Powerlab 4 35

Surface EMG was recorded from the first dorsal interosseous (FDI) muscle of the right hand. Wet gel 10 mm Ag/AgCl electrodes were applied in a belly-tendon montage, grounded on the ulnar styloid process. EMG signals were amplified (× 1000; Bio Amp–ADInstruments New Zealand), bandpass filtered (1 Hz–2 kHz), digitized (10 kHz; PowerLab 4/35; ADInstruments), and recorded (LabChart 8.0—ADInstruments) from 200 ms before to 300 ms after TMS pulses^{26 (link)}.

The extent of the abdominal contractions (VMR) due to colorectal distension was measured by performing electromyography (EMG) on the abdominal muscles and used as a quantitative measure of visceral sensitivity in the rats. Two EMG electrodes were sutured into the external oblique abdominal muscles of the animals under anesthesia and exteriorized dorsally [25 (link)]. VMR assessment was carried out under light anesthesia (2% isoflurane). A lubricated latex balloon (length: 4.5 cm) was attached on an embolectomy catheter and connected to a water-filled syringe used to perform colorectal distension (CRD). The syringe was used to fill the balloon placed into the colon with increasing volumes of water (0.5, 1, 2, and 3 mL, referred to as the distension volume). After colorectal stimulation, the EMG signal was recorded, amplified and filtered (Animal Bio Amp, ADInstruments, Colorado Springs, CO, USA), digitized (PowerLab 4/35, ADInstruments), analyzed, and quantified using LabChart 8 (ADInstruments). To quantify the VMR magnitude under each distension volume, the area under the curve (AUC) immediately before distension (30 s) was subtracted from the AUC during balloon distension (30 s), and the responses were expressed as a percentage increase from the baseline. The time elapsed between two consecutive distensions was 5 min. The entire measurement process lasted about 25 min.

CMG was performed with PBS infusion (25 μl/min)⁶⁴ . Mice were anesthetized by a subcutaneous injection of urethane (1.4 g/kg from 250 mg ml⁻¹ solution in PBS) 30–60 min before surgery. At time of surgery, the mouse was further anesthetized with continuous flow isoflurane (3% induction, 1.0% maintenance). Once the pedal reflex was absent, a 1-cm midline abdominal incision was performed. Flame-flanged polyethylene-50 tubing sheathing a 25Gx1.5 in. needle was implanted through the dome of the bladder, then sutured in place (8-0 silk purse string). The incision site was sutured around the tubing using sterile 5-0 silk, and mice were then placed into a Bollman mouse restrainer for 30–60 min stabilization. The catheter was connected to a pressure transducer (and syringe pump by side arm) coupled to data-acquisition devices (WPI Transbridge and AD Instruments Powerlab 4/35) and a computerized recording system (LabChart software). Bladder filling then commenced, after which voiding occurred naturally through the urethra. Repeated voiding cycles were assessed for change of voiding interval (time between peak pressures), basal pressure (minimum pressure after voiding), threshold pressure (pressure immediately before onset of voiding contraction), peak pressure (maximum voiding pressure minus basal pressure) and compliance (volume, in μl required to increase pressure by 1 cm H₂O).

Surface Electromyography (sEMG) was recorded using bipolar Ag-AgCL electrodes. Two electrodes were placed 2 cm apart on the mid belly of the ECR, with a ground strap placed around the wrist as a common reference for electrodes. Cables were fastened with tape to prevent movement artifact. The skin was prepared prior to electrode placement to ensure a clear signal. sEMG signals were amplified (×100–1000), bandpass filtered (high pass at 13 Hz, low pass at 1000 Hz), digitized online at 2 kHz for 500 ms, recorded and analyzed using PowerLab 4/35 (ADInstruments, Bella Vista, NSW, Australia).

Mouse vitals (heart rate, pulse distension, breath rate, breath distension, pO2) were monitored and recorded in anesthetized mice with a MouseOx Plus suite (Starr Life Sciences) with a paw sensor. For ECG measurement, three needle electrodes were placed subdermally: on both sides of the upper chest cavity near the armpits and lead I in the lower abdomen. Signals were amplified (BioAmp) and digitized using a PowerLab 4/35 and were analyzed in LabChart software (ADInstruments).

After the final of training period, the animals were anesthetized with ketamine (50 mg/kg) and xylazine (10 mg/kg) for left ventricle catheterization. Briefly, the right common carotid artery was separated from connective tissue and catheterized with a fluid-filled polyethylene catheter (PE50). The catheter was connected to a pressure transducer (FE221 Bridge amp, ADInstruments, Australia) and a digital system (Powerlab 4/35, ADInstruments, Australia). After arterial systolic and diastolic blood pressures were recorded, the catheter was advanced into the left ventricle to obtain the following measurements: heart rate (HR), left ventricular systolic pressure (LVSP), end-diastolic pressure (LVEDP), and the maximum rate of pressure rise (+dP/dt) and fall (-dP/dt). It was not possible to measure other parameters related to cardiac function such as cardiac output and ejection fraction because we not evaluate the ventricular volume. However, other studies have been demonstrated that LVEDP presents as an important parameter for the assessment of ventricular function, and his increase is associated with ventricular dysfunction. [21] The heart, soleus muscle, abdominal fat, uterus and a lung were removed immediately after hemodynamic evaluation and weighed.