Mar 20, 2026
Single-Electrode, Bidirectional Control of Heart Rate via Vagus Nerve Modulation in Rat Model
- Shane Bender1,2
- 1Case Western Reserve University;
- 2The MetroHealth System
Protocol Citation: Shane Bender 2026. Single-Electrode, Bidirectional Control of Heart Rate via Vagus Nerve Modulation in Rat Model. protocols.io https://dx.doi.org/10.17504/protocols.io.8epv5yo56l1b/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 12, 2026
Last Modified: March 20, 2026
Protocol Integer ID: 313177
Keywords: vagus nerve modulation in rat model electrical stimulation, vagus nerve modulation, rat model electrical stimulation, bidirectional control of heart rate, electrical nerve block, frequency electrical nerve block, vagus proximal to the electrode, activity on the nerve, side rat vagus nerve, heart rate, somatic nerve, stimulation parameter, precise control of an autonomic system, stimulation, single nerve, target nerve, single electrode, nerve, distal electrode, perturbing stimulus ramp, autonomic system, proximal electrode, oscillations in the heart rate, electrode, bimodal neuromodulation, fuzzy logic controller, powerful treatment tool against autonomic dysregulation, loop fuzzy logic controller, vasovagal syncope
Funders Acknowledgements:
NIH NHLBI
Grant ID: R01HL150136
NIH NHLBI
Grant ID: R01HL162921
NIH NINDS (SPARC)
Grant ID: U41NS129436
Abstract
Electrical stimulation of somatic nerves has long been used as a treatment method for a wide range of diseases by increasing activity on a target nerve. Electrical nerve block is an emerging therapy that can provide the same targeted treatment by decreasing activity on the nerve. Here, we demonstrate that both of these techniques can be applied synergistically via a single electrode to achieve precise control of an autonomic system. Two electrodes were placed on the right-side rat vagus nerve. The vagus proximal to the electrodes was crushed, and the left side was cut to isolate the system. The proximal electrode was used to give a perturbing stimulus ramp (0 Hz → 30 Hz → 0 Hz) over 10 minutes to roughly mimic the vagal activity seen in an episode of vasovagal syncope. The distal electrode was used to apply either stimulation or kilohertz-frequency electrical nerve block (KHFAC) to the vagus to keep the heart rate at a specified setpoint. The stimulation parameters were decided by a closed-loop fuzzy logic controller. Three different gains for the controller were tried. While each gain showed success in controlling the heart rate, lower gain was sometimes not responsive enough for effective control, and high gain was seen to induce oscillations in the heart rate; a medium gain was seen to be effective without either of these issues. This demonstrates that a single electrode can deliver bimodal neuromodulation of a single nerve, providing a powerful treatment tool against autonomic dysregulation.
Materials
Animals:
Male Sprague-Dawley rats, 7–11 weeks old, 400–600 g, RRID:RGD_734476
Stimulation & blocking hardware
COSMIIC GAMMA V1 (COSMIIC.org, Cleveland, OH, USA), or any pair of computer-controllable constant-current stimulators capable of 10 kHz square waves.
The GAMMA V1 contains two Saturn2 integrated circuits (Cirtec Medical, Brooklyn Park, MN, USA) controlled by an STM32WB microcontroller (STMicroelectronics, Plan-les-Ouates, GE, CH)
Custom bipolar platinum cuff electrodes, 1–1 mm windows, or any electrode fitting the rat vagus nerve
ECG recording
CED 1902 Isolated Amplifier + 1902-10 headstage (CED, Milton, EN, UK)
CED Power 1401 data acquisition unit (CED, Milton, EN, UK)
Spike2 v10 software (CED, Milton EN UK; RRID:SCR_000903)
Controller hardware & software
NI USB-6363 BNC data acquisition device (NI, Austin, TX, USA), or any LabVIEW-compatible DAQ
LabVIEW 2024, Windows PC (NI, Austin, TX, USA)
Ventilation & surgical equipment
Harvard Apparatus Model 683 ventilator (Harvard Apparatus, Holliston, MA ,USA), or any rodent ventilator (max 5 mL stroke volume, 60 breaths/min)
14-gauge angiocatheter (16-gauge alternative)
Laryngoscope
Large forceps (for tongue retraction during intubation)
Fine forceps (nerve isolation and crushing)
Intubation board
Heated surgical pad
Anesthesia induction chamber
Anesthesia nose cone
22-gauge needle + syringe (euthanasia)
Razor/clippers (skin preparation)
Drugs & reagents
Isoflurane (3–5% for induction; maintenance; 5% ramp-up for euthanasia)
Sodium pentobarbital (or institution-approved euthanasia agent)
Troubleshooting
Animal Procurement
Procure animals for experimentation
House according to institutional vivarium standards prior to experimentation.
This study used male Sprague-Dawley rats, approximately 7–11 weeks old, weighing 400–600 g
Assemble Equipment
Stimulating/blocking source: open-source COSMIIC project's GAMMA module
Any pair of computer-controllable constant-current stimulators capable of 10kHz square waves can be used with modest changes to the software
Stimulating/blocking electrodes: custom-fabricated bipolar platinum electrodes, 1x1 mm windows
Any electrode that fits on the rat vagus nerve can be used
ECG recording:
Amplifier: CED 1902 Isolated Amplifier with 1902-10 headstage
Data Recording: CED Power 1401 and Spike2v10 software
Recording system of your choice can be substituted
Controller Implementation: NI USB-6363 BNC
Any LabVIEW-compatable data acquisition device can be used with modest changes to the software
Controller Software: Windows PC with LabVIEW 2024
Replacement software should follow a similar data flow:
ECG acquisition → Heart Rate Calculation → Fuzzy Logic Controller → Output to current source
Ventilator: Harvard Apparatus Model 683
Any equivalent rodent vetilator will suffice (max 5 mL stroke volume, 60 breaths per minute)
Surgical Preparation
Induce Rat Anesthesia
Place rat in induction chamber
Deliver isoflurane at 3-5% for 2-5 minutes, until rat is unconcious
Weigh rat and place in anesthesia nose cone on heated pad
Prepare animal skin while anesthesia gets deeper
Continuously monitor rat vital signs until appropriate anesthesia depth for intubation is reached
Minimum breathing rate is 20 breaths per minute
Minimum heart rat is 200 beats per minute
While leaving the animal at 3-5% isoflurane, shave the ventral cervical neck, over the trachea
Intubate the animal
Ensure the animal is sufficiently sedated, such that they will not wake up without isoflurane for the duration of the intubation process, ~1-2 minutes
Lay out your intubation board, large forceps, and laryngoscope, and 14-gauge angiocatheter (16-gauge may be used as well, but may result in air escaping through the larynx in larger animals
Take the rat out of the nose cone, and lay on your intubation board. Secure nose/upper jaw in place
Use one hand to grab the tongue with the large forceps, and use your other hand to place the laryngoscope in the mouth, and visualize the larynx
Once you can see the opening to the trachea, put down the forceps and use that hand to insert the angiocatheter into the trachea
Connect the angiocatheter to a ventillator, hook the ventilator up to the anesthesia, and begin ventillation at 4-5 mL stroke volume, 60 breaths per minute.
Transfer rat back to heated pad in a supine position, being careful not to pull out the angiocatheter
Vagus Exposure
Expose the vagus on each side
Make two rostral-caudal incisions in the skin, one on each side of the trachea over the vagus nerves
Connect the incisions across to form an "H" shape
Blunt dissect to isolate the vagus nerves, often bundled with the carotid artery and jugular vein
Cuff the Vagus
Place two cuff electrodes on the right vagus nerve, leaving space between them and ensuring that they do not pull on or kink the nerve
Isolate the system and remove tonic activity
Use fine forceps to crush the right side nerve proximal to the electrodes. This will keep the nerve structurally intact to prevent the nerve from getting pulled out of the electrodes
The left vagus can be similarly crushed, or cut completely
Calibration
Calibrate stimulation slope for the control (distal) electrode
Find an amplitude for your stimulation that elicits a maximal heart rate decrease (typically tested at 30 Hz, 50 μs per phase, currents 0.1 mA to 5 mA peak-to-peak expected)
If AV block is seen (heart rate drops roughly in half suddenly), back off of the amplitude until normal conduction resumes
Record the baseline heart rate with no stimulation
Increase the control electrode stimulation in steps of 1 Hz, and plot the change in heart rate
Once the heart rate stops changing, fit a linear regression to the data and obtain the slope in bpm/Hz
Find the stimulation amplitude for the perturbation (proximal) electrode
Using the same procedure as above, find the maximal stimulation amplitude at 30 Hz, 50 μs per phase for the proximal electrode
Calibrate the block start and slope for the control (distal) electrode
Apply maximal stimulation to elicit bradycardia on the perturbation electrode
Increase the amplitude of a 10 kHz square wave blocking waveform on the control electrode until you see the heart rate start to increase towards baseline. This is you block starting current.
Continue increasing the amplitude of the block, and plotting the resultant heart rate
Once the heart rate reaches the baseline heart rate (pre-perturbation-stimulation), fit a line to your data and record the blocking slope in bpm/mA
Enter these calibration values into the controller
Perform Experiment
Begin recording data for each trial
Before using the controller, briefly apply 30 Hz stimulation to the control electrode (~30 s)
Compare the stimulation bradycardia heart rate (0%) and the post-stimulation baseline heart rate (100%)
Your controller setpoint will be halfway between these values (50%)
Allow for at least 2 minutes of baseline recording before turning controller on
Set the desired controller Gain (2x, 5x, or 10x)
Turn the controller on and allow the controller to apply stimulation to the control electrode to bring the heart rate down to the 50% setpoint for 2 minutes
After two minutes of the controller being on, start ramping the stimulation frequency on the perturbation electrode
Ramp up to 30 Hz in 5 minutes, then ramp back down to 1 Hz in another 5 mintues
After the perturbation ramp is done, allow the controller to run for an additional 2 minutes
At the end of the trial, turn off the controller, and let the heart rate return to baseline
Repeat this protocol for the other two gain values
End of Experiment
To euthanize the animal, turn your anesthesia up to 5%, and inject sodium pentobarbital into the heart with a 22-gauge needle
This should be substituted with whatever euthanasia method your institution has approved for terminal experiments