The authors have no acknowledgements.

All studies were completed in accordance with regulatory and institutional guidelines. The animal aspects of this study were evaluated by the Kent Scientific Corporation Animal Use Program, approved by the Purdue Animal Care and Use Committee (PACUC), and performed in accordance with the Guide for the Care and Use of Laboratory Animals22.
Note: The low-flow digital anesthesia system used in this protocol is equipped with an integrated pulse oximeter.
1. Set Up the Low-flow Anesthesia System with Integrated Pulse Oximeter
<ol>
	<li>For Isoflurane Delivery
	<ol>
		<li>Select a carrier gas source. To utilize the internal air pump, unscrew Inlet Port on back, allowing the internal pump to intake room air.</li>
		<li>Connect charcoal canister to exhaust port.</li>
		<li>Connect Y adapter to front of the low flow, digital anesthesia system. Use the color-coded clips to connect the white branches to the nose cone, and blue branches to the induction chamber.</li>
		<li>Close white clip clamps and open blue clip clamps to direct airflow to the chamber.</li>
		<li>Select 2 ml syringe.</li>
	</ol>
	</li>
	<li>For Physiological Monitoring, Using the Integrated Pulse Oximeter
	<ol>
		<li>Connect the pulse oximeter paw sensor to port on the back of the low-flow anesthesia system, labeled &#34;AUX&#34;.</li>
	</ol>
	</li>
</ol>
2. Configure the Settings
<ol>
	<li>For Anesthesia
	<ol>
		<li>Power on the anesthesia system, and access the Set Up menu. Press Set Up to access Main Menu &#62; Anesthesia &#62; Setup in red.</li>
		<li>Choose the anesthetic agent. Press Set Up to highlight Type Anest in red. Use the Up and Down arrows to select Isoflurane.</li>
		<li>Set the syringe size. Press Set Up to highlight Syr Size. Use the Up and Down arrows to select a 2 ml syringe.</li>
		<li>Set the Empty Position. Press Set Up to highlight Set Empty in red. Secure empty, fully depressed glass syringe in the Syringe Retention Block by placing the syringe so that the Syringe Holding Clamp is seated on the metal collar of the syringe. Press the Up or Down arrows to move the Pusher Block so that the Pusher Block makes light contact with the top of the syringe plunger. Press select to set empty position.</li>
		<li>Press Set Up to highlight Remove in red. Remove the syringe and fill the syringe with isoflurane using the bottle top adapter.</li>
		<li>Connect the Syringe to the anesthesia system.</li>
		<li>Prime the anesthetic delivery tubing. Press Set Up to highlight Prime Tube in red. Press Down until the anesthetic travels through the syringe and into the black fitting on the vaporizer block.</li>
		<li>Enable anesthesia. Press Set Up to highlight Enable in red. Use the Up and Down arrows to select Yes. Press Run/Back to return to the Main Menu.</li>
		<li>Select the air supply and minute volume. Press Set Up to highlight Air Supply in red. Use the Up and Down arrows to select Internal Pump. Press Set Up to highlight Minute Vol in red. Set flow rate to 250 ml/min.</li>
	</ol>
	</li>
	<li>For Physiological Monitoring
	<ol>
		<li>Set the minimum detected heart rate to Mouse (240). Press Set Up to access Main Menu &#62; MouseStat. Use up and down arrows to set minimum heart rate.</li>
	</ol>
	</li>
</ol>
3. Begin Anesthesia Delivery
<ol>
	<li>Induce the Mouse
	<ol>
		<li>Press Run/Back twice to enter run mode and begin airflow.</li>
		<li>Place the mouse in the induction chamber, closing the lid tightly. Adjust the Anesthetic Agent Concentration knob to 3%.</li>
		<li>Monitor until the mouse has reached the desired plane of anesthesia, determined by loss of righting reflex. Adjust the Anesthetic Agent Concentration as necessary.</li>
		<li>Once the animal loses its righting reflex and is sufficiently anesthetized, turn the Anesthesia concentration dial to 0%. The authors have previously found that allowing the airflow to flush the induction chamber 30-60 sec is sufficient to purge the chamber without reversing the anesthetic depth1.</li>
		<li>Quickly open the white clamps to direct the air to the facemask, and close the blue clamps leading to the induction chamber.</li>
		<li>Open the chamber away from researcher, remove the mouse, and immediately fit the nose cone.</li>
		<li>Center the animal on an infrared warming pad, set to maintain body temperature at 37 &#176;C via a rectal probe on a feedback loop.</li>
		<li>When the mouse is stable on the nose cone, adjust the concentration of isoflurane to 1.5% or as needed for maintenance by turning the Anesthetic Agent Concentration knob.</li>
		<li>Reduce minute volume for maintenance. The minimum flow rate to support the animal is equal to 1.5-2.2 times the animal&#39;s minute volume (for a 30 g mouse, a minimum of 52 ml/min). Refer to the manufacturer&#39;s instructions for a recommended flow rate setting specific to the nose cone style and adjust as necessary. Press Set Up to access the Main Menu, and then press Set Up until Minute Vol is highlighted in red. Use the Up and Down arrows to adjust target flow. Press Run/Back to return to the main screen.</li>
		<li>Confirm depth of anesthesia as determined by a lack of withdrawal reflex during an interdigital pinch. Apply ophthalmic ointment to the eyes to prevent dryness during anesthesia.</li>
	</ol>
	</li>
</ol>
4. Begin Physiological Monitoring
<ol>
	<li>Place sensor over pad of hind paw. Position the sensor so that the red light is beneath the paw and illuminates the paw. Use the up and down arrows on the screen to display the oxiwave. The animal is now safely anesthetized using a low flow, syringe-driven, digital vaporizer.</li>
</ol>
5. Remove the Animal
<ol>
	<li>Turn Off the Anesthesia Delivery.
	<ol>
		<li>To stop the delivery of anesthetic, turn Anesthetic Agent Concentration knob to Min (or 0%) and remove animal from facemask.</li>
		<li>Monitor the mouse during anesthesia recovery. Once the mouse has become fully ambulatory, return it to the cage.</li>
	</ol>
	</li>
</ol>

<ol>
	<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> JAALAS. 54 (6), 756-762 (2015).</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 J Anaesth. 57 (5), 464-471 (2013).</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>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>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> Am J Vet Res. 67 (6), 936-940 (2006).</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>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. , 335 (2011).</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> Brit J Anaesth. 26 (5), 323-332 (1954).</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=Physical+principles+and+maintenance.">Physical principles and maintenance.</a> Anaesthetic equipment. , (1985).</li><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> Br. J. Clin. Pharmacol. 12 (2), 281-282 (1971).</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> Anes Analg. 111 (5), 1176-1179 (2010).</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> Anes Analg. 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> Exp Anim. 56 (2), 131-137 (2007).</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> Biol Pharm Bull. 30 (9), 1716-1720 (2007).</li><li>Voightsverger, S., 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=Sevoflurane+ameliorates+gas+exchange+and+attenuates+lung+damage+in+experimental+lipopolysaccharide-induced+lung+injury.">Sevoflurane ameliorates gas exchange and attenuates lung damage in experimental lipopolysaccharide-induced lung injury.</a> Anesthesiology. 111 (6), 1238-1248 (2009).</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. , (2011).</li></ol>

This project was supported with equipment and funding by Kent Scientific Corporation, the American Heart Association to CJ Goergen (SDG18220010), and Purdue University. The authors Krista Bigiarelli and Irina Toore are employees of Kent Scientific Corporation that produces equipment used in this article. Open access publication of this article is sponsored by Kent Scientific Corporation.

The digital low-flow anesthesia system allows the user to effectively anesthetize mice at very low flow rates without the use of any compressed gas. This differs greatly from standard passive vaporizers, most of which require a compressed gas source at minimum flow rates of about 500 ml/min. Standard vaporizers utilize dials that lack precision between gradations, and they must be serviced annually to maintain accuracy. A syringe driven anesthetic system can provide a specific concentration of anesthetic at the set flow rate to calculate the exact necessary speed of the syringe pump. Routine calibrations are unnecessary, resulting in additional cost and time savings.
The recommended minimum flow rate to maintain an animal on a non-rebreathing circuit is 1.5-2.2 times the animal&#39;s minute volume. The flow rate of 100 ml/min used in this study exceeded this minimum to deliver sufficient anesthetic to the animals. The flow rate settings are critical for this anesthetic delivery technique, as the flow rate is directly related to the amount of isoflurane used for a given timeframe. When used at low flow rates, this technique can greatly reduce the amount of isoflurane required during use, while the animal is still anesthetized effectively1,19-21.
New equipment costs between traditional vaporizers and low-flow digital vaporizers are comparable. However, the digital low-flow anesthesia system has the ability to deliver either isoflurane or sevoflurane. This eliminates the need for designated isoflurane and sevoflurane precision vaporizers, reducing initial equipment costs for groups using both anesthetic agents. Recently published comparisons between vaporizer technologies have suggested cost savings over time when using a low-flow digital vaporizer1,19,20. The results of these comparisons could be used to approximate potential cost savings over the course of a year. Assuming typical usage settings performed in 2 hr increments, 5 days a week for 52 weeks, a traditional isoflurane vaporizer will consume 3.8 L of isoflurane, or twelve 250 ml bottles. A low-flow digital vaporizer used at the same frequency would consume just 0.32 L, or two 250 ml bottles. Charcoal canister consumption is also reduced. Assuming that each canister holds 50 g of scavenged waste gas, a traditional vaporizer will fill approximately 21 charcoal canisters over the course of a year. In comparison, a low-flow digital vaporizer will require 6 or fewer. A traditional vaporizer would require approximately 5 large gas cylinders per year, each with a capacity of 9,500 L. The internal air pump, available in some models of digital low-flow vaporizers, eliminates the requirement for compressed gas. If compressed gas were to be used, the system would use only 1 cylinder per year1.
The technique can be modified based on need. Low-flow digital vaporizers allow the user to adjust anesthetic depth quickly and precisely. If the anesthetic depth must be increased or decreased, the user can increase the anesthetic concentration in 0.1% increments using the dial on the top of the system. The flow rate can also be adjusted as needed throughout the procedure. This protocol utilizes a 2 ml syringe, though larger syringe sizes are available for longer procedures. The internal air pump offers users the option to anesthetize animals without requiring a compressed gas source. For procedures requiring compressed gas or supplemental oxygen, the user has the option to connect a gas source to the low-flow system rather than using surrounding air. The user can continue to deliver the selected air source throughout the procedure, or can switch between the internal pump and a compressed gas source as needed. For example, the user may set the system to deliver room air via the internal pump during induction and maintenance, but deliver supplemental oxygen during recovery.
Though there are many advantages to using a low-flow digital vaporizer, there are limitations as well. Because a flush valve is not included, manually flushing the chamber with clean air before opening is the only way to purge the induction chamber. This system is designed to operate at low flow rates only and does not deliver anesthesia above flow rates of 800 ml/min, where traditional vaporizers can be used with flow rates up to 10 L/min. This particular system is therefore only suitable for small animal species. Additionally, the system holds less anesthetic agent compared to a traditional vaporizer. There may be situations where the syringe must be refilled during a procedure. However, delays during refilling can be reduced by pre-filling a second syringe nearby to replace the empty syringe. Syringe sizes up to 10 ml are available to reduce the need to refill syringes mid-procedure. Finally, unlike a traditional vaporizer, the low-flow digital vaporizer requires electricity. Batteries are available for use in instances where electrical power is unavailable or in the event of a power outage.
Previous studies have shown that low-flow digital systems consume less isoflurane, carrier gas, and charcoal canisters compared to a traditional anesthesia system1,19,20. The reduction in scavenged anesthetic gas also could identify a reduction in waste anesthetic gas, though further work is needed in these areas. Infrared gas spectroscopy can be used to monitor waste isoflurane production, and dosimeter badges can be used to quantify isoflurane exposure to laboratory personnel in future comparisons.
In summary, this technique for anesthetic delivery will be beneficial to groups performing rodent anesthesia due to improved safety, efficacy, and precision over traditional systems.

There are many precision vaporizers available for veterinary use that operate between flow rates of 0.5-10 L/min2. These flow rates are not ideal for rodents, as the range is high compared to their small respiratory minute volume. High flow rates are not recommended in veterinary practice due to their promotion of hypothermia and drying of the respiratory tract3,4. Furthermore, many common veterinary vaporizer manufacturer manuals warn that high flow rates may cause an increase in the occurrence of backpressure fluctuations. It has also been shown that many standard vaporizers become inaccurate below flow rates of 500 ml/min, and this rate is regarded as a minimum flow rate in the veterinary field5-7.
An animal can be maintained on a T-piece circuit or modified bain circuit using a flow rate as low as 1.5-2.2 times the animal&#39;s minute volume8-10. These flow rates are considered sufficient to prevent rebreathing of expired gases and prevent an increase in blood carbon dioxide concentrations8. Using this flow rate recommendation, a 30 g mouse could be maintained at a flow rate as low as 52 ml/min, nearly ten times less than the accepted 500 ml/min minimum of a traditional vaporizer.
While a traditional vaporizer depends on gas flow and atmospheric pressure for passive anesthetic vaporization, a direct injection vaporizer measures the total fresh gas flow and injects the vapor directly into the gas stream2. Some direct injection vaporizers utilize a syringe pump to administer anesthetic into the gas stream. Computerized controls allow these systems to automatically adjust the syringe pump speed to inject the volume of liquid agent required to reach the desired concentration of anesthetic. Syringe driven vaporizers are available and approved for clinical and pediatric use, and many similar configurations are regarded as anesthetic conserving devices in clinical practice11-16. Shortly after their approval, anesthetic conserving devices with syringe pump vaporizers were adapted for use in animal studies8,17,18. Unlike a traditional vaporizer, a direct injection system utilizing a syringe pump is not limited by a minimum flow rate in order to maintain accuracy. For this reason, this technology is ideal for use in rodent anesthesia and other instances where low flow rates are necessary. The benefits and potential cost savings associated with this vaporizer design inspired the development of new anesthesia systems designed specifically for rodents1,19,20. This new system also incorporates a built-in air pump, allowing the user to administer anesthesia without requiring a compressed gas source. As an additional benefit, the system is pre-calibrated for use with both isoflurane and sevoflurane. With the introduction of this vaporizer technology in the laboratory animal field, it is now possible to anesthetize small laboratory animals at flow rates closer to recommended levels without the need for compressed gas.

<table border="0" cellpadding="0" cellspacing="0" width="873">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Anesthetic Equipment</td>
			<td style="width:131px;"></td>
			<td style="width:119px;"></td>
			<td style="width:340px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">SomnoSuite Low-Flow Digital Anesthesia System</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">SOMNO</td>
			<td style="width:340px;">Includes anti-spill, anti-vapor bottle top adapter; Y adapter tubing; charcoal scavenging filter</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">MouseSTAT Pulse Oximeter &#38; Heart Rate Monitor</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">SS-MSTAT-Module</td>
			<td style="width:340px;">Integrated into SomnoSuite</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">MouseSTAT Mouse Paw Sensor</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">MSTAT-MSE</td>
			<td style="width:340px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">2mL Glass Syringe</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">SOMNO-2ML</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Low-Cost Induction Chamber, 0.5L</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">SOMNO-0705</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Low Profile Facemask, x-small</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">SOMNO-0304</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Animal Warming</td>
			<td style="width:131px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:283px;">PhysioSuite Physiological Monitoring System with RightTemp Homeothermic Warming</td>
			<td style="width:131px;">Kent Scientific Corporation</td>
			<td style="width:119px;">PS-RT</td>
			<td style="width:340px;">Includes infrared warming pad, rectal probe, and pad temperature probe</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Anesthetic Agents 
			and Medications</td>
			<td style="width:131px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Isoflurane (250mL bottle)</td>
			<td style="width:131px;">Piramal Healthcare</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Puralube Opthalmic Ointment</td>
			<td style="width:131px;">Perrigo</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Animals
3 adult C57/BL6NTac female mice (Taconic, age 6-7 weeks; weight 15+/-1 g) were anesthetized and maintained with 1.3-1.5% isoflurane while heart rate, oxygen saturation, and respiration rate were monitored. All mice were Murine Pathogen Free as determined by routine vendor testing before arrival to the facility. The animals were group-housed in microisolation caging and provided free-access to standard rodent chow and water by bottle.
Isoflurane Usage
The low-flow anesthesia system measures the amount of anesthetic agent remaining in the syringe during use. The volume in the syringe, as measured by the anesthesia system, was noted as the animal was transferred to the nose cone, and again at the end of the maintenance period. The final volume was subtracted from the initial volume to quantify the amount of anesthetic consumed during the maintenance period (Figure 1).
Physiological Parameters
Heart rate, SpO2, and respiration rate were monitored during maintenance via pulse oximetry (Figures 2-4). Body temperature was maintained at 37.5 &#176;C via an infrared warming pad. Each mouse was successfully maintained at low flow rates of 100 ml/min of room air under a surgical plane of anesthesia for 60 min, as determined by a lack of withdrawal reflex from an interdigital pinch. The mouse did not awake or respond to interdigital pinches applied intermittently during the maintenance period. The animals&#39; heart rates (Figure 2), blood oxygen (Figure 3), and respiratory rate (Figure 4) remained relatively stable throughout the study. Due to animal and sensor positioning, the respiratory rate signal from Mouse 1 and Mouse 3 was intermittent and the measurement was interrupted. When the animal&#39;s positioning was adjusted, the signal improved and the measured respiratory rate was comparable to others at similar time points. The low-flow digital anesthesia system used an average of 0.63 ml of isoflurane during the 60 min of maintenance period (Figure 1).
<img alt="Figure 1" src="/files/ftp_upload/54436/54436fig1.jpg" /> 
Figure 1: Isoflurane Usage. The amount of isoflurane used in ml for three different mice over 1 hr of anesthesia maintenance using the digital low-flow anesthesia system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54436/54436fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54436/54436fig2.jpg" /> 
Figure 2: Heart Rate. The heart rate of three mice in beats per min (bpm) 5-60 min after initial anesthetic induction with the digital low-flow anesthesia system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54436/54436fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54436/54436fig3.jpg" /> 
Figure 3: Oxygen Saturation. The blood oxygen saturation levels (%) of three mice 5-60 min after initial anesthetic induction with the digital low-flow anesthesia system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54436/54436fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54436/54436fig4.jpg" /> 
Figure 4: Respiration Rate. The respiration rate of three mice in breaths per min (bpm) 5-60 min after initial anesthetic induction with the digital low-flow anesthesia system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54436/54436fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Use of a Low-flow Digital Anesthesia System for Mice and Rats at JoVE.com

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

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-low-flow-digital-anesthesia-system-for-mice-and-rats

Nanyang Technological University

Research

JoVE Journal

Medicine

17.8K Views.  Purdue University. 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. 

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

A Novel Rescue Technique for Difficult Intubation and Difficult Ventilation

We describe a novel non surgical technique to maintain oxygenation and ventilation in a case of difficult intubation and difficult ventilation, which works especially well with poor mask fit.
Can not intubate, can not ventilate" (CICV) is a potentially life threatening situation. In this video we present a simulation of the technique we used in a case of CICV where oxygenation and ventilation were maintained by inserting an endotracheal tube (ETT) nasally down to the level of the naso-pharynx while sealing the mouth and nares for successful positive pressure ventilation.
A 13 year old patient was taken to the operating room for incision and drainage of a neck abcess and direct laryngobronchoscopy. After preoxygenation, anesthesia was induced intravenously. Mask ventilation was found to be extremely difficult because of the swelling of the soft tissue. The face mask could not fit properly on the face due to significant facial swelling as well. A direct laryngoscopy was attempted with no visualization of the larynx. Oxygen saturation was difficult to maintain, with saturations falling to 80%. In order to oxygenate and ventilate the patient, an endotracheal tube was then inserted nasally after nasal spray with nasal decongestant and lubricant. The tube was pushed gently and blindly into the hypopharynx. The mouth and nose of the patient were sealed by hand and positive pressure ventilation was possible with 100% O2 with good oxygen saturation during that period of time. Once the patient was stable and well sedated, a rigid bronchoscope was introduced by the otolaryngologist showing extensive subglottic and epiglottic edema, and a mass effect from the abscess, contributing to the airway compromise. The airway was secured with an ETT tube by the otolaryngologist.This video will show a simulation of the technique on a patient undergoing general anesthesia for dental restorations.

We describe a technique to maintain oxygenation and ventilation using an endotracheal tube inserted nasally to the level of the naso-pharynx while sealing the mouth and nares for successful positive pressure ventilation.

We describe a novel non surgical technique to maintain oxygenation and ventilation in a case of difficult intubation and difficult ventilation, which ...

Guidelines for Elective Pediatric Fiberoptic Intubation

Fiberoptic intubation in pediatric patients is often required especially in difficult airways of syndromic patients i.e. Pierre Robin Syndrome. Small babies will desaturate very quickly if ventilation is interrupted mainly to high metabolic rate. We describe guidelines to perform a safe fiberoptic intubation while maintaining spontaneous breathing throughout the procedure. Steps requiring the use of propofol pump, fentanyl, glycopyrrolate, red rubber catheter, metal insuflation hook, afrin, lubricant and lidocaine spray are shown.

We describe guidelines to perform a safe and efficient elective fiberoptic intubation in pediatric patients while maintaining spontaneous ventilation.

Fiberoptic intubation in pediatric patients is often required especially in difficult airways of syndromic patients i.e. Pierre Robin Syndrome. Small ...

Use of a Hanging Weight System for Coronary Artery Occlusion in Mice

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. Cardioprotection by ischemic preconditioning (IP) remains an area of intense investigation. To further elucidate its molecular basis, the use of knockout mouse studies is particularly important 7, 14, 30, 39. Despite the fact that previous studies have already successfully performed cardiac ischemia and reperfusion in mice, this model is technically very challenging. Particularly, visual identification of the coronary artery, placement of the suture around the vessel and coronary occlusion by tying off the vessel with a supported knot is technically difficult. In addition, re-opening the knot for intermittent reperfusion of the coronary artery during IP without causing surgical trauma adds additional challenge. Moreover, if the knot is not tied down strong enough, inadvertent reperfusion due to imperfect occlusion of the coronary may affect the results. In fact, this can easily occur due to the movement of the beating heart.
Based on potential problems associated with using a knotted coronary occlusion system, we adopted a previously published model of chronic cardiomyopathy based on a hanging weight system for intermittent coronary artery occlusion during IP 39. In fact, coronary artery occlusion can thus be achieved without having to occlude the coronary by a knot. Moreover, reperfusion of the vessel can be easily achieved by supporting the hanging weights which are in a remote localization from cardiac tissues.
We tested this system systematically, including variation of ischemia and reperfusion times, preconditioning regiments, body temperature and genetic backgrounds39. In addition to infarct staining, we tested cardiac troponin I (cTnI)
as a marker of myocardial infarction in this model. In fact, plasma levels of cTnI correlated with infarct sizes (R2=0.8). Finally, we could show in several studies that this technique yields highly reproducible infarct sizes during murine IP and myocardial infarction6, 8, 30, 40, 41. Therefore, this technique may be helpful for researchers who pursue molecular mechanisms involved in cardioprotection by IP using a genetic approach in mice with targeted gene deletion. Further studies on cardiac IP using transgenic mice may consider this technique.

A murine model for myocardial ischemia and ischemic preconditioning is an important tool study cardioprotective mechanisms in vivo. Here, we report an easy applicable in situ model for cardiac IP using a hanging-weight system for coronary artery occlusion.

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. ...

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 ...

Use of a Hanging-weight System for Liver Ischemia in Mice

Acute liver injury due to ischemia can occur during several clinical procedures e.g. liver transplantation, hepatic tumor resection or trauma repair and can result in liver failure which has a high mortality rate1-2. Therefore murine studies of hepatic ischemia have become an important field of research by providing the opportunity to utilize pharmacological and genetic studies3-9. Specifically, conditional mice with tissue specific deletion of a gene (cre, flox system) provide insights into the role of proteins in particular tissues10-13 . Because of the technical difficulty associated with manually clamping the portal triad in mice, we performed a systematic evaluation using a hanging-weight system for portal triad occlusion which has been previously described3. By using a hanging-weight system we place a suture around the left branch of the portal triad without causing any damage to the hepatic lobes, since also the finest clamps available can cause hepatic tissue damage because of the close location of liver tissue to the vessels. Furthermore, the right branch of the hepatic triad is still perfused thus no intestinal congestion occurs with this technique as blood flow to the right hepatic lobes is preserved. Furthermore, the portal triad is only manipulated once throughout the entire surgical procedure. As a result, procedures like pre-conditioning, with short times of ischemia and reperfusion, can be easily performed. Systematic evaluation of this model by performing different ischemia and reperfusion times revealed a close correlation of hepatic ischemia time with liver damage as measured by alanine (ALT) and aspartate (AST) aminotransferase serum levels3,9. Taken together, these studies confirm highly reproducible liver injury when using the hanging-weight system for hepatic ischemia and intermittent reperfusion. Thus, this technique might be useful for other investigators interested in liver ischemia studies in mice. Therefore the video clip provides a detailed step-by-step description of this technique.

We established a novel murine model of a hanging weight system for portal triad occlusion. This technique may be useful for future investigations of ischemia in murine hepatic models.

Acute liver injury due to ischemia can occur during several clinical procedures e.g. liver transplantation, hepatic tumor resection or trauma repair and ...

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 ...

Intraoperative Detection of Subtle Endometriosis: A Novel Paradigm for Detection and Treatment of Pelvic Pain Associated with the Loss of Peritoneal Integrity

Endometriosis is a common disease affecting 40 to 70% of reproductive-aged women with chronic pelvic pain (CPP) and/or infertility. The purpose of this study was to demonstrate the use of a blue dye (methylene blue) to stain peritoneal surfaces during laparoscopy (L/S) to detect the loss of peritoneal integrity in patients with pelvic pain and suspected endometriosis. Forty women with CPP and 5 women without pain were evaluated in this pilot study. During L/S, concentrated dye was sprayed onto peritoneal surfaces, then aspirated and rinsed with Lactated Ringers solution. Areas of localized dye uptake were evaluated for the presence of visible endometriotic lesions. Areas of intense peritoneal staining were resected and some fixed in 2.5% buffered gluteraldehyde and examined by scanning (SEM) electron microscopy. Blue dye uptake was more common in women with endometriosis and chronic pelvic pain than controls (85% vs. 40%). Resection of the blue stained areas revealed endometriosis by SEM and loss of peritoneal cell-cell contact compared to normal, non-staining peritoneum. Affected peritoneum was associated with visible endometriotic implants in most but not all patients. Subjective pain relief was reported in 80% of subjects. Based on scanning electron microscopy, we conclude that endometrial cells extend well beyond visible implants of endometriosis and appear to disrupt the underlying mesothelium. Subtle lesions of endometriosis could therefore cause pelvic pain by disruption of peritoneal integrity, allowing menstrual or ovulatory blood and associated pain factors access to underlying sensory nerves. Complete resection of affected peritoneum may provide a better long-term treatment for endometriosis and CPP. This simple technique appears to improve detection of subtle or near invisible endometriosis in women with CPP and minimal visual findings at L/S and may serve to elevate diagnostic accuracy for endometriosis at laparoscopy.

Loss of peritoneal integrity provides a new paradigm to understand and treat chronic pelvic pain in women with mild forms of endometriosis and can be easily detected using intraoperative instillation of dye at the time of laparoscopy.

Endometriosis is a  common disease affecting 40 to 70% of reproductive-aged women with chronic  pelvic pain (CPP) and/or infertility. The purpose of this ...

A Model of Glaucoma Induced by Circumlimbal Suture in Rats and Mice

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known risk factor for glaucoma. This protocol demonstrates a step-by-step guide on this technique in Long Evans rats and C57BL/6 mice. Under general anesthesia, a "purse-string" suture is applied on the conjunctiva, around the equator and behind the limbus of the eye. The fellow eye serves as an untreated control. Over the duration of our study, which was a period of 8 weeks for rats and 12 weeks for mice, IOP remained elevated, as measured regularly by rebound tonometry in conscious animals without topical anesthesia. In both species, the sutured eyes showed electroretinogram features consistent with preferential inner retinal dysfunction. Optical coherence tomography showed selective thinning of the retinal nerve fiber layer. Histology of the rat retina in cross-section found reduced cell density in the ganglion cell layer, but no change in other cellular layers. Staining of flat-mounted mouse retinae with a ganglion cell specific marker (RBPMS) confirmed ganglion cell loss. The circumlimbal suture is a simple, minimally invasive and cost-effective way to induce ocular hypertension that leads to ganglion cell injury in both rats and mice.

Chronic ocular hypertension is induced by applying a circumlimbal suture in rats and mice, leading to functional and structural deterioration of the retinal ganglion cells consistent with glaucoma.

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known ...

A Pre-Clinical Porcine Model of Orthotopic Heart Transplantation

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart failure. Multiple small-animal models of heart transplantation have been used to study the acute and long-term effects of novel therapies. However, few are tested and demonstrated success in clinical trials. It is of critical importance to evaluate new therapies in a clinically relevant large-animal model for efficient and reliable translation of basic studies&#39;&#160;findings. Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and previously used to investigate novel cardioprotective strategies. This procedure focuses on acute ischemia-reperfusion injury and is a reliable method to investigate novel interventions which have been tested and validated in smaller experimental models, such as the murine model. We demonstrate its usefulness in assessing cardiac performance during the early post-transplantation period and other potential possibilities enabled by the model.

Here, we describe a pre-clinical large-animal (porcine) model of orthotopic heart transplantation that has been firmly established and utilized to investigate novel cardioprotective strategies.

Fifty-years following the first successful report, cardiac transplantation remains the gold-standard treatment for eligible patients with advanced heart ...

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.

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 ...