Direct Recording of Renal Sympathetic Nerve Activity in
Unrestrained, Conscious Mice
Shereen M. Hamza, John E. Hall
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Abstract—Renal sympathetic nerve activity (RSNA) has been measured in anesthetized mice. However, anesthesia and
acute surgical preparation cause poor cardiovascular stability and unphysiological blood pressures. This compromised
physiological state confounds proper interpretation of experimental results considering the inseparable link between
cardiovascular status and autonomic nervous tone. We, therefore, developed a surgical and experimental protocol for
measuring RSNA in conscious, unrestrained mice. Male C57Bl/6J mice were chronically instrumented with blood
pressure radiotelemeters, an indwelling jugular venous catheter and a bipolar electrode for recording RSNA. Mice were
placed in a home cage and left to recover for 48 to 72 hours. Survival rate was 100%; all of the mice exhibited normal
behavior with no sign of distress 24 hours after surgery. RSNA was successfully recorded in 80% of the mice at 48 and
72 hours postsurgery; viable RSNA was reduced to 70% and 50% at 4 and 5 days postsurgery, respectively. Mean
arterial pressure (116⫾2 mm Hg; n⫽10) was consistent with values reported previously for conscious mice. RSNA
increased with the normal physical activities of eating and grooming and was validated by ganglionic blockade and
pharmacological manipulation of blood pressure; reduction in blood pressure to 62⫾3 mm Hg with nitroprusside
increased RSNA by 77⫾9% above baseline (n⫽5; P⬍0.05), whereas an increase in blood pressure to 137⫾6 mm Hg
with phenylephrine reduced RSNA by 79⫾2% compared with baseline (n⫽5; P⬍0.05). Thus, we demonstrate an
accessible and effective method for direct assessment of RSNA in conscious, unrestrained mice. (Hypertension. 2012;
60:00-00.)
Key Words: autonomic tone 䡲 physiological blood pressure 䡲 kidney
M
ice are now routinely used in many areas of biomedical
research, especially with the increasing generation of
genetically engineered animals. For physiologists, use of this
species has posed special technical challenges that have
required development of equipment and devices designed for
measuring critical physiological parameters in mice. An
important focus of research is elucidating the contribution of
the brain and autonomic nervous system to aspects of health
and disease; to date, the function of the autonomic nervous
system in mice has been assessed mainly by indirect methods,
such as measurement of plasma/urine catecholamine levels,
cardiovascular responses to pharmacological autonomic
blockade, or analysis of high- and low-frequency variations in
blood pressure and heart rate.1 These methods yield a rough
estimate of overall autonomic tone and do not reveal the
contribution of specific nerves to a particular phenomenon.
Direct recordings from autonomic nerves are often performed
in anesthetized mice. This approach is rife with concerns,
because relevant hemodynamic parameters, such as blood
pressure, are often not measured or are reported at alarmingly
low levels (60 – 80 mm Hg).2 The relative lack of cardiovascular stability of the anesthetized mouse preparation is of
particular concern when attempting to assess sympathetic
nervous tone considering the intimate link between arterial
blood pressure and sympathetic nerve activity.
To address this issue, we developed a surgical and experimental method for directly measuring renal sympathetic
nerve activity (RSNA) in conscious, freely moving mice.
Animals are simultaneously instrumented for telemetric measurement of arterial blood pressure and are implanted with
indwelling intravenous catheters for direct infusion of agents
of interest. As early as 24 hours after successful surgical
preparation, mice exhibit normal behavior and show no sign
of pain or distress. Recordings can be completed 48 to 72
hours after surgery while animals are resting quietly in their
home cage with free access to food and water at all times. We
demonstrate clear RSNA, which responds appropriately to the
natural physical activity of the animal, as well as pharmacological manipulation of blood pressure with sodium nitroprusside and phenylephrine. The postganglionic nature of the
RSNA signal was verified by ganglionic blockade with
hexamethonium.
Materials and Methods
All of the experimental procedures are in accordance with the
National Institutes of Health Guide for the Care and Use of
Received October 21, 2011; first decision November 12, 2011; revision accepted July 3, 2012.
From the Department of Physiology and Biophysics, University of Mississippi Medical Center, Jackson, MS.
Correspondence to Shereen M. Hamza, Department of Paediatrics, University of Alberta, Edmonton, Alberta, Canada T6G 2S2. E-mail
[email protected]
© 2012 American Heart Association, Inc.
Hypertension is available at http://hyper.ahajournals.org
DOI: 10.1161/HYPERTENSIONAHA.111.186577
1
2
Hypertension
September 2012
A
D
B
pin connectors
PE90 tubing
ground
wire
renal nerve
C
bipolar leads
PE10 tubing
g
bipolar leads
parafilm
groundwire
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
groundwire
Figure 1. Schematic representation of construction and placement of the implantable renal sympathetic nerve activity (RSNA) electrode. A,
Bipolar leads fitted with pin connectors and a third ground wire. B, Wires are threaded through polyethylene (PE) 90 tubing for protection. C,
Construction of the electrode tip to separate the bipolar leads from the ground wire. D, Electrode tip is bent at a 90° angle; renal nerve bundle is positioned perpendicular to the bipolar leads. Parafilm separates the leads from the ground wire touching underlying tissue.
Laboratory Animals and were approved by the institutional animal
care and use committee of the University of Mississippi Medical
Center.
Animals and Housing
Twelve Male C57BL/6J mice (24 –35 g, Jackson Laboratories) were
housed for ⱖ1 week in the laboratory animal facility on arrival at the
University of Mississippi Medical Center. They were maintained on
standard rodent chow and tap water ad libitum in a temperature- and
humidity-controlled environment before and after surgery.
Construction of the Implantable RSNA Electrode
Three lengths (250-mm each) of Teflon-coated stainless steel,
multiple stranded wire (0.001-in bare, 0.0055-in coated; A-M Systems, Inc), were cut, and ⬇15 mm of the Teflon coating was stripped
away from one end with a #11 scalpel blade to expose the metal
wires. The bared ends of 2 of the lengths of wire were each soldered
to a single male pin connector (brass with gold plating; A-M
Systems, Inc) to create the bipolar electrode leads (Figure 1A). A
20-mm piece of heat shrink tubing (1.6-mm diameter, RadioShack)
was slipped over the pin connector and wire to cover the newly
soldered joint while exposing the pin tip. The tubing was shrunk over
a heat gun to insulate the connection between the wire and pin
connector. The unexposed ends of the 2 electrode leads and the third
length of Teflon-coated wire (ground wire) were then threaded
together through a length of polyethylene tubing (Figure 1B;
200-mm long, polyethylene (PE) 90, ID 0.86 mm, OD 1.27 mm,
Braintree Scientific). Of the 3 wires, the ground wire was identified
and pulled through the PE 90 sheath a little further to differentiate it
from the bipolar electrode leads.
Under a dissecting microscope, the 3 loose ends of the electrode
were threaded through a 5-mm–long piece of smaller polyethylene
tubing (PE 10, ID 0.28 mm, OD 0.61 mm, Braintree Scientific; the
purpose of which is to bind the leads together). Similarly, a 1.5-mm
piece of this PE tubing was threaded onto the 3 wires to rest 2 mm
from the first piece. A second 1.5-mm piece of PE 10 was threaded
onto the 2 electrode leads to cover the tips and to separate them from
the ground wire (Figure 1C). The wires were trimmed such that 2
mm of the bipolar leads would form the surface for contact with the
nerve. All of the pieces of PE 10 tubing were secured to the electrode
leads with a small drop of Super Glue (liquid; Loctite). Once the
super glue was dry (overnight), the Teflon coating was stripped from
the bipolar electrode tip and ground wire tip with a #11 scalpel blade,
with care taken not to fray the multiple stranded wires. The electrode
tip was then bent at a 90° angle at the junction of the 5.0-mm and
1.5-mm PE 10 anchors (Figure 1D). A pedestal to help stabilize the
electrode leads to the animal was constructed from 1 inch of
polyethylene tubing (2.70 mm ID⫻4.00 mm OD, Scientific Commodities, Inc), which had been melted at one end to create a flange.
This pedestal was threaded onto the electrode and left until needed.
This pedestal plus the PE 90 cover protected the electrode so that a
spring was not needed and instead used to protect the jugular venous
catheter (see below). The electrodes were then packaged and ozone
sterilized (TSO3) before implantation.
Anesthesia and Surgery
Anesthesia was induced with 4% isoflurane and subsequently maintained with 1.5% to 2.0% isoflurane. Body temperature was maintained with Deltaphase isothermal heat pads (Braintree Scientific).
All of the surgical procedures were conducted under aseptic conditions. Immediately on induction of anesthesia, glycopyrrolate
(50 –70 ␮g/kg, SC) was administered to prevent production of
excessive airway secretions. This dose of glycopyrrolate was administered once again at the midpoint of the surgical procedure.
Implantation of the RSNA Electrode
The mouse was positioned on his right side; the rostral end at the
surgeon’s left, exposing the left flank. A small incision (⬇5 mm) was
made in the skin dorsally, 1 cm below the scapulae; this is the exit
site for the renal sympathetic nerve electrode leads. A second
incision (⬍20 mm) was made perpendicular to the spine in the skin
overlying the left flank, 2 mm caudal to the ribcage. A 13G
stainless-steel needle was tunneled subcutaneously from this flank
incision to the dorsal exit site. The RSNA electrode was threaded
into the 13G needle, which was pulled back to leave the electrode tip
on the abdominal muscle of the left flank, with leads tunneled
subcutaneously to exit dorsally. The electrode tip was placed to the
side and an incision was made in the abdominal muscle. Using
small-tipped cotton applicators, the fat and connective tissue along
the back muscle were gently separated to retroperitoneally expose
the left kidney. Steel microretractors were used to gently open the
surgical field and retract the kidney, with great care taken to avoid
Blood Pressure
(mmHg)
Hamza and Hall
RSNA in Conscious Mice
3
120
100
80
60
40
20
RSNA (µV.s)
RSNA (µV)
30
25
20
15
10
5
0.18
0.17
1 sec
0.1 sec
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Figure 2. Representative recording of systemic arterial blood pressure, renal sympathetic nerve activity (RSNA), and integrated RSNA in
a conscious, resting mouse 48 hours after surgery.
stretching the renal neurovascular bundle or damaging the kidney
itself.
The renal neurovascular bundle was visualized with a high-power
dissecting microscope (Leica M80, Leica Microsystems). A renal
nerve bundle was identified coursing alongside the renal artery and
vein and was gently isolated from the surrounding tissues with fine,
straight forceps (#5 Medical and #5 Medical Biology, FST by
Dumont, Fine Science Tools). It is imperative that the nerve is not
stretched or picked up with forceps during this isolation. In addition,
damage to the renal lymph duct generally running along the renal
vessels was also avoided, because this will lead to continuous
leakage of lymph fluid in the area, which will inevitably interfere
with the nerve signal. The renal nerve bundle was left intact and not
sectioned to maintain long-term viability of the nerve and to help
maintain stable contact between the nerve and electrode. At this
point, the electrode tip was introduced into the abdomen and its
position adjusted such that the bipolar electrode tip and ground wire
laid perpendicular to the renal nerve bundle; the ground wire must
have good contact with the underlying tissues and the electrode must
not hamper the renal circulation (Figure 1D). The renal nerve bundle
was gently lifted with angled forceps (#5/45, FST by Dumont, Fine
Science Tools), and the electrode tip was carefully slipped underneath the nerve, leaving it in direct contact with both electrode wires.
A small piece of Parafilm was placed between the nerve/bipolar
wires and the third (ground) wire (Figure 1D); all of the fluid around
the nerve and electrode was removed with small absorbent spears
(Fine Science Tools). A 2-component silicone elastomer (Kwik-Sil,
World Precision Instruments) was applied to the nerve/electrode
unit, ensuring that the silicone pooled under and around the nerve for
full electric insulation (not just on the surface of the nerve). After the
silicone cured completely (⬇1–2 minutes), the very edges of the
silicone “blob” were carefully lifted with forceps and fixed to the
surrounding tissue with a small amount of surgical adhesive (Vetbond, 3M). The abdominal muscle was then closed with discontinuous absorbable sutures (5-0 Polysorb absorbable suture, Tyco
Healthcare); the overlying skin was similarly sutured closed.
Implantation of Radiotelemeter and Jugular
Venous Catheter
A midline incision was made in the skin of the neck region, and the
underlying glandular tissue was separated by blunt dissection to
expose the muscles of the neck. The left carotid artery was exposed
by blunt dissection and carefully separated from the vagus nerve.
Three 6-0 silk sutures (Deknatel, Braintree Scientific) were passed
underneath the artery with the rostral-most suture occlusively tied
around the vessel. The middle and caudal-most sutures were loosely
tied. Rostral and caudal sutures were retracted and a small incision
made in the artery for introduction of the radiotelemeter catheter
(Model TA11-PAC10, Data Sciences International). The catheter
was inserted 10 mm and sutured in place. The telemeter body was
tunneled to a subcutaneous pocket along the right flank.
The right jugular vein was exposed by blunt dissection, and 2
pieces of 6-0 silk suture were threaded around the vessel. The rostral
suture was tied to completely occlude the vessel, and the caudal
suture was used for temporary retraction. The vein was then
catheterized with heat-stretched rena-pulse tubing (RPT-040, Braintree Scientific); the catheter was advanced ⬇8 mm into the vessel
and secured to the surrounding tissues by tying the 6-0 sutures
around the catheter and also application of an adhesive containing
cyanoacrylate (SuperGlue Gel, Loctite). The mouse was turned onto
the left side, and a 13G stainless-steel needle was used to tunnel the
catheter from the neck to its exit point in the midscapular region. The
neck incision was closed with discontinuous sutures (5-0 Polysorb,
Tyco Healthcare); with the animal in the prone position, a small
subcutaneous button (Instech Solomon) covered in Dacron material
was threaded onto the venous catheter and secured under the skin
with sutures. A stainless-steel spring was threaded over the venous
catheter and secured to the skin button to protect the catheter.
The polyethylene pedestal of the electrode was then secured to the
underlying muscle with surgical adhesive (Vetbond, 3M) with
overlying skin sutured over the flange for further support. Antibiotic
ointment was applied to all of the incisions (Triple Antibiotic
Ointment, Fougera). Carprofen was administered (1 mg/kg SC) for
analgesia. The mice were placed in metabolic cages lined with wood
chip bedding and paper towel to recover. Electrode leads were coiled
outside the cage until the time of the experiment. The cages were
placed on a warm heat pad for the first 24 hours of recovery. Once
the mice recovered fully from anesthesia, food and water were
provided ad libitum.
Experimental Setup
A stainless-steel top antivibration table was electrically grounded
and fitted with a simple Faraday cage constructed from a wooden
frame and aluminum screen mesh. Forty-eight to 72 hours postsurgery, the home cage housing the mouse was placed on a radiotelemetry receiver (PhysioTel Receiver, model RPC-1, Data Sciences
International) within the Faraday cage. This receiver was connected
to a Pressure Output Adapter (model R11CPA, Data Sciences
International), which was, in turn, connected to a PowerLab data
acquisition system for recording of blood pressure (LabChart 7
software, ADInstruments). To record RSNA, the electrode leads
were uncoiled and the pin connectors were plugged into complementary female pin connectors (brass with gold plating, A-M Systems
Inc), which had been soldered previously to the ends of paired,
4
Hypertension
September 2012
MAP (mmHg)
20
0
-20
6
4
2
Figure 3. Representative recording of
systemic arterial blood pressure, renal
sympathetic nerve activity (RSNA), and
integrated RSNA in 2 conscious mice 48
to 72 hours postsurgery, while (A)
actively grooming or (B) eating. Block
arrow denotes commencement of activity from rest.
0
MAP (mmHg)
160
140
120
100
80
60
RSNA (μV)
B
20
Integration (μVs)
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Integration (μVs)
160
140
120
100
80
60
RSNA (μV)
A
0
-20
6
4
2
0
1 sec
shielded PVC insulated cable (PVC Audio Connection Cable, 32
AWG, Belden). The opposite ends of this paired cable were soldered
to banana plugs, which were connected to a preamplifier (⫻10
amplification, model 4002, Dagan Corporation). The ground wire
was connected to the ground pole of this preamplifier. The preamplifier was connected to a differential amplifier (EX4-400, Dagan
Corporation), and the nerve signal was amplified (⫻10 000) and
filtered (low cut, 100 Hz; high cut, 1000 Hz). Signals were displayed
and recorded on a PC online, simultaneously with blood pressure
data.
pressure and RSNA were recorded for 2 to 5 minutes, at which time
a bolus of phenylephrine (20 ␮g/g of body weight in 25 ␮L of saline)
was similarly administered in the infusion line. Blood pressure and
RSNA were recorded for an additional 10 to 15 minutes. During this
procedure, mice were quiet and still. To verify the postganglionic
nature of the nerve signal, we also injected the ganglionic blocker
hexamethonium (50 ␮g/g IV in 25 ␮L of saline). The background
noise for the nerve activity trace was estimated from the residual
activity remaining after ganglionic blockade with hexamethonium or
during postmortem recording of RSNA for 30 minutes after the mice
were euthanized with an overdose of isoflurane.
Experimental Protocol
Mice were unrestrained and housed in their home cages with free
access to food and water at all times. After surgery, mice were
housed in the same temperature- and humidity-controlled room in
which RSNA recording was to take place. At the time of the
experiment (48 hours after surgery), 30 minutes of stabilization was
allowed before recording 1 hour of baseline blood pressure and
RSNA data. At this time, a bolus of sodium nitroprusside (2.5 ␮g/g
of body weight in a volume of 25 ␮L of saline) was slowly
administered in the infusion line followed by 50 ␮L of saline. Blood
Data Analysis
LabChart 7 software was used to analyze raw blood pressure and
RSNA traces. Blood pressure and RSNA data were recorded simultaneously using LabChart 7 software and a PowerLab data acquisition system at 1000 samples per second. Although we used Data
Sciences International radiotelemeters to measure blood pressure, we
used a Blood Pressure Output Adapter with Ambient Pressure
Reference (models R11CPA and APR-1, Data Sciences International) to feed the signal directly to the PowerLab system for
Hamza and Hall
140
120
100
80
60
40
140
120
100
80
60
40
30
20
RSNA (μV)
RSNA (μV)
30
10
0
10
20
10
0
10
D
MAP (mmHg)
MAP(mmHg)
20
20
C
140
120
100
80
60
40
140
120
100
80
60
40
RSNA (μV)
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
30
30
RSNA (μV)
5
B
MAP (mmHg)
MAP (mmHg)
A
RSNA in Conscious Mice
20
10
0
20
10
0
10
10
20
20
0.5 sec
Figure 4. Sequential representative traces of blood pressure and renal sympathetic nerve activity (RSNA) in 1 conscious, resting mouse
several days after surgical preparation, (A) 2 days, (B) 3 days, (C) 4 days, and (D) 5 days postsurgery.
simultaneous recording with RSNA. The raw trace was digitally
integrated and full-wave rectified using this software: Absolute
Integral was selected for the integral settings with a time constant
decay of 0.1 seconds.3 The integrated RSNA signal (in ␮V · s) was
analyzed for each portion of the experimental protocol, with ⱖ3
measurements taken for baseline and experimental periods. RSNA
was analyzed at the maximum blood pressure response to PE and
sodium nitroprusside. These individual measurements were averaged
for each portion of the experimental protocol to give a single value.
Quantification of the experimental responses of RSNA was achieved
by calculating the percentage change of RSNA from baseline, which
Table. Baseline Mean Arterial Pressure and Heart Rate Values in
Instrumented Mice Over 5 Consecutive Days Postsurgery
Animal ID
2d
3d
4d
5d
mm Hg
112
110
108
109
bpm
657
551
626
616
mm Hg
115
107
111
110
bpm
582
652
662
668
mm Hg
115
118
113
111
bpm
591
599
689
664
mm Hg
114
115
116
110
bpm
457
513
599
531
mm Hg
109
109
103
105
bpm
632
687
699
689
A
B
C
D
E
we designated as 100%.4 Statistical analysis of the response of
RSNA to sodium nitroprusside and phenylephrine was completed
with a Student t test; significance was accepted with P values ⬍0.05.
Results
Of the 12 mice instrumented for this study, all survived and
tolerated the surgical procedure well to give a survival rate of
100%. By 24 hours after successful surgery, these animals
showed no sign of pain or distress and clearly exhibited
normal curiosity, as well as typical grooming and feeding
behavior. Ten of these 12 mice showed a true, high-quality
RSNA signal 48 hours after surgery. The RSNA signal
remained 3 days after surgery in all 10 of the mice; however,
the number of animals with a recordable RSNA was reduced
to 7 (70%) and 5 (50%) by 4 days and 5 days postsurgery,
respectively. Although the remaining animals did not show a
reasonable RSNA signal because of excessive noise or
contamination by ECG signals, they were all in good health
until euthanization.
Mean arterial pressure for animals with viable RSNA was
116⫾2 mm Hg, with an average heart rate of 596⫾22 bpm
(n⫽10) 48 hours after surgery. A representative recording of
blood pressure and RSNA 48 hours after recovery shows
rhythmic bursts of RSNA clearly visible above the background noise (Figure 2). Renal sympathetic activity also
increased with activity as shown in mice that were actively
grooming or eating (Figure 3). RSNA was also clearly
discernable for ⱕ5 days after surgery in half of the mice
studied (Figure 4). Blood pressure and heart rate values for
these mice over the course of the 5 days was stable and did
6
Hypertension
September 2012
MAP (mmHg)
A
150
100
50
30
RSNA (μV)
25
20
15
10
B
150
100
50
Figure 5. Representative recording of
blood pressure and renal sympathetic
nerve activity (RSNA) in a conscious
mouse at rest during (A) baseline; (B)
administration of sodium nitroprusside;
and (C) administration of phenylephrine.
30
RSNA (μV)
25
20
15
10
C
MAP (mmHg)
5
150
100
50
30
25
RSNA (μV)
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
MAP (mmHg)
5
20
15
10
5
1 sec
not differ from values obtained previously in our hands in
mice allowed 8 to 10 days of recovery from surgery (Table).5
Verification of true RSNA entrained with the arterial
baroreflex was completed by manipulating blood pressure
with sodium nitroprusside and phenylephrine; RSNA increased in response to the reduction in blood pressure induced
by sodium nitroprusside and was effectively silenced after
elevation of blood pressure with phenylephrine (Figure 5).
Reduction of blood pressure to 62⫾3 mm Hg with sodium
nitroprusside increased RSNA to 77⫾9% above baseline
(n⫽5; P⬍0.05); subsequent elevation of blood pressure to
137⫾6 mm Hg with phenylephrine reduced RSNA to
79⫾2% below baseline values (n⫽5; P⬍0.05; Figure 6).
Ganglionic blockade with hexamethonium eliminated the
RSNA signal, which was no different from the residual
postmortem recording (Figure 7).
Discussion
We have described and validated a method for directly
recording RSNA in conscious, freely moving mice. Animals
instrumented with a telemetric blood pressure recording
device, an indwelling catheter for intravenous infusion, and
exteriorized electrode leads for transmission of RSNA were
allowed to recover from surgery, undisturbed for 48 to 72
hours in their home cage. Subsequent experimental manipulations by the investigator were remote and did not perturb
the animals, which remained in the home cage with free
access to food and water at all times. Thus, this method
eliminated the physiological effects of anesthesia and sources
of stress to the animal, which unavoidably confound interpretation of sympathetic nerve activity data.
All of the mice instrumented for RSNA and blood pressure
recording exhibited normal behavioral characteristics, such as
Hamza and Hall
A
160
B
100
80
140
*
60
120
80
60
∆ RSNA (%)
AP (mmHg)
40
100
20
0
-20
-40
40
-60
20
0
-80
-100
*
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Figure 6. Quantitative response of blood pressure and renal
sympathetic nerve activity (RSNA) to sodium nitroprusside and
phenylephrine. A, Mean arterial pressure at baseline (; 116⫾2
mm Hg) and after sodium nitroprusside (s; 62⫾3 mm Hg) and
phenylephrine (e; 137⫾6 mm Hg). B, Simultaneous RSNA
response during sodium nitroprusside (s; 77⫾9%; n⫽5) or
phenylephrine (e; ⫺79⫾2%; n⫽5). RSNA is expressed as percentage of change from baseline, mean⫾SEM. *Significant difference from baseline (P⬍0.05).
eating, grooming, alertness, and curiosity, 24 hours after
surgical instrumentation. Although the recovery time required
for full restoration of normal cardiovascular parameters, such
as blood pressure, after implantation of telemetric recording
devices has been reported to be as long as 7 days,6 in our
hands normal blood pressure is restored much sooner, and the
values for blood pressure and heart rate reported here are
comparable to similarly instrumented animals in our hands
that were allowed to recover for 8 to 10 days.5,7 The use of
telemetric devices for blood pressure measurement is advantageous not only for the reduction of stress in our animals but
also for improved quality of the blood pressure signal with
preserved pulse pressures and reliable heart rate and blood
pressure values.8 The use of telemetry also eliminates the
need for periodic catheter flushing (and use of the heparinized
fluid that this often entails), as well as unnecessary perturbation of the animal during the experimental protocol. The
indwelling venous catheter and renal sympathetic nerve
electrode leads were also fully exteriorized at the time of
surgery rather than temporarily stored in a subcutaneous
pocket9 to avoid reanesthesia of the animals, however brief,
and any pain or distress directly before the time of the
experiment.
We have shown bursts of RSNA that are clearly visible
above background noise in a resting animal; this RSNA
subsequently increased during natural behavior, such as
eating and grooming, by the animal as expected.10,11 For
analytic purposes, we deliberately focused on periods of time
when the animal was quietly resting or sleeping and excluded
periods of movement to avoid possible misinterpretation of
experimental results because of natural increases in sympathetic tone during eating, physical activity, or even curiosity
and alertness by the animal. Other potential sources of
misinterpretation include signal contamination with ECG
pulses or excessive movement of the electrode leads (which
appears as a “wavering” baseline); these periods should also
be excluded from analysis. Importantly, we demonstrated that
RSNA in Conscious Mice
7
we have recorded true RSNA, which is entrained by the
arterial baroreflex, as shown by increased RSNA on reduction
of blood pressure with sodium nitroprusside and the virtual
elimination of RSNA on an ensuing phenylephrine-induced
rise in blood pressure. That the arterial baroreflex was clearly
intact in our model also directly addresses concerns regarding
the occlusive implantation of the blood pressure transmitter
catheter in the left carotid artery and any potential cardiovascular disturbance that this may cause. Elimination of the
nerve signal after ganglionic blockade also verified the
postganglionic nature of the signal.
Although our ability to obtain a reliable RSNA signal
diminished over several days after surgical preparation, we
were able to reliably record RSNA for 3 days in all of the
mice and for ⱕ5 days in some of the animals. This allows the
possibility to record multiple trials in 1 animal (ie, experimental versus time control) provided that the order of the
trials is randomized and baseline parameters are always
recorded at the start of each individual trial.12 Inherent
limitations to this technique that may preclude simple dayto-day and between-animal comparisons include natural differences in the number of isolated renal nerve fibers between
animals, as well as physical changes that occur over time,
such as shifting contact between the nerve bundle and
electrode and potential loss of viable nerve fires within the
nerve bundle. Nevertheless, we strive to improve this technique to be able to record viable RSNA over several weeks,
as has been demonstrated by the excellent work of Miki and
colleagues in the rat13,14 and briefly outlined by this group in
mice.15
The growing popularity of genetically modified mouse
models in biomedical research has driven a need for the
development of methods that enable reliable, feasible, and
accurate cardiovascular and autonomic phenotyping of these
animals. To our knowledge, there has been only 1 report of
measurement of sensory nerve activity in conscious mice.9 In
this study, bladder sensory nerve activity was recorded, and
the mice were subjected to anesthesia and surgical exteriorization of indwelling catheters immediately before the experiment and were placed in physical restrainers for the duration
of the protocol.9 Although the mild anesthesia and surgical
manipulation before the recording are not ideal and may not
impact the final results, restraining the rodents for the
duration of the recording is a known source of stress. In
addition, only afferent rather than intact nerve activity, which
would include efferent traffic, was measured,9 in contrast to
our intact renal nerve preparation. To date, examination of the
contribution of the sympathetic nerves to various aspects of
physiology and disease in mice has been exclusively completed in anesthetized models. Careful examination of the
literature reveals a plethora of types of anesthesia (injectable,
inhalational), anesthetic combinations, and doses used in
mice. One factor in common with these studies is either the
complete absence of blood pressure measurement or reporting of low blood pressures,2 often despite careful maintenance of core body temperature and artificial cardiovascular
support with intravenous fluid infusion (saline or volume
expanders) or other measures, such as direction of a stream of
oxygen to the animal’s nose.16–21 When attempting to eluci-
Hypertension
RSNA (μV)
A
MAP (mmHg)
8
September 2012
150
100
50
0
10
0
-10
-20
Integration
(μVs)
-30
1.0
0.8
0.6
0.4
0.2
0.0
150
100
50
0
Integration
RSNA (μV)
(μVs)
10
0
-10
-20
-30
1.0
0.8
0.6
0.4
0.2
0.0
RSNA(μV)
MAP(mmHg)
C
Integration
(μVs)
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
MAP (mmHg)
B
150
100
50
0
10
0
-10
-20
-30
1.0
0.8
0.6
0.4
0.2
0.0
Figure 7. Representative trace of blood pressure and renal sympathetic nerve activity (RSNA) at (A) baseline, (B) immediately after ganglionic blockade with hexamethonium (50 ␮g/g), and (C) postmortem.
date the role of the autonomic nervous system in various
mouse models, a physiologically normal blood pressure is
imperative. Blood pressure and autonomic tone are intimately
linked, with increases or decreases in blood pressure resulting
in concomitant alterations in sympathetic tone. Some types of
anesthesia, however, may markedly reduce sympathetic activity. Although these experiments in anesthetized mice
typically compare all experimental values to a recorded
baseline nerve activity, it may be difficult to study factors that
result in subtle changes in neural tone considering the altered
state of the autonomic system.
Although refined surgical skill is necessary for the preparation of mice for conscious nerve recordings, use of this
method avoids the limitations of several indirect assessments
of autonomic control. Measurement of plasma catecholamine
levels in mice is hampered by the wide variation in catecholamine levels in this species, as well as changes related to
sampling from anesthetized animals or the route of blood
sampling (ie, tail versus indwelling catheter).22 Akin to
plasma catecholamine levels, use of pharmacological agents
to block the autonomic nervous system reveals global but not
discrete contributions of autonomic tone.1 Although spectral
analysis of blood pressure and heart rate variability is
commonly used to estimate autonomic tone in humans and is
now often used for rodents, it appears that this may not be as
useful in the evaluation of autonomic balance in mice.22
Hamza and Hall
Direct measurement of nerve activity in conscious mice
comfortable at rest preserves the natural state of the animal as
much as possible and allows for the evaluation of specific
populations of nerves to different physiological phenomena.
Perspectives
It is now possible to routinely measure RSNA directly in
conscious mice chronically instrumented for telemetric measurement of blood pressure and intravenous infusion. The
construction and implantation of the RSNA electrode involve
materials that are easily accessible and relatively inexpensive
compared with telemetric nerve recording systems designed
for rats and larger animals but which are not presently
available for mice. By avoiding anesthesia and restraint, mice
with physiological blood pressures in a comfortable, unstressed environment will undoubtedly yield meaningful
measures of autonomic control in this species and open many
avenues of research using transgenic mouse models.
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Sources of Funding
This research was supported by the National Heart, Lung, and Blood
Institute grant PO1HL-51971. S.M.H. is supported by a postdoctoral
fellowship award from the Canadian Institutes for Health Research.
Disclosures
None.
References
1. Young CN, Davisson RL. In vivo assessment of neuro-cardiovascular
regulation in the mouse: principles, progress and prospects. Am J Physiol
Heart Circ Physiol. 2011;301:H654–H662.
2. Kass DA, Hare JM, Georgakopoulos D. Murine cardiac function: a
cautionary tail. Circ Res. 1998;82:519–522.
3. DeBeck LD, Petersen SR, Jones KE, Stickland MK. Heart rate variability
and muscle sympathetic nerve activity response to acute stress: the effect
of breathing. Am J Physiol Regul Integr Comp Physiol. 2010;299:
R80–R91.
4. Krowicki ZK, Kapusta DR. Microinjection of glycine into the hypothalamic paraventricular nucleus produces diuresis, natriuresis, and inhibition of central sympathetic outflow. J Pharmacol Exp Ther. 2011;337:
247–255.
5. do Carmo JM, da Silva AA, Cai Z, Lin S, Dubinion JH, Hall JE. Control
of blood pressure, appetite, and glucose by leptin in mice lacking leptin
receptors in proopiomelanocortin neurons. Hypertension. 2011;57:
918–926.
RSNA in Conscious Mice
6. Brockway BP, Mills PA, Azar SH. A new method for continuous chronic
measurement and recording of blood pressure, heart rate and activity in
the rat via radio-telemetry. Clin Exp Hypertens A. 1991;13:885–895.
7. Tallam LS, da Silva AA, Hall JE. Melanocortin-4 receptor mediates
chronic cardiovascular and metabolic actions of leptin. Hypertension.
2006;48:58–64.
8. Van Vliet BN, Chafe LL, Antic V, Schnyder-Candrian S, Montani JP.
Direct and indirect methods used to study arterial blood pressure.
J Pharmacol Toxicol Methods. 2000;44:361–373.
9. Zvara P, Wright AJ, Roach K, Ursiny M, Shapiro B, Dagrosa LM, Nelson
MT, Heppner TJ. A non-anesthetized mouse model for recording sensory
urinary bladder activity. Front Neurol. 2010;1:127.
10. O’Hagan KP, Bell LB, Mittelstadt SW, Clifford PS. Effect of dynamic
exercise on renal sympathetic nerve activity in conscious rabbits. J Appl
Physiol. 1993;74:2099–2104.
11. Matsukawa K, Ninomiya I. Changes in renal sympathetic nerve activity,
heart rate and arterial blood pressure associated with eating in cats.
J Physiol. 1987;390:229–242.
12. Smith FG. Techniques for recording renal sympathetic nerve activity in
awake, freely moving animals. Methods. 2003;30:122–126.
13. Miki K, Kosho A, Hayashida Y. Method for continuous measurements of
renal sympathetic nerve activity and cardiovascular function during
exercise in rats. Exp Physiol. 2002;87:33–39.
14. Yoshimoto M, Miki K, Fink GD, King A, Osborn JW. Chronic angiotensin II infusion causes differential responses in regional sympathetic
nerve activity in rats. Hypertension. 2010;55:644–651.
15. Yoshimoto MaM, Yoshimoto MaM. Measurement of renal sympathetic
nerve activity in freely moving mice. J Physiol. 2004;560D:1.
16. Alfie ME, Sigmon DH, Pomposiello SI, Carretero OA. Effect of high salt
intake in mutant mice lacking bradykinin-B2 receptors. Hypertension.
1997;29:483–487.
17. Dietz JR, Landon CS, Nazian SJ, Vesely DL, Gower WR Jr. Effects of
cardiac hormones on arterial pressure and sodium excretion in NPRA
knockout mice. Exp Biol Med (Maywood). 2004;229:813–818.
18. Zhang W, Li JL, Hosaka M, Janz R, Shelton JM, Albright GM, Richardson JA, Sudhof TC, Victor RG. Cyclosporine A-induced hypertension
involves synapsin in renal sensory nerve endings. Proc Natl Acad Sci
U S A. 2000;97:9765–9770.
19. Szczesny G, Veihelmann A, Massberg S, Nolte D, Messmer K.
Long-term anaesthesia using inhalatory isoflurane in different strains of
mice-the haemodynamic effects. Lab Anim. 2004;38:64–69.
20. Tank J, Obst M, Diedrich A, Brychta RJ, Blumer KJ, Heusser K, Jordan
J, Luft FC, Gross V. Sympathetic nerve traffic and circulating norepinephrine levels in RGS2-deficient mice. Auton Neurosci. 2007;136:
52–57.
21. Zuurbier CJ, Emons VM, Ince C. Hemodynamics of anesthetized ventilated mouse models: aspects of anesthetics, fluid support, and strain. Am
J Physiol Heart Circ Physiol. 2002;282:H2099–H2105.
22. Janssen BJ, Smits JF. Autonomic control of blood pressure in mice: basic
physiology and effects of genetic modification. Am J Physiol Regul Integr
Comp Physiol. 2002;282:R1545–R1564.
Novelty and Significance
What Is New?
●
●
This is the first description of renal sympathetic nerve recording in
conscious, unrestrained mice.
Viable recordings are possible for 5 days in a single mouse, which
facilitates a powerful scientific approach of conducting a time control
and experimental treatment in the same animal.
What Is Relevant?
●
The autonomic nervous system is implicated in the development of
hypertension and is a focus of investigation in this field; this method is
a valuable and accessible tool for future research in hypertension.
9
Summary
Direct recording of renal nerve activity is possible in conscious,
chronically instrumented and unrestrained mice.
Direct Recording of Renal Sympathetic Nerve Activity in Unrestrained, Conscious Mice
Shereen M. Hamza and John E. Hall
Hypertension. published online July 30, 2012;
Downloaded from http://hyper.ahajournals.org/ by guest on June 15, 2017
Hypertension is published by the American Heart Association, 7272 Greenville Avenue, Dallas, TX 75231
Copyright © 2012 American Heart Association, Inc. All rights reserved.
Print ISSN: 0194-911X. Online ISSN: 1524-4563
The online version of this article, along with updated information and services, is located on the
World Wide Web at:
http://hyper.ahajournals.org/content/early/2012/07/30/HYPERTENSIONAHA.111.186577
Permissions: Requests for permissions to reproduce figures, tables, or portions of articles originally published
in Hypertension can be obtained via RightsLink, a service of the Copyright Clearance Center, not the Editorial
Office. Once the online version of the published article for which permission is being requested is located,
click Request Permissions in the middle column of the Web page under Services. Further information about
this process is available in the Permissions and Rights Question and Answer document.
Reprints: Information about reprints can be found online at:
http://www.lww.com/reprints
Subscriptions: Information about subscribing to Hypertension is online at:
http://hyper.ahajournals.org//subscriptions/