A Merit Grant from the Department of Veteran Affairs and a T32-HL098062 grant from the NHLBI of the National Institutes of Health supported this work. We wish to gratefully acknowledge the advice of Randolph H. Hasting, M.D., Ph.D. and the advice and support of the Veterinary Medical Unit of the VA San Diego Healthcare System.

1. Animal Preparation
<ol>
	<li>Obtain mice that are older than 8 weeks and more than 20 g&#160;(smaller mice can be intubated by an expert).</li>
	<li>Anesthesia
	<ol>
		<li>Inject mice with&#160; 20 mg/kg, each, of ketamine and xylazine intraperitoneally as a preanesthetic. (This dose is insufficient to fully anesthetize the mouse, but facilitates the safe transfer, after intubation, to mechanical ventilation.)&#160;However, adjustment of the dosages may be necessary according to the anesthetic response in consultation with the institutional veterinarian.)</li>
		<li>Induce anesthesia in mice with 3.5% isoflurane/oxygen in an induction chamber for 90-120 sec.</li>
		<li>Observe the respiratory rate carefully. It should slow progressively to not less than 45 bpm.</li>
	</ol>
	</li>
</ol>
2. Preparation for Intubation
<ol>
	<li>Remove the mouse from the induction chamber and scruff tightly.</li>
	<li>Use cushioned fine forceps to gently extend the tongue from the mouth.</li>
	<li>Maintain extension of the tongue by holding the tongue between two fingers and applying gentle force.</li>
	<li>Pull the mouse up on the speculum of the otoscope with a vertical motion.</li>
	<li>Look through the ocular of the otoscope while scruffing tightly and pull the mouse up gently by the tongue and the scruff.</li>
	<li>Look carefully for the vocal cords. They should be easily visible at this time. The laryngis aditus (the structure of the laryngeal opening) should look relatively white. There should be movement of the cords with each breath. If the cords are not moving or are poorly visualized, rotate the animal slightly and gently hyperextend the neck.</li>
	<li>With the dominant hand, hold the 20 G, 1 in&#160;catheter, with a 1 cm length of the PE10 tubing extending through the tip of the catheter, like a pencil and insert it into the side of the speculum. The PE tubing serves as a stylet or bougie.</li>
	<li>Direct the PE10 tubing (stylet) through the vocal cords and advance the 20 G&#160;catheter (endotracheal tube) over the PE tubing until the hub is at the level of the lower incisor. Remove this stylet quickly.</li>
	<li>Gently take the animal off of the speculum and verify the tube location by placing the animal on a mechanical ventilator with continuous 2% isoflurane to maintain general anesthesia throughout the experiment. Visualize expired air (bubbles) going through a PEEP trap to confirm intubation.&#160;Although esophageal intubation may result in a few bubbles, it will not be as pronounced or as consistent with endotracheal intubation.&#160;Also, observe an airway pressure tracing for negative deflections (Figure 1).&#160;These confirm proper placement of the tube.&#160;Alternatively, place a small amount of water into IV tubing and connect it to the ET tube.&#160;Verifying movement to and from the mouse with respiration2.&#160;In the case of esophageal intubation, the pressure tracing will reveal significantly higher pressures and no negative deflections.&#160;Although end-tidal CO2 would be helpful, technically this would be very difficult given the small tidal volumes (~200 &#181;l) and the equipment to do this would be very expensive.&#160;The simple techniques described are fully sufficient and much less expensive.&#160;If isoflurane anesthesia is not available, all procedures can be conducted using ketamine 80-120 mg/kg in conjunction with xylazine 10-20 mg/kg for general anesthesia. &#160;&#160;</li>
</ol>
3. Alternative Technique: Direct Laryngoscopy with the Otoscope Technique is Used Readily for Other Purposes,&#160;Mainly&#160;for Direct Instillation of Study Substances into the Lung.
<ol>
	<li>Induce anesthesia in the animals with isoflurane alone for this procedure at 3.5% for 90-120 sec in the induction chamber.</li>
	<li>Scruff the animal tightly at the base of the skull and extend the tongue gently with fine forceps.</li>
	<li>Holding the tongue gently, pull the animal straight up on the speculum (unmodified) until the animal cannot be pulled up further.</li>
	<li>The vocal cords can be visualized most often by this technique alone, but rotate the animal and hyper-extended the neck to bring them into full view.</li>
	<li>Advance a gel-loading pipette containing the instillate to the glottis and instill the fluid. Because the speculum makes a seal with the hypopharynx the mouse will aspirate any residual liquid that does not go through the cords. This is easily verified by the rhonchorous sounds the mouse now makes when breathing until the fluid is distributed fully in the lungs.</li>
	<li>Alternatively, connect PE10 tubing to a 0.5 ml syringe containing 50 &#181;l of fluid with an air bolus behind the liquid. Advance the PE tubing carefully through the vocal cords for 0.5 cm and discharge the contents into the airway slowly. However, occasional trauma to the lower airway occurs using this technique without better delivery of the sample.</li>
</ol>

<ol>
	<li>Rockman, H. A., 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=Segregation+of+atrial-specific+and+inducible+expression+of+an+atrial+natriuretic+factor+transgene+in+an+in+vivo+murine+model+of+cardiac+hypertrophy.">Segregation of atrial-specific and inducible expression of an atrial natriuretic factor transgene in an in vivo murine model of cardiac hypertrophy.</a> Proc. natl. Acad. Sci. U.S.A. 88, 8277-8281 (1991).</li><li>Watanabe, A., Hashimoto, Y., Ochiai, E., Sato, A., Kamei, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+method+for+confirming+correct+endotracheal+intubation+in+mice.">A simple method for confirming correct endotracheal intubation in mice.</a> Lab. Anim. 43, 399-401 (2009).</li><li>Orebaugh, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Succinylcholine:+adverse+effects+and+alternatives+in+emergency+medicine.">Succinylcholine: adverse effects and alternatives in emergency medicine.</a> Am. J. Emerg. Med. 17, 715-721 (1999).</li><li>Hastings, R. H., Summers-Torres, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+Laryngoscopy+in+Mice.">Direct Laryngoscopy in Mice.</a> Contemp. Lab. Anim. Sci. 38, 33-35 (1999).</li><li>Berul, C. I., Aronovitz, M. J., Wang, P. J., Mendelsohn, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+cardiac+electrophysiology+studies+in+the+mouse.">In vivo cardiac electrophysiology studies in the mouse.</a> Circulation. 94, 2641-2648 (1996).</li><li>Ewart, S. L., Gavett, S. H., Margolick, J., Wills-Karp, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclosporin+A+attenuates+genetic+airway+hyperresponsiveness+in+mice+but+not+through+inhibition+of+CD4++or+CD8++T+cells.">Cyclosporin A attenuates genetic airway hyperresponsiveness in mice but not through inhibition of CD4+ or CD8+ T cells.</a> Am. J. Respir. Cell Mol. Biol. 14, 627-634 (1996).</li><li>Ewart, S. L., Mitzner, W., DiSilvestre, D. A., Meyers, D. A., Levitt, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Airway+hyperresponsiveness+to+acetylcholine:+segregation+analysis+and+evidence+for+linkage+to+murine+chromosome+6.">Airway hyperresponsiveness to acetylcholine: segregation analysis and evidence for linkage to murine chromosome 6.</a> Am. J. Respir. Cell Mol. Biol. 14, 487-495 (1996).</li><li>Spoelstra, E. N., 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+novel+and+simple+method+for+endotracheal+intubation+of+mice.">A novel and simple method for endotracheal intubation of mice.</a> Lab. 41, 128-135 (2007).</li><li>Boll, H., 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=High-speed+single-breath-hold+micro-computed+tomography+of+thoracic+and+abdominal+structures+in+mice+using+a+simplified+method+for+intubation.">High-speed single-breath-hold micro-computed tomography of thoracic and abdominal structures in mice using a simplified method for intubation.</a> J. Comput. Assist. Tomogr. 34, 783-790 (2010).</li><li>Brown, R. H., Walters, D. M., Greenberg, R. S., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+of+endotracheal+intubation+and+pulmonary+functional+assessment+for+repeated+studies+in+mice.">A method of endotracheal intubation and pulmonary functional assessment for repeated studies in mice.</a> J. Appl. Physiol. 87, 2362-2365 (1999).</li><li>Vergari, A., Polito, A., Musumeci, M., Palazzesi, S., Marano, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Video-assisted+orotracheal+intubation+in+mice.">Video-assisted orotracheal intubation in mice.</a> Lab. Anim. 37, 204-206 (2003).</li><li>Vergari, A., 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+new+method+of+orotracheal+intubation+in+mice.">A new method of orotracheal intubation in mice.</a> Eur. Rev. Med. Pharmacol. Sci. 8, 103-106 (2004).</li><li>Hamacher, 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=Microscopic+wire+guide-based+orotracheal+mouse+intubation:+description,+evaluation+and+comparison+with+transillumination.">Microscopic wire guide-based orotracheal mouse intubation: description, evaluation and comparison with transillumination.</a> Lab. Anim. 42, 222-230 (2008).</li><li>Rivera, B., Miller, S., Brown, E., Price, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+for+endotracheal+intubation+of+mice+and+rats+used+in+imaging+studies.">A novel method for endotracheal intubation of mice and rats used in imaging studies.</a> Cont. Top. Lab. Anim. Sci. 44, 52-55 (2005).</li><li>MacDonald, K. D., Chang, H. Y., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+simple+method+of+mouse+lung+intubation.">An improved simple method of mouse lung intubation.</a> J. Appl. Physiol. 106, 984-987 (2009).</li><li>De Vleeschauwer, S. p. I., 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=Repeated+invasive+lung+function+measurements+in+intubated+mice:+an+approach+for+longitudinal+lung+research.">Repeated invasive lung function measurements in intubated mice: an approach for longitudinal lung research.</a> Lab. Anim. 45, 81-89 (2011).</li><li>Zhao, X., 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+technique+for+retrograde+intubation+in+mice.">A technique for retrograde intubation in mice.</a> Lab. Anim. 35, 39-42 (2006).</li><li>Singer, T., 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=Left-sided+mouse+intubation:+description+and+evaluation.">Left-sided mouse intubation: description and evaluation.</a> Exp. Lung Res. 36, 25-30 (2010).</li></ol>

The authors declare that they have no competing financial interests.

In this report we describe a simple, reliable technique to intubate mice that is nontraumatic and can be used repeatedly in the same animal. This technique can be accomplished with simple laboratory or medical equipment that can be purchased for a modest sum. The technique of direct laryngoscopy, originally reported by Hastings and colleagues4, can also be used for a variety of purposes, but mainly to accurately deliver test substances to the lower respiratory tract. We have found this technique superior to those reported by other investigators to date because of its ease, speed, simplicity, reliability, low expense, and potential for repeated measures.
Techniques to intubate mice are not new. One of the earliest reports, used transillumination of trachea followed by oral intubation with a 24 G&#160;catheter5. However, this study and others6,7 did not provide sufficient information to replicate the technique in our laboratory. Recently other investigators have used this transillumination technique with greater success8,9. The first detailed study by Brown and colleagues reported a study in which mice, suspended on a 45&#176; backboard, had their tracheas transilluminated and their trachea intubated orally with PE90 tubing under direct visualization using a fabricated laryngoscope blade10. This technique has proven difficult to replicate and, therefore, has not been utilized extensively. Additional reports describe the use of rigid optical scopes or fiberoptics to enhance the reliability of the techniques. Vergari and colleagues11,12 used a rigid arthroscope 1.7 mm in outer diameter to visualize the vocal cords and then placed a guide wire through the glottis. The authors report 100% success with this technique that they performed with specialized and expensive equipment. The cost of this surgical equipment seems prohibitive for widespread use in mice. Additional techniques have been developed using an operating microscope and a 2f guide wire13. This technique is similar to ours, but requires the use of an operating microscope, and thus we favor the use of an otoscope that is substantially less expensive.
Fiberoptic intubation techniques have also been reported using a relatively inexpensive battery powered fiberoptic source14. We have attempted to use this fiberoptic technique, but have found the light less than useful because there is no corresponding fiberoptic visualization channel. MacDonald and colleagues used a more powerful halogen light source and reported success in a small sample. They were able to repeatedly measure airway resistance15 also. Repeated measures of airway resistance at different time points has been reported by others16. Zhao, et al. used a retrograde technique by puncturing the trachea and feeding a guide wire through the hypo- and oropharynx17. More recently Singer, et al. have performed selective intubation of the left mainstem bronchus in the mouse using an operating microscope and fluoroscopy to document the location of the catheter18. They then used this method to instill test substances selectively into the lung. However, this again would not be a method that could be used without significant expense and highly technical training. Moreover, fluoroscopy would also add a radiation hazard to the study.
In summary, direct laryngoscopy of mice using a pediatric otoscope is a valuable technique for intubation, repeated intubation and test substance delivery without intubation. We find this technique superior to tracheostomy and previously reported methods of noninvasively performing endotracheal intubation in mice.

The laboratory mouse has supplanted virtually all species as the principal mammalian model of biology and pathobiology. The laboratory mouse is the smallest mammalian species that has been clearly and extensively shown to be of value as a model of human disease and has proven invaluable in advances of our understanding of human biology and disease. The short gestation time and substantially lower cost has allowed the development and study of null and transgenic mice as a commonplace tool in biomedical research. However, the size of the average laboratory mouse (20-25 g) has limited their study in physiologically or surgically based studies and, consequently, some investigators study larger mammalian species. An impediment to using mice in these studies is the difficulty encountered with intubation techniques that would allow physiological measurements or extensive surgical procedures under deep anesthesia. Tracheostomy 1 has been used as a standard technique instead of intubation because of the greater ease of performing this technique and modest skill required. However, tracheostomy is not conducive to chronic or recovery surgery studies; thus, it is limited to acute experiments. Tracheostomy can also be a confounding variable in research in which inflammation or sensitive physiologic reflexes are important.
Our laboratory has tried most of the techniques described by other investigators and found them inadequate for a variety of reasons. Tracheostomy is too traumatic and induces bleeding and airway inflammation. Much more problematic is that it cannot be feasibly repeated. Many relatively noninvasive techniques that require a modest investment in equipment are not sufficiently reliable. Other techniques require expensive equipment that is difficult to justify without knowing if the equipment will work in a specific application. Thus, we sought to develop a nontraumatic technique that required no more than a modest investment in specialized equipment, could be accomplished quickly and reliably, could be repeated in chronic models, and could be used in large numbers of animals. Here we report such a technique.

<table border="0" cellpadding="0" cellspacing="0" width="706">
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	<tbody>
		<tr height="17">
			<td height="17" style="height:17px;width:411px;">Operating Otoscope Head</td>
			<td style="width:144px;">Welch Allyn</td>
			<td align="right" style="width:83px;">21700</td>
			<td style="width:69px;">&#160;$188.98</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Otoscope Handle</td>
			<td>Welch Allyn</td>
			<td align="right">71000</td>
			<td>&#160;$112.20</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Reuseable Speculum</td>
			<td>Welch Allyn</td>
			<td align="right">22002</td>
			<td>&#160;$ 3.98</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Fine Forceps</td>
			<td>Miltex</td>
			<td>18-779</td>
			<td>&#160;$107.18</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Small clamp stand to hold otoscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;"></td>
			<td></td>
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			<td height="17" style="height:17px;">Optional Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Isoflurane Vaporizer (multiple circuit with vacuum waste gas handling)</td>
			<td>Summit Medical</td>
			<td></td>
			<td align="right">$3,000</td>
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		<tr height="17">
			<td height="17" style="height:17px;">Flexivent (Animal Ventilator)</td>
			<td>SCIREQ</td>
			<td></td>
			<td align="right">$35,000</td>
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		<tr height="17">
			<td height="17" style="height:17px;"></td>
			<td></td>
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			<td height="17" style="height:17px;">Supplies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Intravenous catheter (20 ga x 1 inch)</td>
			<td>BD</td>
			<td align="right">381233</td>
			<td>&#160;$&#160; 9.62</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Polyethylene tubing (PE10) (0.011 inch I.D., 0.024 inch O.D.) 100 ft</td>
			<td>Intramedic, Clay-Adams</td>
			<td align="right">427401</td>
			<td>&#160;$115.70</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Ketamine 100 mg/ml (10 ml bottle)</td>
			<td>Butler</td>
			<td align="right">23061</td>
			<td>&#160;$10.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Xylazine (100 ml bottle)</td>
			<td>Vedco</td>
			<td align="right">24105</td>
			<td>&#160;$20.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Isoflurane (250 ml bottle)</td>
			<td></td>
			<td></td>
			<td>&#160;$15.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">vecuronium bromide 10 mg/10 ml</td>
			<td>Pfizer</td>
			<td>NDC 0069-0094-01</td>
			<td align="right">$15.00</td>
		</tr>
	</tbody>
</table>

endotracheal intubation in mice via direct laryngoscopy using an otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G&#160;intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Intubation with the above technique is reliable and fast. The appropriate placement of the endotracheal tube is most easily verified by observing expired gas bubbling from the submerged expiratory limb of the of the ventilator circuit (usually in a PEEP trap) and negative deflections on an airway pressure tracing (Figure 1). The negative deflections on airway pressure tracing are the most reliable. Others have used movement of a small fluid droplet in intravenous tubing connected to the expiratory ventilator circuit2. Although we have not used this technique extensively, it should be sufficient to verify tube location. An experienced operator can intubate mice using this method about 90% of the time on the first pass. For those that fail, the tube is immediately removed, the animal is reinduced with isoflurane anesthesia, and the placement again attempted. If the animal is left connected to the ventilator circuit in the case of an esophageal intubation, the stomach will gradually insufflate generating high peak airway pressures, hypoxemia, and ultimately death if there is no intervention. If high peak airway pressures are observed without negative inspiratory efforts, the animal must be promptly removed from the ventilator and extubated. However, removing the animal from the ventilator and removing the endotracheal tube easily reverse this. During the last 300 animal studies requiring intubation an experienced operator intubated 80% with the initial attempt and more than 95% by the second attempt. All animals were intubated by the third attempt of experienced operators. The time taken to intubate mice is up to 5 min for the preanesthetic, 90-120 sec to induce the mouse with isoflurane, and 30 sec to identify and actually intubate the airway. We have found that new trainees should learn to visualize the vocal cords first. If they can successfully demonstrate a full view of the vocal cords ten times, they are ready to begin training to intubate. Once trainees have successfully intubated 10 times, as demonstrated by negative deflections on an airway pressure tracing in a spontaneously breathing animal, they are ready to perform studies in experimental animals. However, a training video will greatly accelerate the learning curve. Only inexperienced operators induce airway trauma by this technique and deaths of animals may occur in this setting while learning the technique. However, loss of animals with an experienced operator is rare. This is our experience in more than 1,000 animals over the last 4 years.
A 20 G&#160;catheter is used as an endotracheal tube in mice because it makes a tight seal with the airway, thus allowing accurate measures of airway pressures and it is of sufficient diameter for measures of airway resistance. However, using the PE tubing as a stylet or bougie allows the 20 G&#160;to pass easily and more reliably through the vocal cords. The 1 in&#160;(2.5 cm) length of the catheter allows the tube to be placed with the hub at the lower incisors for stability, but without causing trauma to the lower airway or intubating the mainstem bronchi.
We use this technique to make measurements of airway resistance, perform methacholine dose-response curves (Figure 2), and to measure pressure-volume relationships in the lung (Figure 3). For all of these measurements the animals must be fully paralyzed to obtain useable, accurate data. For this purpose we have used pancuronium bromide as an inexpensive, nondepolarizing paralytic in nonrecovery experiments. However, pancuronium is no longer available and cannot be used on animals that are going to recover because of its long half-life. Therefore, we have switched to vecuronium (30 min) or rocuronium for paralysis because of their shorter half-life, providing the option of repeated measurements over time. Other investigators have used succinylcholine as a depolarizing paralytic, but its half-life has been too short for the majority of our experiments. Also succinylcholine can cause histamine release3 and would therefore be a potential confounding variable.
We have also used the direct laryngoscopy technique extensively for the delivery of test substances to the lung based on a technique originally reported in rats by Hastings4. Most investigations have used nasal instillation for the delivery of test substances to the lower respiratory tract because mice are obligate nasal breathers and nasal instillation is easy. However, we have found the nasal instillation method to be unreliable for quantitative delivery. By using the direct laryngoscopy technique we have found that quantitative delivery is more reliable and distribution in the lung is more uniform. Our principal use of the direct instillation technique has been to deliver antigen to the lower respiratory tract in a model of allergic inflammation (Figure 4) and airway hyperresponsiveness (Figure 2). We have also delivered other test substances such as cytokines and drugs via this method.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50269/50269fig1highres.jpg" src="/files/ftp_upload/50269/50269fig1.jpg" /> 
Figure 1. Airway pressure tracing. The blue line reflects the airway pressure and the red line reflects the tidal volume tracing in the mouse immediately after intubation. The negative (downward) deflections of the pressure curve (black arrows) indicate the generation of negative intrathoracic pressure in the mouse (C57BL/6, 24 g) by spontaneous respiratory effort and indicates proper placement of the endotracheal tube. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50269/50269fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50269/50269fig2highres.jpg" src="/files/ftp_upload/50269/50269fig2.jpg" /> 
Figure 2. Airway resistance and methacholine dose-response curves. Mice (C57BL/6) intubated by this technique can undergo measures of airway resistance and bronchial reactivity using doubling doses of methacholine delivered via ultrasonic nebulizer. In this representative example, a control animal has a minimal response to methacholine up to 25 mg/ml and ovalbumin immunized and challenged mouse demonstrates hyperresponsiveness to methacholine. These measurements are made with mice intubated with a 20 G&#160;catheter and are indistinguishable from those made in a tracheostomized mouse.
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/50269/50269fig3highres.jpg" src="/files/ftp_upload/50269/50269fig3.jpg" /> 
Figure 3. Pressure-volume curve. This is a representative pressure-volume curve in a 20 g mouse that has been intubated with a 20 G&#160;intravenous catheter, paralyzed with pancuronium, and is mechanically ventilated. It demonstrates that the catheter does not leak up to 30 cm H2O pressure and can therefore be used to make accurate measurements pressure-volume relationships in the mouse.
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/50269/50269fig4highres.jpg" src="/files/ftp_upload/50269/50269fig4.jpg" /> 
Figure 4. Ovalbumin model of allergic inflammation. This is a representative hematoxylin and eosin stained section of mouse lung that has been immunized and challenged with ovalbumin. It demonstrates the generalized effect of instillation of ovalbumin administered via the direct laryngoscopy method (magnification 4X).

Watch this Scientific Journal Video about Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope at JoVE.com

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are ...

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Nanyang Technological University

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JoVE Journal

Medicine

31.0K Views.  VA San Diego Healthcare System. We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Video: Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

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 ...

Using Quantitative Real-time PCR to Determine Donor Cell Engraftment in a Competitive Murine Bone Marrow Transplantation Model

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules that regulate HSCs. In these transplant model systems, the function of HSCs is determined by the ability of these cells to engraft and reconstitute lethally irradiated recipient mice. Commonly, the donor cell contribution/engraftment is measured by antibodies to donor- specific cell surface proteins using flow cytometry. However, this method heavily depends on the specificity and the ability of the cell surface marker to differentiate donor-derived cells from recipient-originated cells, which may not be available for all mouse strains. Considering the various backgrounds of genetically modified mouse strains in the market, this cell surface/ flow cytometry-based method has significant limitations especially in mouse strains that lack well-defined surface markers to separate donor cells from congenic recipient cells. Here, we reported a PCR-based technique to determine donor cell engraftment/contribution in transplant recipient mice. We transplanted male donor bone marrow HSCs to lethally irradiated congenic female mice. Peripheral blood samples were collected at different time points post transplantation. Bone marrow samples were obtained at the end of the experiments. Genomic DNA was isolated and the Y chromosome specific gene, Zfy1, was amplified using quantitative Real time PCR. The engraftment of male donor-derived cells in the female recipient mice was calculated against standard curve with known percentage of male vs. female DNAs. Bcl2 was used as a reference gene to normalize the total DNA amount. Our data suggested that this approach reliably determines donor cell engraftment and provides a useful, yet simple method in measuring hematopoietic cell reconstitution in murine bone marrow transplantation models. Our method can be routinely performed in most laboratories because no costly equipment such as flow cytometry is required.

Determining donor cell engraftment presents a challenge in mouse bone marrow transplant models that lack well-defined phenotypical markers. We described a methodology to quantify male donor cell engraftment in female transplant recipient mice. This method can be used in all mouse strains for the study of HSC functions.

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules ...

A Simple Method of Mouse Lung Intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

Heterotopic Mucosal Engrafting Procedure for Direct Drug Delivery to the Brain in Mice

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high molecular weight compounds from the bloodstream and into brain tissue. Some direct delivery success in humans has been achieved via implantation of transcranial catheters; however this method is highly invasive and associated with numerous complications. A less invasive alternative would be to dose the brain through a surgically implanted, semipermeable membrane such as the nasal mucosa that is used to repair skull base defects following endoscopic transnasal tumor removal surgery in humans. Drug transfer though this membrane would effectively bypass the BBB and diffuse directly into the brain and cerebrospinal fluid. Inspired by this approach, a surgical approach in mice was developed that uses a donor septal mucosal membrane engrafted over an extracranial surgical BBB defect. This model has been shown to effectively allow the passage of high molecular weight compounds into the brain. Since numerous drug candidates are incapable of crossing the BBB, this model is valuable for performing preclinical testing of novel therapies for neurological and psychiatric diseases.

A mouse model of human endoscopic skull base reconstruction has been developed that creates a semipermeable interface between the brain and nose using nasal mucosal grafts. This method allows researchers to study delivery to the central nervous system of high molecular weight therapeutics which are otherwise excluded by the blood-brain barrier when administered systemically.

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

Sleeve Gastrectomy in Mice using Surgical Clips

The number of people who are overweight and obese is continually increasing both in the adult and adolescent populations. This coincides with the increased universal phenomenon of type 2 diabetes (T2D) and other metabolic problems. Bariatric surgery, such as SG, is currently one of the most effective and commonly used long-term treatment for obesity and T2D, but the association between them is not completely explored yet. The mechanisms underlying the outcomes seen after bariatric surgery in humans can be investigated based on preclinical animal studies. The SG reduces body weight, glucose levels and many metabolic parameters, and is easy to perform with a low incidence of complications. The goal of this work is to provide a simple method and an uncomplicated preclinical model of bariatric surgery in animals for researchers.

The prevalence of diabetes and obesity is continuously increasing worldwide. The mechanisms between diabetes, obesity and their associated mortality and co-morbidities need to be further investigated. Here, we present a protocol for sleeve gastrectomy (SG) in animals as an uncomplicated preclinical model of bariatric surgery.

The number of people who are overweight and obese is continually increasing both in the adult and adolescent populations. This coincides with the ...

Repeated Orotracheal Intubation in Mice

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ventral neck for direct confirmation of cannula placement in the trachea. The relative difficulty or the tissue trauma induced to the subject by such procedures can be an impediment to an investigator&#8217;s ability to perform longitudinal studies. This article illustrates a technique in which physical manipulation of the mouse following the use of a depilatory to remove hair from the ventral neck permits transcutaneous visualization of the trachea for orotracheal intubation regardless of degree of skin pigmentation. This method is innocuous to the subject and easily achieved with a limited understanding of murine anatomy. This refined approach facilitates repeated intubation, which may be necessary for monitoring progression of disease or instillation of treatments. Using this method may result in a reduction of the number of animals and technical skill required to measure lung function in mouse models of respiratory disease.

The goal of this article is to describe a refined method of intubation of the laboratory mouse. The method is noninvasive and, therefore, ideal for studies that require serial monitoring of respiratory function and/or instillation of treatments into the lung.

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ...

Blind Endotracheal Intubation in Neonatal Rabbits

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical characteristics, which is further complicated in the case of the newborn. Of the different methods of advanced airway management, endotracheal intubation is less aggressive than tracheostomy, and is more feasible than supraglottic management given the lack of supraglottic devices of such a small size. As direct glottis visualization is very difficult in animals this size, this blind intubation model is presented as an effective alternative, especially for experiments requiring prolonged anesthesia. Using this method, we performed blind intubations with a 90% success rate.

We describe a technique of endotracheal intubation in newborn rabbits after esophageal catheterization with a gastric tube.

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical ...

Direct Drug Delivery to Kidney via the Renal Artery

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research field. Accumulation of drugs in certain organs may give rise to adverse and unwanted effects, depending on the nature of injectate. To minimize spillover and/or accumulation in other tissues, the herein described method directs the formulation into the renal artery bloodstream by inserting a catheter in the infra renal aorta, just below where it branches into the renal artery, resulting in the kidney as first reached organ and distributing of formulation throughout the kidney.
This manuscript provides a detailed description of the method, as well as its challenges and difficulties. It guides the experimenter to become skillful with this type of microsurgery that requires accuracy under sterile conditions. Speed is crucial for minimizing the ischemia and practicing the procedure will increase the chance of successful injections without adverse effects. By modulating the time between injection and reperfusion as well as the injected volume, the risk of spillover to other organs is mitigated. 
Note that this technique is suitable for single dosing strategies.

This manuscript describes a method for targeted delivery to a single kidney via a catheter placed in the infrarenal abdominal aorta in the mouse.

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research ...