Male 8- to 12-week-old mice weighing 22–28 g underwent transverse aortic constriction (TAC) or sham surgery as previously described (51 (link)). Briefly, mice were anesthetized with 4% isoflurane and intubated with a 20-gauge cannula. Ventilation was initiated and continued under 2.5% isoflurane at 135 breaths per minute by a small-animal TOPO ventilator (Kent Scientific). The aortic arch was exposed via a left thoracotomy and by careful separation of the thymus. A constriction of the transverse aorta was generated by tying of a 6-0 Ethilon ligature against a 27-gauge blunt needle around the aorta between the brachiocephalic and left common carotid arteries. Promptly the needle was removed, lungs were inflated, and the chest and skin were closed by 5-0 polypropylene suture. The animal was removed from ventilation, and sustained-release buprenorphine (subcutaneous, 0.05 mg/kg) and 0.9% saline (i.p., 0.2 mL) were administered for analgesia and hydration, respectively. Sham-operated mice underwent all the same procedures as TAC mice excluding the constriction of the aorta. ATPIF1 F/F and cKO mice were randomly assigned to sham or TAC surgery. Combined mortality (acute and chronic) was less than 10%, and all mice surviving surgery were included in the analysis. Sample size was determined by power analysis based on our previous experiments.
Topo ventilator
Manufactured by Kent Scientific
Sourced in United States
The TOPO ventilator is a compact, portable device designed for respiratory support in medical settings. It provides controlled mechanical ventilation to patients who require assistance with breathing. The TOPO ventilator is capable of delivering precise volumes and pressures to meet the patient's respiratory needs.
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8 protocols using topo ventilator
1
Transverse Aortic Constriction in Mice
2
Cardiac Hemodynamics in DOX-Treated Mice
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BALB/c mice weighing 22–25 g (Jackson Laboratories, Bar Harbor, ME) were separated into DOX, RBC-DOX, and control groups (n = 6 per group). The DOX group received 5 mg/kg IV doses of DOX, the RBC-DOX group received 5 mg/kg equivalent of DOX and the control group received a vehicle infusion of saline. The animals were treated at days 0, 3, 6, and 9. Hemodynamic measurements of heart rate (HR), mean arterial pressure (MAP), and systemic vascular resistance (SVR) were completed 5 days after final administration. Animals were anesthetized using sodium pentobarbital (40 mg/kg IP). Animal preparation included: (i) left femoral artery catheterization, (ii) tracheotomy (polyethylene-90 tube), and (iii) left ventricle conductance catheter introduction through the right carotid artery. Animals were placed in the supine position on a heating pad to maintain core body temperature at 37 °C. Animals were mechanically ventilated (TOPO ventilator, Kent Scientific, CT) using room air (respiration rate of 90 breaths per minute; peak inspiratory pressure of 20 cmH2O). After instrumentation, volatile anesthesia (0.6%/vol Isoflurane, Drägerwerk AG Lübeck) was administered using a vaporizer connected to the ventilator. Deep of anesthesia was continually verified via toe pinch, if needed, isoflurane was increased by 0.1%/vol to prevent animal discomfort.
3
Transverse Aortic Constriction Mouse Model
4
Transverse Aortic Constriction in Mice
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PP2Cm KO mice and their littermate controls at the age of 10–12 weeks underwent TAC or sham surgery56 (link). Briefly, mice were anesthetized with an intraperitoneal injection of 130 mg/kg ketamine and 8.8 mg/kg xylazine in saline. Mice were intubated with 20 G cannula and ventilated 140 breaths per minute by small animal TOPO ventilator (Kent Scientific). The aortic arch was exposed via a left thoracotomy and by carefully separating the thymus. A constriction of the transverse aorta was generated by tying a 6-0 Ethilon ligature against a 27-gauge blunt needle around the aorta between the brachiocephalic and left common carotid arteries. Promptly the needle was removed and the chest and skin were closed by 5-0 polypropylene suture. The animal was removed from ventilation and kept on a heating blanket during recovery from anesthesia. SR buprenorphine (0.05 mg/kg) was administered subcutaneously for analgesia. Sham operated mice underwent all the same procedures as TAC mice excluding the constriction of the aorta. All mice were monitored every 12 h during the first 72 h post-surgery, followed by daily visits over the next 4 weeks.
5
Surgical Induction of Myocardial Infarction in Mice
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Male 8- to 12-week-old mice weighing 22–28 g underwent permanent left anterior descending artery ligation or sham operation as previously described (52 (link)). Briefly, mice were anesthetized with 4% isoflurane and intubated with a 20-gauge cannula. Ventilation was initiated and continued under 2 % isoflurane at 130 breaths per minute by a small-animal TOPO ventilator (Kent Scientific). A small skin cut (~1.2 cm) was made over the left chest. After dissection and retraction of the pectoral major and minor muscle, the fourth intercostal space was exposed. A small hole was made at the fourth intercostal space, and a mosquito clamp was used to open the pleural membrane and pericardium. With the clamp slightly open, the heart was smoothly and gently popped out through the hole. The left coronary artery was located, sutured, and ligated at a site about 3 mm from its origin using a 7-0 Ethilon silk suture. After ligation, the heart was immediately placed back into the intrathoracic space, followed by manual evacuation of air and closure of muscle and the skin, by means of the previously placed purse-string suture. Mice that did not survive the first 24 hours after the surgery were excluded from analysis. Sham-operated animals underwent the same procedure without coronary artery ligation.
6
Golden Syrian Hamster Cardiac Hemodynamic Protocol
7
Cardiac Function Assessment in Rats
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Studies were performed in 200–250g male Sprague-Dawley rats (Harlan Laboratories, Indianapolis, IN). Animal handling and care followed NIH Guide for Care and Use of Laboratory Animals and all protocols were approved by the UC San Diego Institutional Animal Care and Use Committee. Animals were initially anesthetized with isoflurane (Draegerwerk AG, Luebeck, Germany) at 5% to induce deep anesthesia. Anesthetic was subsequently reduced to 2.5% for the remainder of the experiment, and animals were placed in the supine position on a heating pad to maintain core body temperature at . Catheters were placed in the left femoral artery and left jugular vein. Animals were then instrumented with a 2F pressure-volume (PV) conductance catheter (SPR858, Millar Instruments, TX) inserted into the left ventricle (LV) via the right common carotid artery. For certain experimental protocols, a tracheostomy was performed, and animals were mechanically ventilated (TOPO ventilator, Kent Scientific, Torrington, CT).
8
Invasive Hemodynamic Monitoring in Hamsters
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Studies were performed in male Golden Syrian hamsters (80–90 g, Charles River Laboratories, Boston, MA, United States). The experimental protocol is similar to that reported in a previous study by our group (Ao-Ieong et al., 2017 (link)). Animals were initially anesthetized using sodium pentobarbital (40 mg/kg IP). Subsequently, the animals were implanted with left jugular vein and left femoral artery catheters. A LV conductance catheter was introduced through the right carotid artery. A tracheotomy was performed (polyethylene-90 tube), and animals were ventilated (TOPO ventilator, Kent Scientific, Torrington, CT, United States) at a respiration rate of 90 breaths/min and peak inspiratory pressure of 20 cmH2O to aid with breathing in the supine position. Animals were placed on a heating pad to maintain core body temperature at 37 °C and after instrumentation, volatile anesthesia (0.6%/vol Isoflurane, Drägerwerk AG, Lübeck, Germany) was administered using a vaporizer connected to the ventilator to preserve the depth of anesthesia during the experimental protocol. Depth of anesthesia was continually verified via toe pinch, and, if needed, isoflurane concentration was increased by 0.1%/vol to prevent animal discomfort. Experimental setup is presented in Figure 1C .
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