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>

<ol>
	<li>El-Attar, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guided+isoflurane+injection+in+a+totally+closed+circuit.">Guided isoflurane injection in a totally closed circuit.</a> Anaesthesia. 46 (12), 1059-1063 (1991).</li><li>Lockwood, G., Chakrabarti, M. K., Whitwam, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+computer-controller+closed+anaesthetic+breathing+system.">A computer-controller closed anaesthetic breathing system.</a> Anaesthesia. 48 (8), 690-693 (1993).</li><li>Lowe, H. 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. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+circuit+injection+technic+for+syringe-measured+administration+of+methoxyflurane:+a+new+dimension+in+anesthesia.">A new circuit injection technic for syringe-measured administration of methoxyflurane: a new dimension in anesthesia.</a> Anesthesia &amp; Analgesia. 52 (4), 634-642 (1973).</li><li>Enlund, M., Wiklund, L., Lambert, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+device+to+reduce+the+consumption+of+a+halogenated+anaesthetic+agent.">A new device to reduce the consumption of a halogenated anaesthetic agent.</a> Anaesthesia. 56 (5), 429-432 (2001).</li><li>Kelly, J. M., Kong, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accuracy+of+ten+isoflurane+vaporisers+in+current+clinical+use.">Accuracy of ten isoflurane vaporisers in current clinical use.</a> Anaesthesia. 66 (8), 682-688 (2011).</li><li>Matsuda, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NARCOBIT+-+A+newly+developed+inhalational+anesthesia+system+for+mice.">NARCOBIT - A newly developed inhalational anesthesia system for mice.</a> Experimental Animals. 56 (2), 131-137 (2007).</li><li>Soro, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+accuracy+of+the+anesthetic+conserving+device+(Anaconda)+as+an+alternative+to+the+classical+vaporizer+in+anesthesia.">The accuracy of the anesthetic conserving device (Anaconda) as an alternative to the classical vaporizer in anesthesia.</a> Anesthesia &amp; Analgesia. 111 (5), 1176-1179 (2010).</li><li>Ward, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Physical principles and maintenance. Anaesthetic equipment. , (1985).</li><li>Ambrisko, T. D., Klide, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+isoflurane+and+Sevoflurane+vaporizers+over+a+wide+range+of+oxygen+flow+rates.">Evaluation of isoflurane and Sevoflurane vaporizers over a wide range of oxygen flow rates.</a> American Journal of Veterinary Research. 67 (6), 936-940 (2006).</li><li>Damen, F. W., Adelsperger, A. R., Wilson, K. E., Goergen, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+traditional+and+integrated+digital+anesthetic+vaporizers.">Comparison of traditional and integrated digital anesthetic vaporizers.</a> Journal of the American Association for Laboratory Animal Science. 54 (6), 756-762 (2015).</li><li>Adelsperger, A. R., Bigiarelli-Nogas, K. J., Toore, I., Goergen, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+a+Low-flow+Digital+Anesthesia+System+for+Mice+and+Rats.">Use of a Low-flow Digital Anesthesia System for Mice and Rats.</a> Journal of Visualized Experiments. (115), e54436 (2016).</li><li>Flecknell, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Laboratory animal anaesthesia. , (2009).</li><li>Mapleson, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+elimination+of+rebreathing+in+various+semiclosed+anaesthetic+systems.">The elimination of rebreathing in various semiclosed anaesthetic systems.</a> British Journal of Anaesthesia. 26 (5), 323-332 (1954).</li><li>Chakravarti, S., Basu, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modern+Anaesthesia+Vapourisers.">Modern Anaesthesia Vapourisers.</a> Indian Journal of Anaesthesia. 57 (5), 464-471 (2013).</li><li>Mullin, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+anesthesia+carrier+gas+on+in+vivo+circulation+times+of+ultrasound+microbubble+contrast+agents+in+rats.">Effect of anesthesia carrier gas on in vivo circulation times of ultrasound microbubble contrast agents in rats.</a> Contrast Media &amp; Molecular Imaging. 6 (3), 126-131 (2011).</li><li>Flores, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+anesthetic+agent+and+carrier+gas+on+blood+glucose+and+tissue+uptake+in+mice+undergoing+dynamic+FDG-PET+imaging:+sevoflurane+and+isoflurane+compared+in+air+and+in+oxygen.">The effects of anesthetic agent and carrier gas on blood glucose and tissue uptake in mice undergoing dynamic FDG-PET imaging: sevoflurane and isoflurane compared in air and in oxygen.</a> Molecular Imaging and Biology. 10 (4), 192 (2008).</li><li>Carroll, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Small Animal Anesthesia and Analgesia. , (2008).</li><li>Thomas, J., Lerche, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Anesthesia and Analgesia for Veterinary Technicians, 4th ed. 335, (2011).</li><li>McKelvey, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Veterinary Anesthesia and Analgesia. , (2003).</li><li>Tranquilli, W. J., Thurmon, J. C., Grimm, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Lumb and Jones' veterinary anesthesia and analgesia. , 23-86 (2013).</li><li>Matsuda, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+newly+developed+inhalation+anesthesia+system+and+intraperitoneal+anesthesia+on+the+hemodynamic+state+in+mice.">Comparison of newly developed inhalation anesthesia system and intraperitoneal anesthesia on the hemodynamic state in mice.</a> Biological and Pharmaceutical Bulletin. 30 (9), 1716-1720 (2007).</li><li>Garber, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Guide for the Care and Use of Laboratory Animals. 8th edn. , (2011).</li><li>Zarndt, B. 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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 O2 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://www.jove.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://www.jove.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://www.jove.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://www.jove.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://www.jove.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://www.jove.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://www.jove.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 ...

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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

Use of a Hanging-weight System for Isolated Renal Artery Occlusion

In hospitalized patients, over 50% of cases of acute kidney injury (AKI) are caused by renal ischemia 1-3. A recent study of hospitalized patients revealed that only a mild increase in serum creatinine levels (0.3 to 0.4 mg/dl) is associated with a 70% greater risk of death than in persons without any increase 1. Along these lines, surgical procedures requiring cross-clamping of the aorta and renal vessels are associated with a renal failure rates of up to 30% 4. Similarly, AKI after cardiac surgery occurs in over 10% of patients under normal circumstances and is associated with dramatic increases in mortality. AKI are also common complications after liver transplantation. At least 8-17% of patients end up requiring renal replacement therapy 5. Moreover, delayed graft function due to tubule cell injury during kidney transplantation is frequently related to ischemia-associated AKI 6. Moreover, AKI occurs in approximately 20% of patients suffering from sepsis 6.The occurrence of AKI is associated with dramatic increases of morbidity and mortality 1. Therapeutic approaches are very limited and the majority of interventional trials in AKI have failed in humans. Therefore, additional therapeutic modalities to prevent renal injury from ischemia are urgently needed 3, 7-9. 
To elucidate mechanisms of renal injury due to ischemia and possible therapeutic strategies murine models are intensively required 7-13. Mouse models provide the possibility of utilizing different genetic models including gene-targeted mice and tissue specific gene-targeted mice (cre-flox system). However, murine renal ischemia is technically challenging and experimental details significantly influence results. We performed a systematic evaluation of a novel model for isolated renal artery occlusion in mice, which specifically avoids the use of clamping or suturing the renal pedicle 14. This model requires a nephrectomy of the right kidney since ischemia can be only performed in one kidney due to the experimental setting. In fact, by using a hanging-weight system, the renal artery is only instrumented once throughout the surgical procedure. In addition, no venous or urethral obstruction occurs with this technique. We could demonstrate time-dose-dependent and highly reproducible renal injury with ischemia by measuring serum creatinine. Moreover, when comparing this new model with conventional clamping of the whole pedicle, renal protection by ischemic preconditioning is more profound and more reliable. Therefore his new technique might be useful for other researchers who are working in the field of acute kidney injury.

A precise murine model for acute kidney injury (AKI) due to ischemia is an important tool to investigate acute kidney injury and possibly find therapeutic tools to treat renal injury. The hanging weight system offers a tool for immediate and reliable renal artery occlusion and reperfusion without causing renal congestion.

In hospitalized patients, over 50% of cases of acute kidney injury (AKI) are caused by renal ischemia 1-3. A recent study of hospitalized patients ...

The Use of Cystometry in Small Rodents: A Study of Bladder Chemosensation

The lower urinary tract (LUT) functions as a dynamic reservoir that is able to store urine and to efficiently expel it at a convenient time. While storing urine, however, the bladder is exposed for prolonged periods to waste products. By acting as a tight barrier, the epithelial lining of the LUT, the urothelium, avoids re-absorption of harmful substances. Moreover, noxious chemicals stimulate the bladder's nociceptive innervation and initiate voiding contractions that expel the bladder's contents. Interestingly, the bladder's sensitivity to noxious chemicals has been used successfully in clinical practice, by intravesically infusing the TRPV1 agonist capsaicin to treat neurogenic bladder overactivity1. This underscores the advantage of viewing the bladder as a chemosensory organ and prompts for further clinical research. However, ethical issues severely limit the possibilities to perform, in human subjects, the invasive measurements that are necessary to unravel the molecular bases of LUT clinical pharmacology. A way to overcome this limitation is the use of several animal models2. Here we describe the implementation of cystometry in mice and rats, a technique that allows measuring the intravesical pressure in conditions of controlled bladder perfusion.
	After laparotomy, a catheter is implanted in the bladder dome and tunneled subcutaneously to the interscapular region. Then the bladder can be filled at a controlled rate, while the urethra is left free for micturition. During the repetitive cycles of filling and voiding, intravesical pressure can be measured via the implanted catheter. As such, the pressure changes can be quantified and analyzed. Moreover, simultaneous measurement of the voided volume allows distinguishing voiding contractions from non-voiding contractions3.
	Importantly, due to the differences in micturition control between rodents and humans, cystometric measurements in these animals have only limited translational value4. Nevertheless, they are quite instrumental in the study of bladder pathophysiology and pharmacology in experimental pre-clinical settings. Recent research using this technique has revealed the key role of novel molecular players in the mechano- and chemo-sensory properties of the bladder.

Cystometry is an efficient technique to measure bladder function of small animals in vivo. The bladder is continuously infused at rates controlled through an intravesical catheter, whereas the urethra is left free for micturition. This allows for repetitive filling and emptying of the bladder, while intravesical pressure and voided volume are recorded.

	The lower urinary tract (LUT) functions as a  dynamic reservoir that is able to store urine and to efficiently expel it at a  convenient time. While ...

Use of Animal Model of Sepsis to Evaluate Novel Herbal Therapies

Sepsis refers to a systemic inflammatory response syndrome resulting from a microbial infection. It has been routinely simulated in animals by several techniques, including infusion of exogenous bacterial toxin (endotoxemia) or bacteria (bacteremia), as well as surgical perforation of the cecum by cecal ligation and puncture (CLP)1-3. CLP allows bacteria spillage and fecal contamination of the peritoneal cavity, mimicking the human clinical disease of perforated appendicitis or diverticulitis. The severity of sepsis, as reflected by the eventual mortality rates, can be controlled surgically by varying the size of the needle used for cecal puncture2. In animals, CLP induces similar, biphasic hemodynamic cardiovascular, metabolic, and immunological responses as observed during the clinical course of human sepsis3. Thus, the CLP model is considered as one of the most clinically relevant models for experimental sepsis1-3.
Various animal models have been used to elucidate the intricate mechanisms underlying the pathogenesis of experimental sepsis. The lethal consequence of sepsis is attributable partly to an excessive accumulation of early cytokines (such as TNF, IL-1 and IFN-&gamma;)4-6 and late proinflammatory mediators (e.g., HMGB1)7. Compared with early proinflammatory cytokines, late-acting mediators have a wider therapeutic window for clinical applications. For instance, delayed administration of HMGB1-neutralizing antibodies beginning 24 hours after CLP, still rescued mice from lethality8,9, establishing HMGB1 as a late mediator of lethal sepsis. The discovery of HMGB1 as a late-acting mediator has initiated a new field of investigation for the development of sepsis therapies using Traditional Chinese Herbal Medicine. In this paper, we describe a procedure of CLP-induced sepsis, and its usage in screening herbal medicine for HMGB1-targeting therapies.

Sepsis refers to a systemic inflammatory response syndrome resulting from a microbial infection, and can be simulated by a surgical technique termed cecal ligation and puncture (CLP). Here we describe a method to use CLP-induced animal model to screen medicinal herbs for therapeutic agents.

Sepsis refers to a systemic inflammatory response syndrome resulting from a microbial infection. It has been routinely simulated in animals by several ...

The Use of Primary Human Fibroblasts for Monitoring Mitochondrial Phenotypes in the Field of Parkinson's Disease

Parkinson's disease (PD) is the second most common movement disorder and affects 1% of people over the age of 60 1. Because ageing is the most important risk factor, cases of PD will increase during the next decades 2. Next to pathological protein folding and impaired protein degradation pathways, alterations of mitochondrial function and morphology were pointed out as further hallmark of neurodegeneration in PD 3-11.
After years of research in murine and human cancer cells as in vitro models to dissect molecular pathways of Parkinsonism, the use of human fibroblasts from patients and appropriate controls as ex vivo models has become a valuable research tool, if potential caveats are considered. Other than immortalized, rather artificial cell models, primary fibroblasts from patients carrying disease-associated mutations apparently reflect important pathological features of the human disease.
Here we delineate the procedure of taking skin biopsies, culturing human fibroblasts and using detailed protocols for essential microscopic techniques to define mitochondrial phenotypes. These were used to investigate different features associated with PD that are relevant to mitochondrial function and dynamics. Ex vivo, mitochondria can be analyzed in terms of their function, morphology, colocalization with lysosomes (the organelles degrading dysfunctional mitochondria) and degradation via the lysosomal pathway. These phenotypes are highly relevant for the identification of early signs of PD and may precede clinical motor symptoms in human disease-gene carriers. Hence, the assays presented here can be utilized as valuable tools to identify pathological features of neurodegeneration and help to define new therapeutic strategies in PD.

Fibroblasts from patients carrying mutations in Parkinson's disease-causing genes represent an easily accessible ex vivo model to study disease-associated phenotypes. Live cell imaging gives the opportunity to study morphological and functional parameters in living cells. Here we describe the preparation of human fibroblasts and subsequent monitoring of mitochondrial phenotypes.

Parkinson's disease (PD) is the second most  common movement disorder and affects 1% of people over the age of 60 1. Because ageing is  the most important ...

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 ...

Use of a Piglet Model for the Study of Anesthetic-induced Developmental Neurotoxicity (AIDN): A Translational Neuroscience Approach

Anesthesia cannot be avoided in many cases when surgery is required, particularly in children. Recent investigations in animals have raised concerns that anesthesia exposure may lead to neuronal apoptosis, known as anesthesia-induced developmental neurotoxicity (AIDN). Furthermore, some clinical studies in children have suggested that anesthesia exposure may lead to neurodevelopmental deficits later in life. Nonetheless, an ideal animal model for preclinical study has yet to be developed. The neonatal piglet represents a valuable model for preclinical study, as they share a striking number of developmental similarities with humans.
The anatomy and physiology of piglets allow for implementation of rigorous human perioperative conditions in both survival and non-survival procedures. Femoral artery catheterization allows for close monitoring, thus enabling prompt correction of any deviation of the piglet's vital signs and chemistries. In addition, there are multiple developmental similarities between piglets and human neonates. The techniques required to use piglets for experimentation will require experience to master. A pediatric anesthesiologist is a critical member of the investigative team. We describe, in a general sense, the appropriate use of a piglet model for neurodevelopmental study.

Use of a Piglet Model for the Study of Anesthetic-induced Developmental Neurotoxicity AIDN: A Translational Neuroscience Approach

Anesthesia-induced developmental neurotoxicity (AIDN) research has focused on rodents, which are not broadly applicable to humans. Non-human primate models are more relevant, but are cost-prohibitive and difficult to use for experimentation. The piglet, in contrast, is a clinically relevant, practical animal model ideal for the study of anesthetic neurotoxicity.

Anesthesia cannot be avoided in many cases when surgery is required, particularly in children.  Recent investigations in animals have raised concerns that ...

Sucrose Preference and Novelty-Induced Hypophagia Tests in Rats using an Automated Food Intake Monitoring System

The prevalence and incidence of depressive disorders are rising worldwide, affecting about 322 million individuals, underlining the need for behavioral studies in animal models. In this protocol, to study depression-like and anhedonic behavior in rats, the established sucrose preference and novelty-induced hypophagia tests are combined with an automated food and liquid intake monitoring system. Prior to testing, in the sucrose preference paradigm, male rats are trained for at least 2 days to consume a sucrose solution in addition to tap water. During the test, rats are again exposed to water and sucrose solution. Consumption is registered every second by the automated system. The ratio of sucrose to total water intake (sucrose preference ratio) is a surrogate parameter for anhedonia. In the novelty-induced hypophagia test, male rats undergo a training period in which they are exposed to a palatable snack. During training, rodents show a stable baseline snack intake. On test day, the animals are transferred from home cages into a fresh, empty cage representing a novel unknown environment with access to the known palatable snack. The automated system records the total intake and its underlying microstructure (e.g., latency to approaching the snack), providing insight into anhedonic and anxious behaviors. The combination of these paradigms with an automated measuring system provides more detailed information, along with higher accuracy by reducing measuring errors. However, the tests use surrogate parameters and only depict depression and anhedonia in an indirect manner.

Presented here is a protocol to study depression-like and anhedonic behavior in rats. It combines two well-established behavioral methods, the sucrose preference and novelty-induced hypophagia tests, with an automated food and liquid intake monitoring system, to indirectly investigate rodent behavior using surrogate parameters.

The prevalence and incidence of depressive disorders are rising worldwide, affecting about 322 million individuals, underlining the need for behavioral ...

TBase - an Integrated Electronic Health Record and Research Database for Kidney Transplant Recipients

TBase is an electronic health record (EHR) for kidney transplant recipients (KTR) combining automated data entry of key clinical data (e.g., laboratory values, medical reports, radiology and pathology data) via standardized interfaces with manual data entry during routine treatment (e.g., clinical notes, medication list, and transplantation data). By this means, a comprehensive database for KTR is created with benefits for routine clinical care and research. It enables both easy everyday clinical use and quick access for research questions with highest data quality. This is achieved by the concept of data validation in clinical routine in which clinical users and patients have to rely on correct data for treatment and medication plans and thereby validate and correct the clinical data in their daily practice. This EHR is tailored for the needs of transplant outpatient care and proved its clinical utility for more than 20 years at Charit&#233; - Universit&#228;tsmedizin Berlin. It facilitates efficient routine work with well-structured, comprehensive long-term data and allows their easy use for clinical research. To this point, its functionality covers automated transmission of routine data via standardized interfaces from different hospital information systems, availability of transplant-specific data, a medication list with an integrated check for drug-drug interactions, and semi-automated generation of medical reports among others. Key elements of the latest reengineering are a robust privacy-by-design concept, modularity, and hence portability into other clinical contexts as well as usability and platform independence enabled by HTML5 (Hypertext Markup Language) based responsive web design. This allows fast and easy scalability into other disease areas and other university hospitals. The comprehensive long-term datasets are the basis for the investigation of Machine Learning algorithms, and the modular structure allows to rapidly implement these into clinical care. Patient reported data and telemedicine services are integrated into TBase in order to meet future needs of the patients. These novel features aim to improve clinical care as well as to create new research options and therapeutic interventions.

TBase combines an electronic health record with an innovative research database for kidney transplant recipients. TBase is built upon an in-memory database platform, connected to different hospital systems, and used for regular outpatient care. It automatically integrates all relevant clinical data including transplantation-specific data creating a unique research database.

TBase is an electronic health record (EHR) for kidney transplant recipients (KTR) combining automated data entry of key clinical data (e.g., laboratory ...

Multi-system Monitoring for Identification of Seizures, Arrhythmias and Apnea in Conscious Restrained Rabbits

Patients with ion channelopathies are at a high risk of developing seizures and fatal cardiac arrhythmias. There is a higher prevalence of heart disease and arrhythmias in people with epilepsy (i.e., epileptic heart.) Additionally, cardiac and autonomic disturbances have been reported surrounding seizures. 1:1,000 epilepsy patients/year die of sudden unexpected death in epilepsy (SUDEP). The mechanisms for SUDEP remain incompletely understood. Electroencephalograms (EEG) and electrocardiograms (ECG) are two techniques routinely used in the clinical setting to detect and study the substrates/triggers for seizures and arrhythmias. While many studies and descriptions of this methodology are in rodents, their cardiac electrical activity differs significantly from humans. This article provides a description of a non-invasive method for recording simultaneous video-EEG-ECG-oximetry-capnography in conscious rabbits. As cardiac electrical function is similar in rabbits and humans, rabbits provide an excellent model of translational diagnostic and therapeutic studies. In addition to outlining the methodology for data acquisition, we discuss the analytical approaches for examining neuro-cardiac electrical function and pathology in rabbits. This includes arrhythmia detection, spectral analysis of EEG and a seizure scale developed for restrained rabbits.

Using simultaneous video-EEG-ECG-oximetry-capnography, we developed a methodology to evaluate the susceptibility of rabbit models to develop provoked arrhythmias and seizures. This novel recording system establishes a platform to test the efficacy and safety of therapeutics and can capture the complex cascade of multi-system events that culminate in sudden death.

Patients with ion channelopathies are at a high risk of developing seizures and fatal cardiac arrhythmias. There is a higher prevalence of heart disease ...

Standardized Rat Coronary Ring Preparation and Real-Time Recording of Dynamic Tension Changes Along Vessel Diameter

As a key event of cardiovascular system diseases, coronary artery disease (CAD) has been widely regarded as the main culprit of atherosclerosis, myocardial infarction, and angina pectoris, which seriously threaten the life and health of people all over the world. However, how to record the dynamic biomechanical characteristics of isolated blood vessels has long puzzled people. Meanwhile, precise positioning and isolation of coronary arteries to measure in vitro dynamic vascular tension changes have become a trend in CAD drug development. The present protocol describes the macroscopic identification and microscopic separation of rat coronary arteries. The contraction and dilation function of the coronary artery ring along the vessel diameter was monitored using the established multi myograph system. The standardized and programmed protocols of coronary ring tension measurement, from sampling to data acquisition, tremendously improve the repeatability of the experimental data, which ensures the authenticity of vascular tension records after physiological, pathological, and drug intervention.

The present protocol describes the wire myograph technique for measuring vascular reactivity of the rat coronary artery.