type channels, allowing the subsequent rebound in membrane
potential to activate them (Cheong & Shin, 2013). Channel activation leads to a positive
feedback that activates CaV1 and CaV2 channels, resulting in greater depolarization and a
train of action potentials. Another important role for CaV3 channels are in the heart’s
sino-atrial node, where the channel helps set the pacemaker current necessary for proper
heartbeat generation (Catterall, 2011; Mangoni et al., 2006). Interestingly, despite their
non-axonal localization, CaV3 channels may facilitate low-voltage neurotransmitter
release through the formation of nanodomains consisting of CaV3.1 and CaV3.2 channels
in association with syntaxin and SNAP-25, likely establishing basal release at resting
conditions or mild activation (Carbone, Calorio, & Vandael, 2014).
In spite of their marked differences with their CaV1 and CaV2 counterparts, CaV3
channels share many similar forms of regulation. CaV3 channels undergo inactivation like
CaV1 and CaV2 channels, but their threshold for inactivation is much more rapid than the
high-voltage activated channels and is not particularly voltage-dependent (Catterall,
2011; Cribbs et al., 1998; L Lacinová, 2005). In a similar fashion, classical CDI has never
been described in T-type channels. However, CaV3.2 channels do exhibit Ca2+-dependent
facilitation via CaMKII (Christel & Lee, 2012). In this pathway, CICR activates CaMKII,
resulting in phosphorylation of conserved serine residues on the CaV3.2 α1-subunit and
shifting the voltage required for activation in a more negative direction (Christel & Lee,
2012). Blocking CaMKII through the use of inhibitors, or expressing a constitutively
active form of the kinase also blocks this effect (Christel & Lee, 2012; L Lacinová,
2005). Also notable is the observation that CaV3.2 channels can be inhibited by a
particular Gβγ complex through direct interaction with the channel α1-subunit, much like
15
the form of negative regulation present in CaV2 channels (DePuy et al., 2006). While
these initial observations demonstrate inactivation and regulation in these channel
subtypes, the masking of CaV3 currents by high-voltage activated counterparts and lack
of selective blockers will continue to make these enigmatic channels promising subjects
for future study for years to come.
1.2 Intracellular Ca2+ Receptors: IP3Rs and RyRs
While VGCCs on the plasma membrane are critical for cellular Ca2+ signaling,
intracellular Ca2+ receptors, represented by the inositol 1, 4, 5-trisphosphate receptor
(IP3R) and ryanodine receptor (RyR) are equally necessary for signal amplification and
second messenger systems. Like VGCCs, IP3Rs and RyRs form complexes with other
proteins to modulate intracellular Ca2+ levels and signaling, but differ in terms of
activation, inactivation, and regulation. A brief summary of the structure, function, and
modulation of each of these channel classes will be described in the coming section.
1.2.1 IP3 Receptors
Inositol 1, 4, 5-trisphosphate (IP3) is a ubiquitous signaling molecule in cells,
having critical roles in the modulation of development, synaptic plasticity, gene
expression, peptide release, and lipid metabolism (Goto & Mikoshiba, 2011; Hagenacker,
Ledwig, & Büsselberg, 2008; Subramanian, Jayakumar, Richhariya, & Hasan, 2013). As
mentioned previously, IP3 is produced by the cleavage of PIP2 by PLC, often downstream
16
of Gαq activation. Unlike Ca2+ which has a limited range of action, IP3 diffuses far from
its place of synthesis (a maximum distance of about 24 µm), allowing the molecule to act
as a global intracellular signaling molecule (Allbritton, Meyer, & Stryer, 1992; Goto &
Mikoshiba, 2011; Parys & Smedt, 2012). The only known signaling role for IP3 is to
activate the IP3R, binding to the receptor and leading to intracellular Ca2+ release from
intracellular compartments like the endoplasmic reticulum via CICR (Baylis, Furuichi, &
Yoshikawa, 1999; Hagenacker et al., 2008; Parys & Smedt, 2012; D. Walker, Gower, &
Ly, 2002). In mammals, there are three IP3R genes: IP3R1, IP3R2, and IP3R3, each with
differing affinities to IP3 and unique signaling roles (Parys & Smedt, 2012). IP3Rs have
multiple splice isoforms, and different subunits can homo- and heterotetramerize to form
functional receptors, but the configurations of which subunits can and cannot associate
are not well understood (Joseph, Bokkala, Boehning, & Zeigler, 2000). Each IP3R
subunit contains six C-terminal transmembrane regions, two ligand-binding domains, and
an N-terminal suppressor domain, with the channel’s pore being located between
transmembrane domains 5 and 6 (Goto & Mikoshiba, 2011; Parys & Smedt, 2012;
Wojcikiewicz, Pearce, Sliter, & Wang, 2009). Unlike the VGCCs, the crystal structure of
IP3R1 has been partially solved, demonstrating multiple binding regions for Ca2+, IP3,
and other interacting proteins (Goto & Mikoshiba, 2011; Joseph et al., 2000; Parys &
Smedt, 2012). Interestingly, IP3Rs undergo Ca2+ activation and inactivation, as Ca2+ ions
bind the receptor as a co-agonist with IP3 with a bell-shaped affinity curve based on the
amount of Ca2+ present in both the cytoplasm and ER lumen (Allbritton et al., 1992; Goto
& Mikoshiba, 2011; Parys & Smedt, 2012). The IP3Rs can also be inhibited by various
drugs including heparin, 2-APB, and xestospongins, although IP3R-targeting drugs are
17
wide-spectrum and do not act as specific receptor subtype antagonists (Saleem, Tovey,
Molinski, & Taylor, 2014). Like the VGCCs mentioned above, IP3Rs are extensively
modulated by interacting proteins. Over 50 proteins have been implicated in having
positive and negative interactions with the mammalian IP3R1 and its immediate partners,
including kinases such as PKA, Akt/PKB, PKC, CDK1, ERK, and Fyn, phosphatases
such as PP1, PP2A, and calcineurin (CaN), as well as scaffolding proteins such as
AKAP9 and RACK1 (Parys & Smedt, 2012). Taken together, these data suggest that
IP3Rs are critical regulators of intracellular Ca2+ signaling, and that tight spatiotemporal
regulation of Ca2+ entry into the cytoplasm is required to prevent aberrant pathway
activation.
1.2.2 Ryanodine Receptors (RyRs)
In contrast to IP3Rs, RyRs are another class of intracellular Ca2+ receptors found
in excitable cells. RyRs are massive homotetrameric channels that are the largest ion
channels known to date, with an average molecular weight of over 2.2 megadaltons (Van
Petegem, 2012). There have been three distinct RyR isoforms discovered to date: RyR1,
the channel often found in skeletal muscle and the first channel of this subtype cloned;
RyR2, commonly expressed in the heart; and RyR3, which are most commonly expressed
in the brain (Lanner, Georgiou, Joshi, & Hamilton, 2010; Van Petegem, 2012). The
primary method for RyR activation is the presence of extracellular Ca2+ leading to
conformational changes in the channel pore resulting in channel opening. Interestingly,
non-Ca2+-dependent mechanisms of activation have also been described, including a
18
physical interaction with the skeletal CaV1.1 channel, as well as store overload-induced
Ca2+ release, which can open the channel via changes in membrane potential or when
internal stores reach a certain Ca2+ concentration, respectively (Fill et al., 1989; Van
Petegem, 2012). Notably, while RyRs often associate with other channels in vivo,
crystallization studies have revealed that RyR1 channels will align themselves in a
checkerboard pattern in the absence of other proteins, and can link up with each other for
simultaneous activation (Van Petegem, 2012; Yin, D’Cruz, & Lai, 2008; Yin, Han, Wei,
& Lai, 2005).
Like the other Ca2+ channels discussed previously, RyRs are targeted by multiple
signaling molecules and pathways. RyRs are strongly inhibited by ryanodine, an alkaloid
antagonist produced by Ryania speciosa and the drug for which these receptors are
named (Fill et al., 1989; Maryon, Coronado, & Anderson, 1996; Van Petegem, 2012).
The antagonist has two different effects depending on threshold: at nanomolar
concentrations, ryanodine locks the RyR into a half-open state, still allowing the channel
to pass some Ca2+; however, when the concentration reaches the millimolar range, the
channel is blocked completely (Fill et al., 1989; Gomez & Yamaguchi, 2014; Maryon et
al., 1996; Van Petegem, 2012). RyRs are also heavily modulated by interacting proteins
as well, with CaM (both apo- and Ca2+/CaM), PKA, CaMKII, PP1, PP2A, calsequestrin,
and Homer1c all binding the massive RyR tetramer to modulate channel function
positively or negatively depending on context, isoform, or location (Lanner et al., 2010;
Rodney, Williams, Strasburg, Beckingham, & Hamilton, 2000; Van Petegem, 2012; J.
Wang & Best, 1992). In closing, RyRs, along with their VGCC and IP3R counterparts, all
19
play critical roles of amplifying Ca2+ signals as well as the regulation of complex Ca2+
signaling cascades.
1.3 Caenorhabditis elegans: A model organism for studying sensory neuron
modulation
While the physiological properties of VGCCs have been extensively studied,
much less is known about the role of channel modulation in generating animal behavior.
Additionally, much of the work studying VGCC modulation has been in the context of
the cardiovascular system, and not much is known about the effects of channel
modulation in neurons. One difficulty in higher systems is specificity: as VGCCs are
omnipresent in excitable cells, ascribing channel function to behavior in the context of
one cell or a group of cells has proven challenging. Additionally, mammalian brains
contain an estimated 86 billion neurons and hundreds of trillions of synapses, making
studies at a single-neuron resolution effectively impossible given current technologies
(Azevedo et al., 2009). Therefore, to study the effects of sensory inputs on VGCCs in the
context of animal behavior, we have chosen the model organism Caenorhabditis elegans
for our experimentation. C. elegans is a small free-living, soil-dwelling nematode
(roundworm) that grows up to 1 mm long as an adult. Worms come in two sexes, males
and hermaphrodites, which allow for easy genetic crosses and propagation of genetic
clones (Brenner, 1974). Notably, C. elegans has a short generation time, developing
from the embryo to adult in about three and a half days under standard laboratory
conditions (grown on OP50 E. coli at 20 ºC) , and hermaphrodites can self-fertilize for
20
easy passage of genetically identical animals (Brenner, 1974; Corsi, 2015). Adult C.
elegans hermaphrodites have 959 somatic cells that arise in a developmentally invariant
pattern, allowing for easy identification of cell types in individual animals (Altun & Hall,
2015; Kaletta & Hengartner, 2006; Sulston, Schierenberg, White, & Thomson, 1983;
White et al., 1986). Worms are also amenable to genetic analysis, with a fully-mapped
genome and widely-established genome-editing methods including RNA interference,
TALENs, and CRISPR/Cas9-mediated homologous recombination (Esposito, Di Schiavi,
Bergamasco, & Bazzicalupo, 2007; Fire, Xu, Montgomery, & Kostas, 1998; FrøkjærJensen, 2013; Lo et al., 2013; Waaijers & Boxem, 2014). Finally, there is appreciable
conservation in proteins from C. elegans and vertebrates: about 38% of C. elegans genes
have identified orthologs in humans; conversely, a predicted 60-80% of human genes
also have functional orthologs in C. elegans (Corsi, 2015; Kaletta & Hengartner, 2006;
Shaye & Greenwald, 2011). Here, we utilize C. elegans as a genetic model system for the
dissection of sensory neuron signaling, and to evaluate the significance of neuronal Ca2+
signals in the context of nociceptive sensory circuit function.
1.3.1 The C. elegans Nervous System: Small but Capable of Complex Behavior
Of the 959 somatic cells in the C. elegans hermaphrodite, 302 of these cells are
neurons (White et al., 1986). Despite the relatively small size, the worm’s nervous system
is incredibly complex, containing more than 7000 chemical and electrical synapses and a
fully-mapped connectome revealing extensive feedback between neurons (Corsi, 2015;
White et al., 1986). Notably, C. elegans can perform multiple complex sensory-evoked
21
behaviors, including positive and negative chemotaxis, aerotaxis, thermotaxis, responses
to mechanical stimuli, and osmotic stress (Bretscher et al., 2011; Ezcurra, Tanizawa,
Swoboda, & Schafer, 2011; Glauser et al., 2011; G. Harris et al., 2010; Hilliard,
Bargmann, & Bazzicalupo, 2002; Kodama-Namba et al., 2013; Oda, Tomioka, & Iino,
2011). Remarkably, these behaviors can be integrated, resulting in complicated
behavioral decisions based on nutritional state, past experience, and other environmental
factors (Kodama-Namba et al., 2013; Sengupta, 2013). The C. elegans nervous system is
also extensively modulated by small molecule neurotransmitters such as 5-HT, γaminobutyric acid (GABA), glutamate, ACh, DA, octopamine (OA), tyramine (TA), and
neuropeptides signaling through GPCRs and ligand-gated ion channels to activate second
messenger pathways (C I Bargmann & Kaplan, 1998; C. I. Bargmann, 1998; Chalasani et
al., 2010; Troemel, Chou, Dwyer, Colbert, & Bargmann, 1995). The interplay between
neurotransmitters and external stimuli is critical for the animal’s behavior, with
neurotransmitter signaling combining with multiple sensory inputs to give rise to
complex sensory-driven behaviors to allow the worm to properly navigate its
environment.
1.3.2 The polymodal ASH nociceptors as a model for sensory modulation
C. elegans senses its environment primarily through 14 pairs of sensory neurons,
concentrated in the head and tail. The sensory neurons in the head are directly exposed to
the external environment at the animal’s nose, in a specialized sensory organ known as
the amphid (Hilliard et al., 2002; D. S. Walker et al., 2009). Of particular interest are the
22
ASH sensory neurons, a pair of polymodal, nociceptive sensory neurons that are part of
the amphid sensory apparatus. The ASHs sense noxious stimuli including harsh nose
touch, high osmolarity, and noxious chemicals (Aoki, Yagami, & Sasakura, 2011;
Colbert, Smith, & Bargmann, 1997; Ezcurra et al., 2011; G. P. Harris et al., 2009; G.
Harris et al., 2010; Hilliard et al., 2005, 2002). Upon sensation of a noxious olfactory
stimulus, the TRPV1-like OSM-9/OCR-2 channel is activated downstream of the
olfactory G-protein ODR-3, depolarizing the neurons (Roayaie, Crump, Sagasti, &
Bargmann, 1998; Tobin et al., 2002). Neuronal depolarization results in the VGCC
activation and glutamate release onto downstream AVA command interneurons, which
then in turn activate motorneurons to initiate a rapid escape response known as a reversal
(Lindsay, Thiele, & Lockery, 2011; Thiele, Faumont, & Lockery, 2009). The modulation
of ASH-mediated circuits, including reversals, has been a powerful tool for unlocking
regulation of sensory-evoked behaviors in the worm.
The ASH nociceptive neurons are commonly studied in the context of their
responses to high osmolarity, light nose touch, and noxious chemicals. Our group focuses
specifically on ASH responses to 1-octanol, a volatile fatty alcohol that is thought to be
present in the animal’s natural environment (Chao et al., 2004; Troemel et al., 1995)(see
Section 1.3.5). As the ASHs are sensory neurons, they signal downstream onto
interneurons and motor neurons, eventually resulting in the escape response. ASHs signal
primarily through two parallel pathways: a stimulatory circuit, where ASHs synapse
directly onto the AVA interneurons to stimulate reversals, and a disinhibitory circuit,
where ASHs are thought to stimulate the AIB interneurons, resulting in a disinhibition of
the RIM interneurons, and ultimately promoting reversals (Piggott, Liu, Feng, Wescott, &
23
Xu, 2011). ASHs use glutamate as a neurotransmitter, as evidenced by the presence of
the vesicular glutamate transporter EAT-4 and the presence of glutamate-gated channels
on downstream interneurons (Hills, Brockie, & Maricq, 2004; R. Y. Lee, Sawin, Chalfie,
Horvitz, & Avery, 1999; Mellem, Brockie, Zheng, Madsen, & Maricq, 2002; Rose, Kaun,
Chen, & Rankin, 2003; Zheng, Brockie, Mellem, Madsen, & Maricq, 1999). ASHs also
express at least four known neuropeptide-encoding genes: NLP-3, NLP-15, FLP-21, and
INS-1, whose peptide products are also thought to act on downstream interneurons to
trigger modulation of the aversive response (G. Harris et al., 2010; Holden-Dye &
Walker, 2013; Mills et al., 2011; Rogers et al., 2003). However, only FLP-21 has a
known receptor (NPR-1), making the deorphanization of these receptors promising
targets for studying future sensory circuit modulation (Holden-Dye & Walker, 2013;
Rogers et al., 2003).
1.3.3 In vivo recordings from C. elegans neurons: Insights from a transparent worm
While behavior provides us with critical information about how an animal
navigates its environment, sensory responses are still a sum total of the activities of
multiple neurons, resulting in complex and emergent circuit states. To isolate the
activities of a single neuron in the context of sensory signaling, many researchers have
turned to optical methods such as Ca2+ imaging and optogenetics to probe cellular
function. An underappreciated benefit of using C. elegans as a model organism for
sensory signaling is that the animals are transparent, providing an effective and noninvasive way to study neuronal function. Indeed, a common way to probe neural circuits
24
in C. elegans is through the use of genetically-encoded Ca2+ indicators such as cameleons
or GCaMP (Choi et al., 2015; Ezcurra et al., 2011; Hilliard et al., 2005; Kerr et al., 2000;
Nakai, Ohkura, & Imoto, 2001; Tian et al., 2009). Ca2+ imaging studies are often
performed by expressing a genetically-encoded Ca2+ indicator cell-specifically and
studying Ca2+ responses in combination with drug treatments, genetic mutants, or
behavioral assays (Ezcurra et al., 2011; Hilliard et al., 2005; Kodama-Namba et al., 2013;
Mills et al., 2011; Oda et al., 2011; Piggott et al., 2011; Zahratka, Williams, Summers,
Komuniecki, & Bamber, 2015; Zimmer et al., 2009). In concert with behavioral studies,
optical recordings of ASH nociceptors provide some cell-based data to contrast with
more indirect behavioral studies. ASHs exhibit robust Ca2+ transients in response to nose
touch and osmotic stresses evoked by 1 M glycerol, as well as noxious chemical stimuli
including Cu2+, denatonium, quinine, SDS, and 1-octanol (Ezcurra et al., 2011; Hilliard et
al., 2005; Krzyzanowski et al., 2013; Mills et al., 2011; Zahratka et al., 2015). In parallel
to behavior, ASH Ca2+ transients are mediated by monoamines, with Cu2+ and glycerol
Ca2+ responses increasing in the presence of DA, and nose touch responses increasing in
the presence of 5-HT (Ezcurra et al., 2011; Hilliard et al., 2005). Similarly, other sensory
neurons are the focus of modulation as well, with other groups studying behavioral
changes in response to oxygen concentration, attractive chemicals, and salt chemotaxis
after conditioning (Oda et al., 2011; Thiele et al., 2009; Troemel, Kimmel, & Bargmann,
1997; Zimmer et al., 2009). Finally, recent advances in Ca2+ imaging technology have
allowed for widespread imaging of the entire C. elegans nerve ring, permitting the
measurement of neuronal Ca2+ signals in the context of entire neural circuits or even
nervous systems (Prevedel et al., 2014; Schrodel, Prevedel, Aumayr, Zimmer, & Vaziri,
25
2013). Together, these advances allow correlation of behavioral activity with measurable
physiological changes in neurons, permitting the study of behavioral circuits at a singleneuron resolution.
Although Ca2+ imaging is a popular technique for neuronal study due to its
relative ease and non-invasive preparation, patch-clamp electrophysiology is still
necessary to understand neuronal excitability. C. elegans presents great challenges for
electrical recordings in neurons, due to their small soma size (3 µm on average) and
pressurized cuticle, making dissection of cells difficult (Goodman, Hall, Avery, &
Lockery, 1998). However, in spite of these difficulties, some work has been done
elucidating basic electrical properties of C. elegans neurons. Unlike mammalian neurons,
C. elegans nerve cells do not fire traditional action potentials, and indeed lack voltagegated Na+ channels (C. I. Bargmann, 1998; Goodman et al., 1998). As a result, the major
conductances in C. elegans neurons are Ca2+, Na+ (often passed through non-specific
cation channels like TRP channels), K+, and Cl-, with multiple voltage- and ligand-gated
channels for each of these ions allowing for graded transmission of neuronal signals (C. I.
Bargmann, 1998; Colbert et al., 1997; Dent, Smith, Vassilatis, & Avery, 2000; Maëlle
Jospin, Jacquemond, Mariol, Ségalat, & Allard, 2002; Montell, 2003; Putrenko,
Zakikhani, & Dent, 2005; Sokolchik, Tanabe, Baldi, & Sze, 2005; Tobin et al., 2002).
Interestingly, C. elegans neurons are thought to be isopotential due to high resistances
and low voltage attenuation (Goodman et al., 1998). A majority of worm
electrophysiology has been performed in the context of muscle physiology and touch
receptor neurons, but little is known about the electrical signaling in other sensory
neurons or interneurons (Arnadóttir, O’Hagan, Chen, Goodman, & Chalfie, 2011; M.
26
Jospin, Mariol, Segalat, & Allard, 2002; Maëlle Jospin et al., 2002; Laine et al., 2011).
Basic characterization of neurotransmitter function, especially glutamate, has been
partially described for the ASH nociceptive circuit, but electrophysiological dissection of
sensory regulation of aversive responses remains elusive (Mellem et al., 2002; Piggott et
al., 2011; Summers et al., 2015; Zheng et al., 1999). Here, we perform in vivo patchclamp recordings to directly measure ASH responses to 1-octanol and their modulation
by 5-HT.
1.3.4 Calcium Channels in C. elegans
While mammals and other higher organisms exhibit multiple Ca2+ channel family
members and isoforms, the C. elegans genome has more simplified contents. Unlike
mammals, worms only encode for one of each major type of VGCC, and one of each
intracellular Ca2+ receptor. C. elegans has one CaV1 homolog, EGL-19, one CaV2
channel, UNC-2, one CaV3 subunit, CCA-1, and one each of the IP3R and RyR, encoded
by ITR-1 and UNC-68, respectively (Baylis et al., 1999; Caylor, Jin, & Ackley, 2013;
Maëlle Jospin et al., 2002; Laine et al., 2011; Maryon et al., 1996; Mathews et al., 2003;
Saheki & Bargmann, 2009; Shtonda & Avery, 2005; Steger, Shtonda, Thacker, Snutch, &
Avery, 2005; D. S. Walker et al., 2009). Additionally, while mammalian channels have
multiple α2δ- and β-subunits for channel modulation, the C. elegans genome only has two
α2δ-subunits, UNC-36 and TAG-180, and two β-subunits, CCB-1 and CCB-2 (Caylor et
al., 2013; Laine et al., 2011; Saheki & Bargmann, 2009). Relatively little is known about
the C. elegans α2δ- and β-subunits, but they are required for proper channel modulation,
27
kinetics, and trafficking similar to their mammalian counterparts (Caylor et al., 2013;
Laine et al., 2011; Saheki & Bargmann, 2009). Each of the α1-subunits and intracellular
Ca2+ receptors will be discussed in greater detail below.
1.3.4.1 EGL-19
As previously stated, the egl-19 locus encodes the single L-type α1 VGCC subunit
in C. elegans. EGL-19 is primarily expressed in the body wall and pharyngeal muscle, as
well as in a subset of head and tail neurons (Maëlle Jospin et al., 2002; Williams &
Waterston, 1994). The egl-19 gene has sequence similarity to both CaV1.2 and CaV1.3
channels, including an N-terminal NSCaTE domain, four ion transport domains, a pre-IQ
domain, an IQ domain, and a C-terminal PDZ-binding domain (Jee et al., 2004; Kwok et
al., 2006, 2008). EGL-19 has three isoforms, with isoforms A and B encoding the entire
channel coding region, while isoform C only encodes the C-terminal regulatory region
without the NSCaTE and ion transport domains (WormBase, 2015b). Notably, EGL-19
channels are inhibited by the dihydropyridine antagonist nemadipine-A (NemA), which
binds EGL-19 channels with great specificity but is not effective on mammalian
orthologs (Kwok et al., 2006, 2008). Conversely, the conventional mammalian
dihydropyridines, nimodipine and nifedipine, are also less efficacious in C. elegans
(Kwok et al., 2006, 2008). Interestingly, egl-19 null animals are embryonic lethal,
exhibiting paralysis at the two-fold developmental stage, implying the channel is
necessary for proper muscle contraction required for development (Williams &
Waterston, 1994). Reduced function and gain-of-function alleles have also been
28
characterized: hypomorphs exhibit flaccid paralysis, feeble pharyngeal pumping, and egg
retention, while hypermorphs are myotonic, nemadipine-A resistant, and lay eggs
constitutively (Kwok et al., 2008; R. Y. N. Lee, Lobel, Hengartner, Horvitz, & Avery,
1997). EGL-19 is often found coupled with the UNC-36 α2δ-subunit and CCB-1 βsubunit in muscle, whereas TAG-180 and CCB-2 do not have any effect on EGL-19
muscle currents (Laine et al., 2011). In neurons, the auxiliary subunit makeup has yet to
be described, potentially allowing the discovery of another promising target of neuronal
modulation.
1.3.4.2 UNC-2
Mammalian CaV2 channels are a diverse group, encoding the N-, P-, Q-, and Rtype channel families (CaV2.1-2.4) often involved in neurotransmitter release. The C.
elegans unc-2 gene is the worm ortholog of a CaV2 channel, having sequence similarity
closest to the mammalian CaV2.1 P/Q-type channel (Estevez, Estevez, Cowie, & Gardner,
2004; Schafer & Kenyon, 1995). The unc-2 gene encodes five isoforms, including four
full-length channels and one truncated product containing only the channel C-terminus
(WormBase, 2015c). UNC-2 is primarily expressed in motor neurons, but also is
expressed in the egg-laying circuit as well as a subset of sensory neurons where the
channel acts as a major facilitator of synaptic neurotransmitter release (Caylor et al.,
2013; Saheki & Bargmann, 2009; Schafer & Kenyon, 1995; WormBase, 2015c; Zahratka
et al., 2015). Interestingly, proper UNC-2 localization requires the novel protein CALF-1,
a neuronal transmembrane trafficking protein that is present throughout the nervous
29
system, suggesting UNC-2 expression may extend beyond sensory and motor neurons
(Saheki & Bargmann, 2009). As their name implies, unc-2 reduced function mutants
demonstrate uncoordinated movement, with severe alleles not moving at all likely due to
hampered synaptic neurotransmitter release (Estevez et al., 2004; Mathews et al., 2003;
Schafer & Kenyon, 1995). Like EGL-19, UNC-2 often forms heteromers with UNC-36
and CCB-1 in muscle, but the neuronal subunit composition has yet to be described
(Caylor et al., 2013; Saheki & Bargmann, 2009). There is no published evidence that the
spider peptide toxin ω-agatoxin IVA inhibits UNC-2 channels, but it is possible that these
studies, in addition to others involving synaptic transmitter release, could identify a role
for UNC-2 in circuit modulation.
1.3.4.3 CCA-1
Like other C. elegans VGCC α1-subunits, cca-1 encodes the singular CaV3
channel in the worm. Unlike the other channels which have been extensively studied,
relatively little is known about CCA-1, other than it is a low-voltage activated channel
that opens at membrane potentials around -40 mV and inactivates rapidly similar to other
T-type channel subunits (Shtonda & Avery, 2005; Steger et al., 2005). CCA-1 has close
sequence homology to the mammalian CaV3.3 channel (42% homology), but is also
similar to the mammalian CaV3.1 (39% homology) and CaV3.2 (37% homology) (Steger
et al., 2005). Like mammalian CaV3 channels, CCA-1 also lacks a β-subunit interaction
domain at its N-terminus and calmodulin-interacting domains at its C-terminus, further
supporting the channel as the C. elegans ortholog of the T-type channel (Steger et al.,
30
2005). CCA-1 has a diverse expression pattern including pharyngeal muscle, head and
tail neurons, ventral nerve cord, and anal ganglia (Steger et al., 2005). CCA-1 has five
isoforms, but unlike EGL-19 and UNC-2, which have isoforms lacking the ion transport
domain, each isoform likely encodes the entire CCA-1 subunit and therefore a functional
channel (Steger et al., 2005; WormBase, 2015a). Ni2+ ions are effective inhibitors of
CCA-1, again much like their mammalian counterparts (Shtonda & Avery, 2005).
Notably, cca-1 mutants exhibit no easily observable phenotypes, but closer studies reveal
roles in shaping the initial depolarization of the pharyngeal action potential, as well as
triggering release at the neuromuscular junction in response to signaling by motor
neurons (Caylor et al., 2013; Shtonda & Avery, 2005; Steger et al., 2005). Recently,
CCA-1 has been implicated in having a role of the serotonergic modulation of egg-laying,
but the exact location of channel activity has yet to be determined (Kara Zang, personal
communication). More work will be required to dissect this enigmatic channel subunit
and its role throughout the C. elegans nervous system.
1.3.4.4 ITR-1
IP3 signaling is conserved in vertebrates and invertebrates, and the IP3R is a
notable modulator in C. elegans. Like in mammals, IP3 is produced downstream of the
Gαq subunit EGL-30 and the phospholipase Cβ ortholog EGL-8 (G. Harris et al., 2010;
Rex et al., 2005; D. S. Walker et al., 2009). Worms have a single IP3R encoded by the itr1 gene, which has six isoforms and homology similar to the mammalian channels (Baylis
et al., 1999). Like its mammalian counterparts, itr-1 has an N-terminal suppressor
31
domain, an IP3-binding domain, and C-terminal ankyrin repeats (Baylis et al., 1999).
Interestingly, itr-1 has conserved binding sites for Homer, chromogranin A, and
FKBP12, although no known Homer or chromogranin A orthologs are known to exist in
the worm (Baylis et al., 1999). ITR-1 channels are expressed throughout the worm, in the
adult intestine, pharynx, excretory cells, in the germ line, but only demonstrate limited
expression in neurons (Baylis et al., 1999; D. S. Walker et al., 2009). Interestingly,
neuronal ITR-1 expression is thought to have a modulatory role in addition to its primary
function of Ca2+ amplification, as disruption of itr-1 in the ASH sensory neurons changes
Ca2+ kinetics in response to nose touch and 1-octanol (D. S. Walker et al., 2009; Zahratka
et al., 2015)(see Chapter 4). While mammalian IP3Rs are often blocked through the use
of drugs like xestospongin C or Ca2+ stores are emptied by exposure to thapsigargin, C.
elegans ITR-1 channel function is often performed by overexpression of the channel’s
own suppressor domain, known as the “IP3 sponge” to inhibit channel function (D.
Walker et al., 2002; D. S. Walker et al., 2009). ITR-1 also interacts directly with the C.
elegans ortholog of Shank, SHN-1, which affects defecation rhythm and mating behavior
through the interaction of the two proteins via ankyrin repeats (Jee et al., 2004; Oh, Song,
Cho, & Park, 2011). ITR-1’s limited neuronal expression and involvement in multiple
signaling pathways make the receptor a promising target for future studies of modulation.
1.3.4.5 UNC-68
The single C. elegans ryanodine receptor ortholog is encoded by unc-68. UNC68’s major function is the channel’s role in excitation-contraction coupling in muscle,
32
where the receptor acts as a Ca2+ amplifier (Maryon et al., 1996; Maryon, Saari, &
Anderson, 1998). The unc-68 gene has four isoforms, and the channel has notable
homology with the mammalian RYR1, RYR2, and RYR3 subunits (Hamada, Sakube,
Ahnn, Kim, & Kagawa, 2002; Sakube, Ando, & Kagawa, 1997). Despite being a major
Ca2+ amplifier in muscle, UNC-68 is not required for proper excitation-contraction
coupling, only demonstrating a weak uncoordinated movement phenotype in unc-68 nulls
(Hamada et al., 2002; Maryon et al., 1996; Sakube et al., 1997). Much like itr-1, unc-68
has a wide expression pattern, including body wall muscle, some head and tail neurons,
the pharynx, and anal depressor and sphincter muscles (Hamada et al., 2002; Maryon et
al., 1996). Like mammalian RyRs, UNC-68 channels are blocked by ryanodine, and unc68 loss-of-function mutants demonstrate ryanodine resistance (Maryon et al., 1996).
Interestingly, UNC-68 activity is required for normal quantal size at the neuromuscular
junction, implicating this Ca2+ as a potential modulator of synaptic function (Liu et al.,
2005). Despite extensive characterization of UNC-68 function in body wall and
pharyngeal muscle, the neuronal function of this Ca2+ is not well understood, and more
characterization will be required to see how UNC-68 and ITR-1 act in the nervous system
to modulate sensory inputs.
1.3.5 Monoamines and neuropeptides extensively modulate ASH sensory responses
1.3.5.1 Monoaminergic-peptidergic modulation of ASH sensory responses is
complex and requires multiple neurons
33
Monoamines and neuropeptides extensively modulate ASH sensory responses to
give rise to proper behaviors. The ASH neurons are the primary sensors of dilute 1octanol, but the ADL and AWB sensory neurons are likely also involved in sensing
higher concentrations of the odorant (Chao et al., 2004; G. P. Harris et al., 2009; Mills et
al., 2011)(see Chapter 3). When exposed to 30% 1-octanol (dissolved in ethanol,
volume/volume) in the absence of food, wild-type C. elegans elicit a reversal in about 10
seconds (Chao et al., 2004; G. P. Harris et al., 2009; Zahratka et al., 2015). In contrast,
animals on food exhibit a stimulated response, instead initiating a reversal in five seconds
(Chao et al., 2004; G. P. Harris et al., 2009; Wragg et al., 2007). Interestingly, incubating
animals on 4 mM 5-HT for 20 – 30 minutes mimics the food signal, stimulating reversal
and resulting in a five-second response (Chao et al., 2004; G. P. Harris et al., 2009; G.
Harris et al., 2010, 2011; Zahratka et al., 2015). Notably, serotonergic stimulation of the
behavioral response requires the 5-HT6-like GPCR SER-5 receptor expressed in ASH, as
ser-5 null animals are not stimulated behaviorally by 5-HT, and the effect only returns
when the receptor is rescued with a cell-specific transgene (G. P. Harris et al., 2009).
Interestingly, the serotonergic modulation of the ASH reversal circuit extends beyond the
ASHs, with the 5-HT-gated Cl- channel MOD-1 acting in the AIY and AIB interneurons,
and the 5-HT2-like GPCR SER-1 acting in the RIA and RMD interneurons to stimulate
aversive responses in the presence of 5-HT (G. P. Harris et al., 2009). Together, these
findings suggest a role for 5-HT throughout the nervous system in the stimulation of
aversive behaviors.
While 5-HT acts at multiple levels to modulate aversive responses, ASHs are
modulated by multiple other neurotransmitters and receptors. OA, the invertebrate
34
homolog of norepinephrine, directly antagonizes 5-HT activity in ASH via the Gαocoupled OCTR-1, preventing 5-HT-mediated stimulation of responses to dilute 1-octanol
resulting in a 10-second reversal (Wragg et al., 2007). Interestingly, a second OAactivated GPCR, the Gαq-coupled SER-3, inhibits OCTR-1 at high concentrations of OA
(Mills et al., 2011). OA has additional roles at high concentrations of 1-octanol, as it
facilitates release of multiple neuropeptide products downstream of its SER-6 GPCR:
NLP-7, and NLP-9 from the ASIs, NLP-7 and NLP-8 from the ADLs, and NLP-9 from
the AWB sensory neurons (Mills et al., 2011). NLP-9 from AWB goes on to activate
NPR-18 on the ASER sensory neuron to inhibit avoidance behaviors, while NLP-7
activates NPR-15 on AWC sensory neurons to stimulate avoidance behaviors (Mills et
al., 2011). TA acts comparably to OA by inhibiting serotonergic stimulation at dilute 1octanol concentrations as well as inhibiting responses to 100% 1-octanol alone
(Komuniecki, Harris, Hapiak, Wragg, & Bamber, 2011; Mills et al., 2011; Wragg et al.,
2007). Like OA, TA also stimulates the release of neuropeptide products at multiple
levels in the ASH nociceptive circuit. Notably, the Gαq-coupled TYRA-3 acts in the ASIs
to facilitate release of NLP-1, NLP-14, and NLP-18 onto downstream interneurons, in the
dopaminergic ADEs and CEPs to facilitate DA release, and in the RICs to facilitate OA
release (Hapiak et al., 2013). NLP-1 peptides act through NPR-11 on the AIA
interneurons to inhibit downstream signaling, where NLP-14 peptides inhibit the ADLs
via NPR-10 (Hapiak et al., 2013). The ASIs also release two other peptides, NLP-6 and
NLP-18, but their method of release and downstream targets remain unknown (Hapiak et
al., 2013; Mills et al., 2011). ASH odorant responses are also mediated bidirectionally by
DA through the inhibitory DOP-3 and excitatory DOP-4 receptors, decreasing and
35
increasing behavioral sensitivity to ASH-sensed repellents, respectively (Ezak & Ferkey,
2010; Ezcurra et al., 2011). In summary, multiple small-molecule inputs to the ASHs can
impact C. elegans behavioral responses to extensively modulate sensory behavior, and
this regulation can be found across multiple neurons and neurotransmitter systems
throughout the ASH locomotory circuit.
1.3.5.2 Calcium Channels in C. elegans as Sensory Modulators
While a majority of the work studying Ca2+ channels in C. elegans has been in the
context of body wall muscle, very little is known about the activity of VGCCs in the
nervous system of the worm and how they interact with monoamines and neuropeptides.
In Chapter 3, I will describe the basic characterization of 1-octanol responses in sensory
neurons. I will also characterize the modulation of the 1-octanol response to monoamines
and neuropeptides. In Chapter 4, I will discuss a novel situation where 5-HT inversely
modulates 1-octanol-evoked Ca2+ signals and depolarization. Finally, in Chapter 5, I will
discuss the Ca2+-driven negative feedback loop that directly inhibits the EGL-19 channel
when the animal is exposed to 5-HT.
36
Chapter 2
Materials and Methods
2.1 Worm strains and maintenance. Strains were maintained on standard nematode
growth media (NGM) plates at 20 °C seeded with E. coli OP50 bacteria according to
standard protocols (Brenner, 1974). The following strains were used: N2, CX10979
kyEx2865[Psra-6::GCaMP3]; FY907 grIs17[Psra-6::GCaMP3]; FY867 ser-5(tm2654)
I, kyEx2865[Psra-6::GCaMP3]; FY882 itr-1(sa73) IV, grIs17[Psra-6::GCaMP3];
FY933 egl-19(n582) IV, grIs17[Psra-6::GCaMP3]; FY934 egl-30(n686sd) I,
grIs17[Psra-6::GCaMP3]; FY935 unc-68(e540) V, grIs17[Psra-6::GCaMP3]; FY883
unc-2(e55) X, grIs17[Psra-6::GCaMP3]; FY905 shn-1(ok1241) II, grIs17[Psra6::GCaMP3], FY975 tax-6(p675) IV, grIs17[Psra-6::GCaMP3], FY960 aka-1(ok2520)
II, grIs17[Psra-6::GCaMP3], FY937 osm-9(ky10) IV, grIs17[Psra-6::GCaMP3], ser5(tm2654) I, JT73 itr-1(sa73) IV, CB540 unc-68(e540) V, RB1196 shn-1(ok1241) II,
PR675 tax-6(p675) IV, VC1939 aka-1(ok2520) II, CX10 osm-9(ky10) IV.
2.2 RNA interference. RNAi transgenes were constructed as previously described using
PCR and the Phusion DNA polymerase (New England BioLabs, Ipswich, MA) (Esposito
et al., 2007). Psra-6::egl-19RNAi arrays were expressed in the ASH sensory neurons of
37
FY907 animals. The transgene was created by fusing the sra-6 promoter (expressed in
ASH, ASI, and PVQ neurons) to DNA fragments of exon-rich regions of egl-19. A sra-6
forward primer (sra-6F) was amplified from genomic DNA with Psra-6::egl-19Prs
(sense) and Psra-6::egl-19Pra (antisense) reverse primers to generate RNAi templates B
and C, respectively. Template A was amplified from genomic DNA using primers
targeted against the egl-19 coding region with the egl-19Tf (forward) and egl-19Tr
(reverse) primers. The sense RNAi construct was created by amplification of templates A
and B with the sra-6F* and egl-19Tr* primers, and the antisense construct was created by
amplification of templates A and C with the sra-6F* and egl-19Tf* primers. Sense and
antisense transgenes were microinjected along with Punc-122::GFP into the gonad at
concentrations of 100ng/µl. Lines were analyzed within four generations of original
injection. The primers used for transgene construction were: sra-6F: 5’CTTTTCATCTCGACCAGACGGTG-3’; sra-6F*: 5’CAATGTCCACTGATGTACCTTTCTATC-3’; egl-19 Tf: 5’GTTCCGTGTGATGCGTCTCGTG-3’; egl-19 Tf*: 5’GCGTCTCGTGAAGCTGCTTTC-3’; egl-19 Tr: 5’-CTTGCCAGGCTTCTCCAGTTG3’; egl-19 Tr*: 5’-CAGGCTTCTCCAGTTGCTGATC-3’; Psra-6::egl-19 Prs: 5’CACGAGACGCATCACACGGAACGGCAAAATCTGAAATAAATAAATATTAAAT
TCTGCG-3’; Psra-6::egl-19 Pra: 5’CAACTGGAGAAGCCTGGCAAGGGCAAAATCTGAAATAATAAATATTAAATTC
TGC-3’.
38
2.3 Behavioral assays. 1-octanol assays were performed as previously described (Chao
et al., 2004; Harris et al., 2009; Zahratka et al., 2015) with slight modification. 30 – 40 L4
and young adult animals were picked the night before assays to fresh NGM plates seeded
with OP50. Two hours before the assay, fresh NGM plates were prepared, with 4 mM
serotonin creatinine sulfate monohydrate (Sigma-Aldrich, St. Louis, MO) being added to
the molten media before pouring and cooling (~50 °C). A glass capillary (Sutter
Instruments) was dipped in 30% 1-octanol (dissolved in 100% ethanol, volume/volume)
was placed in front of a forward-moving animal. In the absence of 5-HT, animals were
picked to an intermediate plate without food for 1 – 5 minutes, picked to the unseeded
assay plate, and then assayed after 10 minutes. For plates containing 5-HT, animals were
picked to an intermediate plate without food for 1 – 5 minutes, picked to unseeded 5-HT
assay plates, and then assayed after 30 minute 5-HT incubation. Assays were performed
between 12 – 4 PM and in a temperature range of 20 – 24 ºC to control for potential
environmental variability.
2.4 Calcium imaging. There were two different protocols for calcium imaging studies,
one using 1-octanol or other odorants as a stimulus, and the other using variable
potassium concentrations in external solution to artificially induce depolarization.
1-octanol calcium imaging. Young adult animals expressing Psra-6::GCaMP3 were
glued to 1.5 mm coverslips coated with Sylgard (Dow Corning, Midland, MI) immersed
in electrophysiology external solution (ingredients listed below) using WormGlu
cyanoacrylate glue (GluStitch, Delta, British Columbia, Canada). Prepared coverslips
39
were placed in a Warner RC26G laminar flow chamber (Warner Instruments, Hamden,
CT) and continuously perfused with external solution. Saturated 1-octanol solution (2.373
µM), 1 mM DHCA, 10 mM primaquine, or 100 µM IAA was dissolved in external
solution and delivered by gravity feed under control of solenoid valves through a
Perfusion Pencil (AutoMate Scientific, Berkeley, CA) or an equivalent device produced
in-house for 20 seconds. Odorant solutions also contained 1 µM of the fluorescent tracer
sulforhodamine 101 (SR101, Sigma-Aldrich), which was used to confirm stimulus
application. Odorant responses were only evoked in ASH, not ASI or PVQ, and only
evoked responses in the presence of odorant, not SR101 or external alone (data not
shown). Recordings were performed on an Axioskop FS Plus 2 upright compound
microscope (Zeiss ×40 Achroplan water immersion objective, GFP filter set number 38)
with an Orca ER CCD camera (Uniblitz, Vincent Associates, Rochester, NY).
Fluorescent illumination was limited to 30% of maximum intensity to limit GCaMP3
photobleaching. There was no significant difference in bleaching rates across recording
days or genotypes (data not shown). 5-HT treatment was performed by incubating
animals on NGM plates containing 4 mM 5-HT for 30 – 45 minutes prior to recording.
For nemadipine-A (NemA) experiments, 5 µM NemA was spread across the top of an
NGM plate, and animals were incubated for 30 minutes. While recording, NemA was
also present in the bath perfusion to keep the animals constantly exposed to the ligand.
Animals were recorded for about 110 s, with fluorescent light coming on at 10 s to
establish baseline levels. 1-octanol or other odorants were applied at 30 s into the
recording, with drug removal occurring at 50 s. Fluorescent images and video were
40
acquired with MetaVue 7.6.5 (MDS Analytical Technologies, Sunnyveil, CA). Exposure
times were 50 ms with 4 × binning and a length of 1500 frames.
High K+ calcium imaging. Young adult animals expressing Psra-6::GCaMP3 were
prepared as above. However, unlike delivery of 1-octanol or DHCA solutions, Highpotassium electrophysiology external (115 mM NaCl, 30 mM KCl, 5mM CaCl2, 1 mM
MgCl2, 10 mM glucose, and 15 mM HEPES; pH 7.27 – 7.33, 327 – 333 mOsm.) was
delivered using a four-barreled drug delivery apparatus controlled by a SF-77B Fast-Step
step motor (Warner Instruments, step size 300 µm) and syringe pump (KD Scientific,
Holliston, MA). For other concentrations of K+, the KCl and NaCl concentrations were
balanced against one another to maintain proper osmolarity. Animals were dissected
using a polished electrode (TW150-3, World Precision Instruments, Sarasota, FL) shaped
into a sharp point on a Narishige MF-83 microforge (Narishige, Setagaya-ku, Japan),
mounted on a micromanipulator (Sutter MP285, Sutter Instruments, Novato, CA) slitting
the cuticle and exposing the ASH neurons to the external bath. Animals with visible
blebbing of the axon, dendrite, or soma were discarded. ASH neurons were then bathed
in K+ external for 20 seconds as described above, artificially depolarizing the neuron
independently of amphid stimulation. 5-HT and NemA treatments and video acquisition
were performed as above.
2.5 Electrophysiology. For patch-clamp electrophysiology experiments, young adult
animals expressing Psra-6::GCaMP3 were glued to coverslips as described above.
Animals were dissected as described above, exposing the ASH neurons to the external
41
bath. Whole-cell recordings were performed using pressure-polished patch pipettes with
10-30 MΩ resistance (Goodman and Lockery, 2000). Electrophysiological external
solution contained 150 mM NaCl, 5 mM KCl, 5 mM CaCl2, 1 mM MgCl2, 10 mM
glucose, and 15 mM HEPES; pH 7.27 – 7.33, 327 – 333 mOsm. Electrophysiological
internal solution contained 115 mM potassium gluconate, 15 mM KCl, 10 mM HEPES, 5
mM MgCl2, 0.25 mM CaCl2, 5 mM EGTA, 20 mM sucrose, 5 mM MgATP, 0.25 mM
NaGTP, pH 7.20, 315 mOsm. 1-octanol was applied with the Perfusion Pencil as
described previously, but was mounted to the Fast Step motor for precise spatial and
temporal control of odorant application in these experiments. A second pipette 20 – 30
µm in diameter driven by the syringe pump was placed in the bath along with the
Perfusion Pencil to tightly control drug application to the animal’s nose and keep the 1octanol solution away from the exposed ASH neurons. After recording, animals were
checked for SR101 staining, and any animals without visible SR101 fluorescence on the
nose or visible SR101 fluorescence on the ASH neurons were excluded.
Electrophysiological transients were recorded with an Axopatch 200B amplifier
(Molecular Devices, Sunnyvale, CA) in current clamp with 0 pA injected current, 10 kHz
sampling and 2 kHz filtering. Samples were digitized with a Digidata 1440A digitizer,
and analyzed with pCLAMP10 software (Molecular Devices). 5-HT treatment was
performed as above.
2.6 Statistical analysis. ROI selection. Square regions of interest were chosen for
calcium imaging experiments by centering the ASH soma, and enclosing an area equal to
42
roughly 100 soma areas. For ASH axons, regions equal in size to soma ROIs were
chosen, but the ASH dendrite and soma were excluded.
Quantification of calcium imaging experiments. After recording, videos were analyzed
using Jmalyze (Rex Kerr) and output files were analyzed in Microsoft Excel. The change
in fluorescence relative to baseline, ΔF/F0, was calculated as follows:
Where F is the fluorescence value at the current frame, and F0 is the baseline
fluorescence, which is the frame immediately prior to drug application. Any recording
with baseline variance greater than 5% of the average of frames five seconds prior to
drug application were excluded from analysis. ΔF/F0 values were calculated for frames
corresponding to 20 – 60 s.
Kinetic analysis. Calcium kinetic rise times were calculated by subtracting the time at
which a calcium transient was at 90% of its peak by the time at which the same transient
was at 10% of its peak. Slopes were calculated by taking two short timeframes, one
during drug application (43 – 48 s) and one after (52 – 57 s) and measuring the change in
fluorescence values over time (Δy/Δx).
Statistical tests and graphs. All statistical analysis was performed using unpaired, twotailed Student’s t-tests unless otherwise specified. Graphs were constructed using
GraphPad Prism (GraphPad, La Jolla, CA) and presented as mean ± SEM.
43
All other reagents were obtained from Fisher Scientific (Pittsburgh, PA) or SigmaAldrich (St. Louis, MO) unless otherwise specified. Worm strains were obtained from the
Caenorhabditis Genetics Center (University of Minnesota, Minneapolis, MN) or National
BioResource Project of Japan (Tokyo Women’s Medical University, Shinjuku-ku, Tokyo,
Japan).
44
Chapter 3
Monoamines and neuropeptides cell-specifically
modulate ASH-mediated aversive behavior in
Caenorhabditis elegans
3.1 Results
3.1.1 ASH neurons, but not ADL or AWB neurons, are required for saturating 1octanol sensation
Monoamines modulate C. elegans olfactory behaviors as a response to nutritional
status and environmental factors. C. elegans respond to multiple noxious stimuli,
including harsh nose touch, osmotic stress, unfavorable concentrations of oxygen and
carbon dioxide, strongly alkaline pH, and volatile odorants (Aoki et al., 2011; Colbert et
al., 1997; Ezcurra et al., 2011; Hilliard et al., 2005, 2002; Kodama-Namba et al., 2013;
Sassa, Murayama, & Maruyama, 2013; Zimmer et al., 2009). The ASH sensory neurons
are polymodal, sensing nose touch, volatile odorants, and osmotic stresses (Ezcurra et al.,
2011; G. Harris et al., 2010; Hilliard et al., 2005; Mills et al., 2011; Zahratka et al., 2015).
ASH-mediated sensory responses have been extensively studied using 1-octanol, a
volatile odorant that elicits escape behavior (Chao et al., 2004; Ezak & Ferkey, 2010; G.
P. Harris et al., 2009; G. Harris et al., 2010; Mills et al., 2011; Wragg et al., 2007).
45
Previous behavioral studies have implicated the AWB and ADL sensory neurons in 1octanol sensation, but only at high concentrations and only when the ASH is ablated
(Chao et al., 2004). However, because behavior is the sum total of multiple neurons
throughout a sensory circuit, cell-specific assays are required to elucidate a single
neuron’s involvement in the pathway. To determine which sensory neurons are required
for 1-octanol sensation, we performed Ca2+ imaging experiments using the GCaMP3
indicator expressed in ASHs, ADLs, and AWBs (Tian et al., 2009) (Figure 3-1).
Interestingly, only ASH neurons, but not ADLs and AWBs, demonstrate measurable
responses to 2.4 µM 1-octanol (Figure 3-1). Together, these results confirm that in intact
animals, the ASHs are the primary sensors of saturating 1-octanol and initiators of
reversals, with ADLs and AWBs likely only acting at higher concentrations to initiate the
aversive response.
46
*
*
*
Figure 3-1 1-octanol responses of ASH, ADL, and AWB sensory neurons. (A – C) Acute
1-octanol exposure triggers a rise in intracellular Ca2+ level in ASH neurons, but not in
ADL or AWB neurons. Representative traces of GCaMP3 fluorescence intensity from
single neurons as shown, relative to baseline. Grey boxes indicate 30-second stimulus
application. (D) Quantitative comparison of 1-octanol responses from ASH, ADL, and
AWB sensory neurons. Bars represent mean ± SEM. Figure and figure legend adapted
from Mills et al., 2012. * denotes P < 0.001 compared to ASH.
47
3.1.2 Basic characterization of ASH 1-octanol responses: Dose response and kinetics
Since the ASHs are the primary sensors of saturating 1-octanol in our system, we
decided to perform basic characterization of Ca2+ imaging responses. Notably, ASHs
respond to multiple concentrations of 1-octanol, with fluorescent intensity changes
relative to baseline increasing linearly from 3% of saturating (72 nM) to 100% saturating
(2.4 µM) (Figure 3-2a). ASH Ca2+ responses to 1-octanol demonstrate slow kinetics
compared to nose touch stimuli, starting to rise around 730 ms (±110 ms, n = 8),
suggestive of activation of an olfactory G-protein coupled receptor (Figure 3-2b). In
summary, these data indicate that ASH Ca2+ responses to 1-octanol are graded and
increase with stimulus intensity, and that 1-octanol likely triggers a sensory signal
transduction cascade through an olfactory G-protein coupled receptor. Finally, the
TRPV1 channel ortholog osm-9 is necessary for ASH-mediated sensory responses to
Cu2+ ions and osmotic stress induced by 1 M glycerol (Colbert et al., 1997; Hilliard et al.,
2005; Liedtke, 2007; Tobin et al., 2002). In response to these stimuli, olfactory Gproteins, such as ODR-3 or GPA-3, recruit polyunsaturated fatty acids which are then
thought to activate OSM-9 to depolarize the neuron and initiate sensory transduction
(Colbert et al., 1997; Hilliard, Bergamasco, Arbucci, Plasterk, & Bazzicalupo, 2004;
Kahn-Kirby, Dantzker, Apicella, & Schafer, 2004; Roayaie et al., 1998; Tobin et al.,
2002). To test if the OSM-9 channel is necessary for 1-octanol-induced Ca2+ responses,
we Ca2+ imaged 1-octanol responses in osm-9 null mutant animals. Consistent with other
ASH-mediated stimuli, osm-9 nulls exhibited undetectable 1-octanol-evoked Ca2+
responses (Figure 3-2c). These data suggest that ASH sensory neurons can respond to a
48
range of 1-octanol concentrations, and that 1-octanol signal transduction likely occurs
through the coupling of an olfactory GPCR resulting in the activation of OSM-9.
49
Figure 3-2 Basic characterization of the 1-octanol Ca2+ response in ASH neurons. (A) 1octanol presentation evokes Ca2+ increases at multiple water-soluble concentrations,
including 3% of saturating (72 nM), 10% of saturating (240 nM), and 100% saturating
(2.4 µM). Behavioral data suggest ASH can respond to higher concentrations of 1octanol, but 2.4 µM is the highest concentration dissolvable in our electrophysiological
external solution. (B) Focused view of the first four seconds of 1-octanol exposure in
ASH neurons. Sensory transduction cascades result in a visual Ca2+ increase after an
average of 730 ms (±110 ms, n = 8). (C) Quantitative representation of ASH Ca2+
responses in wild-type and osm-9(ky10) null mutant animals. Bars represent mean ±
SEM. ***: P < 0.001 compared to wild-type. n ≥ 5 for each group.
50
3.1.3 ASH-mediated 1-octanol responses are modulated by monoamines and
neuropeptides
The ASH sensory neurons are extensively modulated by monoamines and
neuropeptides, including 5-HT, DA, OA, and NLP-3 (Ezcurra et al., 2011; G. P. Harris et
al., 2009; G. Harris et al., 2010; Mills et al., 2011; Wragg et al., 2007). Additionally, 5HT and OA bidirectionally modulate ASH-mediated behavioral responses to 1-octanol,
dependent on the SER-5 and OCTR-1 receptors in ASH (G. P. Harris et al., 2009; Mills
et al., 2011). To examine the role of monoamines and neuropeptides on 1-octanol-evoked
Ca2+ responses in ASH, we incubated animals on 4 mM 5-HT, DA, OA, and TA
individually while exposing animals to 10% or 100% saturating 1-octanol solution
(Figure 3-3). Surprisingly, incubation on 4 mM 5-HT dramatically decreased 1-octanolevoked Ca2+ transients at dilute and saturating concentrations, a surprising result contrary
to previous behavioral and optical imaging studies (Figure 3-3a, b)(G. P. Harris et al.,
2009; Hilliard et al., 2005; Zahratka et al., 2015). Behavioral studies suggest a role for 5HT acting as a “food-at-hand” signal on three distinct receptors in the ASH locomotory
circuit, with SER-5 in the ASHs stimulating behavioral responses (Chao et al., 2004; G.
P. Harris et al., 2009). Similarly, treatment with 2.5 mM 5-HT stimulated sensory-evoked
Ca2+ transients in response to nose touch (Hilliard et al., 2005). Our results, taken with
previous observations, illustrate a unique role for 5-HT in the modulation of ASH-evoked
sensory responses to chemorepellents including 1-octanol.
In contrast to 5-HT above, treatment with 4 mM OA, DA, or TA alone had no
significant effect on ASH 1-octanol-evoked Ca2+ signals (Figure 3-3b). However,
51
treatment with 4 mM 5-HT and 4 mM OA simultaneously resulted in no significant
difference in Ca2+ transient amplitudes compared to wild-type, suggesting that OA
directly antagonizes 5-HT directly in ASH (Figure 3-3b).
In addition to monoamine neurotransmitters, neuropeptides are also important
modulators in C. elegans. To study the role of neuropeptide modulation in ASH, we
examined Ca2+ transients in egl-3 reduced function mutants. EGL-3 is a proprotein
convertase required for proper neuropeptide processing in C. elegans, so egl-3
hypomorphs likely have inhibited neuropeptide signaling (G. Harris et al., 2010; Kass,
Jacob, Kim, & Kaplan, 2001; Mellem et al., 2002; Mills et al., 2011). 1-octanol-evoked
Ca2+ transients were significantly reduced in egl-3(n150) hypomorphs without
monoamine treatment, suggesting that neuropeptide signaling also modulates ASH
sensory responses (Figure 3-3c). Curiously, 5-HT treatment had no effect on 1-octanolevoked Ca2+ transients in egl-3 mutants, while treatment with OA further depressed the
Ca2+ signal (Figure 3-3c). In closing, these experiments suggest that ASH sensory
neurons are extensively modulated by monoamines and neuropeptides, and the
neuromodulators can bidirectionally modulate each other to form a complex web of
sensory regulation.
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Figure 3-3 Monoamines and neuropeptides modulate the 1-octanol response. (A)
Quantification of saturating 1-octanol-evoked ASH Ca2+ responses after 30-minute
exposure to 4 mM monoamines. (B) 4 mM 5-HT also reduces 1-octanol-evoked Ca2+
transients at the lower 10% of saturating concentration. (C) Saturating 1-octanol-evoked
ASH Ca2+ responses in wild-type and egl-3(n150) animals treated with 4 mM 5-HT or
OA. Bars represent mean ± SEM. *: P < 0.05 compared to wild-type. ‡: P < 0.05
compared to egl-3(n150) untreated. n ≥ 7 for each group.
53
3.1.4 5-HT modulates ASH sensory responses to multiple ASH-sensed stimuli
5-HT inhibition of ASH Ca2+ in response to 1-octanol was very surprising,
considering that 5-HT treatment enhances behavioral responses, and increases in
intracellular Ca2+ are generally taken to be indicative of increased neuronal activity
(Grienberger & Konnerth, 2012; G. P. Harris et al., 2009; G. Harris et al., 2010; Mills et
al., 2011; Zimmer et al., 2009). To examine if 5-HT modulated other ASH-sensed
noxious stimuli, we performed Ca2+ imaging experiments on other odorants, including
dihydrocaffeic acid (DHCA), primaquine, and isoamyl alcohol (IAA) (Aoki et al., 2011;
Ezcurra et al., 2011; Hilliard et al., 2004; Yoshida et al., 2012). DHCA, primaquine, and
IAA each elicit ASH Ca2+ transients, although each of these odorants had significantly
smaller transients compared to saturating 1-octanol (Figure 3-4). 4 mM 5-HT
significantly decreased evoked Ca2+ transients in DHCA-exposed animals, consistent
with 1-octanol exposure (Figure 3-4). Conversely, 5-HT treatment did not inhibit
sensory-evoked Ca2+ transients in animals exposed to primaquine (Figure 3-4). IAAexposed animals were not assayed with 5-HT. In summary, multiple odorants evoke Ca2+
signals in the ASHs that are negatively regulated by 5-HT, suggesting that serotonergic
signaling is a modulator of a subset of olfactory stimuli in the ASH sensory neurons.
54
Figure 3-4 5-HT modulates ASH activity in response to multiple odorants.
Quantification of evoked ASH Ca2+ responses to saturating 1-octanol, 1 mM DHCA, 10
mM primaquine, or 100 µM IAA in the presence or absence of 4 mM 5-HT treatment.
Bars represent mean ± SEM. *: P < 0.05 compared to untreated. n ≥ 6 for each group.
55
3.2 Discussion
In the present chapter, we have isolated a role for the ASH sensory neurons, but not
ADLs or AWBs, in the sensation of saturating 1-octanol in our Ca2+ imaging setup. We
also have elucidated roles for 5-HT, OA, and neuropeptides in direct modulation of ASH
Ca2+ responses in response to 1-octanol. Finally, we demonstrate that ASHs can respond
to multiple odorants, such as DHCA, which are in turn modulated by 5-HT.
3.2.1 ASHs are the primary sensors of saturating 1-octanol
Behaviorally, ASH neurons are the primary sensors of dilute 1-octanol, where
sensation higher concentrations also requires ADLs and AWBs (Chao et al., 2004; G. P.
Harris et al., 2009). To examine the cell-specific activity of these neurons in vivo, we
performed Ca2+ imaging experiments in response to 1-octanol dissolved in our
electrophysiological external solution. ASH neurons elicited large Ca2+ transients in
response to 1-octanol exposure, but ADL and AWB neurons failed to respond (Figure 31). Given these data, we can infer that the saturating 1-octanol solution is closer to the
30% dilute 1-octanol concentrations used in behavioral experiments than 100% 1-octanol
assuming that these assays are equivalent. This is not surprising, due to 1-octanol’s poor
solubility in water. Additionally, this inference explains the lack of ADL and AWB
responses, as the saturating 1-octanol concentration is likely not great enough to trigger
the activation of these neurons resulting in downstream OA-mediated, SER-6-dependent
neuropeptide release (Mills et al., 2011). While the exact 1-octanol concentrations
56
reaching an animal’s nose in behavioral assays is not known, other groups have modeled
the diffusion of another volatile odorant, 2-nonanone, which could illustrate the
gaseous/liquid interface where an animal would encounter the chemical in its
environment (Yamazoe-Umemoto, Fujita, Iino, Iwasaki, & Kimura, 2015). Importantly,
the ASHs also responded to lower concentrations of 1-octanol as well including 3% and
10% of our saturating solution (Figure 3-2). ASH neurons increased their sensory-evoked
Ca2+ transients as the concentration of 1-octanol increased. These data are important
because they suggest that the animal has the capability to distinguish between multiple
concentrations of an aversive stimulus, and allow us the potential ability to study
modulation of ASH responses if the saturating concentration is too strong and bypasses
regulation.
In addition to basic information about the ASH 1-octanol response, our
characterization was also able to dissect some of the sensory transduction required for 1octanol sensation. 1-octanol-evoked ASH Ca2+ transients began about 730 ms after drug
exposure on average, a slow response suggestive of the activity of an olfactory GPCR
(Figure 3-2b). The 1-octanol stimulus likely transduces a signal through a GPCR due to
730 ms being a long activation time compared to the activation of mechanosensitive ion
channels, which activate on a time scale of about 3 – 5 ms (Geffeney et al., 2011).
Secondly, the TRPV1 ortholog OSM-9 is required for 1-octanol-evoked Ca2+ transients
(Figure 3-2c). These findings are consistent with previous studies, which implicate osm-9
in sensation of multiple ASH-sensed olfactory and osmotic stimuli (Colbert et al., 1997;
Hilliard et al., 2005; Lindy et al., 2014; Sokolchik et al., 2005; W. Wang et al., 2015).
Interestingly, OSM-9 is thought to be downstream of the olfactory G-protein ODR-3,
57
which is activated in response to sensory stimuli (Roayaie et al., 1998). In this pathway,
1-octanol would bind to the currently undiscovered 1-octanol-sensing GPCR at the
amphid, which would activate ODR-3. ODR-3 in turn would recruit polyunsaturated fatty
acids to activate OSM-9, leading to neuronal activation and downstream signaling,
eventually resulting in downstream neurotransmitter release (Colbert et al., 1997; KahnKirby et al., 2004; Roayaie et al., 1998). However, because ASH senses multiple noxious
stimuli, some discrimination is required for proper behavioral outputs. ASH also
expresses multiple atypical chemosensory G-alpha proteins and an olfactory-specific Ggamma subunit, GPC-1, allowing for great complexity and specificity for sensory
transduction in this polymodal neuron (C I Bargmann & Kaplan, 1998; C. I. Bargmann,
1998; Chao et al., 2004; Jansen et al., 2002; Roayaie et al., 1998; Yamada et al., 2009).
Therefore, there is likely considerable crosstalk between G-protein signaling pathways
for odorant discrimination in the ASHs.
3.2.2 5-HT stimulates behavior, but inhibits Ca2+ transients in response to 1-octanol
ASHs are extensively modulated by monoamines and neuropeptides, as chronic
exposure to these neurotransmitters can directly affect behavioral responses (Ezcurra et
al., 2011; G. P. Harris et al., 2009; G. Harris et al., 2010; Mills et al., 2011; Wragg et al.,
2007). To demonstrate the effects of monoamines and neuropeptides in our Ca2+ imaging
system, we incubated animals on 4 mM of the respective monoamine for 30 minutes as
one would in behavioral experiments and then exposed them to saturating 1-octanol. 5HT and OA are known modulators of ASH behaviorally, with effects reliant upon the
58
GPCRs SER-5 and OCTR-1 (G. P. Harris et al., 2009; Mills et al., 2011; Zahratka et al.,
2015). 5-HT, acting through SER-5, stimulates behavioral responses to dilute 1-octanol,
while OA, acting through OCTR-1, antagonizes 5-HT-mediated stimulation (G. P. Harris
et al., 2009; Mills et al., 2011). Surprisingly, 5-HT inhibits 1-octanol-evoked ASH Ca2+
signals, which was unexpected given the stimulating effect the modulator has
behaviorally (Figure 3-3a). Moreover, this effect is demonstrated at multiple
concentrations, as 5-HT inhibits Ca2+ transients at both saturating and 10% of saturating
concentrations (Figure 3-3b). These data raise the interesting conundrum where the
behavioral data suggest 5-HT has a stimulatory effect on ASH cellular activity, whereas
the Ca2+ data predicts that 5-HT should inhibit ASH cellular activity. The dissection of
this apparent uncoupling is the major topic of this work, with Chapter 4 resolving the
cell-specific effects of 5-HT on ASHs, and Chapter 5 describing a novel Ca2+-driven
negative feedback loop modulated by 5-HT. Interestingly, simultaneous treatment with 5HT and OA resulted in no significant difference in 1-octanol-evoked ASH Ca2+
transients, further supporting that OA suppresses 5-HT behaviorally and at the level of
ASH Ca2+ responses (Figure 3-3b). Behavioral experiments implicate SER-5 and OCTR1 as 5-HT and OA receptors, respectively, and our data confirm a cell-specific role for
SER-5 in ASH (see Chapters 4 and 5)(G. P. Harris et al., 2009; Mills et al., 2011).
Behavioral experiments utilizing whole-animal mutants, cell-specific RNAi, and cellspecific rescue suggest SER-5 is acting through an excitatory G-protein (either Gαs or
Gαq), and OCTR-1 is inhibitory via Gαo (Cunningham et al., 2012; G. Harris et al., 2010;
Wragg et al., 2007). Therefore, there could be G-protein network crosstalk via these
monoamines, with 5-HT stimulating behavior via Gαs/ Gαq, and OA inhibition via Gαo.
59
Interestingly, the atypical chemosensory G-protein α-subunit GPA-11 is also required for
5-HT-mediated stimulation of behavioral responses to 1-octanol, suggesting even more
crosstalk between conventional and sensory-specific G-protein signaling pathways (Chao
et al., 2004).
In contrast to 5-HT’s surprising decrease of 1-octanol-evoked Ca2+ signals, OA,
TA, and DA had no measurable effects by themselves (Figure 3-3b). These results are
also consistent with behavioral data, where OA and TA only act to antagonize 5-HT in
ASH at dilute 1-octanol concentrations (G. P. Harris et al., 2009; G. Harris et al., 2010;
Mills et al., 2011). Interestingly, the ASHs express another OA receptor, SER-3, which
acts to inhibit OCTR-1 signaling via Gαq at high concentrations of exogenous OA (Mills
et al., 2011). Our data demonstrate that application of 4 mM OA alone is not sufficient to
significantly change 1-octanol-evoked Ca2+ levels, suggesting that the effects of OCTR-1
can only be seen in the presence of 5-HT, and that SER-3 is likely not involved at this
lower concentration (Figure 3-3b). A similar effect may be responsible for the lack of
responses seen to TA, as the ASHs express a single TA receptor, TYRA-2, which is
thought to also couple to Gαo (Rex et al., 2005). Notably, TA also inhibits 5-HTmediated stimulation of aversive responses at both 30% and 100% 1-octanol, but this
inhibition is likely dependent on another TA receptor, TYRA-3, that is not expressed in
the ASH neurons, demonstrating that monoaminergic modulation of the 1-octanol
sensory circuit takes place over multiple neurons (Hapiak et al., 2013; Wragg et al.,
2007). Like OA, ASHs express two DA receptors, DOP-3 and DOP-4, which may
interact with each other to modulate behavioral responses (Ezak & Ferkey, 2010; Ezcurra
et al., 2011). Interestingly, DOP-3 likely inhibits sensory responses to 1-octanol, as dop-3
60
mutants are hypersensitive behaviorally to the odorant at all concentrations (Ezak &
Ferkey, 2010). One possibility is, as before, that DA signaling through DOP-3 sets the
tone for modulation by decreasing behavioral sensitivity to 1-octanol, and DOP-4 could
act to counterbalance these effects depending on nutritional state based on data for other
ASH-sensed chemorepellents (Ezak & Ferkey, 2010; Ezcurra et al., 2011). While we saw
no effects on the 1-octanol-evoked Ca2+ signal in the presence of 4 mM DA, other groups
have demonstrated marked increases of sensory-evoked Ca2+ transients at 10 mM DA
that are DOP-4-and nutritional state-dependent (Ezcurra et al., 2011). Taken together,
these data suggest that multiple monoamines can modulate ASH sensation at a variety of
concentrations, demonstrating the need for complex pathways that ultimately give rise to
behavior.
3.2.3 Neuropeptides interact with monoamines to modulate ASH-mediated aversive
behaviors
While multiple monoamines are known to modulate ASH signaling, the effects of
neuropeptides have not been well explored. To examine if neuropeptides had an effect on
1-octanol-evoked Ca2+ transients, we imaged egl-3(n150) reduced-function mutants
(Figure 3-3c). egl-3 encodes a proprotein convertase necessary for proper synthesis of
neuropeptides, resulting in egl-3 mutants having severe defects in their neuropeptide
signaling (C I Bargmann & Kaplan, 1998; C. I. Bargmann, 1998; Kass et al., 2001;
Mellem et al., 2002). In our system, egl-3(n150) animals demonstrate significantly
reduced 1-octanol-evoked Ca2+ transients compared to wild-type counterparts, suggesting
61
that neuropeptides, like 5-HT and OA above, help set the tone for behavioral sensitivity
in the 1-octanol circuit (Figure 3-3c). Unlike their wild-type counterparts, 5-HT treatment
did not further reduce 1-octanol-evoked Ca2+ signals, implying that neuropeptides act
downstream of 5-HT to modulate sensory-evoked ASH Ca2+. However, as egl-3(n150) is
a global mutation, there is also a possibility of neuropeptides acting upstream of 5-HT to
set a comparatively higher level of sensory-evoked Ca2+ as well. The ASH neurons
express three known neuropeptide receptors, NPR-1, NPR-2, and NTR-1, which could
possibly modulate ASH sensory activity (Beets et al., 2012; Garrison et al., 2012; Glauser
et al., 2011; Luo, Xu, Tan, Zhang, & Ma, 2014; Rogers et al., 2003). Curiously, an NPR1 ligand, FLP-21, is also expressed in the ASHs, allowing for potential autoregulation of
sensory activity by neuropeptides (Glauser et al., 2011; Rogers et al., 2003). Notably,
NPR-1 also responds to peptides encoded by FLP-18, which has an expression pattern
including AVA, AIY, RIG, RIM, and pharyngeal neurons, but not the ASHs (Rogers et
al., 2003). Together, these expression data suggest two potential NPR-1 mediated
pathways for ASH regulation: a FLP-18-mediated pathway activating due to feedback
from interneurons, and an autoregulatory pathway driven by FLP-21.
In addition to NPR-1, the ASHs also express the less-commonly studied NPR-2
and NTR-1 receptors. Little is known about the role of NPR-2 in sensory signaling, but
the receptor may be involved in C. elegans responses to methyl salicylate, although these
effects are likely not mediated via the ASHs (Luo et al., 2014). Similarly, NTR-1 encodes
a vasopressin/oxytocin receptor ortholog whose mutation leads to defects in NaCl
sensation and male mating behavior (Beets et al., 2012; Garrison et al., 2012). NTR-1’s
ligand precursor NTC-1 is expressed in AFD, AVK, NSM, DVA, and the pharyngeal M5
62
neurons (Beets et al., 2012). One possibility for ASH sensory regulation is the release of
NTC-1 from NSMs acting as a regulator, much like NSM release of 5-HT can modulate
SER-5 activity (G. Harris et al., 2011). However, ntr-1 null animals are stimulated by 5HT behaviorally, and do not have altered Ca2+ transients in the presence or absence of 5HT, suggesting that NTR-1 may be involved in some ASH-mediated behaviors, but not
all of them (Data not shown, Paul Williams, personal communication). Finally, in
addition to these receptors, there could be many more neuropeptide receptors on the
ASHs that have yet to be discovered, making complex peptidergic regulation a promising
target for future studies for the role of small molecule neurotransmitters in sensory
signaling.
In addition to expression of neuropeptide receptors, the ASHs are also known to
produce products from four neuropeptide-encoding genes, but the individual roles of each
of these neuropeptides in sensory modulation are mostly unknown. One peptide-coding
gene, NLP-3, is known to be released in response to 5-HT stimulation, resulting in a fast
behavioral response to 1-octanol via its potential receptor NPR-17 (G. Harris et al.,
2010). However, the roles of NLP-15, FLP-21, and INS-1, the other ASH-expressed
peptides, remain unclear (G. Harris et al., 2010; Kodama et al., 2006; Mills et al., 2011;
Nathoo, Moeller, Westlund, & Hart, 2001). Interactions between other monoamines and
neuropeptides in ASHs seem likely, but it is difficult to speculate on downstream
signaling events when imaging a single neuron pair upstream of neuropeptide release.
More specifically, there is a distinct possibility that OA, DA, and TA could potentially
inhibit 5-HT-mediated neuropeptide release, leading to inhibition of NLP-3 release and a
delayed 1-octanol response. Our data support this hypothesis, as treatment of egl-3
63
reduced-function mutants with OA significantly decreased the 1-octanol-evoked ASH
Ca2+ signal, where treatment with 5-HT had no visible effect (Figure 3-3c). One potential
explanation of these data are that OA is inhibiting another signaling pathway that sets the
basal evoked Ca2+ response alongside neuropeptides, and this inhibition is only
noticeable in the absence of neuropeptide signaling. However, it is also important to
realize that egl-3 mutants have hampered neuropeptide signaling globally, so peptidergic
pathways upstream and downstream of the ASHs are likely to be affected on a global
scale and developmentally. Given that this mutation is so widespread and developmental
compensation is likely, examining behavior and Ca2+ levels in a cell-specific egl-3 RNAi
construct may be more appropriate for further study of peptide signaling. Additionally,
this interpretation also assumes that sensory-evoked Ca2+ levels will directly correlate
with neuronal excitability and neurotransmitter release, which is likely not a warranted
assumption (see Chapters 4 & 5). These results once again demonstrate the complex
interplay between monoamines and neuropeptides in the modulation of aversive
behavior.
3.2.4 5-HT inhibits sensory-evoked Ca2+ signals from multiple chemorepellents
The ASH sensory neurons are polymodal, sensing a variety of stimuli including
noxious chemicals, osmotic stresses, light nose touch, alkaline pH, and possibly
temperature (Ezcurra et al., 2011; Geffeney et al., 2011; Hilliard et al., 2005; KodamaNamba et al., 2013; Sassa et al., 2013; Thiele et al., 2009; W. Wang et al., 2015). To
examine the role of 5-HT modulation in the context of other sensory modalities, we Ca2+
64
imaged ASH neurons in response to the soluble chemorepellents DHCA, primaquine, and
IAA. Without monoamine treatment, 1 mM DHCA, 10 mM primaquine, and 100 µM
IAA all elicited sensory-evoked ASH Ca2+ transients that were significantly decreased
compared to transients evoked by saturating 1-octanol (Figure 3-4). Interestingly, other
groups have demonstrated that IAA is attractive to C. elegans at concentrations lower
than 100 µM, suggesting the animal can discriminate attractive and repulsive
concentrations of a single odorant (Yoshida et al., 2012). Like 1-octanol, incubation on 4
mM 5-HT for 30 min significantly decreased Ca2+ transients evoked by DHCA,
suggesting serotonergic modulation of chemorepellents is not specific to 1-octanol
(Figure 3-4). However, 5-HT inhibition was not observed in animals exposed to 10 mM
primaquine, suggesting that there may be some divergence of serotonergic modulation
based on modality-specific sensory proteins, such as qui-1 in the case of primaquine
(Hilliard et al., 2004). From these data, it may be possible to generalize serotonergic
modulation as a feature of acute, low-concentration chemosensation, with some stimulusspecific exceptions. While we have studied serotonergic regulation of multiple
chemorepellents, we have not looked at other ASH-sensed modalities, including nose
touch or osmotic stress. Other labs have looked at these modalities previously, with 5-HT
treatment giving mixed results (Hilliard et al., 2005). Therefore, if 5-HT is a general
modulator of low-concentration ASH chemosensation, it may not be a general modulator
of all ASH-sensed stimuli due to differences in sensory transduction (Colbert et al., 1997;
Geffeney et al., 2011; Hart, Kass, Shapiro, & Kaplan, 1999). The dissection of individual
pathways, in addition to their crosstalk, will be of extreme importance to understanding
how multiple sensory transduction pathways can work in a polymodal sensory neuron.
65
In the present chapter, I have discussed the basics of the ASH response to 1octanol and other chemorepellents in terms of amplitude and kinetics, and described how
the responses are modulated by monoamines and neuropeptides. During our studies, we
observed that 4 mM 5-HT increased behavioral sensitivity to dilute 1-octanol, but
decreased 1-octanol-evoked Ca2+ transients. In the next chapter, I will dissect the ASH
Ca2+ signal genetically, focusing on VGCCs and intracellular Ca2+ stores; explore the 5HT receptors required for serotonergic activity in ASHs; and examine the relationships
between 5-HT’s effects on ASH depolarization and ASH Ca2+ signals.
66
Chapter 4
Serotonin differentially modulates calcium transients
and depolarization in a C. elegans nociceptor
Note: The majority of this work has been published in Zahratka JA, Williams PDE,
Summers PJ, Komuniecki RW, and Bamber BA. (2015) Serotonin differentially
modulates Ca2+ transients and depolarization in a C. elegans nociceptor. J Neurophysiol
113: 1041 – 1050.
4.1 Results
4.1.1 Distinct Ca2+ pools mediate ASH-mediated aversive responses
The ASH sensory neurons are necessary for responses to dilute 1-octanol and
other noxious odorants, and these responses can be heavily modulated by monoamines
and neuropeptides (see Chapter 3). Curiously, we found that incubating animals on 4 mM
5-HT stimulates behavioral responses, but decreases the sensory-evoked Ca2+ transient
(Figure 3-3). To examine the relationship between 5-HT, Ca2+ levels, depolarization, and
behavior, we first identified the VGCCs acting in ASH using a combination of genetics
and pharmacology. In untreated conditions, Ca2+ signals were observed in the dendrites,
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somas, and axons of imaged animals (Figure 4-1a). Somal signals were drastically
inhibited by treatment with 5 µM nemadipine-A (NemA), a dihydropyridine antagonist
that specifically targets the C. elegans L-type VGCC EGL-19 (Figure 4-1b)(Kwok et al.,
2006, 2008). Evoked somal Ca2+ levels were also significantly reduced in egl-19(n582)
hypomorphs and animals expressing cell-specific egl-19 RNAi transgenes, but not in unc2(e55) null animals, demonstrating that the L-type EGL-19 is the predominant VGCC in
the soma (Figure 4-1b). In contrast, 1-octanol-evoked axonal Ca2+ signals were less
sensitive to NemA than their somal counterparts, but were still significantly decreased
compared to their untreated counterparts (Figure 4-1c). Null mutants for the unc-2 P/Qtype VGCC also significantly decreased 1-octanol-evoked Ca2+ levels, and only NemA
treatment in unc-2 mutants was able to abolish the sensory-evoked Ca2+ signal (Figure 41c). Taken together, these results indicate subcellular localization of VGCCs in the ASH
sensory neuron, with the L-type EGL-19 being the major channel in the soma but both
EGL-19 and UNC-2 acting in the axon.
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Figure 4-1 Voltage-gated Ca2+ channels mediate 1-octanol-evoked Ca2+ influx into ASH
somas and axons. (A) 1-octanol stimulates Ca2+ transients in the soma and axon; NemA
inhibits somal signals. (B) Somal Ca2+ is inhibited in egl-19 hypomorphs and in NemAtreated animals, but not in unc-2 mutants. (C) Axonal Ca2+ is mediated by both EGL-19
and UNC-2. Bars represent mean ± SEM, *: P < 0.05 compared to wild-type.
‡ indicates significantly different from unc-2 untreated.
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4.1.2 Intracellular stores receptors help shape ASH Ca2+ kinetics
In addition to VGCCs, intracellular Ca2+ stores can play critical roles in shaping
kinetics and intracellular signaling pathways. The C. elegans encodes two intracellular
Ca2+ channels: the IP3R ortholog itr-1, and the RyR ortholog unc-68 (Baylis et al., 1999;
Maryon et al., 1996). Upon stimulation, VGCCs can pass Ca2+ into the cell, triggering
activation of intracellular stores receptors via CICR (Hagenacker et al., 2008; Lian et al.,
2007). CICR does appear to occur in both ASH compartments we studied, acting via
ITR-1 and UNC-68 in the soma, but only ITR-1 is involved in the axon (Figure 4-2b, d).
itr-1 mutants demonstrate reduced evoked Ca2+ amplitudes, slow sensory-evoked Ca2+
transient rise times, and altered decay kinetics. Wild-type Ca2+ signals begin to
desensitize after about 10 s, but then fall to baseline. However, itr-1 hypomorphs fail to
desensitize, instead plateauing at their maximal evoked level until stimulus removal,
upon which the transient drops to baseline (Figure 4-2a, c, e-h). In contrast to itr-1, unc68 mutants do not demonstrate altered rise times or decay kinetics, but still exhibit
reduced Ca2+ amplitudes (Figure 4-2a, c, e-h). These data illustrate the importance of
intracellular stores as Ca2+ amplifiers in the context of sensory-evoked ASH responses,
and add an additional layer of complexity between sensory signals and visible Ca2+
transients.
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Figure 4-2 Intracellular stores receptors amplify and shape sensory-evoked ASH Ca2+
responses in somas and axons. (A) 1-octanol-evoked Ca2+ transients in wild-type, itr-1,
and unc-68 mutant animals in the ASH somas and (C) axons. (B) Quantification of peak
amplitudes of 1-octanol-evoked Ca2+ transients in wild-type, itr-1, and unc-68 mutants
shown in somas and (D) axons. (E) 10-90% rise times of 1-octanol-evoked Ca2+ signals
in somas and (G) axons. (F) Decay kinetic measurements of Ca2+ in the presence (black
bars) and upon removal (white bars) of 1-octanol in somas and (H) axons. Bars indicate
mean ± SEM. *: P < 0.05 compared to wild-type.
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4.1.3 RNAi-mediated knockdown of EGL-19 has no behavioral effect
Since EGL-19 is the major VGCC in the soma, we wanted to explore a direct link
between Ca2+ levels and behavior. To meet this aim, we performed behavioral and 1octanol-evoked Ca2+ imaging experiments in animals expressing RNAi transgenes
targeting egl-19 specifically in ASHs. Interestingly, RNAi knockdown of egl-19 had no
significant effect on initiation of aversive behavior, with both wild-type and transgenic
animals responding to the stimulus in about 10 s (Figure 4-3a). In contrast, egl-19 RNAi
transgenic animals exhibited significantly decreased 1-octanol-evoked somal Ca2+ levels
compared to wild-type (Figure 4-3b). Together, these data suggest that large increases in
ASH somal Ca2+ are not necessary for ASH-mediated aversive responses to dilute 1octanol off food.
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Figure 4-3 1-octanol responses are not affected by reduction of Ca2+ influx through
EGL-19 channels. (A) ASH-specific RNAi knockdown of EGL-19 (driven by the sra-6
promoter) did not significantly affect behavioral responsiveness to 1-octanol. (B) Peak
amplitudes of 1-octanol-evoked Ca2+ transients are significantly inhibited in animals
expressing the Psra-6::egl-19RNAi transgene. Bars indicate mean ± SEM. *: P < 0.05
compared to wild-type. Some behavioral assays were performed by Philip Summers.
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4.1.4 5-HT likely inhibits ASH Ca2+ transients via SER-5 and the EGL-30/Gαq
Pathway
Food and 5-HT both enhance aversive responses to dilute 1-octanol through a
complex serotonergic pathway involving multiple neurons and receptors, including SER5 in the ASH sensory neurons (G. P. Harris et al., 2009; G. Harris et al., 2010, 2011). In
the previous chapter, we also demonstrate that 5-HT treatment decreases 1-octanolevoked ASH Ca2+ transients (see Figure 3-3). To see if GCaMP3 expression alone can
affect aversive responses, we performed behavioral assays in Psra-6::GCaMP3
expressing animals. Treatment with 4 mM 5-HT continued to stimulate aversive
responses similar to non-transgene expressing animals, suggesting GCaMP3 expression
alone does not affect behavioral responses (Figure 4-4a). To further explore where 5-HT
is acting in the ASHs to mediate modulation of aversive responses, we performed Ca2+
imaging assays in response to 1-octanol. Once again, although 5-HT stimulated aversive
responses, sensory-evoked Ca2+ transients were dramatically inhibited in both ASH
somas. (Figure 4-4b, c). However, 5-HT treatment did not significantly decrease ASH
Ca2+ transients in the axon, suggesting 5-HT acts primarily in the soma to mediate
aversive responses (Figure 4-4d). Finally, SER-5 is thought to be the critical 5-HT
receptor in ASHs for modulation of aversive behavior, potentially acting through the Gαq
subunit EGL-30 to stimulate reversals in response to dilute 1-octanol (G. P. Harris et al.,
2009). As predicted, 5-HT-mediated inhibition of ASH somal Ca2+ was not present in
ser-5, egl-30, or itr-1 (a downstream effector of EGL-30) mutant animals, suggesting that
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5-HT inhibits ASH somal Ca2+ transients through a Gαq-mediated pathway involving the
intracellular Ca2+ receptor ITR-1 (Figure 4-4c).
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Figure 4-4 Modulation of 1-octanol-evoked ASH Ca2+ transients by 5-HT. (A)
Behavioral responses to dilute 1-octanol were unchanged in Psra-6::GCaMP3 animals.
(B) 5-HT significantly inhibits ASH Ca2+ transients in ASH somas. Solid line indicates
mean over time, dashed lines indicate SEM, and grey box indicates saturating 1-octanol
application time. (C) Serotonergic modulation of ASH aversive responses requires the
SER-5 GPCR, EGL-30 G protein α-subunit, and ITR-1 internal stores receptor. (D) ASH
axonal Ca2+ transients were not significantly reduced by 5-HT. Bars indicate mean ±
SEM. * denotes P < 0.05 compared to untreated.
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4.1.5 5-HT inversely modulates ASH somal Ca2+ transients and depolarization
The behavioral and Ca2+ imaging data presented previously raise an interesting
dilemma, where the behavioral data predict that ASH depolarization should increase due
to serotonergic stimulation of the aversive response, but the somal Ca2+ data predict the
opposite. To resolve this disconnect, we turned to patch-clamp electrophysiology.
However, because 1-octanol is a long-chain alcohol that could interfere directly with cell
membranes and hamper cellular function, we had to modify existing electrophysiological
protocols for our setup to ensure that 1-octanol does not directly contact the dissected
ASH soma (see Chapter 2, Figure 4-5a). Using this system, we measured 1-octanolevoked depolarizations that were activated slowly compared to mechanical responses
described by another group (Geffeney et al., 2011) (Figure 4-5b). Sensory-evoked
depolarizations were measured at the cell soma, and are likely to represent changes in
membrane potential throughout the neuron, as C. elegans neurons are thought to be
isopotential (Goodman et al., 1998). Surprisingly, 5-HT treatment (performed identically
to behavioral and Ca2+ imaging experiments) increased 1-octanol-evoked depolarization
amplitudes, in spite of 5-HT decreased 1-octanol-evoked Ca2+ transients (Figure 4-5b, c).
To ensure that this difference was not an artifact due to neuronal dissection, we also
performed Ca2+ imaging experiments on dissected ASHs (Figure 4-5d). Treatment with
5-HT still knocked down ASH somal Ca2+ transients, suggesting that 5-HT stimulates
ASH-mediated aversive responses, potentiates sensory-evoked ASH depolarization, but
inhibits sensory-evoked Ca2+ transients.
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Figure 4-5 5-HT potentiates 1-octanol-evoked ASH depolarization. (A) Diagram of
modified electrophysiological recording setup. Left panel: location of drug delivery
(above) and auxiliary pipettes (below) before 1-octanol exposure. Center panel:
Placement of pipettes during 1-octanol exposure. Note that the 1-octanol stream (pink) is
not in contact with the dissected ASH neuron. Right panel: After drug exposure, the
pipettes go back to their original positions, and recording continues. (B) Representative
traces of 1-octanol-evoked ASH depolarization in untreated and 5-HT-treated worms. (C)
Quantification of the effects of 5-HT on 1-octanol-evoked ASH depolarization. (D)
Quantification of 1-octanol-evoked Ca2+ transients in dissected ASH neurons. Bars
represent mean ± SEM, * denotes P < 0.05 compared to untreated. Electrophysiological
experiments largely performed by Dr. Bruce Bamber, dissected Ca2+ imaging in (D) by
Paul Williams.
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4.2 Discussion
The two ASH sensory neurons are necessarily and sufficient for responses to
dilute 1-octanol, and are heavily modulated by monoamines and neuropeptides. ASH
aversive responses can be measured primarily in two ways: through behavioral assays
and Ca2+ imaging. In the present chapter, we have established the subcellular localization
of VGCCs and intracellular Ca2+ stores using these techniques to probe neuromodulation
in the ASHs. Specifically, we have elucidated a unique role for 5-HT as a modulator that
decreases sensory-evoked Ca2+ signals, but increases sensory-evoked ASH depolarization
via a Gαq signaling pathway.
4.2.1 The ASH soma and axon have distinct Ca2+ channel configurations
In the previous chapter, we established that 5-HT decreases sensory-evoked Ca2+
transients in response to 1-octanol. To study potential targets for serotonergic regulation,
we first identified the relevant Ca2+ channels in the ASHs using genetic mutants and
pharmacology. Treatment with 5 µM NemA abolished somal Ca2+ transients, suggesting
that EGL-19 is the only major VGCC in the cell soma (Figure 4-1a). Conversely, 5 µM
NemA was not sufficient to completely knock down axonal Ca2+, and further
experimentation revealed that both EGL-19 and P/Q-type UNC-2 channels were required
in the axons to account for the entire Ca2+ signal (Figure 4-1a). The presence of UNC-2
in the axon is not surprising, as the CaV2 ortholog has been widely implicated in
neurotransmitter release (Estevez et al., 2004; Saheki & Bargmann, 2009; WormBase,
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2015c). In addition to EGL-19 and UNC-2, the C. elegans genome also contains a CaV3
homolog, CCA-1 (Shtonda & Avery, 2005; Steger et al., 2005; WormBase, 2015a).
While we did not test for CCA-1 activity directly, the channel would likely not contribute
significantly to the sensory-evoked Ca2+ signal, as T-type channels pass relatively small
currents that desensitize rapidly before activation of high voltage activated currents
(Cribbs et al., 1998; Tuckwell, 2012; Welling, 2009). That being said, there is still a
possibility for CCA-1 activity to pass a transient amount of Ca2+ at the initiation of
sensory transduction, allowing for an increase in voltage to activate the EGL-19 and
UNC-2 high voltage activated channels. Finally, the ASHs contain two other major
subcellular locales that were not as intensely studied: the amphid and the dendrite. The
amphid is a sensory organ at the animal’s nose where sensory neurons, including the
ASHs, are exposed to the external environment (C. Bargmann & Horvitz, 1991; Hilliard
et al., 2002; Kaplan & Horvitz, 1993). The TRPV1-like OSM-9 channel is thought to be
acting in the amphid compartment of ASHs, though this does not rule out VGCCs being
localized in this compartment as well (Colbert et al., 1997; Tobin et al., 2002). The ASH
dendrites are the long processes that extend from the amphid to the ASH soma, and
respond to NemA and genetic mutation comparably to the soma, suggesting that these
two cellular compartments contain similar VGCC configurations.
VGCCs are a primary driver of Ca2+ entry into cytoplasm, but intracellular stores
receptors are also important for amplification and signaling. To examine the role of the
IP3Rs and RyRs in ASHs, we studied Ca2+ signaling in hypomorphs for each of the C.
elegans internal stores receptors. ITR-1 appears to act in both the soma and the axon,
affecting kinetic waveforms as well as rise times (Figure 4-2). Importantly, itr-1
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hypomorphs exhibited delayed rise times and less desensitization compared to wild-type,
suggesting that ITR-1 acts as more than just a Ca2+ signal amplifier, but also shapes the
kinetic signature of the ASH response. Given the lack of desensitization observed in itr-1
hypomorphs, the receptor is also likely important for proper feedback of Ca2+ dynamics,
suggesting a possible disruption of intracellular signaling (Chapter 5 explores this
concept in greater detail). In contrast, UNC-68 only appears to act in the soma, and
mutation does not significantly affect Ca2+ kinetics (Figure 4-2). One possibility for the
unique subcellular localization of UNC-68 in the soma could be a role for somal peptide
release, where activation of VGCCs is tightly controlled along with RyR activation
resulting in dense core vesicle release (Ludwig et al., 2002; Sabatier, Shibuya, &
Dayanithi, 2004). However, even though ASH is known to produce neuropeptide gene
products, the relationship between intracellular Ca2+ and neuropeptide release remains
unclear (G. Harris et al., 2010). A likely sequence of events is that upon depolarization at
the amphid by OSM-9, EGL-19 channels are activated in the dendrite, soma, and axon,
with additional Ca2+ amplification from ITR-1 and UNC-68. Meanwhile, in the axon,
UNC-2 is also activated, and along with intracellular stores, leads to glutamate release
onto downstream neurons in the reversal circuit.
4.2.2 Sensory-evoked ASH Ca2+ levels are not predictive of behavior
The further probe the function of EGL-19 in ASH Ca2+ signaling, we examined
behavioral sensitivity and Ca2+ amplitudes in animals expressing RNAi transgenes
against egl-19 cell-specifically in ASHs. RNAi knockdown of egl-19 in ASHs resulted in
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decreased 1-octanol-evoked Ca2+ transients, but had no effect on aversive behavior
(Figure 4-3). These data suggest there is not a straightforward linear relationship between
sensory-evoked Ca2+ levels and behavior, and in the case of 5-HT modulation (see
below), inhibition of the Ca2+ signal alone is likely necessary, but not sufficient in and of
itself to stimulate aversive responses.
4.2.3 5-HT decreases sensory-evoked Ca2+ influx but increases sensory-evoked
depolarization via an EGL-30/Gαq signaling pathway in the soma
Monoaminergic modulation of sensory responses is a common regulatory tool for
ensuring an organism behaves appropriately to a given stimulus. In C. elegans, 5-HT acts
in the ASH sensory neurons to stimulate behavioral responses to noxious stimuli such as
1-octanol (Chao et al., 2004; G. P. Harris et al., 2009). We discovered that 5-HT inhibits
sensory-evoked Ca2+ in the soma, but does not significantly reduce evoked signals in the
axon (Figure 4-4). A parsimonious explanation for why axonal Ca2+ is likely not fully
inhibited is that 5-HT modulates Ca2+ signals by directly inhibiting EGL-19. The ASH
axon, containing both EGL-19 and UNC-2, would demonstrate a larger sensory-evoked
Ca2+ response due to UNC-2 continuing to act uninhibited by serotonergic modulation.
Consistently with behavioral data, 5-HT also seems to be acting through the SER-5
GPCR to modulate ASH activity, which acts upstream of EGL-30 to directly modulate
the Ca2+ signal. One critical downstream effector of the Gαq pathway is ITR-1, which is
activated by IP3 produced by PLC (in C. elegans, EGL-8) downstream of Gαq signaling.
Therefore, ITR-1 likely acts as a major Ca2+ modulator in addition to an amplifier, as
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CICR downstream of Gαq signaling could have broader implications on intracellular
signaling (see Chapter 5 for greater detail). Like RyR coupling, Gαq signaling also
facilitates neuropeptide release in other systems, so another possible function of the
somal Ca2+ signal is to trigger release of neuropeptides to further modulate downstream
responses to aversive behavior (Bergquist & Ludwig, 2008; Ludwig et al., 2002).
Serotonergic modulation of ASH-mediated behavioral responses and Ca2+ signals
also presented an interesting conundrum, where behavioral responses suggested that 5HT would increase sensory-evoked depolarization, but Ca2+ transients predicted
decreased depolarization. Interestingly, 5-HT treatment increased sensory-evoked
depolarization, consistent with behavioral data (Figure 4-5). These data show that the
amplitude of ASH somal Ca2+ transients need not accurately reflect depolarization
strength. What then, does the decreased Ca2+ signal represent, if it doesn’t accurately
reflect the neuronal depolarization status? While the ASH Ca2+ transient is complex and
involving multiple inputs, there are a few possibilities. The most likely possibility is the
existence of a Ca2+-driven negative feedback loop explaining the apparent disconnect
between increased depolarization and decreased Ca2+ transient amplitudes. Upon
sensation of 1-octanol, the stimulus binds its olfactory receptor, leading to the activation
of the Gαolf ODR-3. Next, ODR-3 activates OSM-9, depolarizing the neuron, and leading
to activation of VGCCs and Ca2+ influx (Roayaie et al., 1998). In the soma and axon, the
combination of VGCC and internal stores activation via CICR could pass a threshold to
activate an inhibitory conductance, such as a Ca2+-gated K+ channels or Ca2+-activated
Cl- channels, to temper neuronal excitability. Indeed, C. elegans expresses at least one
Ca2+-gated K+ channel (SLO-1), and multiple genes of Ca2+-activated Cl- channels: two
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anoctamin orthologs (ANOH-1, ANOH-2), and at least 24 bestrophin orthologs (BEST-1
to BEST-24) (Hobert, 2013; Hongkyun Kim et al., 2009; Y. Wang et al., 2013).
Additionally, the Ca2+ could then be downregulated by some form of CDI, previously
described in mammals, which is studied in great detail in Chapter 5 (Budde et al., 2002;
Oliveria et al., 2012; Olson et al., 2005).
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Chapter 5
Serotonin negatively regulates C. elegans L-VGCCs via
a calcium-dependent negative feedback loop
5.1 Results
5.1.1 5-HT modulates ASH Ca2+ signals downstream of sensory transduction
Sensory modulation is a complex process, with multiple regulatory steps to ensure
an animal senses a stimulus appropriately. In Chapters 3 and 4, we previously isolated 5HT as a major regulator of ASH Ca2+ signaling and depolarization in the cell soma. To
assess at what point 5-HT was modulating Ca2+ signaling in ASHs, we performed Ca2+
imaging experiments on dissected ASHs in response to electrophysiological external
solution containing increasing concentrations of K+. Our extracellular K+ concentration is
5 mM under standard conditions, but increasing this amount can reverse the
electrochemical gradient for the ion, resulting in neuronal depolarization bypassing
sensory transduction. Treatment with 15 mM, 30 mM, and 150 mM K+-containing
external solution all successfully evoked increases in ASH somal Ca2+ levels, but not in a
linear fashion (Figure 5-1). To probe the role of the L-type channel in depolarizationevoked Ca2+ transients, we targeted the EGL-19 channel using the specific antagonist
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NemA. Importantly, depolarization-evoked ASH Ca2+ transients were significantly
decreased in the presence of NemA, suggesting that EGL-19 is a major charge carrier in
our artificial depolarization setup (Figure 5-1). Next, we wanted to examine where 5-HT
acted in the ASHs to modulate the Ca2+ signal. If 5-HT acted at the level of the sensory
transduction, 5-HT treatment would demonstrate no effect on depolarization-evoked Ca2+
transients; if 5-HT acted on a downstream regulator of signal transduction, the signal
should be knocked down. Since EGL-19 is a major charge carrier in these experiments
(see above) and we know that EGL-19 is the major VGCC acting in the cell soma (see
Chapter 4), 5-HT-mediated knockdown of the Ca2+ signal would directly target
serotonergic regulation to the L-type channel or another associated protein (e.g. ITR-1).
To examine if 5-HT acts downstream of sensory transduction to modulate depolarizationevoked Ca2+ levels, animals were incubated on 4 mM 5-HT for 30 min, then assayed.
Interestingly, 5-HT incubation caused knockdown of depolarization-evoked Ca2+
transients at 30 mM, but not at 150 mM (Figure 5-1). Taken together, these data implicate
a direct role for 5-HT in negative regulation of ASH Ca2+ signaling downstream of
sensory transduction, but only at low levels of depolarization.
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B
Figure 5-1 ASH sensory neurons are modulated downstream of sensory transduction by
5-HT. (A) A representative trace of ASH exposure to 30 mM K+. (B) Quantification of
ASH Ca2+ responses to electrophysiological external solutions containing increasing
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concentrations of K+, in the presence and absence of 4 mM 5-HT or 5 µM NemA. Bars
represent mean ± SEM. *: P < 0.05 compared to 30 mM untreated. ‡: P < 0.05 compared
to 150 mM untreated. n ≥ 5 for each group.
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5.1.2 5-HT inhibits ASH Ca2+ transients via SER-5 and EGL-30/Gαq
Previous behavioral experiments have demonstrated a role for SER-5 in the
modulation of ASH-mediated aversive behavior (G. P. Harris et al., 2009). In Chapter 4,
we demonstrate the involvement of SER-5 and Gαq subunit EGL-30 as key modulators of
the ASH Ca2+ response. To examine if SER-5 is the receptor necessary for 5-HT
modulation downstream of sensory transduction, we performed depolarization-evoked
Ca2+ imaging experiments using 30 mM K+ as a stimulus in ser-5(tm2654) receptor null
mutants as well as mutants for two downstream effectors: egl-30(n686sd) and itr1(sa73)(Figure 5-2). Consistently with 1-octanol experiments, 4 mM 5-HT was unable to
significantly decrease K+-evoked Ca2+ transients in ser-5, egl-30, or itr-1 mutant
backgrounds (Figure 5-2). Once again, itr-1(sa73) animals demonstrate diminished Ca2+
transients in the absence of 5-HT, indicative of the IP3R Ca2+ acting as a major source of
somal Ca2+ (see Chapter 4)((Zahratka et al., 2015). These data suggest 5-HT modulates
ASH Ca2+ signaling via a Gαq signaling pathway which acts downstream of sensory
transduction to modulate sensory signaling.
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Figure 5-2 5-HT inhibits ASH Ca2+ downstream of sensory transduction via Gαq.
Quantification of ASH sensory responses to electrophysiological external solution
containing 30 mM K+ in ser-5(tm2654), egl-30(n686sd), and itr-1(sa73) mutants in the
presence and absence of 4 mM 5-HT. Bars represent mean ± SEM. *: P < 0.05 compared
to wild-type. n ≥ 3 for each group.
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5.1.3 CDI may be acting in the ASH nociceptive circuit to modulate sensory
responses
As SER-5 is likely acting through a Gαq signaling pathway in order to modulate
ASH Ca2+ responses and behavior, we wanted to find a potential mechanism for how Gαq
signaling could shut down Ca2+ signaling. One potential mechanism is CDI, where Ca2+
influx from EGL-19 and ITR-1 could be enough to activate CaN, a Ca2+-dependent
phosphatase, which dephosphorylates a conserved serine residue on the L-type channel’s
C-terminus leading to inactivation (Budde et al., 2002; Oliveria et al., 2012). CDI
requires five major components: the L-type channel, an IP3R, CaN, and two scaffolds,
Shank and an AKAP (Marshall et al., 2011; Oliveria et al., 2012; Zhang et al.,
2005)(Figure 5-3a). Intriguingly, C. elegans have orthologs for each of these genes: egl19 is the L-type channel, itr-1 is the IP3R, tax-6 is CaN, shn-1 is Shank, and aka-1 is an
AKAP (Baylis et al., 1999; Jee et al., 2004; Maëlle Jospin et al., 2002; Kuhara, Inada,
Katsura, & Mori, 2002; Oh et al., 2011; Pastok et al., 2013). To examine the role of these
putative CDI components in ASH-mediated behaviors, we performed dilute 1-octanol
behavioral assays with and without 5-HT. In contrast with wild-type animals, which
exhibited 5-HT-mediated stimulation and 5-second response times, mutants for itr-1, tax6, shn-1, and aka-1 exhibited 10-second response times on 5-HT, suggesting that 5-HT
regulation is disrupted in each of these genotypes (Figure 5-3b). Notably, egl-19(n582)
hypomorphs contain movement defects making them unsuitable for behavioral
experiments, so instead we used cell-specific RNAi knockdown of the channel in ASHs
(Psra-6::egl-19RNAi), which also elicited 10-second response times on 5-HT (Figure 591
3b)(R. Y. N. Lee et al., 1997). In summary, these results indicate that each of the putative
CDI pathway members are acting throughout the ASH avoidance circuit to regulate
serotonergic effects on avoidance behaviors.
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Figure 5-3 A canonical CDI pathway may be acting in the ASH nociceptive circuit
downstream of 5-HT. (A) Diagram of canonical CDI pathway, including mammalian and
C. elegans genes of interest. Note that C. elegans does not contain a Homer ortholog. (B)
5-HT is unable to stimulate behavioral responses in itr-1(sa73), tax-6(p675), shn1(ok1241), aka-1(ok2520) mutants, nor in cell-specific Psra-6::egl-19RNAi transgene93
expressing animals. Bars represent mean ± SEM. *: P < 0.05 compared to wild-type
untreated. n ≥ 50 for each group, analyzed over at least two experimental days.
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5.1.4 The canonical CDI pathway is not present in ASH sensory neurons
Whole-animal mutations are useful for knowing if a given gene of interest is
involved in a signaling pathway, but are not as useful for localization. To see if each of
the CDI components were acting cell-specifically in the ASH sensory neurons, we
examined mutants for shn-1, tax-6, and aka-1 in our saturating 1-octanol Ca2+ imaging
setup (see Figure 4-4 for itr-1). Notably, shn-1, tax-6, and aka-1 mutants all had
significantly decreased Ca2+ levels in the untreated condition, demonstrating that each of
these proteins have a significant role in setting baseline sensory-evoked Ca2+ signals in
ASHs (Figure 5-4). As expected, shn-1 and tax-6 mutant animals were immune to 5-HTmediated knockdown of the Ca2+ signal, suggesting that each of these gene products were
acting specifically in ASH to modulate serotonergic regulation of aversive responses.
Curiously, aka-1 mutant animals retained 5-HT-mediated Ca2+ inhibition, implying this
gene is not likely involved in serotonergic regulation in ASHs. Together, these data
suggest a role for tax-6 in modulating the ASH Ca2+ signal via 5-HT, but not via a
canonical CDI pathway.
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Figure 5-4 A canonical CDI pathway is not present in the ASHs to mediate aversive
responses. Quantification of ASH Ca2+ responses to saturating 1-octanol solution (2.4
µM) in shn-1(ok1241), tax-6(p675), or aka-1(ok2520) mutants ± 4 mM 5-HT. Bars
represent mean ± SEM. *: P < 0.05 compared to wild-type untreated. ‡: P < 0.05
compared to aka-1(ok2520) untreated. n ≥ 5 for each group.
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5.1.5 5-HT activates TAX-6/CaN via SER-5 and EGL-30/Gαq to inhibit EGL-19
Despite our finding that a traditional CDI pathway was not acting in the ASHs to
modulate aversive behavior, we nonetheless discovered a role for TAX-6/CaN in ASH
sensory regulation. To examine if TAX-6 acts downstream of the sensory transduction
apparatus, we performed Ca2+ imaging experiments in response to 30 mM K+ as
previously described in tax-6(p675) reduced function mutants. Consistently with other
data, tax-6(p675) mutants demonstrated 30 mM K+-evoked Ca2+ signals, and these
responses were not inhibited by 4 mM 5-HT (Figure 5-5). These data, along with the data
previously presented in Figure 5-2, present a scenario where SER-5 acting through EGL30 eventually leads to the recruitment of TAX-6 to EGL-19, which shuts the channel
down and inhibits the evoked Ca2+ signal.
97
Figure 5-5 The C. elegans TAX-6 phosphatase is downstream of 5-HT activity in ASHs.
Quantification of ASH sensory responses to electrophysiological external solution
containing 30 mM K+ in tax-6(p675) mutants in the presence and absence of 4 mM 5-HT.
Bars represent mean ± SEM. *: P < 0.05 compared to wild-type untreated. n ≥ 5 for each
group.
98
5.2 Discussion
5.2.1 High concentrations of extracellular K+ can depolarize ASHs bypassing
sensory transduction
The ASH sensory neurons have tightly compartmentalized Ca2+ signaling, with
the EGL-19 VGCC and internal stores receptors ITR-1 and UNC-68 acting to shape the
somal Ca2+ signal. In the previous chapter, we established a case where 5-HT acts to
inhibit sensory-evoked Ca2+ signals but increases neuronal depolarization. To probe
where 5-HT was acting in ASHs, we established a protocol for depolarizing ASHs
downstream of sensory transduction. By changing the extracellular concentration of K+,
we were able to depolarize dissected ASHs bypassing the activation of the OSM-9
signaling apparatus. Interestingly, increasing extracellular K+ did not increase the evoked
Ca2+ signal in a linear fashion, suggesting there may be a ceiling to the amount of Ca2+ a
given stimulus can provoke (Figure 5-1). However, NemA was able to depress somal K+evoked Ca2+ transients at concentrations of both 30 mM and 150 mM, once again
demonstrating that EGL-19 is the primary driver of somal Ca2+ (Figure 5-1).
Surprisingly, 5-HT only knocked down K+-evoked Ca2+ transients at 30 mM, but not at
150 mM (Figure 5-1). One potential explanation for these data is that 5-HT modulation
only acts on weaker stimuli, and that beyond a certain threshold the 5-HT response is
inhibited. This is consistent with ASH behavioral data, as responses to 100% 1-octanol
are not affected by 5-HT and the animal will respond in about 3 seconds with or without
its presence (Chao et al., 2004; G. P. Harris et al., 2009). Therefore, perhaps 30 mM K+ is
99
a similar stimulation level to 30% 1-octanol behaviorally and saturating 1-octanol in Ca2+
imaging, as each of these modes are modulable by 5-HT (see Chapters 3 and 4). Since
TA and OA can also modulate responses to 100% 1-octanol by facilitating neuropeptide
release and inhibit 5-HT modulation of dilute 1-octanol, each of these monoamines may
be promising targets for future study of ASH responses at higher concentrations of
extracellular K+ (Chapter 3)(Hapiak et al., 2013; Mills et al., 2011; Wragg et al., 2007).
5.2.2 5-HT acts through EGL-30/Gαq to directly modulate Ca2+ signaling
In the previous chapter, we implicate the SER-5 GPCR acting through Gαq as the
source of serotonergic regulation of ASH Ca2+ responses. To test where 5-HT was acting
in ASHs to modulate Ca2+, we examined ASH Ca2+ responses to 30 mM K+ in ser-5, egl30, and itr-1 mutant animals. Consistently with our 1-octanol behavioral data, each of
these mutants exhibited high levels of evoked Ca2+ in the presence of 5-HT, suggesting
each of these genes act in the serotonergic pathway to modulate behavior (Figure 5-2).
These data illuminate two important points: First, that 5-HT is acting downstream of the
sensory transduction apparatus, implying that it does not directly modulate OSM-9 (or
any other part of the sensory transduction pathway); and second, that 5-HT is likely
modulating one of the Ca2+ channels directly (likely EGL-19 or ITR-1), or modulating a
protein that directly regulates the Ca2+ channels. Unfortunately, these data are still
probing the question indirectly, and do not give direct evidence for a linkage between 5HT and its target. Furthermore, each of the genes we have studied in this assay are wholeanimal knockouts, bringing forth the possibility of off-target effects or developmental
100
compensation. One possible clue in the direct effector of 5-HT modulation is the
experiment performed in Figure 5-5, where the C. elegans CaN subunit TAX-6 also
demonstrates high levels of sensory-evoked Ca2+ in response to 30 mM K+. The
involvement of TAX-6 heavily implicates EGL-19 as 5-HT’s direct target, as the L-type
channel has a conserved serine residue that is likely a target for the phosphatase (Figure
5-3). However, as previously stated, these experiments involve using whole-animal
knockouts and are indirect. A potential way forward from this conundrum is to directly
mutagenize likely regulatory sites on each of the channel proteins, including the
conserved serine, perhaps using some of the promising technologies such as
CRISPR/Cas9-mediated homologous recombination (Frøkjær-Jensen, 2013; Heesun Kim,
Ishidate, Ghanta, & Seth, 2014; Zhao, Zhang, Ke, Yue, & Xue, 2014).
5.2.3 The classical CDI pathway is not acting in ASHs, but each of the proteins are
still important for the basal evoked Ca2+ signal and behavior
Ca2+-dependent inactivation is a critical regulatory process described in the
mammalian heart and brain, where Ca2+ acts as a signaling molecule to initiate a negative
feedback loop to prevent further increases in Ca2+ levels (Blaich et al., 2012; Budde et al.,
2002; Oliveria et al., 2012; Olson et al., 2005; Poomvanicha et al., 2011). There are five
primary components to the CDI complex, and each of them have a C. elegans ortholog:
an L-type channel (EGL-19), an IP3R (ITR-1), CaN (TAX-6), and two scaffold proteins
(SHN-1 and AKA-1). To see if CDI was acting in ASH to modulate evoked Ca2+ levels
in response to 5-HT, we performed a screen using behavioral responses to 30% 1-octanol
101
in whole-animal CDI gene hypomorphic and null mutants (Figure 5-3b). Notably, each of
the C. elegans CDI complex ortholog genes, itr-1, shn-1, tax-6, and aka-1 were not
stimulated by 5-HT, suggesting that each of these genes are involved in serotonergic
regulation of behavioral responses at some point in the 1-octanol avoidance circuit
(Figure 5-3b). However, as each of these mutants were whole-animal hypomorphs or
nulls, we also performed 1-octanol-evoked Ca2+ imaging experiments to confirm cell
specificity (Figure 5-4). Interestingly, even though shn-1(ok1241) and tax-6(p675)
mutants failed to have 5-HT-mediated inhibition of Ca2+ signals, aka-1(ok2520) mutants
retained 5-HT inhibition, suggesting that this gene product is not directly involved in the
5-HT regulatory pathway (Figure 5-4). These results suggest one of two possible
outcomes: first, that CDI is not occurring in the ASHs to modulate EGL-19 through the
complex seen in mammals. Second, although AKA-1 is not necessary for 5-HT
inhibition, another related protein could be taking its place as a key scaffold for CDI.
However, there are no other predicted AKAP-like proteins in C. elegans, suggesting the
second outcome to be less likely. Additionally, AKA-1 is divergent from mammalian
AKAPs, showing a greater affinity for RI regulatory subunits instead of the RII subunit
binding partners of mammalian AKAP5 (Angelo & Rubin, 2000; Pastok et al., 2013).
Therefore, it is likely that C. elegans use a divergent mechanism for channel inactivation
that differs from mammals, but results in the same end point of L-type channel
dephosphorylation mediated by CaN.
Although CDI as we know it in mammals is unlikely to be occurring in the ASHs,
our data illustrate that individually knocking down each of the proteins involved
dramatically decreases sensory-evoked Ca2+ signals even without 5-HT treatment
102
(Figures 4-4, and 5-4). These results indicate that each of the proteins are critical for
basal evoked Ca2+ levels, even in the absence of modulation. While reduced function
EGL-19 and ITR-1 channels would intuitively decrease evoked Ca2+ signals due to
reduced Ca2+ entry into the cytoplasm, the effects of the scaffold proteins and TAX-6 are
less clear. SHN-1 is likely tethering EGL-19 and ITR-1 together, so one possibility for
decreased Ca2+ levels would be due to mislocalization of these two channels to one
another, resulting in an inability for EGL-19 Ca2+ to activate ITR-1 (Jee et al., 2004; Oh
et al., 2011). Likewise, AKA-1 could be playing a role in proper trafficking of PKA to its
cellular targets, and the lack of this protein could separate the kinase from potential
partners involved in Ca2+ release or generation of aversive behavior (Angelo & Rubin,
2000; Pastok et al., 2013). Indeed, one possibility is that PKA can phosphorylate L-type
VGCCs to increase their activity, a phenomenon known as Ca2+-dependent facilitation
(Bers & Morotti, 2014; Christel & Lee, 2012; Lian et al., 2007; Tang et al., 2012).
Finally, TAX-6 has multiple diverse targets throughout the animal’s body, so a reduced
function allele could easily have wide-sweeping effects on Ca2+ homeostasis, intracellular
signaling, and gene transcription (Crabtree, 2001; Kuhara et al., 2002; Xu et al., 2010).
Interestingly, one potential CaN-mediated transcriptional target in mammals is IP3R1, the
mammalian IP3R, suggesting a Ca2+-dependent signaling pathway could feed back onto
itself at a transcriptional level (Crabtree, 2001). In the future, single cell knockouts
(through RNAi or CRISPR) or protein-specific inhibitors (cyclosporin A, tacrolimus) can
be used to probe the role of these proteins in intracellular Ca2+ signaling (Crabtree, 2001;
Oliveria et al., 2012).
103
5.2.4 A model for serotonergic modulation of EGL-19 by SER-5, EGL-30, and TAX6
In this chapter, we have described genetic evidence for a Ca2+-dependent negative
feedback loop acting in the ASH sensory neurons to modulate aversive behavior. 5-HT,
acting through the SER-5, activates the Gαq subunit EGL-30. EGL-30 then leads to the
cleavage of PIP2, resulting in IP3 production to activate ITR-1. Endoplasmic reticulumstored Ca2+, combined with cytoplasmic Ca2+ carried by the EGL-19 VGCC, reaches a
threshold to activate TAX-6, which feeds back onto EGL-19, leading to
dephosphorylation and inactivation (Figure 5-6). What, then, is the role of ASH somal
Ca2+? In Chapter 4, we established one hypothesis that somal Ca2+ is acting to recruit a
Ca2+-activated K+ channel, such as SLO-1, to modulate cell excitability and maintenance
of behavioral circuits. Similarly, we have elucidated a pathway for the maintenance of
ASH intracellular Ca2+ after response to a noxious challenge. However, these
explanations do not explain the significance of the somal Ca2+ signal. We can, however,
speculate on the function of somal Ca2+ with clues from other work performed in C.
elegans and other systems. For example, 5-HT is already known to be necessary for the
release of neuropeptides encoded by the nlp-3 gene expressed in ASHs (G. Harris et al.,
2010). It is possible that ASH somal Ca2+ release is leading directly to the release of
NLP-3 peptides in the short term (i.e. before TAX-6-mediated downregulation),
stimulating behavioral responses. This scenario is not unlike vasopressin and oxytocin
release from magnocellular neurons in the mammalian supraoptic nucleus, which trigger
peptide release from the soma and dendrites of peptidergic neurons in response to PLC
104
activation downstream of Gαq (Bergquist & Ludwig, 2008; Ludwig et al., 2002; Sabatier
et al., 2004). A possible time course of action is upon sensory stimulation, 5-HT binds
SER-5, resulting in NLP-3 release in the short term to stimulate aversive responses. Over
time, increases in Ca2+ trigger TAX-6 activation, shutting down EGL-19 to prevent
aberrant peptide release. Additionally, EGL-19 and TAX-6 could also be working at a
transcriptional level, as each of these proteins demonstrate a role in excitationtranscription coupling (Brittain et al., 2012; Christel & Lee, 2012; Crabtree, 2001;
Marshall et al., 2011; Simms et al., 2013; Tuckwell, 2012; Wheeler et al., 2008). L-type
channels can result in downstream CREB activation in mammals, allowing Ca2+ to
directly modulate gene transcription (Brittain et al., 2012; Morad & Soldatov, 2005;
Simms et al., 2013; Weick et al., 2003; Wheeler et al., 2008). In C. elegans, a CREB
transcriptional coactivator, CRTC-1, can also act downstream of TAX-6 to regulate
transcription and lifespan (Burkewitz et al., 2015; Mair et al., 2011). Preliminary
characterization of crtc-1 and the C. elegans CREB ortholog crh-1 in 1-octanol
behavioral assays exhibit delayed responses on 5-HT, suggesting each of these proteins
may be involved in the regulation of aversive responses somewhere in the ASH sensory
circuit, but the whole-animal nature of the mutations cannot rule out developmental
compensation or off-target effects (data not shown). At any rate, further work will be
necessary to dissect the meaning of somal Ca2+ in the context of ASH aversive responses.
105
Figure 5-6 Model of a hypothetical Ca2+-driven negative feedback loop controlling ASH
Ca2+ influx and excitability. The activation of ASHs by a noxious stimulus will
depolarize the neuron (ΔV), which spreads to the soma, dendrite, and axon, leading to
Ca2+ entry through VGCCs including EGL-19. Meanwhile, 5-HT released from
neighboring neurons will bind SER-5, initiating a Gαq signaling cascade through EGL30. The IP3R ITR-1 activates downstream of EGL-30, which allows for even more Ca2+
influx into the cytoplasm. The ITR-1 and EGL-19 Ca2+ pools together are likely enough
to recruit the activation of TAX-6, which likely dephosphorylates a key serine residue on
the EGL-19 C-terminus to inactivate the channel and halt Ca2+ influx.
106
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