of 13
ARTICLE
Identi
fi
cation of peripheral neural circuits that
regulate heart rate using optogenetic and viral
vector strategies
Pradeep S. Rajendran
1
, Rosemary C. Challis
2
, Charless C. Fowlkes
3
, Peter Hanna
1
, John D. Tompkins
1
,
Maria C. Jordan
1
, Sarah Hiyari
1
, Beth A. Gabris-Weber
4
, Alon Greenbaum
2
, Ken Y. Chan
2,6
,
Benjamin E. Deverman
2,6
, Heike Münzberg
5
, Jeffrey L. Ardell
1
, Guy Salama
4
, Viviana Gradinaru
2
&
Kalyanam Shivkumar
1
Heart rate is under the precise control of the autonomic nervous system. However, the wiring
of peripheral neural circuits that regulate heart rate is poorly understood. Here, we develop a
clearing-imaging-analysis pipeline to visualize innervation of intact hearts in 3D and
employed a multi-technique approach to map parasympathetic and sympathetic neural cir-
cuits that control heart rate in mice. We identify cholinergic neurons and noradrenergic
neurons in an intrinsic cardiac ganglion and the stellate ganglia, respectively, that project to
the sinoatrial node. We also report that the heart rate response to optogenetic versus
electrical stimulation of the vagus nerve displays different temporal characteristics and that
vagal afferents enhance parasympathetic and reduce sympathetic tone to the heart via
central mechanisms. Our
fi
ndings provide new insights into neural regulation of heart rate,
and our methodology to study cardiac circuits can be readily used to interrogate neural
control of other visceral organs.
https://doi.org/10.1038/s41467-019-09770-1
OPEN
1
Cardiac Arrhythmia Center and Neurocardiology Research Program of Excellence, David Geffen School of Medicine, University of California - Los Ange
les
(UCLA), Los Angeles, CA 90095, USA.
2
Division of Biology and Biological Engineering, California Institute of Technology, Pasadena, CA 91125, USA.
3
Department of Computer Science, University of California - Irvine, Irvine, CA 92697, USA.
4
Department of Cell Biology, University of Pittsburgh, Pittsburgh,
PA 15261, USA.
5
Neurobiology of Nutrition and Metabolism Department, Louisiana State University, Baton Rouge, LA 70808, USA.
6
Present address: Stanley
Center for Psychiatric Research, Broad Institute, Massachusetts Institute of Technology, Cambridge, MA 02142, USA. Correspondence and requests f
or
materials should be addressed to V.G. (email:
viviana@caltech.edu
) or to K.S. (email:
kshivkumar@mednet.ucla.edu
)
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1
1234567890():,;
S
ituation-dependent changes in heart rate are essential for
survival and are under the precise control of the autonomic
nervous system (ANS)
1
. Heart rate reduction during sleep
2
and elevation during exercise
3
result from changes in para-
sympathetic and sympathetic tone. In fact, heart rate variability
has been utilized extensively as an index of ANS function
4
6
.
Although it is well known that the parasympathetic and sympa-
thetic nervous systems innervate the sinoatrial (SA) node
7
,
8
and
regulate heart rate
9
13
, the wiring of these neural circuits in the
periphery is not well characterized. Anatomical and functional
maps of these fundamental cardiac circuits are needed to
understand physiology, characterize remodeling in disease (e.g.,
sick sinus syndrome
14
), and develop novel therapeutics. However,
these efforts have been hindered by a shortage of tools that target
the peripheral nervous system (PNS) with speci
fi
city and
precision.
Cardiac circuit anatomy has traditionally been studied using thin
sections
15
and whole-mount preparations
16
. However, these
methods do not preserve the structure of intact circuits and only
provide 2D information. In contrast, tissue clearing methods render
tissues optically transparent while preserving their molecular and
cellular architecture and can be combined with a variety of labeling
strategies to enable 3D visualization of intact circuits
17
,
18
.Totrace
cardiac circuits, dyes and proteins have historically been used.
However, achieving cell type-speci
fi
city and/or sparse labeling
needed for singe-cell tracing and delineating circuit connectivity is
dif
fi
cult or not possible with these methods
19
. Additionally, many
peripheral neuronal populations such as intrinsic cardiac ganglia are
challenging to access surgically for tracer delivery. Adeno-associated
viruses (AAVs) can address these limitations since they can be used
to express genes of interest (e.g.,
fl
uorescent proteins) in de
fi
ned cell
populations in wild-type and transgenic animals
20
22
. In addition,
intersectional strategies can be used to titrate gene expression to
achieve sparse labeling
21
. AAVs can also be delivered systemically
to target dif
fi
cult-to-reach populations
20
22
.
Functional mapping of cardiac circuits has relied on electrical
or pharmacological manipulation of the ANS with simultaneous
physiological measurements
23
,
24
. However, each of these methods
has disadvantages. Electrical techniques lack spatial precision and
speci
fi
city. Autonomic nerves such as the vagus contain motor
and sensory
fi
bers
25
, and electrical stimulation typically activates
both
fi
ber types
26
,
27
as well as surrounding tissues
28
. Pharma-
cological techniques exhibit improved selectivity but lack
temporal resolution. In contrast, optogenetics, which uses
light-sensitive ion channels (e.g., channelrhodopsin-2 (ChR2),
halorhodopsin), enables precise spatiotemporal control of de
fi
ned
cell populations.
Here, we develop a clearing-imaging-analysis pipeline to
visualize innervation of whole hearts in 3D and employ a multi-
technique approach, which includes AAV-based sparse labeling
and tracing, retrograde neuronal tracing with cholera toxin sub-
unit B (CTB), and optogenetics with simultaneous physiological
measurements, to map peripheral parasympathetic and sympa-
thetic neural circuits that regulate heart rate in mice.
Results
Tissue clearing and computational pipeline to assess cardiac
innervation
. To characterize global innervation of the mouse
heart in 3D, we developed a clearing-imaging-analysis pipeline
(Fig.
1
a). We stained whole hearts with an antibody against the
pan-neuronal marker protein gene product 9.5 (PGP9.5) and
rendered them optically transparent using an immunolabeling-
enabled three-Dimensional Imaging of Solvent-Cleared Organs
(iDISCO) protocol (Fig.
1
b)
29
. We used confocal microscopy to
image large tissue volumes (Fig.
1
c and Supplementary Movie 1)
and both confocal and lightsheet microscopy to image entire
hearts (Supplementary Movie 2)
30
,
31
. We observed cardiac
ganglia surrounding the pulmonary veins and a dense network of
nerve
fi
bers coursing through the atrial and ventricular
myocardium (Fig.
1
c). In contrast to whole-mount stained hearts,
innervation was seen throughout the entire thickness of the
myocardium in iDISCO-cleared hearts, with large-diameter nerve
fi
ber bundles located near the epicardium and smaller
fi
ber
bundles in the mid-myocardium and endocardium (Fig.
1
c,
Supplementary Fig. 1, and Supplementary Movie 3). To analyze
these data, we created a semiautomated computational pipeline to
detect nerve
fi
bers over large tiled volumes and to measure
microanatomical features of
fi
bers such as diameter and orien-
tation (Fig.
1
d and e). Large-diameter nerve
fi
ber bundles typi-
cally entered near the base of the dorsal heart. These bundles
coursed perpendicular to the atrioventricular (AV) groove and
branched into smaller
fi
ber bundles as they progressed towards
the apex. These data from healthy hearts will be important for
future characterization of neural remodeling in cardiovascular
diseases such as myocardial infarction (MI) in which innervation
patterns are disrupted and nerve sprouting occurs
32
,
33
.
While the iDISCO protocol was speci
fi
cally developed for
immunostaining applications, other clearing techniques such as
the PAssive Clarity Technique (PACT) are better suited for
visualizing endogenous
fl
uorescence in large tissue volumes or
whole organs
30
,
34
. We demonstrate that PACT preserves the
fl
uorescence of virally labeled cholinergic and endogenously labeled
noradrenergic neurons and nerve
fi
bers in the heart (Supplementary
Fig. 2 and Supplementary Movies 4
6). Therefore, the application
should dictate the choice of clearing technique.
AAV-based labeling and tracing of cholinergic neurons on the
heart
. After visualizing global cardiac innervation, we assessed
whether we could identify a subset of cholinergic neurons that form
an anatomical circuit with the SA node to potentially regulate heart
rate. We used an AAV-based system to trace
fi
bers, presumably
from cholinergic neurons in intrinsic cardiac ganglia. Multicolor
labeling strategies allow individual cells to be distinguished from
one another (Fig.
2
a) and sparse labeling reduces the fraction of
labeled cells to allow individual
fi
bers to be visualized (Fig.
2
b)
21
.To
demonstrate this, we systemically co-administered Cre-dependent
vectors expressing
fl
uorescent proteins (XFPs) from the
tetracycline-responsive element (TRE)-containing promoter at a
high dose and the tetracycline transactivator (tTA) from the human
synapsin I promoter (hSyn1) at a lower dose in ChAT-IRES-Cre
transgenic mice (Fig.
2
b)
21
. Compared to dense multicolor labeling
(Fig.
2
a), sparse multicolor labeling resulted in a labeling density in
intrinsic cardiac ganglia that was lower and that would more easily
allow for tracing (Fig.
2
b)
21
. To trace cholinergic
fi
ber, we utilized
sparse single-color labeling with tdTomato. Three weeks after viral
delivery, hearts were collected and stained with an antibody for
hyperpolarization-activated cyclic nucleotide-gated potassium
channel 4 (HCN4). HCN4 staining along with anatomical land-
marks were used to identify the SA node, the AV node, and the
conduction system
35
,
36
. We observed cholinergic
fi
bers, presumably
from cardiac ganglia, coursing along the SA node, AV node, and
ventricles (Fig.
2
c), identifying cholinergic neurons that are poten-
tially involved in chronotropic, dromotropic, and ventricular con-
trol, respectively.
Optogenetic stimulation of cholinergic neurons in the inferior
pulmonary vein-ganglionated plexus
. Next, to functionally assess
whether cholinergic neurons in the inferior pulmonary vein-
ganglionated plexus (IPV-GP) regulate heart rate and AV con-
duction, we used an optogenetic approach. We expressed ChR2 in
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cholinergic neurons by crossing transgenic ChAT-IRES-Cre mice
with reporter mice containing a Cre-dependent ChR2-enhanced
yellow
fl
uorescent protein allele (ChR2-eYFP; offspring from this
cross are subsequently referred to as ChAT-ChR2-eYFP mice).
ChR2-eYFP expression in intrinsic cardiac neurons was con
fi
rmed
by staining hearts for GFP (to amplify ChR2-eYFP detection) and
PGP9.5 (Fig.
3
a). All GFP
+
neurons were PGP9.5
+
(100.0 ± 0.0%)
and the majority of PGP9.5
+
neurons were GFP
+
(96.4 ± 1.2%)
(Fig.
3
b). GFP staining was also present in PGP9.5
+
nerve
fi
bers in
the atria and ventricles (Fig.
3
a). These data are consistent with
previous studies reporting that the majority of intrinsic cardiac
neurons are cholinergic
37
and that the ventricles as well as atria
receive cholinergic innervation
37
39
.
After verifying ChR2 expression, we next assessed whether
selective stimulation of cholinergic neurons in the IPV-GP
modulated heart rate and AV nodal conduction using optoge-
netics in ex vivo Langendorff-perfused hearts. A blue laser-
coupled optical
fi
ber was positioned for focal illumination of the
IPV-GP while cardiac electrical activity was simultaneously
recorded (Fig.
3
c, d). Optogenetic stimulation resulted in a
decrease in heart rate that was dependent on light pulse power,
frequency, and pulse width (Fig.
3
e
g and Supplementary Table 1)
but did not change the AV interval (35.7 ± 1.2 ms before
stimulation versus 35.9 ± 1.0 ms during stimulation) (Fig.
3
h).
The lack of change in AV nodal conduction suggests that
fi
bers
from this ganglion may pass through the AV node without
synapsing. In addition, stimulation prolonged the P wave
duration (9.3 ± 1.0 ms before stimulation versus 12.3 ± 2.4 ms
during stimulation) and caused P wave fractionation (Fig.
3
i, j). P
wave fractionation has been reported in humans following
administration of adenosine
40
, which mimics the effects of
acetylcholine released from cholinergic nerve terminals
41
,
42
.
During stimulations at higher frequencies, we occasionally
observed ectopic atrial rhythms (
n
=
3/6 mice) and even asystole
(
n
=
1/6 mice) (Fig.
3
d), demonstrating the profound effect of the
IPV-GP on the SA node and atrial function. The response to
stimulation was abolished by administration of the muscarinic
receptor antagonist atropine (
33.5 ± 11.0% with stimulation
before atropine versus
0.3 ± 0.2% with stimulation after
atropine) (Fig.
3
k and Supplementary Table 1), indicating that
the bradycardic response was indeed mediated by selective
stimulation of cholinergic neurons.
Since ChR2 is expressed in both preganglionic cholinergic
inputs to and postganglionic cholinergic neurons in the IPV-GP
Imaging using
confocal or lightsheet
microscopy
Automated nerve fiber
detection and 3D visualization
Derivation of microanatomical
data on nerve fibers
Perfusion and
fixation
Staining
Clearing
ab
After
Before
iDISCO
c
PGP9.5
Ventral
Dorsal
1
2
Cardiac ganglion
1
Ventricle
2
Nerve diameter
d
Diameter (
μ
m)
0
30
Nerve orientation
e
Orientation
x
y
z
x
y
z
Fig. 1
Tissue clearing and computational pipeline to assess cardiac innervation.
a
Schematic of the clearing-imaging-analysis pipeline for assessing cardiac
innervation.
b
A whole heart (top) was rendered transparent (bottom) using the iDISCO protocol.
c
3D confocal projections of the ventral (1500
μ
mz-
stack) and dorsal side of a cleared heart (1200
μ
m z-stack) with PGP9.5 staining (gray). Inset 1 shows a maximum intensity projection (MIP) confocal
image of a cardiac ganglion. Inset 2 shows 785
μ
m-thick 3D projections of the entire left ventricular wall.
d
,
e
A semiautomated computational pipeline was
used to detect nerve
fi
bers in the dorsal heart image from (
c
) and derive their diameter (
d
) and orientation (
e
). Insets show higher magni
fi
cation images.
Scale bars are 2 mm (
b
), 1 mm (
c
(top),
d
(left),
e
(left)), and 100
μ
m(
c
(bottom),
d
(right),
e
(right))
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in ChAT-ChR2-eYFP mice, we assessed whether we could
stimulate only postganglionic cholinergic neurons in the IPV-
GP using optogenetics and still modulate heart rate. We
fi
rst
evaluated whether we could preferentially deliver transgenes to
peripheral cholinergic neurons in intrinsic cardiac ganglia rather
than central cholinergic neurons in the medulla using systemic
AAVs. We used AAV-PHP.S, a capsid variant that more
ef
fi
ciently transduces the PNS and many visceral organs including
the heart, as compared to AAV9
21
. We packaged a Cre-
dependent genome that expresses eYFP from the ubiquitous
CAG promoter into AAV-PHP.S and systemically administered
the virus to ChAT-IRES-Cre transgenic mice. Three weeks later,
we evaluated eYFP expression in the medulla, vagus nerve, and
cardiac ganglia with GFP staining. Central cholinergic neurons in
the dorsal motor nucleus of the vagus nerve (DMV) (1.5 ± 0.9%
expressed GFP) and
fi
bers in the vagus nerve were weakly
transduced (Supplementary Fig. 3a
c). In contrast, we observed
robust transduction of peripheral cholinergic neurons in cardiac
ganglia (91.0 ± 1.5%) (Supplementary Fig. 3d and e), likely due to
the strong tropism AAV-PHP.S displays for the PNS over the
CNS. Further, GFP expression was highly speci
fi
c for ChAT
+
neurons in cardiac ganglia (100.0 ± 0.0%) (Supplementary Fig. 3e).
For functional studies, we packaged a Cre-dependent genome that
expresses ChR2-eYFP from the ubiquitous CAG promoter in
AAV-PHP.S, systemically administered the virus to ChAT-IRES-
Cre transgenic mice, and evaluated expression 5 weeks later. In
Langendorff-perfused hearts, we were able to optogenetically
stimulate postganglionic cholinergic neurons in the IPV-GP and
decrease heart rate (Supplementary Fig. 4). Taken together, our
anatomical and functional data establish an IPV-GP-SA node
a
Cardiac ganglion
Dense labeling in ChAT-IRES-Cre mouse
ssAAV-PHP.S:CAG-DIO-XFPs (3 × 10
12
vg total)
c
Sparse labeling in ChAT-IRES-Cre mouse
ssAAV-PHP.S:TRE-DIO-XFPs (3 × 10
12
vg total)
ssAAV-PHP.S:ihSyn1-DIO-tTA (1 × 10
10
vg)
Cardiac ganglion
RV
RA
SVC
PV
tdTom
HCN4
Sparse labeling in ChAT-IRES-Cre mouse
ssAAV-PHP.S:TRE-DIO-tdTomato (1 × 10
12
vg)
ssAAV-PHP.S:ihSyn1-DIO-tTA (1 × 10
10
vg)
Dorsal atrium
SA node
IPV-GP
AV node
Tract-tracing
tdTom
HCN4
SA node
IPV-GP
AV node
Dorsal heart
One-component expression system for
cell type-specific dense labeling
Two-component expression system for
cell type-specific sparse labeling
b
+
Variable dose
tTA
High dose
G
R
B
Cre transgenic
mice
High dose
G
R
B
Cre transgenic
mice
SA node
LA
RA
IVC
SVC
PVs
AV node
IPV-GP
RV
LV
Fig. 2
AAV-based labeling and tracing of cholinergic neurons on the heart.
a
Schematic of the one-component expression system for cell type-speci
fi
c,
dense multicolor labeling (top). To densely label cholinergic neurons in cardiac ganglia, ChAT-IRES-Cre mice were systemically co-administered 3
Cre-
dependent vectors expressing either mRuby2, mNeonGreen, or mTurquoise2 from the ubiquitous CAG promoter (ssAAV-PHP.S:CAG-DIO-XFPs) (1 × 10
12
vector genomes (vg) each; 3 × 10
12
vg total). A MIP image of a densely labeled cardiac ganglion (bottom left). Inset shows a higher magni
fi
cation image
(bottom right).
b
Schematic of the two-component expression system for cell type-speci
fi
c, sparse multicolor labeling (top). Expression of XFPs is
dependent on cotransduction of an inducer in Cre-expressing cells. The dose of the inducer vector can be titrated to control extent of XFP labeling. To
sparsely label cholinergic neurons in cardiac ganglia, ChAT-IRES-Cre mice were systemically co-administered 3 Cre-dependent vectors expressing
XFPs
from the TRE-containing promoter (ssAAV-PHP.S:TRE-DIO-XFPs) (1 × 10
12
vg each; 3 × 10
12
vg total) and a Cre-dependent inducer vector expressing the
tTA from the ihSyn1 promoter (ssAAV-PHP.S:ihSyn1-tTA) (1 × 10
10
vg). A MIP image of a sparsely labeled cardiac ganglion (bottom left). Inset shows a
higher magni
fi
cation image (bottom right).
c
To trace cholinergic
fi
bers, presumably from cardiac ganglia, sparse labeling was performed by systemically
co-administering ssAAV-PHP.S:TRE-DIO-tdTomato at a high dose (1 × 10
12
vg) and ssAAV-PHP.S:ihSyn1-DIO-tTA at a lower dose (1 × 10
10
vg). Cartoon of
the dorsal heart depicting the orientation of images (left). A MIP image of the dorsal atrium with native tdTomato
fl
uorescence (red) and HCN4 staining
(green) (middle). Fibers were traced with neuTube and overlaid on a grayscale MIP image (right). Orange
fi
bers coursed along the right atrium (RA)
including the sinoatrial (SA) and atrioventricular (AV) nodes and blue
fi
bers along the ventricles. Scale bars are 50
μ
m(
a
,
b
) and 200
μ
m(
c
). All images
were acquired on tissue collected 3 weeks after intravenous injection. IPV-GP inferior pulmonary vein-ganglionated plexus, IVC inferior vena cava
, LA left
atrium, LV left ventricle, PV pulmonary vein, RV right ventricle, SVC superior vena cava
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circuit involved in heart rate regulation (Fig.
3
l). Furthermore, the
engineered AAV, AAV-PHP.S
21
, can be a powerful tool to dissect
out the roles of peripheral versus central circuits on organ
control.
Optogenetic versus electrical stimulation of the vagus nerve
.
Electrical vagus nerve stimulation (VNS) has been used in
numerous preclinical and clinical studies for the treatment of
cardiovascular diseases
43
and other conditions (e.g., rheumatoid
arthritis
44
). However, the relative contributions of vagal efferent
and afferent
fi
bers on cardiac function are not well understood
because conventional techniques do not allow for
fi
ber type-
speci
fi
c stimulation. To address this limitation, we examined
whether we could selectively stimulate efferent
fi
bers in the vagus
nerve using optogenetics. We also assessed whether there was a
difference between optogenetic and electrical VNS on heart rate
(Fig.
4
a), as a previous study showed that electrical stimulation of
motor nerves results in a non-orderly, non-physiological
recruitment of
fi
bers, with larger
fi
bers activated
fi
rst
45
. The
0
5
10
15
20
P wave duration (ms)
*
Pre-
stimulation
Stimulation
Pre-
stimulation
Stimulation
25
30
35
40
45
AV interval (ms)
IPV-GP
Langendorff preparation
EP catheter
Optical
fiber
ECG
cd
Pre-atropine
stimulation
Post-atropine
stimulation
–80
–60
–40
–20
0
20
*
h
fg
e
10 Hz
20 Hz
1 s
100 ms
a
b
Cardiac ganglion
Ventricle
PGP9.5
GFP
1
2
ChAT-ChR2-eYFP
1
2
GFP
PGP9.5
Dorsal heart
GFP
PGP9.5
k
l
SA node
LA
RA
IVC
SVC
PVs
IPV-GP
Dorsal heart
i
10 Hz
1 s
10 ms
j
0
50
100 150 200 250
–80
–60
–40
–20
0
Power (mW)
Δ
Heart rate (%)
–80
–60
–40
–20
0
Δ
Heart rate (%)
Δ
Heart rate (%)
–80
–60
–40
–20
0
Δ
Heart rate (%)
0
5
10
15
20
Frequency (Hz)
0510
Pulse width (ms)
GFP+ & PGP9.5+/
GFP+
GFP+ & PGP9.5+/
PGP9.5+
0
20
40
60
80
100
Cardiac ganglion
neurons (%)
QRS
P
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vagus and other autonomic nerves contain both motor and sen-
sory
fi
bers that vary in diameter and myelination
28
and non-
electrical techniques such as optogenetics are needed to study
their physiological role.
To con
fi
rm that ChR2-eYFP expression was limited to vagal
efferents in ChAT-ChR2-eYFP mice, we stained for GFP and
PGP9.5 in the nodose/jugular ganglion complex, which contains
the cell bodies of vagal sensory neurons, and the cervical vagus
Fig. 3
Ex vivo optogenetic stimulation of cholinergic neurons in the IPV-GP.
a
A 3D projection (1200
μ
m z-stack) of the dorsal side of a heart from a ChAT-
ChR2-eYFP mouse whole-mount stained with PGP9.5 (red) and GFP (green). Insets 1 and 2 show MIP images of a cardiac ganglion and the ventricle,
respectively. Blue dashed boxes indicate location of higher magni
fi
cation images in blue boxes.
b
Percentage of cardiac ganglion neurons expressing GFP
and PGP9.5 over those expressing GFP or PGP9.5.
c
Langendorff-perfused hearts were used for optogenetic stimulation of cholinergic neurons in a cardiac
ganglion. A blue laser-coupled optical
fi
ber was positioned for focal illumination of the IPV-GP (circle). A surface electrocardiogram (ECG) was recorded
with bath electrodes and intracardiac electrograms with an electrophysiology (EP) catheter.
d
Representative ECGs during stimulation (blue shading) at
10 Hz, 10 ms, and 221 mW (top) and 20 Hz, 10 ms, and 221 mW (bottom). Insets show the ECGs before and during stimulation.
e
g
Dose response
curves summarizing the effects of altering light pulse power (
e
), frequency (
f
), and pulse width (
g
) on heart rate.
h
Summary of the AV interval before and
during stimulation at 10 Hz and 10 ms (
t
4
=
0.1656,
P
=
0.8765).
i
Representative ECG during stimulation (blue shading) at 10 Hz, 10 ms, and 221 mW.
Insets show a single beat before and during stimulation, with gray boxes showing a higher magni
fi
cation of the P wave.
j
The P wave duration before versus
during stimulation (
t
5
=
2.920, *
P
=
0.0330).
k
Summary of the heart rate response to stimulation before versus after atropine administration (
t
4
=
2.993,
*
P
=
0.0402).
l
Cartoon of the dorsal heart depicting the IPV-GP-SA node circuit.
n
=
6 mice (
b
,
e
g
,
j
) and 5 mice (
h
,
k
); mean ± s.e.m.; paired, two-tailed
t
-test. Scale bars are 1 mm (
a
(left),
c
) and 100
μ
m(
a
(right))
g
BVNx cranial RVNS
0
5
10
15
20
Frequency (Hz)
**
h
0
Intact
BVNx rostral
5
10
15
Time to peak heart
rate response (s)
Electrical RVNS
**
i
abc
de
GFP
PGP9.5
GFP
PGP9.5
Nodose/jugular complex
f
Vagus nerve
Trachea
In vivo optical versus electrical
vagus nerve stimulation
1 s
300
400
500
600
700
Heart rate (bpm)
20 Hz intact RVNS
Electrical
Optical
ChAT-ChR2-eYFP
0
5
10
15
20
–80
–60
–40
–20
0
Frequency (Hz)
Δ
Heart rate (%)
Δ
Heart rate (%)
Intact RVNS
0
5
10
15
20
Frequency (Hz)
RVNx caudal RVNS
0
5
10
15
20
–80
–60
–40
–20
0
Frequency (Hz)
BVNx caudal RVNS
RVNx cranial RVNS
0
5
10
15
20
–20
–15
–10
–5
0
5
–80
–60
–40
–20
0
Δ
Heart rate (%)
Δ
Heart rate (%)
Δ
Heart rate (%)
–20
–15
–10
–5
0
5
Frequency (Hz)
**
*
Fig. 4
In vivo optogenetic versus electrical stimulation of the vagus nerve.
a
Cartoon depicting optogenetic and electrical vagus nerve stimulation strategy
in ChAT-ChR2-eYFP mice. The right vagus nerve was surgically exposed in anesthetized mice and either light or electricity was used for stimulation.
b
MIP
images of the right nodose/jugular ganglion complex and vagus nerve whole-mount stained with PGP9.5 (red) and GFP (green).
c
Representative heart
rate responses during optogenetic versus electrical right vagus nerve stimulation (RVNS; magenta shading) at identical frequencies (20 Hz) and pul
se
widths (10 ms) in the intact state. The light pulse power for optogenetic stimulation was 57 mW and the current for electrical stimulation was 5
μ
A.
d
Frequency response curves summarizing the effects of optogenetic versus electrical RVNS on heart rate in the intact state.
e
,
f
Frequency response curves
summarizing the effects of optogenetic versus electrical RVNS of the caudal end on heart rate following right vagotomy (RVNx) (
e
) and bilateral vagotomy
(BVNx) (
f
).
g
,
h
Frequency response curves summarizing the effects of optogenetic versus electrical RVNS of the cranial end on heart rate following RVNx
(
t
4
=
3.576, *
P
=
0.0232 at 10 Hz;
t
4
=
5.229, **
P
=
0.0064 at 20 Hz) (
g
) and BVNx (
t
4
=
8.588, **
P
=
0.0010 at 20 Hz) (
h
). In
d
h
insets show a
schematic of the stimulation protocol.
i
The time to peak heart rate response during electrical RVNS in the intact state versus of the cranial end following
BVNx (
t
4
=
6.335, **
P
=
0.0032).
n
=
8 mice (
d
), three mice (
e
,
f
), and
fi
ve mice (
g
,
h
,
i
); mean ± s.e.m.; paired, two-tailed
t
-test. Scale bar is 200
μ
m(
b
)
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nerve (Fig.
4
b). eYFP was not detected in PGP9.5
+
cell bodies in
the nodose/jugular ganglion complex and was only present in a
subset of PGP9.5
+
vagal
fi
bers (
n
=
5 mice). After verifying
expression, we next performed functional studies in anesthetized
mice in which we positioned a laser-coupled optical
fi
ber for focal
illumination above and a hook electrode underneath the right
vagus nerve. In this context, optogenetic VNS activates only
efferent
fi
bers (GFP
+
), whereas electrical VNS presumably
activates subsets of efferent and afferent
fi
bers (PGP9.5
+
)in
the vagus nerve (Fig.
4
b). With both vagi intact, optogenetic and
electrical right VNS resulted in a similar decrease in heart rate
(Fig.
4
c and d); however, the slopes of the responses were
dramatically different (
n
=
6/8 mice) (Fig.
4
c), likely due to
differential
fi
ber recruitment
45
. The heart rate response to
optogenetic versus electrical stimulation of the caudal end of
the right vagus nerve was also similar following either right or
bilateral vagotomy (Fig.
4
e, f and Supplementary Table 2). In
contrast, optogenetic stimulation of the cranial end of the right
vagus nerve following either right or bilateral vagotomy did not
affect heart rate, whereas electrical stimulation surprisingly
resulted in a decrease in heart rate at 10 Hz (
6.5 ± 1.8% versus
0.1 ± 0.1% following right vagotomy for electrical versus
optogenetic stimulation) and 20 Hz (
9.9 ± 1.8% versus
0.1 ±
0.1% following right vagotomy and
3.2 ± 0.4% versus
0.2 ±
0.1% following bilateral vagotomy for electrical versus optoge-
netic stimulation) (Fig.
4
g, h and Supplementary Table 2). The
decrease in heart rate to electrical stimulation of the cranial end of
the right vagus nerve following right vagotomy was also greater
than that following bilateral vagotomy at 20 Hz (
t
4
=
3.123,
P
=
0.0354, paired, two-tailed
t
-test) (Fig.
4
g, h, red line and
Supplementary Table 2), suggesting that the response following
ipsilateral vagotomy was in part due to a vagal afferent-mediated
increase in parasympathetic efferent out
fl
ow through the intact
contralateral vagus nerve. Furthermore, the decrease in heart rate
to electrical stimulation of the cranial end of the right vagus nerve
following bilateral vagotomy (Fig.
4
h, red line and Supplementary
Table 2) indicates that vagal-afferents mediate a decrease in
sympathetic efferent out
fl
ow, since both vagi were transected. In
addition, there was an increased latency to peak heart rate
response with electrical stimulation of the cranial end of the right
vagus nerve following bilateral vagotomy compared to that of the
intact right vagus nerve (8.7 ± 0.4 ms versus 3.9 ± 0.5 ms) (Fig.
4
i),
which further supports that vagal afferents cause withdrawal of
sympathetic tone
46
. Taken together, our data suggest that
optogenetic stimulation selectively activates vagal efferents in
ChAT-ChR2-eYFP mice and that vagal afferents act centrally to
(1) increase parasympathetic efferent out
fl
ow and (2) decrease
sympathetic efferent out
fl
ow to the heart.
Location and optogenetic stimulation of cardiac-projecting
noradrenergic neurons in the stellate ganglia
. The sympathetic
nervous system, along with the parasympathetic nervous system,
precisely regulates heart rate in normal physiology. To anato-
mically and functionally dissect noradrenergic neurons that form
a circuit with the SA node, we used a retrograde neuronal tracer
and an optogenetic approach. Noradrenergic nerve
fi
bers densely
innervate the heart as shown by staining for tyrosine hydroxylase
(TH), the rate-limiting enzyme in norepinephrine synthesis
(Fig.
5
a). To identify the location of cardiac-projecting sympa-
thetic neurons, we injected the retrograde neuronal tracer CTB
conjugated to Alexa Fluor 488 into the heart (Fig.
5
b). The
ab c
d
0.5
1.0
1.5
2.0
0
0
0.5
1.0
x
(mm)
y
(mm)
RSG
0.5
1.0
1.5
2.0
0
2.5
0
0.5
1.0
1.5
x
(mm)
y
(mm)
LSG
0
0.5
1.0
1.5
×10
–3
CTB+ cells/
μ
m
2
CTB-488
Ansa
SG
T2G
MCG
Left paravertebral chain ganglia
Heart
SG
T2G
CTB-488
LV
RV
Paravertebral
chain
TH
Dorsal heart
Fig. 5
Cardiac-projecting neurons in stellate ganglia (SG) are clustered craniomedially.
a
A3Dprojection(1200
μ
m z-stack) of the dorsal side of a C57BL/6J
mouse heart whole-mount stained with TH (green).
b
Cartoon depicting cholera toxin subunit B (CTB)-Alexa Fluor 488 (CTB-488) injections into the heart to
retrogradely trace neurons in the paravertebral chain ganglia that project to the heart.
c
A MIP image of the left paravertebral chain from the middle cervical
ganglion (MCG) to the second thoracic ganglion (T2G) showing the location of CTB-488
+
neurons that project to the heart.
d
Summary heat map of the right
stellate ganglion (RSG) and the left stellate ganglion (LSG).
n
=
5mice(
d
). Scale bars are 1 mm (
a
,
c
). CTB injections were performed in C57BL/6J mice
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7
majority of CTB
+
neurons were located in the stellate ganglia of
the paravertebral chain, with fewer labeled neurons in the middle
cervical and second thoracic (T2) ganglia (Fig.
5
c and Supple-
mentary Movie 7). On average, 236 ± 39 neurons were labeled in
the right and 261 ± 34 neurons labeled in the left stellate ganglion
(
n
=
5 mice). Heat maps of the right stellate ganglion (RSG) and
left stellate ganglion show that cardiac-projecting sympathetic
neurons are clustered in the craniomedial aspect (Fig.
5
d) and
suggest that these ganglia may have a viscerotopic organization.
Next, we assessed whether we could selectively stimulate
noradrenergic neurons in the paravertebral chain using optoge-
netics and modulate heart rate (Fig.
6
a). We expressed ChR2 in
noradrenergic neurons by crossing transgenic TH-IRES-Cre mice
with reporter mice containing a Cre-dependent ChR2-eYFP allele
(offspring from this cross are subsequently referred to as TH-
ChR2-eYFP mice). ChR2-eYFP expression in stellate ganglion
neurons was con
fi
rmed by staining for GFP and TH (Fig.
6
b). The
majority of GFP
+
neurons were TH
+
(98.7 ± 0.4%) and all TH
+
neurons were GFP
+
(100.0 ± 0.0%) (Fig.
6
c). After verifying
expression, we performed functional studies in open-chest
anesthetized mice in which we positioned a laser-coupled optical
fi
ber for focal illumination above the craniomedial RSG or right
T2 ganglion (Fig.
6
a). Optogenetic stimulation of the craniomedial
RSG resulted in a frequency- and pulse width-dependent increase
in heart rate (Fig.
6
d
f and Supplementary Table 3). Although a
small number of cardiac-projecting neurons were located in the
T2 ganglion (Fig.
5
c), there was no heart rate response to
stimulation of this ganglion and/or preganglionic sympathetic
fi
bers coursing through this region (0.1 ± 0.1% with right T2
ganglion stimulation versus 9.5 ± 1.8% with RSG stimulation)
(Fig.
6
g and Supplementary Table 3). The response to craniome-
dial RSG stimulation was abolished by administration of the
β
-
adrenergic receptor antagonist propranolol (5.6 ± 1.4% with
stimulation before propranolol versus 0.1 ± 0.1% with stimulation
after propranolol) (Fig.
6
h and Supplementary Table 3), indicating
that the tachycardic response was indeed mediated by selective
GFP+ &TH+/
GFP+
TH+ & GFP+/
TH+
0
20
40
60
80
100
Stellate ganglion
neurons (%)
Pre-propranolol
RSGS
Post-propranolol
RSGS
–5
0
5
10
15
20
*
def
g
TH-ChR2-eYFP
ac
400
450
500
550
600
Heart rate (bpm)
RSGS
1 s
b
Heart
SG
T2G
In vivo optical stimulation of
paravertebral ganglia
SG
T2G
TH
GFP
TH
GFP
TH
GFP
Right paravertebral chain ganglia
h
Ventral heart
RSG
SA node
SVC
RA
RSGS
RT2GS
–5
0
5
10
15
20
**
0
5
10
15
20
0
5
10
15
20
Frequency (Hz)
Δ
Heart rate (%)
Δ
Heart rate (%)
Δ
Heart rate (%)
Δ
Heart rate (%)
0510
0
5
10
15
20
Pulse width (ms)
i
Fig. 6
In vivo optogenetic stimulation of noradrenergic neurons in the RSG.
a
Cartoon depicting optogenetic SG and T2G stimulation strategy in TH-ChR2-
eYFP mice. The right paravertebral chain was surgically exposed in anesthetized mice and light was used for stimulation.
b
A MIP image of the right
paravertebral chain whole-mounted stained with TH (red) and GFP (green) and showing the SG and the T2G. Inset shows single-plane images of the SG.
Blue dashed boxes indicate location of higher magni
fi
cation images in blue boxes.
c
Percentage of stellate ganglion neurons expressing GFP and TH over
those expressing GFP or TH.
d
Representative heart rate response during 10 Hz, 10 ms, and 126 mW craniomedial RSG stimulation (RSGS).
e
,
f
Dose
response curves summarizing the effects of altering craniomedial RSGS frequency (
e
) and pulse width (
f
) on heart rate.
g
Summary of the heart rate
response to craniomedial RSGS versus right T2G stimulation (RT2GS) (
t
6
=
5.435, **
P
=
0.0016).
h
Summary of the heart rate response to craniomedial
RSGS before versus after propranolol administration (
t
3
=
3.951, *
P
=
0.0289).
i
Cartoon of the ventral heart depicting the craniomedial RSG-SA node
circuit
.n
=
6 mice (
c
), seven mice (
e
,
g
),
fi
ve mice (
f
), and four mice (
h
); mean ± s.e.m.; paired, two-tailed
t
-test. Scale bars are 200
μ
m(
b
(left) and 50
μ
m
(
b
(right)
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stimulation of noradrenergic neurons. Taken together, our
anatomical and functional data establish a craniomedial RSG-SA
node circuit involved in heart rate regulation (Fig.
6
i).
Discussion
We developed a clearing-imaging-analysis pipeline to visualize
innervation of whole hearts and employed a multi-technique
approach to dissect fundamental parasympathetic and sympa-
thetic neural circuits involved in heart rate regulation. We report
several novel
fi
ndings: (1) cholinergic neurons in the IPV-GP and
noradrenergic neurons in the craniomedial RSG project to the SA
node and modulate its function; (2) the evoked cardiac response
to optogenetic versus electrical stimulation of the vagus nerve
displays different temporal characteristics; and (3) vagal afferents
enhance parasympathetic and reduce sympathetic efferent out-
fl
ow to the heart via central mechanisms.
Despite advances in tissue clearing
33
,
47
and imaging
techniques
30
,
31
, high-resolution, 3D datasets of global cardiac
innervation do not exist. We show, for the
fi
rst time, innervation
of entire cleared mouse hearts with cardiac ganglia located
around the pulmonary veins and a dense network of nerve
fi
bers
throughout the myocardium. To analyze these data, we developed
a semiautomated computational pipeline to detect nerve
fi
bers
and to measure microanatomical features such as diameter and
orientation. These analytical tools and resulting measurements
are needed to build a reference atlas of cardiac innervation and
for quantitative descriptions of innervation in healthy versus
diseased states such as MI. Following MI, innervation around the
infarct scar and of remote regions of the heart is altered
32
,
33
,
48
,
and this neural remodeling can modulate the arrhythmia sub-
strate
49
. Understanding changes in innervation post-MI can
provide new insights into arrhythmia mechanisms. Furthermore,
our clearing-imaging-analysis pipeline can be readily applied to
assess innervation of other visceral organs including endogen-
ously and virally labeled tissues.
It is well known that the parasympathetic and sympathetic
nervous systems are critical for heart rate regulation. The SA node
and conduction system are densely innervated
7
,
8
, and stimulation
of the vagus nerve
9
, stellate ganglia
10
, and noradrenergic
fi
bers
11
,
12
modulates heart rate. However, the precise wiring of
the underlying neural circuits has not been delineated. We used a
novel sparse AAV labeling system
21
and an optogenetic approach
to anatomically and functionally characterize cholinergic neurons
that regulate heart rate. Although we were unable to directly
visualize synapses, we identi
fi
ed cholinergic
fi
bers, presumably
from cardiac ganglia, that coursed along the SA node, the AV
node, and the ventricles. Selective optogenetic stimulation of
cholinergic neurons in the IPV-GP modulated heart rate, con-
sistent with a recent study that stimulated cholinergic
fi
bers in the
right atrium also using optogenetics
13
. Previous studies showed
that electrical stimulation of pulmonary vein ganglia results in
biphasic heart rate responses (initial bradycardia followed by
tachycardia)
23
,
24
. However, since electrical stimulation is non-
speci
fi
c, it is dif
fi
cult to interpret whether the biphasic response
was due to activation of a mixed population of neurons contained
in cardiac ganglia (i.e., parasympathetic, sympathetic, and sen-
sory)
15
and/or pass through
fi
bers. Our
fi
ndings demonstrate an
IPV-GP-SA node circuit and highlight the importance of using
techniques such as optogenetics, which confer cell type-speci
fi
-
city, to dissect cardiac neural circuity. Furthermore, electrical and
optogenetic techniques (using traditional transgenic and AAV-
based approaches for ChR2 delivery) stimulate both central
preganglionic inputs to and postganglionic neurons in cardiac
ganglia. Therefore, we used a novel engineered AAV, AAV-PHP.
S
21
, that has a strong tropism for the PNS to preferentially deliver
ChR2 to postganglionic cholinergic neurons on the heart rather
than preganglionic cholinergic neurons in the medulla. Future
studies of peripheral neural circuits should use AAV-PHP.S and
other engineered AAVs
20
,
22
to dissect the role of central versus
peripheral neuronal populations on organ function.
To map noradrenergic neurons that regulate heart rate, we
used a retrograde neuronal tracer and optogenetic approach. A
previous study in canines using horseradish peroxidase showed
that sympathetic postganglionic neurons that innervate the heart
are primarily located in the middle cervical ganglia of the para-
vertebral chain
50
. However, we report, using CTB and con
fi
rm
with optogenetic stimulation, that the stellate ganglia have a
viscerotopic organization with cardiac-projecting neurons clus-
tered in the craniomedial aspect, consistent with a study in cats
51
.
In addition to the heart, the stellate ganglia project to many other
thoracic structures, including the sweat glands in the forepaw
52
,
the lung and trachea
53
, the esophagus
51
, and brown fat
54
. Char-
acterizing stellate ganglia target innervation and cell-type speci-
fi
cation is an area of ongoing investigation
55
,
56
that is of interest
from a developmental, physiological, and therapeutic perspective.
Current understanding of the role of vagal efferent and afferent
fi
bers on cardiac function is largely based on studies using elec-
trical stimulation, which is non-speci
fi
c and results in non-
orderly, non-physiological recruitment of
fi
bers
45
. Electrical sti-
mulation of the vagus nerve typically activates large-diameter
myelinated A
fi
bers, followed by medium-diameter myelinated B
fi
bers and then small-diameter unmyelinated C
fi
bers
28
. In vivo
we report that optogenetic stimulation of motor
fi
bers in the
vagus nerve results in a heart rate response that has a slower onset
than electrical stimulation of motor and sensory
fi
bers, likely due
to differential
fi
ber recruitment
45
. Our
fi
ndings suggest that non-
electrical techniques such as optogenetics are needed to char-
acterize neural control of cardiac physiology. Although there was
robust expression of ChR2-eYFP as con
fi
rmed by immunohis-
tochemistry, we cannot rule out that incomplete expression or
optical capture affected our results. We also show that activation
of vagal afferents decreases heart rate by enhancing para-
sympathetic efferent out
fl
ow and reducing sympathetic efferent
out
fl
ow centrally, consistent with a prior study showing that
global activation of vagal afferents in Vglut2-ChR2 mice and
selective activation of Npy2r-ChR2 vagal afferents causes a pro-
found bradycardia
57
. In support of our functional data, anato-
mical tracing studies have previously shown that cardiac vagal
afferent neurons in the nodose/jugular ganglion complex project
to neurons in the nucleus tractus solitarii of the medulla
58
. These
neurons then project to the nucleus ambiguus and the DMV in
the medulla to modulate parasympathetic efferent out
fl
ow
58
and
to the paraventricular nucleus of the hypothalamus to modulate
sympathetic efferent out
fl
ow
59
. Future studies aimed at identify-
ing cardiac-speci
fi
c vagal efferent and afferent
fi
bers, similar to
those performed in the lungs
57
and the gastrointestinal system
60
,
are needed to better understand vagal control of cardiac phy-
siology and to design next-generation VNS therapies.
Overall, our data highlight the complexity of cardiac neural
circuitry and demonstrate that a multi-technique approach is
needed to delineate circuit wiring. Understanding the neural
control of organ function in greater detail is critical as neuro-
modulation therapies are emerging as promising approaches to
treat a wide range of diseases. Tools such as optogenetics and
AAVs are already providing new scienti
fi
c insights into the
structure and function of peripheral neural circuits
57
,
60
. A com-
bination of these approaches will help disentangle neural control
of autonomic physiology and enable a new era of targeted neu-
romodulation approaches.
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