This work was funded by a grant from the Center for Innovation in Prevention and Treatment of Airway Diseases, NINDS grant (NS052574), and from the Sandler Program for Asthma Research.

1. Equipment
The major components of a contraction measurement device are shown schematically in Figure 2A).
<ol>
 <li>A tissue bath. The tissue bath maintains an oxygenated physiological solution at warm temperature. For mice trachea rings, we use a 10 ml tissue bath that contains a water jacket for circulating a warming solution, a fritted glass inlet to bubble oxygen (95%/5% O2/CO2 mixture) and inlet and outlet ports for changing solutions. A reservoir PSS solution is stored with constant bubbling of 95%/5% O2/CO2 mixture in a 37 &deg;C water bath (not shown). For solution exchanges, PSS solution is pumped from the reservoir to the tissue bath inlet (bottom port) at approximately 100 mls per minute to allow relatively fast solution exchange. The solution outlet is through an overflow port (top port) that allows constant volume (~10 ml) in the tissue bath during solution exchange. We use a Haake heating circulator to pump warm water through the tissue bath jacket (to maintain 37 &deg;C). Tissue baths can be obtained from a number of suppliers and come in a variety of sizes and styles to fit the experimental needs of the investigator. </li>
 <li>A force transducer. To measure isometric tension, the trachea tube is threaded over L-shaped ends of two stainless steel rods (Figure 2A). Care should be taken to use a stainless steel type that is compatible with biological material. The top rod is connected via a clip to an isometric force transducer. The bottom rod holds the trachea at a fixed position, and is mounted on a micrometer for adjustment of passive tension and/or muscle length. Contraction of the trachea creates tension on the force transducer, which is converted to a voltage signal at the preamplifier. The bottom rod may also be configured to include two rectangular platinum plates (4 mm apart) that flank the trachea (Figure 2B). The platinum plates are wired to a Grass S88 stimulator that allows delivery of an electric field across the trachea. Open wires and solder are coated with Sylgard (Sylgard 184 Silicone Elastomer, Dow Corning Corp, Midland, MI), to prevent leaching of metals into the bath solution.</li>
 <li>A/D converter, computer and acquisition software. Signals from the preamplifiers are recorded on a MacLab 8 A/D system. This is an older version of the current ADInstrument Powerlab hardware. We use the program Chart (ADInstruments) that allows continuous recording of tension throughout the experiment. Tension generation of trachea muscle is quite slow, and therefore we find that acquisition of 100 points per second is adequate. The tension measurement is calibrated using known weights (up to 5 grams) before each experiment. Similar systems are available from other venders (e.g., BioPac, GW Instruments).</li>
</ol>
2. Trachea Isolation
<ol>
 <li>Before tissue isolation the force transducer is calibrated with known weights, and the tissue bath is filled with normal PSS (see Table I). The air inlet is adjusted to obtain a light stream of O2/CO2.</li>
 <li>Tracheas from two month or older mice are optimal. Younger animals may be used but tracheas obtained from these require greater skill to mount on the force transducer wires because of their small size. Prior to dissection, mice are deeply sedated with isoflurane. The proper level of sedation is reached when a toe-pinch with forceps is unable to elicit a response. Mice are immediately sacrificed by cervical dislocation. An important note: We have observed that Avertin (Tribromoethanol), a sedative commonly used in mice, has strong relaxant effects on airway smooth muscle and therefore should not be used for trachea contraction studies.</li>
 <li>The skin (and fur) is removed from thorax to throat. Ribs are cut from the base of the sternum, laterally (on both sides) to the top of the heart. The sternum and ribs are then pulled forward to the throat to reveal the heart/lungs, thymus, trachea (ventral) and esophagus (attached to - and dorsal to trachea).</li>
 <li>The trachea is excised by cutting below the bronchial bifurcation and above the pharynx. The trachea is placed into an ice-cold oxygenated (95/5) PSS solution (composition given in Table I).</li>
 <li>The trachea is dissected clean of surrounding tissues. During cleaning, the trachea can be held at the pharynx or below the bifurcation. However, caution should be taken not to directly apply the forceps to the trachea itself. Fine scissors may be used to cut off surrounding tissues, but the cut should always be made parallel to the trachea to avoid damage. This part of the procedure is facilitated if the trachea preparation is pinned below the bifurcation and above the pharynx onto a Sylgard-coated dish (Sylgard 184 Silicone Elastomer, Dow Corning Corp, Midland, MI).</li>
 <li>After removing surrounding tissue, the trachea is cut below the pharynx and above the bronchial bifurcation and gently mounted on the force transducer wires.</li>
 <li>The trachea is threaded over two L-shaped metal prongs (Figure 2A). One prong is connected to a force-displacement transducer for continuous recording of isometric tension. Another prong is connected to a micrometer. The tissue bath is then raised so that the trachea is immersed in PSS. Mounting of the trachea should be done as quickly as possible to minimize the time that trachea are held outside of PSS. With practice, mounting of the trachea can be done within one minute but we generally avoid times longer than 3 minutes to avoid loss of viability.</li>
 <li>The micrometer is adjusted slowly to obtain a passive tension of ~10 mN (~ 1 gram-force). The optimal resting tension was determined empirically and we have found that passive tension of ~ 5 - 10 mN results in an equivalent, maximal response to high potassium stimulation. This is consistent with a number of other studies that utilize a passive tension in this range16,17,18. Over the first 5-10 minutes, trachea passive tension tends to decline somewhat (stress-relaxation phenomenon) and the micrometer is used to adjust the passive tension to ~10 mN during equilibration. The trachea is allowed to equilibrate for at least 1 hour before experimental challenges.</li>
</ol>
3. High Potassium Stimulation
Following equilibration, the trachea is challenged twice with a high potassium PSS solution (67 mM KCl, Table I). The contraction generally requires ~ 5-10 minutes to reach steady-state at which time the tissue bath is rinsed several times with normal PSS to completely relax the trachea. The potassium contraction is repeated a second time, and a third time (if necessary) until reproducible contractions are obtained.
4. Contraction Evoked by Electrical Stimulation
The trachea is flanked by two rectangular platinum plates (electrodes) that allow electric field stimulation (EFS) to the preparation. The contractile response to EFS is a function of frequency and voltage. It is also affected by physical parameters such as the area of the electrodes and the distance between them. The power characteristics of the stimulator also affect responses such that at higher voltages and current outputs the stimulator can reach its maximum. The characteristics of any EFS system should be determined by examining the muscle contractile responses at differing stimulus durations, frequencies, voltages and pulse durations. For our experimental setup, we have found that electrodes separated by ~ 4 mm, and stimulation amplitude of 44 V (0.5 ms pulses) and 30 Hz are optimal to achieve reproducible near-maximal contractile responses.
5. Contraction Evoked by Cholinergic Stimulation
The responsiveness of the trachea to exogenously applied compounds is evaluated either by addition of a single dose of the drug of interest or, by multiple additions of the drug in a cumulative dose fashion. For the trachea, our laboratory routinely used carbachol to activate cholinergic receptors because, unlike acetylcholine, carbachol is not degraded by acetylcholinesterase. A reasonable dose-response range is from 10-8 to 10-5 M carbachol. Fitting of the Log[carbachol]-contractile response curve with a Hill-type equation allows an estimation of EC50 (half-maximal effective concentration) that is a measure of sensitivity of tracheal contraction to the cholinergic agonist 19. It is worth noting that a given dose of carbachol will give a slightly greater response as a single dose than as part of a cumulative dose response curve.
6. Representative Results
An example of a contractile response to high potassium is shown in Figure 3A. The contraction reaches a maximum within approximately 10 minutes but may show a small decline thereafter. During early washout of high potassium, the muscle may show a transient increase in contraction that is due to a drop in temperature as the small volume of unheated PSS solution in the solution lines transiently perfuses the preparation. This can be minimized by having a minimum dead volume in the tubing connecting the heated PSS reservoir and tissue bath, and also by exchanging solution relatively quickly (generally we pump solutions at 100 ml/min). Each preparation will have some differences in contractile response due to differences in muscle mass or damage incurred during the dissection. Figure 3B shows two tracheas of different muscle mass challenged with high potassium and carbachol. Although the cholinergic-evoked contractions differ, they are similar after normalization to the response with high potassium solution (Figure 3C).
Figure 4 shows an example of a carbachol (cholinergic) evoked contraction using single doses (A) and a cumulative increase (B). The carbachol solutions are added directly to the bath and the bubbled gas helps in rapid mixing. It is worth noting that addition of a single dose (i.e. 1 &mu;M, Figure 4A) has a slightly larger response than the equivalent concentration during a cumulative dose-response curve (1 &mu;M, Figure 4B). Figure 4C shows a plot of contraction force as a function of carbachol concentrations using data from Figure 4B. Effects of carbachol saturate at 10-5 M concentration. Although cholinergic agonists initiates contraction through calcium release mechanisms, a substantial component of the contraction is also mediated by depolarization and activation of voltage-dependent calcium channels 20.
Figure 5A shows an example of EFS-evoked contractions. Tracheas are stimulated using 0.5 millisecond duration, 40 volt pulses until contractions reach a plateau (see inset a1). An increase in stimulation frequency causes an increased contractile response (frequency-response curve is plotted in Figure 5B). Electric field stimulation has been shown to evoke contractions predominantly by activating presynaptic nerves. This is evidenced by the effect of botulinum toxin, a blocker of neurotransmitter release that blocks the majority of the EFS-evoked contractions of trachea 21. Moreover, tetrodotoxin, an agent that blocks Na+ channels also inhibits nerve activity and eliminates the response of trachea to EFS.
<img alt="Figure 1" src="/files/ftp_upload/3703/3703fig1.jpg" /> 
Figure 1. Diagram of the major signaling pathways in an isolated trachea preparation. Shown is a cholinergic axon terminal innervating a tracheal smooth muscle cell. The major signaling pathways are M3- and M2- muscarinic acetylcholine receptor activation (mACHR) that cause calcium release through IP3 receptors (M3) and reduction of cAMP (M2). M2 receptors (and some contribution of M3 receptors) also cause cholinergic-evoked depolarization that activates L-type voltage-dependent calcium channels and calcium influx. Common contractile agents and their effectors are 1. high potassium (depolarize smooth muscle cell and cholinergic axon), 2. electric field stimulation (EFS, depolarizes cholinergic axon) and 3. exogenous application of cholinergic agents such as acetylcholine or carbachol (activates muscarinic receptors directly).
<img alt="Figure 2" src="/files/ftp_upload/3703/3703fig2.jpg" /> 
Figure 2. Diagram of equipment used to measure tracheal contractions. A. The force transducer, micrometer and tissue bath are mounted on supporting rods via screw clamps. The tracheal ring is threaded over the top and bottom rods. In the diagram, the tissue bath is positioned below the preparation (i.e. during mounting of the trachea onto the force transducer). During contraction studies, the tissue bath is moved vertically to bathe the preparation. B. For electric field stimulation the bottom rod is modified to include two platinum plates that are mounted laterally to the trachea holding wire. The platinum plates are connected by electrical wires to a stimulator.
<img alt="Figure 3" src="/files/ftp_upload/3703/3703fig3.jpg" /> 
Figure 3. Examples of high potassium (67 mM) contractile response of trachea. (A) shows duplicate, reproducible responses to high potassium. (B) Examples of contractile response to carbachol of two different tracheas. (C) Responses to the tracheas in B are similar when normalized to the high potassium response.
<img alt="Figure 4" src="/files/ftp_upload/3703/3703fig4.jpg" /> 
Figure 4. Examples of carbachol-induced contractions. (A) Carbachol-induced contractions using single doses followed by washout. (B) Example cumulative dose-response curve for trachea in A. (C) Peak contractions from B are plotted as a function of carbachol concentration.
<img alt="Figure 5" src="/files/ftp_upload/3703/3703fig5.jpg" /> 
Figure 5. Examples of contractions evoked by electric field stimulation. (A) Electric field stimulation of trachea contraction using 0.5 ms pulse, 40 volts, and various stimulation frequencies as indicated. Inset is time-expanded contraction at 30 Hz. (B) Peak contractions from A are plotted as a function of stimulation frequency.
Normal PSS
<table border="1">
 <tbody>
 <tr>
 <td>Salt</td>
 <td>Conc.(mM)</td>
 <td>Amount(g/2 L)</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>119</td>
 <td>13.91</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>4.7</td>
 <td>0.7</td>
 </tr>
 <tr>
 <td>KH2PO4</td>
 <td>1.18</td>
 <td>0.32</td>
 </tr>
 <tr>
 <td>MgSO4 x 7H2O</td>
 <td>1.17</td>
 <td>0.58</td>
 </tr>
 <tr>
 <td>NaHCO3</td>
 <td>18</td>
 <td>3.02</td>
 </tr>
 <tr>
 <td>EDTA</td>
 <td>0.026</td>
 <td>0.1 ml of 0.5 M</td>
 </tr>
 <tr>
 <td>Glucose</td>
 <td>11</td>
 <td>3.96</td>
 </tr>
 <tr>
 <td>Sucrose</td>
 <td>12.5</td>
 <td>8.56</td>
 </tr>
 <tr>
 <td>CaCl2</td>
 <td>2</td>
 <td>400 ml of 10 mM</td>
 </tr>
 </tbody>
</table>
High K+ PSS (NaCl and KCl adjustments)
<table border="1">
 <tbody>
 <tr>
 <td>Salt </td>
 <td>Conc. (mM) </td>
 <td>Amount (g/2 L)</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>56.7</td>
 <td>6.628</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>67</td>
 <td>9.991</td>
 </tr>
 </tbody>
</table>
Table 1. Recipe for PSS solutions. Note: Solutions are made fresh weekly, with ultrapure quality water, and are stored in a refrigerator for no longer than 5 days to avoid contaminating growth.

<ol>
	<li>Lloyd, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Building+better+mouse+models+of+asthma.">Building better mouse models of asthma.</a> Curr. Allergy Asthma Rep. 7, 231-236 (2007).</li><li>Hausding, M., Sauer, K., Maxeiner, J. H., Finotto, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenic+models+in+allergic+responses.">Transgenic models in allergic responses.</a> Curr. Drug Targets. 9, 503-510 (2008).</li><li>Eglen, R. M., Hegde, S. S., Watson, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscarinic+receptor+subtypes+and+smooth+muscle+function.">Muscarinic receptor subtypes and smooth muscle function.</a> Pharmacol Rev. 48, 531-565 (1996).</li><li>Ehlert, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contractile+role+of+M2+and+M3+muscarinic+receptors+in+gastrointestinal,+airway+and+urinary+bladder+smooth+muscle.">Contractile role of M2 and M3 muscarinic receptors in gastrointestinal, airway and urinary bladder smooth muscle.</a> Life Sci. 74, 355-366 (2003).</li><li>Hall, I. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Second+messengers,+ion+channels+and+pharmacology+of+airway+smooth+muscle.">Second messengers, ion channels and pharmacology of airway smooth muscle.</a> Eur. Respir. J. 15, 1120-1127 (2000).</li><li>Berridge, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inositol+trisphosphate+and+calcium+signalling.">Inositol trisphosphate and calcium signalling.</a> Nature. 361, 315-325 (1993).</li><li>Ehlert, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacological+analysis+of+the+contractile+role+of+M2+and+M3+muscarinic+receptors+in+smooth+muscle.">Pharmacological analysis of the contractile role of M2 and M3 muscarinic receptors in smooth muscle.</a> Receptors Channels. 9, 261-277 (2003).</li><li>Sankary, R. M., Jones, C. A., Madison, J. M., Brown, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscarinic+cholinergic+inhibition+of+cyclic+AMP+accumulation+in+airway+smooth+muscle.+Role+of+a+pertussis+toxin-sensitive+protein.">Muscarinic cholinergic inhibition of cyclic AMP accumulation in airway smooth muscle. Role of a pertussis toxin-sensitive protein.</a> Am. Rev. Respir Dis. 138, 145-150 (1988).</li><li>Widdop, S., Daykin, K., Hall, I. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+muscarinic+M2+receptors+in+cultured+human+airway+smooth+muscle+cells.">Expression of muscarinic M2 receptors in cultured human airway smooth muscle cells.</a> Am. J. Respir. Cell Mol. Biol. 9, 541-546 (1993).</li><li>Karaki, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+movements,+distribution,+and+functions+in+smooth+muscle.">Calcium movements, distribution, and functions in smooth muscle.</a> Pharmacol. Rev. 49, 157-230 (1997).</li><li>Somlyo, A. V., Somlyo, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electromechanical+and+pharmacomechanical+coupling+in+vascular+smooth+muscle.">Electromechanical and pharmacomechanical coupling in vascular smooth muscle.</a> J. Pharmacol Exp. Ther. 159, 129-145 (1968).</li><li>Fryer, A. D., Jacoby, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscarinic+receptors+and+control+of+airway+smooth+muscle.">Muscarinic receptors and control of airway smooth muscle.</a> Am. J. Respir. Crit. Care Med. 158, 154-160 (1998).</li><li>Fernandez-Rodriguez, S., Broadley, K. J., Ford, W. R., Kidd, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+muscarinic+receptor+activity+of+airway+smooth+muscle+isolated+from+a+mouse+model+of+allergic+asthma.">Increased muscarinic receptor activity of airway smooth muscle isolated from a mouse model of allergic asthma.</a> Pulm. Pharmacol. Ther. 23, 300-307 (2010).</li><li>Garssen, J., Loveren, H. V. a. n., Van Der Vliet, H., Nijkamp, F. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+isometric+method+to+study+respiratory+smooth+muscle+responses+in+mice.">An isometric method to study respiratory smooth muscle responses in mice.</a> J. Pharmacol. Methods. 24, 209-217 (1990).</li><li>Vass, G., Horvath, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adenosine+and+adenosine+receptors+in+the+pathomechanism+and+treatment+of+respiratory+diseases.">Adenosine and adenosine receptors in the pathomechanism and treatment of respiratory diseases.</a> Curr. Med. Chem. 15, 917-922 (2008).</li><li>Borchers, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methacholine-induced+airway+hyperresponsiveness+is+dependent+on+Galphaq+signaling.">Methacholine-induced airway hyperresponsiveness is dependent on Galphaq signaling.</a> Am. J. Physiol. Lung Cell Mol. Physiol. 285, 114-120 (2003).</li><li>Sausbier, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduced+rather+than+enhanced+cholinergic+airway+constriction+in+mice+with+ablation+of+the+large+conductance+Ca2+-activated+K++channel.">Reduced rather than enhanced cholinergic airway constriction in mice with ablation of the large conductance Ca2+-activated K+ channel.</a> Faseb. J. 21, 812-822 (2007).</li><li>Scheerens, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+topical+exposure+to+toluene+diisocyanate+in+mice+leads+to+antibody+production+and+in+vivo+airway+hyperresponsiveness+three+hours+after+intranasal+challenge.">Long-term topical exposure to toluene diisocyanate in mice leads to antibody production and in vivo airway hyperresponsiveness three hours after intranasal challenge.</a> Am. J. Respir. Crit. Care. Med. 159, 1074-1080 (1999).</li><li>Kenakin, T. 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> A pharmacology primer : theory, applications, and methods. , (2009).</li><li>Semenov, I., Wang, B., Herlihy, J. T., Brenner, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BK+Channel+{beta}1+Subunits+Regulate+Airway+Contraction+Secondary+to+M2+Muscarinic+Acetylcholine+Receptor+Mediated+Depolarization.">BK Channel {beta}1 Subunits Regulate Airway Contraction Secondary to M2 Muscarinic Acetylcholine Receptor Mediated Depolarization.</a> J. Physiol. , 1803-1817 (2011).</li><li>Moffatt, J. D., Cocks, T. M., Page, C. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+the+epithelium+and+acetylcholine+in+mediating+the+contraction+to+5-hydroxytryptamine+in+the+mouse+isolated+trachea.">Role of the epithelium and acetylcholine in mediating the contraction to 5-hydroxytryptamine in the mouse isolated trachea.</a> Br. J. Pharmacol. 141, 1159-1166 (2004).</li><li>Bachar, O., Adner, M., Uddman, R., Cardell, L. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nerve+growth+factor+enhances+cholinergic+innervation+and+contractile+response+to+electric+field+stimulation+in+a+murine+in+vitro+model+of+chronic+asthma.">Nerve growth factor enhances cholinergic innervation and contractile response to electric field stimulation in a murine in vitro model of chronic asthma.</a> Clin. Exp. Allergy. 34, 1137-1145 (2004).</li><li>Weigand, L. A., Myers, A. C., Meeker, S., Undem, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mast+cell-cholinergic+nerve+interaction+in+mouse+airways.">Mast cell-cholinergic nerve interaction in mouse airways.</a> J. Physiol. 587, 3355-3362 (2009).</li><li>Kao, J., Fortner, C. N., Liu, L. H., Shull, G. E., Paul, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ablation+of+the+SERCA3+gene+alters+epithelium-dependent+relaxation+in+mouse+tracheal+smooth+muscle.">Ablation of the SERCA3 gene alters epithelium-dependent relaxation in mouse tracheal smooth muscle.</a> Am. J. Physiol. 277, 264-270 (1999).</li><li>Krane, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aquaporin+5-deficient+mouse+lungs+are+hyperresponsive+to+cholinergic+stimulation.">Aquaporin 5-deficient mouse lungs are hyperresponsive to cholinergic stimulation.</a> Proc. Natl. Acad. Sci. U.S.A. 98, 14114-14119 (2001).</li><li>Semenov, I., Wang, B., Herlihy, J. T., Brenner, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BK+channel+beta1-subunit+regulation+of+calcium+handling+and+constriction+in+tracheal+smooth+muscle.">BK channel beta1-subunit regulation of calcium handling and constriction in tracheal smooth muscle.</a> Am. J. Physiol. Lung. Cell Mol. Physiol. 291, L802-L810 (2006).</li><li>Fortner, C. N., Breyer, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EP2+receptors+mediate+airway+relaxation+to+substance+P+ATP,+and+PGE2.">EP2 receptors mediate airway relaxation to substance P ATP, and PGE2.</a> Am. J. Physiol. Lung Cell Mol. Physiol. 281, 469-474 (2001).</li><li>Hay, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+modulation+of+endothelin+ligand-induced+contraction+in+isolated+tracheae+from+endothelin+B+(ET(B))+receptor+knockout+mice.">Differential modulation of endothelin ligand-induced contraction in isolated tracheae from endothelin B (ET(B)) receptor knockout mice.</a> Br. J. Pharmacol. 132, 1905-1915 (2001).</li><li>Stengel, P. W., Yamada, M., Wess, J., Cohen, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=M(3)-receptor+knockout+mice:+muscarinic+receptor+function+in+atria,+stomach+fundus,+urinary+bladder,+and+trachea.">M(3)-receptor knockout mice: muscarinic receptor function in atria, stomach fundus, urinary bladder, and trachea.</a> Am. J. Physiol. Regul. Integr. Comp Physiol. 282, R1443-R1449 (2002).</li><li>Trevisani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+in+vitro+expression+of+B1+receptor+in+the+mouse+trachea+and+urinary+bladder.">Evidence for in vitro expression of B1 receptor in the mouse trachea and urinary bladder.</a> Br. J. Pharmacol. 126, 1293-1300 (1038).</li><li>Mehats, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PDE4D+plays+a+critical+role+in+the+control+of+airway+smooth+muscle+contraction.">PDE4D plays a critical role in the control of airway smooth muscle contraction.</a> FASEB J. 17, 1831-1841 (2003).</li><li>Kumar, R. K., Herbert, C., Foster, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+"classical"+ovalbumin+challenge+model+of+asthma+in+mice.">The "classical" ovalbumin challenge model of asthma in mice.</a> Curr. Drug Targets. 9, 485-494 (2008).</li></ol>

No conflicts of interest declared.

The protocol presented here provides a physiological preparation to assess airway muscle function. We generally operate 3-4 organ bath preparations simultaneously, however, prepackaged systems are available from a number of suppliers that allow simultaneous measurements of up to 8 preparations (ADInstruments, World Precision Instruments, and Harvard Apparatus). We have utilized a number of force transducers and tissue organ baths with equivalent results. However, we find that electric field stimulation provides som...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Analogue-Digital Converter</td>
 <td>ADInstruments</td>
 <td>PowerLab 4/35</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Carbachol
 (Carbamoylcholine Chloride)</td>
 <td>Sigma-Aldrich</td>
 <td>C4832
 </td>
 <td>10-2 M in water (aliquots can be stored at -20&deg;C)</td>
 </tr>
 <tr>
 <td>Charting Software </td>
 <td>ADInstrtuments</td>
 <td>LabChart</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Heating Circulator</td>
 <td>Haake</td>
 <td>Mixer Mill MM400</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Isometric Force Transducer</td>
 <td>Kent Scientific</td>
 <td>TRN001</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Stimulator</td>
 <td>Grass Technologies</td>
 <td>S88 Dual Output Square Pulse Stimulator</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tissue Bath</td>
 <td>WPI</td>
 <td>47264</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

in vitro measurements of tracheal constriction using mice

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of isolated tracheal preparations has proven to be a useful assay of airway smooth muscle (ASM) contractile response in genetically modified mice. These in vitro tracheal preparations are relatively simple, provide a robust response, and retain both functional cholinergic nerve endings and muscle responses, even after long incubations.
Tracheal tensometry also provides a functional assay to study a variety of second messenger signaling pathways that affect contraction of smooth muscle. Contraction in trachea is primarily mediated by parasympathetic, cholinergic nerves that release acetylcholine onto ASM (Figure 1). The major ASM acetylcholine receptors are muscarinic M2 and M3 which are Gi/o and Gq coupled receptors, respectively3,4,5. M3 receptors evoke contraction by coupling to Gq to activate phospholipase C, increase IP3 production and IP3-mediated calcium release from the sarcoplasmic reticulum3,6,7. M2/Gi/o signaling is believed to enhance contractions by inhibition of adenylate cyclase leading to a decrease in cAMP levels5,8,9,10. These pathways constitute the so called "pharmaco-contraction coupling" of airway smooth muscle11. In addition, cholinergic signaling through M2 receptors (and modulated by M3 signaling) involves pathways that depolarize the ASM which in turn activate L-type, voltage-dependent calcium channels (Figure 1) and calcium influx (so called "excitation-contraction coupling")4,7. More detailed reviews on signaling pathways controlling airway constriction can be found4,12. The above pathways appear to be conserved between mice and other species. However, mouse tracheas differ from other species in some signaling pathways. Most prominent is their lack of contractile response to histamine and adenosine13,14, both well-known ASM modulators in humans and other species5,15.
Here we present protocols for the isolation of murine tracheal rings and the in vitro measurement of their contractile output. Included are descriptions of the equipment configuration, trachea ring isolation and contractile measurements. Examples are given for evoking contractions indirectly using high potassium stimulation of nerves and directly by depolarization of ASM muscle to activate voltage-dependent calcium influx (1. high K+, Figure 1). In addition, methods are presented for stimulations of nerves alone using electric field stimulation (2. EFS, Figure 1), or for direct stimulation of ASM muscle using exogenous neurotransmitter applied to the bath (3. exogenous ACH, Figure 1). This flexibility and ease of preparation renders the isolated trachea ring model a robust and functional assay for a number of signaling cascades involved in airway smooth muscle contraction.

Watch this Scientific Journal Video about In vitro Measurements of Tracheal Constriction Using Mice at JoVE.com

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic mice have been extremely useful in ascribing physiological function to genes. As such, research in general, and functional studies of airway, in particular, have undergone a remarkable shift toward murine models. Here we provide protocols for in vitro trachea constriction studies to evaluate smooth muscle function in murine airway.

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of ...

in-vitro-measurements-of-tracheal-constriction-using-mice

Research

JoVE Journal

Medicine

20.7K 조회수.  UT Health Science Center, San Antonio. 형질 전환 생쥐는 유전자로 생리 기능을 ascribing에서 매우 유용한되었습니다. 일반적으로 같은, 연구하고,기도의 기능적 연구로서 특히, murine 모델 향한 놀라운 변화를받은. 여기에 대한 프로토콜을 제공 체외에서기도의 수축 연구 murine기도의 평활근 기능을 평가합니다.

비디오: In vitro Measurements of Tracheal Constriction Using Mice

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.

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.

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

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  ...

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly hydrated, forming a substantial endothelial surface layer (ESL) that contributes to the maintenance of endothelial function. As the endothelial glycocalyx is often aberrant in vitro and is lost during standard tissue fixation techniques, study of the ESL requires use of intravital microscopy. To best approximate the complex physiology of the alveolar microvasculature, pulmonary intravital imaging is ideally performed on a freely-moving lung. These preparations, however, typically suffer from extensive motion artifact. We demonstrate how closed-chest intravital microscopy of a freely-moving mouse lung can be used to measure glycocalyx integrity via ESL exclusion of fluorescently-labeled high molecular weight dextrans from the endothelial surface. This non-recovery surgical technique, which requires simultaneous brightfield and fluorescent imaging of the mouse lung, allows for longitudinal observation of the subpleural microvasculature without evidence of inducing confounding lung injury.

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx/endothelial surface layer is ideally studied using intravital microscopy. Intravital microscopy is technically challenging in a moving organ such as the lung. We demonstrate how simultaneous brightfield and fluorescent microscopy may be used to estimate endothelial surface layer thickness in a freely-moving in vivo mouse lung.

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly ...

A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated procedures to obtain such measurements in the same animal are generally not feasible.  Here, we demonstrate protocols for obtaining from mice repeated measurements of AHR and bronchoalveolar lavage fluid samples.  Mice were challenged intranasally seven times over 14 days with a potent allergen or sham treated.  Prior to the initial challenge, and within 24 hours following each intranasal challenge, the same animals were anesthetized, orally intubated and mechanically ventilated.  AHR, assessed by comparing dose response curves of respiratory system resistance (RRS) induced by increasing intravenous doses of acetylcholine (Ach) chloride between sham and allergen-challenged animals, were determined.  Afterwards, and via the same intubation, the left lung was lavaged so that differential enumeration of airway cells could be performed.  These studies reveal that repeated measurements of AHR and BAL fluid collection are possible from the same animals and that maximal airway hyperresponsiveness and airway eosinophilia are achieved within 7-10 days of initiating allergen challenge.  This novel technique significantly reduces the number of mice required for longitudinal experimentation and is applicable to diverse rodent species, disease models and airway physiology instruments.  

Repeated measurements of rodent respiratory physiology and sampling of airway inflammatory cells are desirable, but generally not feasible. Here we describe a repeatable method for orally intubating mice that permits repeated measurements of airway hyperreactivity and sampling of airway inflammatory cells.

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated ...

Biology

Transverse Aortic Constriction in Mice

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart failure.1 TAC initially leads to compensated hypertrophy of the heart, which often is associated with a temporary enhancement of cardiac contractility. Over time, however, the response to the chronic hemodynamic overload becomes maladaptive, resulting in cardiac dilatation and heart failure.2 The murine TAC model was first validated by Rockman et al.1, and has since been extensively used as a valuable tool to mimic human cardiovascular diseases and elucidate fundamental signaling processes involved in the cardiac hypertrophic response and heart failure development. When compared to other experimental models of heart failure, such as complete occlusion of the left anterior descending (LAD) coronary artery, TAC provides a more reproducible model of cardiac hypertrophy and a more gradual time course in the development of heart failure. Here, we describe a step-by-step procedure to perform surgical TAC in mice. To determine the level of pressure overload produced by the aortic ligation, a high frequency Doppler probe is used to measure the ratio between blood flow velocities in the right and left carotid arteries.3, 4 With surgical survival rates of 80-90%, transverse aortic banding is an effective technique of inducing left ventricular hypertrophy and heart failure in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model to study mechanisms underlying cardiac hypertrophy and heart failure development. Here, we describe procedures to constrict the aorta to create a reproducible degree of cardiac hypertrophy in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart ...

Automated Measurement of Pulmonary Emphysema and Small Airway Remodeling in Cigarette Smoke-exposed Mice

COPD is projected to be the third most common cause of mortality world-wide by 2020(1). Animal models of COPD are used to identify molecules that contribute to the disease process and to test the efficacy of novel therapies for COPD. Researchers use a number of models of COPD employing different species including rodents, guinea-pigs, rabbits, and dogs(2). However, the most widely-used model is that in which mice are exposed to cigarette smoke. Mice are an especially useful species in which to model COPD because their genome can readily be manipulated to generate animals that are either deficient in, or over-express individual proteins. Studies of gene-targeted mice that have been exposed to cigarette smoke have provided valuable information about the contributions of individual molecules to different lung pathologies in COPD(3-5). Most studies have focused on pathways involved in emphysema development which contributes to the airflow obstruction that is characteristic of COPD. However, small airway fibrosis also contributes significantly to airflow obstruction in human COPD patients(6), but much less is known about the pathogenesis of this lesion in smoke-exposed animals. To address this knowledge gap, this protocol quantifies both emphysema development and small airway fibrosis in smoke-exposed mice. This protocol exposes mice to CS using a whole-body exposure technique, then measures respiratory mechanics in the mice, inflates the lungs of mice to a standard pressure, and fixes the lungs in formalin. The researcher then stains the lung sections with either Gill&#8217;s stain to measure the mean alveolar chord length (as a readout of emphysema severity) or Masson&#8217;s trichrome stain to measure deposition of extracellular matrix (ECM) proteins around small airways (as a readout of small airway fibrosis). Studies of the effects of molecular pathways on both of these lung pathologies will lead to a better understanding of the pathogenesis of COPD.

The goal of this protocol is to provide automated methods to quantify chronic lung pathologies in a murine model of COPD. The protocol includes exposing mice to cigarette smoke (CS), measuring pulmonary function, inflating the lungs, and using morphometry methods to measure emphysema and small airway remodeling in mice.

COPD is projected to be the third most common cause of mortality world-wide by 2020(1).  Animal models of COPD are used to identify molecules that ...

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 ...

Minimally Invasive Transverse Aortic Constriction in Mice

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard open-chest transverse aortic constriction (TAC). Although transverse aortic constriction is a highly functional method for the induction of high pressure in the left ventricle, it is a more difficult and lengthy procedure due to its use of artificial ventilation with tracheal intubation. TAC is oftentimes also less survivable, as the newer method, MTAC, neither requires the cutting of the ribs and intercostal muscles nor tracheal intubation with a ventilation setup. In MTAC, as opposed to a thoracotomy to access to the chest cavity, the aortic arch is reached through a midline incision in the anterior neck. The thyroid is pulled back to reveal the sternal notch. The sternum is subsequently cut down to the second rib level, and the aortic arch is reached simply by separating the connective tissues and thymus. From there, a suture can be wrapped around the arch and tied with a spacer, and then the sternal cut and skin can be closed. MTAC is a much faster and less invasive way to induce left ventricular hypertension and enables the possibility for high-throughput studies. The success of the constriction can be verified using high-frequency trans-thoracic echocardiography, particularly color Doppler and pulsed-wave Doppler, to determine the flow velocities of the aortic arch and left and right carotid arteries, the dimension of the blood vessels, and the left ventricular function and morphology. A successful constriction will also trigger significant histopathological changes, such as cardiac muscle cell hypertrophy with interstitial and perivascular fibrosis. Here, the procedure of MTAC is described, demonstrating how the resulting flow changes in the carotid arteries can be examined with echocardiography, gross morphology, and histopathological changes in the heart.

Minimally invasive transverse aortic constriction (MTAC) conserves the essentials of regular transverse aortic constriction (TAC) while eliminating the use of a ventilator with tracheal intubation. It proves to be a highly desirable method for high-throughput studies on left ventricular overload, particularly in translational studies.

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard ...

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to perform a partial thoracotomy to visualize the transverse aortic arch. However, the surgical trauma caused by the thoracotomy in open-chest models changes the respiratory physiology as the ribs are dissected and left unattached after chest closure. To prevent this, we established a minimally invasive, closed chest approach via lateral thoracotomy. Herein we approach the aortic arch via the 2nd intercostal space without entering the chest cavities, leaving the mouse with a less traumatic injury to recover from. We perform this operation using standard laboratory settings for open chest TAC procedures with equal survival rates. Apart from maintaining physiological breathing patterns due to the closed chest approach, the mice seem to benefit by showing rapid recovery, as the less invasive technique appears to facilitate a fast healing process and to reduce immune response after trauma.

Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to ...

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 ...