Name: use of a low-flow digital anesthesia system for mice and rats
Uploaded: 2016-09-07T13:00:00.000+00:00
Duration: 8 min 6 s
Description: Watch this Scientific Journal Video about Use of a Low-flow Digital Anesthesia System for Mice and Rats at JoVE.com

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>

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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><tbody><tr><td>&lt;em&gt;Anesthetic Equipment&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>SomnoSuite Low-Flow Digital Anesthesia System</td><td>Kent Scientific Corporation</td><td>SOMNO</td><td>Includes anti-spill, anti-vapor bottle top adapter; Y adapter tubing; charcoal scavenging filter</td></tr><tr><td>MouseSTAT Pulse Oximeter &amp;amp; Heart Rate Monitor</td><td>Kent Scientific Corporation</td><td>SS-MSTAT-Module</td><td>Integrated into SomnoSuite</td></tr><tr><td>MouseSTAT Mouse Paw Sensor</td><td>Kent Scientific Corporation</td><td>MSTAT-MSE</td><td></td></tr><tr><td>2 ml Glass Syringe</td><td>Kent Scientific Corporation</td><td>SOMNO-2ML</td><td></td></tr><tr><td>Low-Cost Induction Chamber, 0.5 L</td><td>Kent Scientific Corporation</td><td>SOMNO-0705</td><td></td></tr><tr><td>Low Profile Facemask, x-small</td><td>Kent Scientific Corporation</td><td>SOMNO-0304</td><td></td></tr><tr><td>&lt;em&gt;Animal Warming&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>PhysioSuite Physiological Monitoring System with RightTemp Homeothermic Warming</td><td>Kent Scientific Corporation</td><td>PS-RT</td><td>Includes infrared warming pad, rectal probe, and pad temperature probe</td></tr><tr><td>&lt;em&gt;Anesthetic Agents&lt;br/&gt; and Medications&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Isoflurane (250 ml bottle)</td><td>Piramal Healthcare</td><td></td><td></td></tr><tr><td>Puralube Opthalmic Ointment</td><td>Perrigo</td><td></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/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/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/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/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 ...

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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

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Neuroscience

Proper Positioning and Restraint of a Rat Hind Limb for Focused High Resolution Imaging of Bone Micro-architecture Using In Vivo Micro-computed Tomography

The use of in vivo micro-computed tomography (&#181;CT) is a powerful tool which involves the non-destructive imaging of internal structures at high resolutions in live animal models. This allows for repeated imaging of the same rodent over time. This feature not only reduces the total number of rodents required in an experimental design and thereby reduces the inter-subject variation that can arise, but also allows researchers to assess longitudinal or life-long responses to an intervention. To acquire high quality images that can be processed and analyzed to more accurately quantify outcomes of bone micro-architecture, users of in vivo &#181;CT scanners must properly anesthetize the rat, and position and restrain the hind limb. To do this, it is imperative that the rat be anesthetized to a level of complete relaxation, and that pedal reflexes are lost. These guidelines may be modified for each individual rat, as the rate of isoflurane metabolism can vary depending on strain and body size. Proper technique for in vivo &#181;CT image acquisition enables accurate and consistent measurement of bone micro-architecture within and across studies.

Proper Positioning and Restraint of a Rat Hind Limb for Focused High Resolution Imaging of Bone Micro-architecture Using In Vivo Micro-computed Tomography

This paper instructs users of in vivo micro-computed tomography (&#181;CT) scanners how to anesthetize, correctly position and restrain the hind limb of a rat for minimal movement during high-resolution imaging of the tibia. The result is high quality images that can be processed to accurately quantify bone micro-architecture.

The use of in vivo micro-computed tomography (&#181;CT) is a powerful tool which involves the non-destructive imaging of internal structures at high ...

Bioengineering

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

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

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

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

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for erectile dysfunction. However, the methods used vary between laboratories, and pitfalls still exist. The goal of this study was to describe a surgical technique that would provide a reliable and reproducible model. By exposing the ischiocavernosus muscle at its point of insertion on the ischial tuberosity, the penile crus could be cannulated with minimal dissection and injury to the structures involved in erectile function. Repeated stimulation of the CN, without the need for lifting and drying, was achieved by using a 125 &#181;m bipolar silver electrode and biocompatible silicon glue to isolate the electrode-nerve complex. This method prevents neuropraxia by reducing stretching and drying the nerve and provides complete isolation of the nerve, negating electrical leakage and preventing stimulation of alternative pathways.

This study describes a simplified surgical procedure and technique for performing cavernous nerve stimulation with the isolation of the nerve-electrode complex using silicone glue and intracavernous pressure measurement.

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

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

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

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

Direct Intrabronchial Administration to Improve the Selective Agent Deposition Within the Mouse Lung

Intratracheal (IT) administration of experimental agents is an essential technique in murine models of diffuse lung diseases, such as bleomycin-induced pulmonary fibrosis.&#160; However, distribution of intratracheally-administered agents to the distal mouse lung is often asymmetric, with lung parenchymal concentrations increased in the smaller (but equally accessible) left lung of the mouse.&#160; Described in this report is a novel intrabronchial (IB) approach to cannulate the left and/or right lungs of living mice non-operatively.&#160; It is also demonstrated how this approach can be used to selectively administer agents to one lung or adapted (via dose-adjusted IB delivery) to improve the left-right symmetry of lung delivery of experimental agents, thereby improving models of diffuse lung disease such as bleomycin-induced pulmonary fibrosis.

Intratracheal (IT) administration of experimental agents in mice often results in asymmetric delivery to the distal lungs.&#160; In this report, we describe a direct intrabronchial (IB) approach to cannulate each lung in living mice non-operatively.&#160; This approach can be used to selectively administer agents to one lung or may be adapted to improve the symmetric agent delivery to both lungs.

Intratracheal (IT) administration of experimental agents is an essential technique in murine models of diffuse lung diseases, such as bleomycin-induced ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

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 ...

Intracerebroventricular Delivery of Gut-Derived Microbial Metabolites in Freely Moving Mice

The impact of gut microbiota and their metabolites on host physiology and behavior has been extensively investigated in this decade. Numerous studies have revealed that gut microbiota-derived metabolites modulate brain-mediated physiological functions through intricate gut-brain pathways in the host. Short-chain fatty acids (SCFAs) are the major bacteria-derived metabolites produced during dietary fiber fermentation by the gut microbiome. Secreted SCFAs from the gut can act at multiple sites in the periphery, affecting the immune, endocrine, and neural responses due to the vast distribution of SCFAs receptors. Therefore, it is challenging to differentiate the central and peripheral effects of SCFAs through oral and intraperitoneal administration of SCFAs. This paper presents a video-based method to interrogate the functional role of SCFAs in the brain via a guide cannula in freely moving mice. The amount and type of SCFAs in the brain can be adjusted by controlling the infusion volume and rate. This method can provide scientists with a way to appreciate the role of gut-derived metabolites in the brain.

Gut-derived microbial metabolites have multifaceted effects leading to complex behavior in animals. We aim to provide a step-by-step method to delineate the effects of gut-derived microbial metabolites in the brain via intracerebroventricular delivery via a guide cannula.

The impact of gut microbiota and their metabolites on host physiology and behavior has been extensively investigated in this decade. Numerous studies have ...

Measuring Pressure Volume Loops in the Mouse

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

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

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

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

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Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Proper Positioning and Restraint of a Rat Hind Limb for Focused High Resolution Imaging of Bone Micro-architecture Using In Vivo Micro-computed Tomography

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

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

Laminectomy and Spinal Cord Window Implantation in the Mouse

Direct Intrabronchial Administration to Improve the Selective Agent Deposition Within the Mouse Lung

In vivo Calcium Imaging in Mouse Inferior Olive

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

Intracerebroventricular Delivery of Gut-Derived Microbial Metabolites in Freely Moving Mice

Measuring Pressure Volume Loops in the Mouse