PWV was acquired from the cardiovascular pulse, distance between locations d, and transit time Δt for the pulse to travel distance d. After isoflurane anesthesia, the mice were placed supine, and distance d between the supraclavicular notch and the ankle of the left hindlimb was measured. Three acupuncture needle electrodes were inserted subcutaneously into the right and left forelimbs and the left hindlimb. The other ends of the electrodes were connected to an amplifier cable for electrocardiography (ECG) of lead II (Powerlab/8sp, BIO Amp, ADinstruments, Dunedin, New Zealand). The pulse oximeter (MouseOx, Starr Life Sciences, Oakmont, PA, USA) was placed on the left ankle. The transit time (Δt) was acquired using the pulse wave between the initial peak of the ECG R-wave and arrival peak of pulse oximeter wave (LabChart software v7, Adinstruments, New Zealand). PWV was calculated as follows: PWV = d (meters)/Δt (seconds).
Mouseox
Manufactured by Starr Life Sciences
Sourced in United States
MouseOx is a non-invasive monitoring system designed for small animals. It measures key physiological parameters such as heart rate, breathing rate, oxygen saturation, and body temperature. The device provides real-time data through a sensor that is placed on the animal's body.
Automatically generated - may contain errors
Lab products found in correlation
Labchart software v7, by ADInstruments (1 mentions)
Bio amp, by ADInstruments (1 mentions)
Powerlab 8sp, by ADInstruments (1 mentions)
Mp150, by Biopac (1 mentions)
I stat, by Abbott (1 mentions)
Stereotaxic frame, by Kopf Instruments (1 mentions)
Mousestat, by Kent Scientific (1 mentions)
Infrared thermometer, by Thermo Fisher Scientific (1 mentions)
Stereo lumar v12 fluorescence microscope, by Zeiss (1 mentions)
Micromotor drill, by Stoelting (1 mentions)
Eyfp wpre hgh, by Addgene (1 mentions)
52 protocols using mouseox
1
Measuring Pulse Wave Velocity in Mice
2
Inducing Moderate Hypothermia in Rats
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Temperatures were recorded using both a rectal probe and the epidural probe implanted at surgery. Within 5 min of trauma, the rats were placed on a warming pad and the epidural temperature was maintained at 37±1°C for 4 h. Subsequently, moderate hypothermia (epidural temperature, 33.0±0.3°C) was induced for 4 h as described previously.21 (link) Rats were transferred to the cooling system, comprised of a water blanket and pump (HTP WD-020 and HTP-1500, Adroit; Medical Systems Corp., Loudon, TN) with the pump reservoir filled with ice water. The rats were covered with a thermal blanket to stabilize their temperature. During this time, they were anesthetized with low-dose isoflurane (0.75%) via face mask, to prevent them from shivering or moving and dislodging the temperature probe. They reached the target epidural temperature for hypothermia within 30 min of starting cooling. Blood oxygen saturation and pulse rate (MouseOx, Starr, Life Sciences Corp, Holliston, MA), as well as respiratory rate, were monitored continuously and recorded every 30 min. After 4 h, the animals were gradually rewarmed at 1°C/h until they were normothermic. After completing the hypothermia treatment, the epidural probes were removed by gentle traction, and the rats were returned to their cages, where food and water were easily accessible.
3
Oxycodone Antinociception and Respiratory Depression
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Groups of 16 rats were immunized with OXY-dKLH or control dKLH vaccine containing 60 μg immunogen and 90 μg aluminum as Alhydrogel. Vaccine was administered intramuscularly every 3 weeks for a total of 4 doses. One week after the final vaccine dose baseline antinociception (prior to oxycodone administration) was assessed by placing rats on a 54°C hotplate and measuring the latency to respond, defined as the time until a response of hindpaw lick or jumping [18 (link)]. Trials were terminated at 60 sec if rats had not responded. Immediately following the hotplate trial a MouseOx (STARR Life Sciences Corp., Oakmont, PA) arterial oxygen saturation (SaO2) monitor was placed via neck collar and baseline SaO2 was measured. Rats then received oxycodone every 17 minutes s.c. so that their cumulative oxycodone dose at successive intervals was 1.1, 2.2, 4.5 and 9.0 mg/kg. This oxycodone dose range produced a wide range of antinociception, and the 9.0 mg/kg dose was the highest that could be administered without producing excessive respiratory depression. Fifteen minutes after each oxycodone dose antinociception and SaO2 was again measured; this took 2 minutes, resulting in the 17 minutes oxycodone dosing interval. Heart rate was obtained from the oximeter.
4
Arterial and Venous Catheterization for Physiological Monitoring
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Arterial and venous catheters (PE 50, Instech Laboratories, Inc., Plymouth Meeting, PA, USA) were inserted for monitoring blood pressure, blood collection and infusion of dyes and other fluids, as described [23 (link), 24 (link)]. Arterial blood samples were collected to measure hematocrit, pH, lactate, base excess (BE), creatinine, blood urea nitrogen (BUN), Na+, Cl-, and K+ (I-stat, Abbott, Chicago, IL, USA), syndecan-1, and viscosity. Plasma syndecan-1 was analyzed using a commercial ELISA kit for rats (Antibodies Online, Atlanta, GA, USA). Whole blood viscosity was measured at 37 °C and a shear rate of 225 sec-1 using a cone-plate viscometer (Brookfield LV, Middleboro, MA, USA). Total plasma protein was measured using a clinical refractometer (model 300005, Sper Scientific, Scottsdale, AZ, USA). Hemoglobin O2 saturation, and respiratory rate were continuously recorded (MouseOx, Starr Lifesciences, Oakmont, PA, USA), along with arterial pressure and heart rate using a data acquisition system (MP150, Biopac, Goleta, CA, USA).
5
Monitoring Peripheral Oxygen Saturation in Rats
6
Murine Anesthesia and Monitoring Protocol
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Anesthesia was initiated by 80 mg/kg pentobarbital injected i.p. The mouse was placed on a support heated by a flat pad (Fig. 1). After cannulation of the jugular vein by a catheter with an outer diameter of 0.61 mm (Fig. 2), anesthesia was continued intravenously with 40-60 mg methohexital-sodium per kg and h. A tracheotomy was performed and a tube inserted for artificial ventilation (Fig. 2). Active respiratory movements were abolished by paralysis with pancuronium (800 µg per kg supplemented i.p. every hour) and artificial ventilation was performed with a gas mixture of CO 2 (2.5 %), O 2 (47.5 %), and N 2 (50 %) at 120 strokes/min (120-160 µl/stroke depending on the weight). Rectal body temperature, heart rate (ECG: platinum wires of 0.3 mm diameter inserted into both forelegs and connected via a differential amplifier with an oscilloscope) and O 2 blood saturation (MouseOx ® , Starr Life Sciences, Oakmont, USA) were continuously monitored. Appropriate dose of anesthesia was initially established by the absence of corneal and pinna reflexes, later after paralysis, by falling body temperature and heart rate.
7
Anesthesia Induction and Monitoring for Rat Studies
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An induction chamber with 4–5% isoflurane (in 70% nitrogen and 30% oxygen) was used to induce light anaesthesia prior to intraperitoneal injections of urethane (i.p. 1.25 g/kg, divided into four doses delivered at 30-min intervals). Isoflurane was discontinued after the first urethane injection, and rats remained anaesthetized until euthanasia. Throughout the experiment, temperature was maintained at 36.5–37.5 °C with a thermostatically controlled warming pad. Pulse oximetry (MouseOx, STARR Life Sciences) was used to monitor heart rate, oxygen saturation, and breath rate.
8
Oxygen Saturation and Blood Chemistry in Mice
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Oxygen saturation was measured in adult mice on the day before the first tamoxifen injection by pulse oxymetry (MouseOx; Starr Life Sciences Corp.), and over the subsequent 11 days. The mouse was anesthetised by isoflurane mixed with medical air, and the sensor was placed on a hind limb. Blood chemistry parameters were obtained using blood harvested by cardiac puncture from anesthetised adult mice (10 days after the first tamoxifen injection) and analysed on a CC8+ cartridge (Abaxis) using an i-STAT portable reader (Abbott Laboratories, Princeton, NJ).
9
Multiphoton Intravital Microscopy Skin Flap
10
Vital Signs Monitoring in Mice
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Some mice were recorded with an additional commercial pulse-oximeter (MouseOx, StarrLife Sciences) to measure heart rate, respiratory rate, and additionally track SpO2, either using the manufacturer’s thigh sensor (n = 3 mice, 3 recordings) or throat collar (n = 7 mice, 12 recordings). For placement, the hair on the thigh (for mice receiving epinephrine injections) or neck (for mice undergoing isoflurane variation or ketamine/xylazine administration) was shaved and followed by treatment with depilatory cream (Nair). Vitals were recorded at either 1, 5, or 15 Hz with files timestamped for synchronization to ECG and PZT acquisition.
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