The authors have no acknowledgments.

All animal studies were approved by the Purdue Animal Care and Use Committee.
1. Setup of the low-flow vaporizer
<ol>
	<li>Isoflurane or sevoflurane delivery
	<ol>
		<li>Select a carrier gas source. To utilize the internal air pump, remove the red cap from the Inlet port on the back of the system, allowing the system to intake room air. To use compressed gas, use a pressure regulator or pressure reducer set to 15 PSI, and connect to the Compressed Gas port on the back of the system.</li>
		<li>Connect the charcoal canister to the exhaust port.</li>
		<li>Connect the Accessory Connector to the Inspiratory and Expiratory ports on the front of the system. Connect the induction chamber to branches with blue clips and the nose cone to branches with white clips (Figure 1).</li>
	</ol>
	</li>
	<li>For mechanical ventilation
	<ol>
		<li>Connect the intubation connector tubing to the yellow coded clips (Figure 2).</li>
		<li>Calibrate the ventilator by performing a deadspace calibration. From the Vent Run Screen, touch Setup, and then Calib &#38; Tests. Select Deadspace Calibration and press Dial B.</li>
	</ol>
	</li>
	<li>For pulse oximetry
	<ol>
		<li>Connect the sensor to the port on the back of the system, labeled MouseSTAT.</li>
	</ol>
	</li>
	<li>For warming
	<ol>
		<li>Connect the warming pad to the &#8216;Pad Power&#8217; port on the front of the system.</li>
		<li>Connect one sensor to the &#8220;Body Sensor&#8217; port, and the other to the &#8216;Pad Sensor&#8217; port. Secure the Pad Sensor to the warming pad.</li>
	</ol>
	</li>
</ol>
2. Configure the settings
<ol>
	<li>For anesthesia
	<ol>
		<li>Power on the anesthesia system. From the Anest Run Screen, touch Set Up.</li>
		<li>Choose the anesthetic agent. Touch Type Anest, and then turn Dial B to select Isoflurane or Sevoflurane.</li>
		<li>Set the syringe size. Touch Syringe Size, and then turn Dial B to select a size.</li>
		<li>Touch Back to return to the Anest Run Screen.</li>
		<li>Using the bottle top adapter, fill the syringe with anesthetic.</li>
		<li>Connect the syringe to the anesthesia system. Touch Remove to move the pusher block backwards if needed.</li>
		<li>Prime the syringe. Touch and hold Prime to move the pusher block forward until the pusher block touches the top of the syringe plunger. Turning B while holding the Prime button regulates the pusher block speed.</li>
	</ol>
	</li>
	<li>For mechanical ventilation
	<ol>
		<li>Touch the Vent Run Screen tab, and then Setup.</li>
		<li>Touch Body Weight and enter the weight of the animal.</li>
		<li>Touch Priority to choose volume or pressure-controlled ventilation. The Body Weight setting automatically sets appropriate respiratory rate and tidal volumes.</li>
	</ol>
	</li>
	<li>For pulse oximetry
	<ol>
		<li>Touch the Oxi Run Screen tab, and then Setup.</li>
		<li>Touch HR and turn Dial B to set the minimum allowed heart rate reading. Presets are available.</li>
	</ol>
	</li>
	<li>For warming
	<ol>
		<li>From the Warm Run Screen, touch Setup. Choose a warming method and target temperature setting.</li>
	</ol>
	</li>
</ol>
3. Begin anesthesia delivery
<ol>
	<li>Anesthetize the mouse
	<ol>
		<li>From the Anest Run Screen, touch Start Induction to begin airflow. The default Induction flow rate is 500 mL/min. Turning Dial A adjusts the flow rate as needed.</li>
		<li>Place the mouse in the induction chamber, closing the lid tightly. Adjust the Anesthetic Agent Concentration dial to 3% for isoflurane.</li>
		<li>Monitor until the mouse has reached the desired anesthetic plane, determined by a decrease in respiration rate and a loss of righting reflex when the chamber is tipped. Adjust the Anesthetic Agent Concentration dial as necessary.</li>
		<li>Once the animal has lost righting reflex and is sufficiently anesthetized, touch Stop Induction.</li>
		<li>If desired, touch Flush Chamber to empty the chamber of residual anesthetic gas.</li>
		<li>Open the clamps leading to the nose cone, and close the clamps leading to the chamber.</li>
		<li>Touch Start Nose Cone. The Body Weight setting determines the nose cone flow rate, though it can be manually adjusted by turning Dial A.</li>
		<li>Immediately fit the nose cone, and center the animal on the infrared warming pad.</li>
		<li>Insert the Animal Sensor as a rectal probe.</li>
	</ol>
	</li>
</ol>
4. Begin mechanical ventilation
<ol>
	<li>Intubate the animal.
	<ol>
		<li>Transfer the animal to the intubation stage while keeping the animal anesthetized.</li>
		<li>Suspend the animal from its upper incisors using a thread fixed onto the vertical intubation stage (Figure 3).</li>
		<li>Gently displace the animal&#8217;s tongue to the side and visualize the trachea using the lights provided in the intubation kit.</li>
		<li>Carefully insert tracheal tube and verify correct placement by connecting small air bladder to the tube and checking if the lungs inflate.</li>
	</ol>
	</li>
	<li>Connect the endotracheal tube to the ventilation tubing.</li>
	<li>Touch Stop Nose Cone, and then touch Start Ventilator. 
	NOTE: The Body Weight setting automatically determines proper respiration rate and tidal volumes. To perform pressure-controlled ventilation, set the target inspiratory pressure between 15-18 cm H2O. Make adjustments to the ventilator settings as needed per surgical protocols.</li>
</ol>
5. Begin physiological monitoring
<ol>
	<li>Place the sensor over the hind paw of the animal (Figure 4). The Pulse Oximeter will begin reading HR and SpO2 automatically. Touch the Oxi Run Screen Tab to view pulse oximetry data.</li>
</ol>

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J., Cupic, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dose-regulated+automated+anesthesia+(Abstract).">Dose-regulated automated anesthesia (Abstract).</a> British Journal of Clinical Pharmacologyl. 12 (2), 281-282 (1971).</li><li>Walker, T. J., Chackrabarti, M. K., Lockwood, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uptake+of+desflurane+during+anaesthesia.">Uptake of desflurane during anaesthesia.</a> Anaesthesia. 51 (1), 33-36 (1996).</li><li>Weingarten, M., Lowe, H. 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This project was supported with equipment and funding by Kent Scientific Corporation. The authors Krista Bigiarelli and Dave FitzMiller are employees of Kent Scientific Corporation that manufactures equipment used in this article. Open access publication of this article is sponsored by Kent Scientific Corporation.

This digital low-flow anesthesia system integrates anesthesia, ventilation, warming, and physiological monitoring systems into a single piece of equipment. Additionally, the system contains an internal pump, allowing it to draw in ambient air for use as a carrier gas, eliminating the need for a source of compressed gas.
In this procedure, the system is used as a sole piece of equipment to replace an anesthetic vaporizer, mechanical ventilator, pulse oximeter, and warming pad. We previously demonstrated anesthetic delivery at a flow rate of 100mL/min2. The flow rate settings are critical for this anesthetic delivery technique, as the flow rate directly controls the volume of liquid anesthetic used. We also previously demonstrated how using low flow rates save anesthetic liquid1,2. When a traditional vaporizer is connected to a mechanical ventilator, the vaporizer must run continuously while the ventilator inlet samples from the gas stream. In the case of the digital vaporizer with integrated ventilator, only the gas necessary for ventilation is output by the ventilator. This reduces the costs associated with anesthetic liquid, carrier gases, and charcoal filters.
Though there are many advantages to using a low-flow digital vaporizer, there are limitations as well. This system is designed to operate at low flow rates ideal for rodents and other small mammals, but does not deliver anesthesia above flow rates of 1000 mL/min. This particular system is therefore only suitable for small animal species. The integrated pulse oximeter includes a sensor for paw use only. The sensor is not recommended for use on the tail, which may be a limitation for certain surgical procedures. Further, while respiration rate can be monitored through this system via the paw sensor, it can be difficult to obtaining consistent respiratory recordings over an extended period of time. Finally, unlike a traditional vaporizer, this digital system requires electricity. Batteries are available for use in instances where electrical power is unavailable or in the event of a power outage, and can power the system through several hours of usage.
This setup and protocol demonstrate safe and effective use of a digital, low flow anesthesia system with integrated ventilator and physiological monitoring modules. This setup will be useful for any laboratories with limited bench spaces, or where it is not feasible to house multiple pieces of equipment and tubing near a surgical field. There are numerous benefits to an all-in-one system, including the elimination of compressed gas tanks and separate physiological monitoring equipment. Overall, this integrated system could be considered by groups where use of a traditional vaporizer is not ideal.

Research involving animal models often requires specialized data collection equipment. There are two common types of anesthetic vaporizer commonly used for small animal surgery. Traditional anesthetic vaporizers rely on the passive vaporization of volatile anesthetics based on atmospheric pressure and gas flow1,2,3,4,5,6,7,8,9,10. They are designed to operate at flow rates of 0.5 L/min to 10 L/min, making them ideal for large animal models11.
We recently demonstrated the effects of a low-flow digital vaporizer compared to a traditional vaporizer12,13. The low-flow digital anesthesia system can be used to maintain an animal on a nose cone at very low flow rates of 1.5-2.2 times the animal&#8217;s minute volume14,15,16.
There are numerous benefits to using a digital anesthesia system. It incorporates a built-in pump, which draws in ambient air to use as a carrier gas. This allows the user to administer anesthesia without the use of compressed gas. Recent studies17,18 have suggested that using air instead of oxygen as a carrier gas may be beneficial for many procedures.
Physiological monitoring and warming capabilities can also be installed into the digital low-flow anesthesia system. In most institutions, animal warming and physiological monitoring are required by Institutional Animal Care and Use Committees19,20,21,22. Studies comparing the physiological effects of anesthetic agents have shown a drastic depression of body temperature, cardiac function, and respiratory function23,24,25. Placing the animal on a warming pad to monitor and maintain a normal body temperature is often required. There are many methods of animal warming available, such as warm water heaters, electric heating pads, and heat lamps, but each of these have significant drawbacks. In studies comparing different methods of animal warming, far infrared warming has been found to be the most beneficial26. The digital vaporizer includes built in homeothermic far infrared warming to maintain a specific animal body temperature. This eliminates the needs for any additional warming pad controllers.
In addition to monitoring body temperature, pulse oximetry is a popular method of monitoring the animal&#8217;s heart rate and oxygen saturation. This noninvasive method is simple, accurate, and provides an overall assessment of the animal&#8217;s ability to regulate blood oxygenation levels. A paw sensor for pulse oximetry can be connected to the anesthesia system, as we have previously demonstrated2.
Mechanical ventilation is often required when the animal is under longer periods of anesthesia, or whenever the animal&#8217;s respiration pattern needs to be controlled. The low-flow digital vaporizer has the capability to deliver controlled breaths in either pressure- or volume-control. An integrated ventilator eliminates the need for an external ventilator and excess tubing setup requirements.
Because all these common monitors and features are combined into a single piece of equipment, the tubing setup is substantially simplified. The purpose of this protocol is to demonstrate the setup and use of an all-in-one digital anesthesia system.

<table><tbody><tr><td>Intubation Kit</td><td>Kent Scientific Corporation</td><td>ETM-MSE</td><td>Includes intubation stage, intubation tube, LED light</td></tr><tr><td>Isoflurane Liquid Inhalation 99.9%</td><td>Henry Schein, Inc.</td><td>1182097</td><td>Glass bottle 250mL</td></tr><tr><td>MouseSTAT Pulse Oximeter</td><td>Kent Scientific Corporation</td><td>SS-03</td><td>Integrated into SomnoSuite</td></tr><tr><td>Oxygen Tank</td><td>Indiana Oxygen Company</td><td>23-160246</td><td>Medical Grade O&lt;sub&gt;2&lt;/sub&gt; 99%</td></tr><tr><td>RoVent Automatic Ventilator</td><td>Kent Scientific Corporation</td><td>SS-04</td><td>Integrated into SomnoSuite</td></tr><tr><td>SomnoSuite Low Flow Digital Anesthesia System</td><td>Kent Scientific Corporation</td><td>SS-01</td><td>Includes RightTemp Homeothermic Warming control, pad, and temperature sensors</td></tr><tr><td>SomnoSuite Mouse Starter Kit</td><td>Kent Scientific Corporation</td><td>SOMNO-MSEKIT</td><td>Includes nose cone, syringes, induction chamber, and charcoal canister</td></tr></tbody></table>

use of an integrated low-flow anesthetic vaporizer, ventilator, and physiological monitoring system for rodents

Low-flow digital vaporizers commonly utilize a syringe pump to directly administer volatile anesthetics into a stream of carrier gas. Per animal welfare recommendations, animals are warmed and monitored during procedures requiring anesthesia. Common anesthesia and physiological monitoring equipment include gas tanks, anesthetic vaporizers and stands, warming controllers and pads, mechanical ventilators, and pulse oximeters. A computer is also necessary for data collection and to run equipment software. In smaller spaces or when performing field work, it can be challenging to configure all this equipment in limited space.
The goal of this protocol is to demonstrate best practices for use of a low-flow digital vaporizer using both compressed oxygen and room air, along with an integrated mechanical ventilator, pulse oximeter, and far infrared warming as an all-inclusive anesthesia and physiological monitoring suite ideal for rodents.

Ten week old, male, wild type C57Bl6j mice weighing 25.41 &#177; 0.8 g were used for this study. The mice were anesthetized and maintained on a nose cone or intubated and maintained on an integrated mechanical ventilator with 1.5-2.5% isoflurane while heart rate and oxygen saturation were monitored. The animals were group-housed in microisolation caging and provided free-access to standard rodent chow and water by bottle.
Heart rate and SpO2 were monitored during maintenance via pulse oximetry (Figure 5, Figure 6, and Figure 7,). Body temperature was maintained at 36.5-37.5 &#176;C via an infrared heating pad and heat lamp. Ventilated animals received continuous delivery of isoflurane during the intubation procedure via intubation stand with integrated nose cone. Each mouse was successfully ventilated or maintained on a nose cone at low flow rates not exceeding 141 mL/min of room air (RA) or oxygen (O2) for 15 minutes. The animals&#8217; heart rates and blood oxygen saturation remained stable with few significant changes in either measurement for all groups. SpO2 remained between 82-99% for all groups, while body temperature was maintained between 36.5-37.5&#160;&#176;C. We observed that both position of the pulse-oximeter and body temperature influenced SpO2 measurements. If we observed an invalid reading from the pulse-oximeter, we adjusted the placement of the sensor and heating level to keep core body temperature stable.
A two-way ANOVA with a Bonferroni correction was performed to determine significance of data in Figure 5, Figure 6 and Figure 7. A p-value less than 0.05 was considered significant.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61311/61311fig01.jpg" /> 
Figure 1: Diagram of tubing setup for anesthetic induction and nose cone maintenance. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61311/61311fig02.jpg" /> 
Figure 2: Diagram of tubing setup for anesthetic induction, intubation, and ventilation. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61311/61311fig03.jpg" /> 
Figure 3: Mice received continuous delivery of isoflurane during the intubation procedure via an intubation stand with an integrated nose cone. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61311/61311fig04.jpg" /> 
Figure 4: Integrated pulse oximeter sensor placement over the hind paw. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61311/61311fig05.jpg" /> 
Figure 5: Average heart rate over 15 minutes &#177; SD with room air (RA) or 100% oxygen (O2) delivered through nose cone or ventilated through tracheal tube (n=5/group).&#160;No significant difference was observed between groups. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61311/61311fig06.jpg" /> 
Figure 6: Heart rate values (bpm) recorded after initial anesthetic induction with the low flow anesthesia system.&#160;Average heart rate values calculated from 30-second time intervals over a 15-minute period. Each data point represents mean &#177; SD of all animals in each group (n=5). No significant changes in heart rate were observed over the 15-minute period in any group. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61311/61311fig07.jpg" /> 
Figure 7: The tissue oxygen saturation levels (%) after initial anesthetic induction with the low flow anesthesia system.&#160;Average SpO2 values calculated from 30-second time intervals over a 15-minute period. Each data point represents mean &#177; SD of all animals in each group (n=5). No significant changes in SpO2 were observed over the 15-minute period in any group. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61311/61311fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Use of an Integrated Low-Flow Anesthetic Vaporizer, Ventilator, and Physiological Monitoring System for Rodents at JoVE.com

Use of an Integrated Low-Flow Anesthetic Vaporizer, Ventilator, and Physiological Monitoring System for Rodents

Here, we present a protocol to safely and effectively administer anesthetic gas to mice using a digital, low flow anesthesia system with integrated ventilator and physiological monitoring modules.

Low-flow digital vaporizers commonly utilize a syringe pump to directly administer volatile anesthetics into a stream of carrier gas. Per animal welfare ...

use-an-integrated-low-flow-anesthetic-vaporizer-ventilator

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

5.8K Views.  Kent Scientific Corporation. Here, we present a protocol to safely and effectively administer anesthetic gas to mice using a digital, low flow anesthesia system with integrated ventilator and physiological monitoring modules.

Video: Use of an Integrated Low-Flow Anesthetic Vaporizer, Ventilator, and Physiological Monitoring System for Rodents

Assessing Changes in Volatile General Anesthetic Sensitivity of Mice after Local or Systemic Pharmacological Intervention

One desirable endpoint of general anesthesia is the state of unconsciousness, also known as hypnosis. Defining the hypnotic state in animals is less straightforward than it is in human patients. A widely used behavioral surrogate for hypnosis in rodents is the loss of righting reflex (LORR), or the point at which the animal no longer responds to their innate instinct to avoid the vulnerability of dorsal recumbency. We have developed a system to assess LORR in 24 mice simultaneously while carefully controlling for potential confounds, including temperature fluctuations and varying gas flows. These chambers permit reliable assessment of anesthetic sensitivity as measured by latency to return of the righting reflex (RORR) following a fixed anesthetic exposure. Alternatively, using stepwise increases (or decreases) in anesthetic concentration, the chambers also enable determination of a population's sensitivity to induction (or emergence) as measured by EC50 and Hill slope. Finally, the controlled environmental chambers described here can be adapted for a variety of alternative uses, including inhaled delivery of other drugs, toxicology studies, and simultaneous real-time monitoring of vital signs.

Loss of the righting reflex has long served as a standard behavioral surrogate for unconsciousness, also called hypnosis, in laboratory animals. Alterations in volatile anesthetic sensitivity caused by pharmacological interventions can be detected with a carefully controlled high-throughput assessment system, which may be adapted for delivery of any inhaled therapeutic.

One desirable endpoint of general anesthesia is  the state of unconsciousness, also known as hypnosis. Defining the hypnotic state in animals is  less ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

Non-restraining EEG Radiotelemetry: Epidural and Deep Intracerebral Stereotaxic EEG Electrode Placement

Implantable EEG radiotelemetry is of central relevance in the neurological characterization of transgenic mouse models of neuropsychiatric and neurodegenerative diseases as well as epilepsies. This powerful technique does not only provide valuable insights into the underlying pathophysiological mechanisms, i.e., the etiopathogenesis of CNS related diseases, it also facilitates the development of new translational, i.e., therapeutic approaches. Whereas competing techniques that make use of recorder systems used in jackets or tethered systems suffer from their unphysiological restraining to semi-restraining character, radiotelemetric EEG recordings overcome these disadvantages. Technically, implantable EEG radiotelemetry allows for precise and highly sensitive measurement of epidural and deep, intracerebral EEGs under various physiological and pathophysiological conditions. First, we present a detailed protocol of a straight forward, successful, quick and efficient technique for epidural (surface) EEG recordings resulting in high-quality electrocorticograms. Second, we demonstrate how to implant deep, intracerebral EEG electrodes, e.g., in the hippocampus (electrohippocampogram). For both approaches, a computerized 3D stereotaxic electrode implantation system is used. The radiofrequency transmitter itself is implanted into a subcutaneous pouch in both mice and rats. Special attention also has to be paid to pre-, peri- and postoperative treatment of the experimental animals. Preoperative preparation of mice and rats, suitable anesthesia as well as postoperative treatment and pain management are described in detail.

Non-restraining EEG radiotelemetry is a valuable methodological approach to record in vivo long-term electroencephalograms from freely moving rodents. This detailed protocol describes stereotaxic epidural and deep intracerebral electrode placement in different brain regions in order to obtain reliable recordings of CNS rhythmicity and CNS related behavioral stages.

Implantable EEG radiotelemetry is of central relevance in the neurological characterization of transgenic mouse models of neuropsychiatric and ...

Neuroscience

Use of a Low-flow Digital Anesthesia System for Mice and Rats

A traditional vaporizer depends on flowing gas and atmospheric pressure for passive anesthetic vaporization. Newly developed direct injection vaporizers utilize a syringe pump to directly administer volatile anesthetics into a gas stream. Unlike a traditional vaporizer, it can be used at very low flow rates, making it ideal for use on mice and rats. 
The equipment's capability to use low flow rates could result in a substantial cost savings due to the reduced need for anesthetic agents, compressed gas, and charcoal scavenging filters1. A lower flow rate means less waste of anesthetic gas and likely reduces the risk of anesthetic exposure to laboratory personnel. Thus, the high levels of precision and safety associated with direct injection vaporizers, along with a reduced need for anesthetic agents, compressed gas, and charcoal filters are beneficial for research requiring small animal anesthesia. 
The goal of this protocol is to demonstrate the use of a syringe-driven direct injection vaporizer as part of a digital, low-flow anesthesia system. The direct injection vaporizer is capable of accurately delivering anesthesia at very low flow rates compared to a traditional vaporizer, making it a promising alternative for controlled gas anesthetic delivery to rodents.

Here, we present a protocol to more safely and efficiently administer anesthetic gas to mice using a digital, low flow anesthesia system utilizing a syringe-driven direct injection vaporizer.

A traditional vaporizer depends on flowing gas and atmospheric pressure for passive anesthetic vaporization. Newly developed direct injection vaporizers ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

The forced oscillation technique (FOT) is a powerful, integrative and translational tool permitting the experimental assessment of lung function in mice in a comprehensive, detailed, precise and reproducible manner. It provides measurements of respiratory system mechanics through the analysis of pressure and volume signals acquired in reaction to predefined, small amplitude, oscillatory airflow waveforms, which are typically applied at the subject's airway opening. The present protocol details the steps required to adequately execute forced oscillation measurements in mice using a computer-controlled piston ventilator (flexiVent; SCIREQ Inc, Montreal, Qc, Canada). The description is divided into four parts: preparatory steps, mechanical ventilation, lung function measurements, and data analysis. It also includes details of how to assess airway responsiveness to inhaled methacholine in anesthetized mice, a common application of this technique which also extends to other outcomes and various lung pathologies. Measurements obtained in na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage are presented to illustrate how this tool can contribute to a better characterization and understanding of studied physiological changes or disease models as well as to applications in new research areas.

The present protocol provides a detailed step-by-step description of the procedures required to execute measurements of respiratory system mechanics as well as the assessment of airway responsiveness to inhaled methacholine in mice using the forced oscillation technique (flexiVent; SCIREQ Inc, Montreal, Qc, Canada).

The  forced oscillation technique (FOT) is a powerful, integrative and translational  tool permitting the experimental assessment of lung function in mice ...

Biology

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) generated by recording both pressure and volume during cardiac catheterization are vital when assessing both systolic and diastolic cardiac function. Left and right heart function are closely related, reflected in ventricular interdependence. Thus, recording biventricular function in the same animal is important to get a complete assessment of cardiac function. In this protocol, a closed chest approach to cardiac catheterization consistent with the way catheterization is performed in patients is adopted in mice. While challenging, the closed chest strategy is a more physiological approach, because opening the chest results in major changes in preload and afterload that create artifacts, most notably a fall in systemic blood pressure. While high-resolution echocardiography is used to assess rodents, cardiac catheterization is invaluable, particularly when assessing diastolic pressures in both ventricles.
Described here is a procedure to perform invasive, closed chest, sequential left and right ventricular pressure-volume (PV) loops in the same animal. PV loops are acquired using&#160;admittance technology with a mouse pressure-volume catheter and pressure-volume system acquisition. The procedure is described, beginning with the neck dissection, which is required to access the right jugular vein and the right carotid artery, to the insertion and positioning of the catheter, and finally the data acquisition. Then, the criteria required to ensure the acquisition of high-quality PV loops are discussed. Finally, the analysis of the left and right ventricular PV loops and the different hemodynamic parameters available to quantify systolic and diastolic ventricular function are briefly described.

Presented here is a protocol to assess biventricular heart function in mice by generating pressure-volume (PV) loops from the right and left ventricle in the same animal using closed chest catheterization. The focus is on the technical aspect of surgery and data acquisition.

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) ...

Measuring Pressure Volume Loops in the Mouse

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the mouse provides great potential to explore cardiac function at the molecular level. The mouse&#39;s small size does present some challenges in regards to performing detailed cardiac phenotyping. Miniaturization and other advancements in technology have made many methods of cardiac assessment possible in the mouse. Of these, the simultaneous collection of pressure and volume data provides a detailed picture of cardiac function that is not available through any other modality. Here a detailed procedure for the collection of pressure-volume loop data is described. Included is a discussion of the principles underlying the measurements and the potential sources of error. Anesthetic management and surgical approaches are discussed in great detail as they are both critical to obtaining high quality hemodynamic measurements. The principles of hemodynamic protocol development and relevant aspects of data analysis are also addressed.

This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the ...

Evaluation of Apoptosis and Lung Injury in DCD Donor Lungs Preserved by EVLP

Establishment of a Novel Ex Vivo Lung Perfusion System for Rat Lungs After Circulatory Death

The increasing scarcity of donor lungs for transplantation has spurred significant interest in the utilization of organs procured from donors after cardiac death (DCD). However, a critical challenge remains in maximizing the number of viable lungs that meet stringent criteria for clinical use. Ex vivo lung perfusion (EVLP) emerges as a promising technology to evaluate and potentially improve the function of donated lungs. This paper describes the development of a cost-effective and reproducible small animal experimental platform specifically designed for EVLP research. This platform offers a high degree of customization, enabling researchers to tailor experimental parameters to address specific research questions within the field of EVLP. The team established a rat model to simulate DCD and implemented a two-stage preservation approach. Lungs were subjected to static cold storage using a perfusate solution at 4 &#176;C for 4 h, followed by normothermic EVLP for another 4 h. Researchers evaluated various lung function parameters, including pulmonary vascular resistance, pulmonary dynamic compliance, blood gas measurements of the perfusate, and apoptotic cell death in lung tissues. The study demonstrates that EVLP could significantly improve the functional performance of DCD lungs compared to simple cold storage at 4 &#176;C, suggesting that EVLP has the potential to mitigate ischemia-reperfusion injury and enhance graft function following circulatory death, thereby increasing the pool of transplantable lungs.

The objective is to develop a new ex vivo lung perfusion (EVLP) model in rats that can sustain consistent lung function for 4 h throughout EVLP and furnish the industry with implementation specifics that will offer a sturdy method for applying EVLP to rodent models.

The increasing scarcity of donor lungs for transplantation has spurred significant interest in the utilization of organs procured from donors after ...

Intravital Imaging Using an Inverted Microscope to Track Skin Cell Dynamics

Streamlined Intravital Imaging Approach for Long-Term Monitoring of Epithelial Tissue Dynamics on an Inverted Confocal Microscope

Understanding normal and aberrant in vivo cell behaviors is necessary to develop clinical interventions to thwart disease initiation and progression. It is therefore critical to optimize imaging approaches that facilitate the observation of cell dynamics in situ, where tissue structure and composition remain unperturbed. The epidermis is the body&#39;s outermost barrier, as well as the source of the most prevalent human cancers, namely cutaneous skin carcinomas. The accessibility of skin tissue presents a unique opportunity to monitor epithelial and dermal cell behaviors in intact animals using noninvasive intravital microscopy. Nevertheless, this sophisticated imaging approach has primarily been achieved using upright multiphoton microscopes, which represent a significant barrier for entry for most investigators. This study presents a custom-designed, 3D-printed microscope stage insert suitable for use with inverted confocal microscopes, streamlining the long-term intravital imaging of ear skin in live transgenic mice. We believe this versatile invention, which may be customized to fit the inverted microscope brand and model of choice and adapted to image additional organ systems, will prove invaluable to the greater scientific research community by significantly enhancing the accessibility of intravital microscopy. This technological advancement is critical for bolstering our understanding of live cell dynamics in normal and disease contexts.

Author Spotlight: Developing a Tool for Using Inverted Confocal Microscopes for In Vivo Intravital Imaging 

The protocol presents a new tool to simplify intravital imaging using inverted confocal microscopy.

Understanding normal and aberrant in vivo cell behaviors is necessary to develop clinical interventions to thwart disease initiation and progression. It ...

Use of an Integrated Low-Flow Anesthetic Vaporizer, Ventilator, and Physiological Monitoring System for Rodents

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Assessing Changes in Volatile General Anesthetic Sensitivity of Mice after Local or Systemic Pharmacological Intervention

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Non-restraining EEG Radiotelemetry: Epidural and Deep Intracerebral Stereotaxic EEG Electrode Placement

Use of a Low-flow Digital Anesthesia System for Mice and Rats

Laminectomy and Spinal Cord Window Implantation in the Mouse

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

Measuring Pressure Volume Loops in the Mouse

Establishment of a Novel Ex Vivo Lung Perfusion System for Rat Lungs After Circulatory Death

Streamlined Intravital Imaging Approach for Long-Term Monitoring of Epithelial Tissue Dynamics on an Inverted Confocal Microscope