The authors thank Brian Johnson of the Histology and Imaging Core at the University of Washington for help with the trichrome staining and analysis. This work was supported by NIH grants HL098067 and HL089455.

The Institutional Animal Care and Use Committees (IACUC) at the University of Washington and Cedars-Sinai Medical Center have approved the animal work necessary for these studies.
1. Preparation
<ol>
	<li>Sterilize both the blunt end forceps and the depressor via autoclave.</li>
	<li>Using a biological safety cabinet, prepare a working stock of BLM in PBS from the lyophilized powder.&#160; Sonicate the solution for 10 min at 35 Khz to assure even mixing. 
	Note: A total volume of between 30 and 45 &#181;l is recommended to prevent pipetting variation on the low end, and suffocation with larger volumes.</li>
	<li>Prepare a clean workspace that includes approximately 1 m2 for the procedure itself, as well as designated locations for cages both before and after the procedure.</li>
	<li>Fix the base of the procedure board to the bench immediately in front of the researcher by laying 2 or 3 strips of laboratory tape across the base and underlying bench. See Figure 1 for further specifications on creating a board.</li>
	<li>Tie a single length of size 4.0 suture thread between the two positioning screws of the procedure board.</li>
	<li>Generate a makeshift spirometer by removing and discarding the plunger from three 1 ml syringes, and depositing 60 &#181;l of PBS into the top of each barrel to form an airtight seal. Secure the hub of the catheter loosely to one of the syringes and place it to one side of the board.</li>
	<li>Aspirate 300 &#181;l of air into a 1 ml syringe and place it to one side of the board.</li>
	<li>Cut an additional piece of tape approximately 6 inches in length and place to one side. This will be used to secure the animal to the board in step 2.4.</li>
	<li>Set up an isoflurane chamber. Attach O2, isoflurane, and vacuum to the appropriate ports on both the exposure chamber and the clearance vacuum. Alternatively, administer anesthetic in an isoflurane-compatible biological safety cabinet.</li>
</ol>
2. Intubation
<ol>
	<li>Anesthetize the mouse with isoflurane in the chamber until it loses consciousness and respiration slows to an appropriate rate. A typical exposure includes 4% isoflurane and 2% O2 for 3 to 4 min, and the ideal outcome is 2 to 2.5 min of sedation. This corresponds to a respiration rate of 1 breath every 2 sec.</li>
	<li>While waiting for sedation to set in, aspirate between 30 and 45 &#181;l of BLM into a pipettor and place to one side.</li>
	<li>When ready, suspend the sedated mouse by its upper incisors from the thread attached to the positioning screws of the procedure platform. Make sure that the animal&#39;s dorsum lies flat against the platform surface.</li>
	<li>Being careful not to restrict ventilation, place a piece of tape loosely across the lower (caudal) portion of the thoracic cavity, just above the diaphragm. Placement should be tight enough to maintain proper alignment during the procedure, but not so tight that it restricts respiration.</li>
	<li>Turn on the illuminator to between 80% and 100% intensity and orient the gooseneck so that it is 1 to 2 cm from the surface of the skin, near the solar plexus. Periodically check the tip of the gooseneck for heat to ensure that it does not injure the mouse.</li>
	<li>Standing behind the platform, use the sterile, blunt end forceps to locate the tongue. Being careful to avoid the lower incisors, gently grip and draw the tongue out of the oral cavity.</li>
	<li>Using the remaining hand, insert the depressor and use it to flatten the tongue against the floor of the oral cavity. Release the forceps, but leave the depressor in place for the next two steps.</li>
	<li>Orient the light so that the trachea is visible by guiding the gooseneck proximally from the level of the solar plexus until it reaches the level of the mainstem bronchi. 
	Note: The trachea can be easily distinguished by the action of respiration, which causes the emitted light to fluctuate in intensity. When correctly positioned, this structure will be discernable in the axial plane as a centrally located pin of light with minimal ambient light in the oral cavity itself.</li>
	<li>Angle the syringe so that it follows the natural path of the trachea, and lower the 22-G catheter tip, with the attached syringe containing the droplet, straight into the lumen. The PBS bubble will begin to rise and fall with each breath upon successful placement. 
	Note: This action may be delayed by several seconds as a result of deep sedation.</li>
	<li>Feed the catheter in an additional 5 mm. Remove the tongue depressor.</li>
	<li>Shift the syringe to the opposite hand, and gripping the hub, gently remove the syringe.</li>
	<li>Deposit between 30 and 45 &#181;l of BLM into the center of the interior of the catheter hub, attach the second syringe and dispense 300 &#181;l of air into the hub.</li>
	<li>Replace the second syringe with the first containing the bubble of PBS. The bubble will continue to rise and fall if the procedure has been performed successfully.</li>
</ol>
3. Post-procedural Care
<ol>
	<li>Remove the catheter and tape, and place the animal in a dry warm place until it regains consciousness - typically within a couple of minutes.</li>
</ol>

<ol><li>Hyde, D. M., Hamid, Q., Irvin, C. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anatomy%2C+pathology%2C+and+physiology+of+the+tracheobronchial+tree%3A+emphasis+on+the+distal+airways%5BTitle%5D)+AND+%22J+Allergy+Clin+Immunol%22%5BJournal%5D)">Anatomy, pathology, and physiology of the tracheobronchial tree: emphasis on the distal airways.</a> J Allergy Clin Immunol. 124, S72-S77 (2009).</li>
<li>Rosenthal, N., Brown, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+mouse+ascending%3A+perspectives+for+human-disease+models%5BTitle%5D)+AND+%22Nat+Cell+Biol%22%5BJournal%5D)">The mouse ascending: perspectives for human-disease models.</a> Nat Cell Biol. 9, 993-999 (2007).</li>
<li>Peters, L. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+mouse+as+a+model+for+human+biology%3A+a+resource+guide+for+complex+trait+analysis%5BTitle%5D)+AND+%22Nat+Rev+Genet%22%5BJournal%5D)">The mouse as a model for human biology: a resource guide for complex trait analysis.</a> Nat Rev Genet. 8, 58-69 (2007).</li>
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<li>Saguil, A., Fargo, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Acute+respiratory+distress+syndrome%3A+diagnosis+and+management%5BTitle%5D)+AND+%22Am+Fam+Physician%22%5BJournal%5D)">Acute respiratory distress syndrome: diagnosis and management.</a> Am Fam Physician. 85, 352-358 (2012).</li>
<li>Wilson, M. S., Wynn, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pulmonary+fibrosis%3A+pathogenesis%2C+etiology+and+regulation%5BTitle%5D)+AND+%22Mucosal+Immunol%22%5BJournal%5D)">Pulmonary fibrosis: pathogenesis, etiology and regulation.</a> Mucosal Immunol. 2, 103-121 (2009).</li>
<li>Wuyts, W. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+pathogenesis+of+pulmonary+fibrosis%3A+a+moving+target%5BTitle%5D)+AND+%22Eur+Respir+J%22%5BJournal%5D)">The pathogenesis of pulmonary fibrosis: a moving target.</a> Eur Respir J. 41, 1207-1218 (2013).</li>
<li>Mouratis, M. A., Aidinis, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Modeling+pulmonary+fibrosis+with+bleomycin%5BTitle%5D)+AND+%22Curr+Opin+Pulm+Med%22%5BJournal%5D)">Modeling pulmonary fibrosis with bleomycin.</a> Curr Opin Pulm Med. 17, 355-361 (2011).</li>
<li>Moeller, A., Ask, K., Warburton, D., Gauldie, J., Kolb, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+bleomycin+animal+model%3A+a+useful+tool+to+investigate+treatment+options+for+idiopathic+pulmonary+fibrosis%3F%5BTitle%5D)+AND+%22Int+J+Biochem+Cell+Biol%22%5BJournal%5D)">The bleomycin animal model: a useful tool to investigate treatment options for idiopathic pulmonary fibrosis?</a> Int J Biochem Cell Biol. 40, 362-382 (2008).</li>
<li>Myllärniemi, M., Kaarteenaho, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pharmacological+treatment+of+idiopathic+pulmonary+fibrosis+-+preclinical+and+clinical+studies+of+pirfenidone%2C+nintedanib%2C+and+N-acetylcysteine%5BTitle%5D)+AND+%22European+Clinical+Respiratory+Journal%22%5BJournal%5D)">Pharmacological treatment of idiopathic pulmonary fibrosis - preclinical and clinical studies of pirfenidone, nintedanib, and N-acetylcysteine.</a> European Clinical Respiratory Journal. 2, (2015).</li>
<li>Reinert, T., Baldotto, C. S. dR., Nunes, F. A. P., Scheliga, A. A. dS. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bleomycin-Induced+Lung+Injury%5BTitle%5D)+AND+%22Journal+of+Cancer+Research%22%5BJournal%5D)">Bleomycin-Induced Lung Injury.</a> Journal of Cancer Research. 2013, 1-9 (2013).</li>
<li>Lakatos, H. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Oropharyngeal+aspiration+of+a+silica+suspension+produces+a+superior+model+of+silicosis+in+the+mouse+when+compared+to+intratracheal+instillation%5BTitle%5D)+AND+%22Exp+Lung+Res%22%5BJournal%5D)">Oropharyngeal aspiration of a silica suspension produces a superior model of silicosis in the mouse when compared to intratracheal instillation.</a> Exp Lung Res. 32, 181-199 (2006).</li>
<li>Helms, M. N., Torres-Gonzalez, E., Goodson, P., Rojas, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Direct+tracheal+instillation+of+solutes+into+mouse+lung%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Direct tracheal instillation of solutes into mouse lung.</a> J Vis Exp. , (2010).</li>
<li>Osier, M., Oberdorster, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Intratracheal+inhalation+vs+intratracheal+instillation%3A+differences+in+particle+effects%5BTitle%5D)+AND+%22Fundam+Appl+Toxicol%22%5BJournal%5D)">Intratracheal inhalation vs intratracheal instillation: differences in particle effects.</a> Fundam Appl Toxicol. 40, 220-227 (1997).</li>
<li>Driscoll, K. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Intratracheal+instillation+as+an+exposure+technique+for+the+evaluation+of+respiratory+tract+toxicity%3A+Uses+and+Limitations%5BTitle%5D)+AND+%22Toxicol+Sci%22%5BJournal%5D)">Intratracheal instillation as an exposure technique for the evaluation of respiratory tract toxicity: Uses and Limitations.</a> Toxicol Sci. 55, 24-35 (2000).</li>
<li>MacDonald, K. D., Chang, H. Y., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+improved+simple+method+of+mouse+lung+intubation%5BTitle%5D)+AND+%22J+Appl+Physiol+%281985%29%22%5BJournal%5D)">An improved simple method of mouse lung intubation.</a> J Appl Physiol (1985). 106, 984-987 (1985).</li>
<li>Spoelstra, E. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+and+simple+method+for+endotracheal+intubation+of+mice%5BTitle%5D)+AND+%22Lab+Anim%22%5BJournal%5D)">A novel and simple method for endotracheal intubation of mice.</a> Lab Anim. 41, 128-135 (2007).</li>
<li>Cai, Y., Kimura, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Noninvasive+intratracheal+intubation+to+study+the+pathology+and+physiology+of+mouse+lung%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Noninvasive intratracheal intubation to study the pathology and physiology of mouse lung.</a> J Vis Exp. , e50601(2013).</li>
<li>Starcher, B., WIlliams, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+method+for+intratracheal+instillation+of+endotoxin+into+the+lungs+of+mice%5BTitle%5D)+AND+%22Lab+Anim%22%5BJournal%5D)">A method for intratracheal instillation of endotoxin into the lungs of mice.</a> Lab Anim. 23, 234-240 (1989).</li>
<li>Daubeuf, F., Frossard, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Performing+bronchoalveolar+lavage+in+the+mouse%5BTitle%5D)+AND+%22Curr+Protoc+Mouse+Biol%22%5BJournal%5D)">Performing bronchoalveolar lavage in the mouse.</a> Curr Protoc Mouse Biol. 2, 167-175 (2012).</li>
<li>Li, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Severe+lung+fibrosis+requires+an+invasive+fibroblast+phenotype+regulated+by+hyaluronan+and+CD44%5BTitle%5D)+AND+%22J+Exp+Med%22%5BJournal%5D)">Severe lung fibrosis requires an invasive fibroblast phenotype regulated by hyaluronan and CD44.</a> J Exp Med. 208, 1459-1471 (2011).</li>
<li>Grazioli, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CYR61+%28CCN1%29+overexpression+induces+lung+injury+in+mice%5BTitle%5D)+AND+%22Am+J+Physiol+Lung+Cell+Mol+Physiol%22%5BJournal%5D)">CYR61 (CCN1) overexpression induces lung injury in mice.</a> Am J Physiol Lung Cell Mol Physiol. 308, L759-L765 (2015).</li>
<li>Redente, E. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tumor+necrosis+factor-alpha+accelerates+the+resolution+of+established+pulmonary+fibrosis+in+mice+by+targeting+profibrotic+lung+macrophages%5BTitle%5D)+AND+%22Am+J+Respir+Cell+Mol+Biol%22%5BJournal%5D)">Tumor necrosis factor-alpha accelerates the resolution of established pulmonary fibrosis in mice by targeting profibrotic lung macrophages.</a> Am J Respir Cell Mol Biol. 50, 825-837 (2014).</li>
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The authors have nothing to disclose.

In instances where aerosolization is impractical due to limited reagent availability, safety, or cost, direct tracheal administration is a superior method for delivery of exogenous agents into the lungs.16 Transtracheal instillation has been widely used to accomplish this; however, as with all surgical intervention, it also carries with it the potential for complications caused by the procedure itself, and not necessarily the agent being instilled.13 For these reasons, it has become...

In spite of some anatomical and physiological differences,1 murine models continue to be invaluable for modeling human biology and disease pathogenesis.2 From a husbandry standpoint, mice are easy to handle, have a low breeding time, an accelerated lifespan, and are relatively inexpensive to house. With the development of diverse genetic strains and strategies (e.g., conditional knock-outs, reporter mice, lineage-tracing approaches, etc.), as well as the wide range of available reagents (e.g., antibodies, recombinant proteins, inhibitors, etc.), mice have become an essential model vertebrate organism to uncover human homeostasis and disease processes.3
Mice have been especially valuable for studying pulmonary conditions, including acute lung injury (ALI) and pulmonary fibrosis.4 ALI in humans can be caused by trauma, injury, or sepsis and is characterized by epithelial and endothelial leak (i.e., edema), inflammation, and nascent fibrosis. In many patients, ALI progresses to its severe form, acute respiratory distress syndrome (ARDS), which often results in fibrosis and death due to respiratory failure.5,6 Pulmonary fibrosis is a progressive, fatal pathology characterized by the excess deposition of extracellular matrix, most notably type I collagen, leading to impaired lung function.7,8 Administration of bleomycin (BLM) is the most widely used and best characterized model for inducing ALI and fibrosis in experimental animals.9 Although BLM-induced pulmonary fibrosis in rodents does not recapitulate fully the human fibrotic phenotypes,10 mouse studies with this model have led to the discovery of many important factors influencing the onset and progression of disease.11
While the exact mechanism(s) behind BLM-induced fibrogenesis are unknown, the initiating injury is thought to arise from contact-dependent DNA strand breaks in the epithelial cells lining the conducting airways and alveoli, and in particular, type 1 pneumocytes.12 The need for direct contact between BLM and the pulmonary epithelium highlights the importance of a robust delivery route, and these concerns are also germane to a broad range of treatments targeted to the distal airways, including recombinant proteins, antibodies, siRNA, virus, bacteria, particulates, and more. Oropharyngeal aspiration (OPA) has been widely used for this purpose13, but a major a shortcoming of OPA is that some portion of the delivered agent may be swallowed into the gastrointestinal tract, thereby leading to imprecision in the administered dose. Another widely used approach is transtracheal instillation, which involves tracheostomy under strong anesthesia to expose the trachea and instillation of an agent directly into the respiratory tract.14 However, not only may such a procedure be undesirable due to its invasivity, but it is also time consuming, requires a fair bit of training, and causes a potent injury to the respiratory tract.15,16 Several protocols have been developed that involve the direct administration of agents into the trachea without the need for surgical intervention,16,17,18,19,20 but these methods involve extended recovery times caused by powerful anesthetics, the use of expensive equipment (i.e., otoscope/laryngoscope, commercially available procedure boards, fiber-optic wires, etc.), an excess of manipulation in the oral cavity, and uncertainty regarding the dosage.
This paper describes a relatively easy method of administration via intubation that allows a researcher to quickly, inexpensively, and reliably instill a reagent into the murine lung with limited risk of residual damage to the surrounding tissues.

<table><tbody><tr><td>Bleomycin For Injection, 30 units/vial</td><td>APP Pharmaceuticals, LLC</td><td>103720</td><td>For best results, BLM should be suspended in PBS, aliquoted, and stored as single use lyophilzed aliquots</td></tr><tr><td>Blunt End Forceps</td><td></td><td></td><td></td></tr><tr><td>Tongue Depressor (i.e. bent Valleylab Blade Electrode, 2.4&quot;)&amp;nbsp;</td><td>Covidien</td><td>E1551G</td><td>Before use, create a 45 degree bend 1.5 cm&amp;nbsp; from the blade tip. A suitable depressor can also be created from any metal implement of similar dimensions.&amp;nbsp;</td></tr><tr><td>Exel Safelet Catheter 22G X 1&quot;</td><td>Exel International</td><td>26746</td><td></td></tr><tr><td>1 ml Slip-tip Disposable Tuberculin Syringe (200/sp, 1600/ca)</td><td>BD</td><td>309659</td><td></td></tr><tr><td>0.2 ml Pipettor and Filter Tips</td><td></td><td></td><td></td></tr><tr><td>Fiber-Lite Illuminator. Model 181-1: Model 180 mated with Standard Dual Gooseneck illuminator</td><td>Dolan Jenner Industries, Inc.</td><td>181-1</td><td>Lower output LED illuminators are not recommended as they fail to suficiently illuminate the trachea.</td></tr><tr><td>Intubation Board</td><td></td><td></td><td>See Diagram 1.</td></tr><tr><td>Colored Label Tape: 0.5&amp;nbsp;in. Wide</td><td>Fisherbrand</td><td>15-901-15A</td><td></td></tr><tr><td>Oxygen&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Phosphate-Buffered Saline, 1X</td><td>Corning</td><td>21-040-CV</td><td>Product should be sterile</td></tr><tr><td>Non-Sterile Silk Black Braided Suture Spool, 91.4 m, Size 4-0</td><td>Harvard Apparatus</td><td>517698</td><td></td></tr><tr><td>Table Top Anesthesia Machine Isoflurane</td><td>Highland Medical Equipment</td><td></td><td>http://www.highlandmedical.net/</td></tr><tr><td>Slide Top Induction Mouse Isoflurane Chamber</td><td>MIP / Anesthesia Technologies</td><td>AS-01-0530-SM</td><td></td></tr><tr><td>FORANE (isoflurane, USP) Liquid For Inhalation 100 mL&amp;nbsp;</td><td>Baxter</td><td>1001936040</td><td></td></tr><tr><td>&amp;nbsp;Nanozoomer Digital Pathology system</td><td>Hamamatsu</td><td></td><td></td></tr><tr><td>IgM ELISA Quantification Kit&amp;nbsp;</td><td>Bethyl Laboratories</td><td>E90-101</td><td></td></tr></tbody></table>

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.

Intubated mice were monitored daily for weight loss and distress, and sacrificed 4, 10 or 17 days later via intraperitoneal injection of 2.5% 2,2,2-tribromoethanol. Bronchoalveolar lavage (BAL) was collected in three washes of PBS as described elsewhere 21, and the right lung was fixed in 10% formalin, paraffin embedded, and stained with Masson&#39;s Trichrome by the University of Washington Histology and Imaging Core 22.
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Watch this Scientific Journal Video about An Improved Method for Rapid Intubation of the Trachea in Mice at JoVE.com

An Improved Method for Rapid Intubation of the Trachea in Mice

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 ...

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25.2K Views.  University of Washington. 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.

Video: An Improved Method for Rapid Intubation of the Trachea in Mice

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged mechanical ventilation, contributes to morbidity and mortality of critical illness, and studies on novel genetic or pharmacological targets are areas of intense investigation (1-3, 5, 8, 26, 30, 33-36). ALI is defined by the acute onset of the disease, which leads to non-cardiac pulmonary edema and subsequent impairment of pulmonary gas exchange (36). We have developed a murine model of ALI by using a pressure-controlled ventilation to induce ventilator-induced lung injury (2). For this purpose, C57BL/6 mice are anesthetized and a tracheotomy is performed followed by induction of ALI via mechanical ventilation. Mice are ventilated in a pressure-controlled setting with an inspiratory peak pressure of 45 mbar over 1 - 3 hours. As outcome parameters, pulmonary edema (wet-to-dry ratio), bronchoalveolar fluid albumin content, bronchoalveolar fluid and pulmonary tissue myeloperoxidase content and pulmonary gas exchange are assessed (2). Using this technique we could show that it sufficiently induces acute lung inflammation and can distinguish between different treatment groups or genotypes (1-3, 5). Therefore this technique may be helpful for researchers who pursue molecular mechanisms involved in ALI using a genetic approach in mice with gene-targeted deletion.

A murine model for ventilator induced lung injury is an important tool to study an acute lung injury in vivo. Here, we report an easy applicable in situ model for acute lung injury using high-pressure mechanical ventilation to induce acute failure of the lung.

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged ...

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  ...

Noninvasive Intratracheal Intubation to Study the Pathology and Physiology of Mouse Lung

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. With the increasing availability of knockout and transgenic derivatives, together with a vast amount of genetic information, mice provide one of the best models to study the molecular mechanisms underlying the pathology and physiology of lung diseases. Inhalation, intranasal instillation, intratracheal instillation, and intratracheal intubation are the most widely used techniques by a number of investigators to administer materials of interest to mouse lungs. There are pros and cons for each technique depending on the goals of a study. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented. This technique was applied to administer bleomycin to mouse lungs as a model to study pulmonary fibrosis.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular ...

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

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 ...

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR ...

Immunology and Infection

In vivo Calcium Imaging in Mouse Inferior Olive

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

In vivo Calcium Imaging in Mouse Inferior Olive

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

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

Neuroscience

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 ...

Tracheotomy-Based Endotracheal Intubation Protocol for Non-Survival Rat Models

Endotracheal Intubation via Tracheotomy and Subsequent Thoracotomy in Rats for Non-Survival Applications

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that require controlled or high ventilation pressures or access to the thoracic cavity and organs. Conventional endoorotracheal intubation using the anatomically existing route through the mouth is well suited for survival experiments. However, this procedure poses some challenges, including generally higher levels of the required experience and technical skill, more advanced equipment, and greater time effort with relevant intubation failure rates and complications such as tracheal perforation, temporary systemic hypooxygenation, and relevant aerial leakage.
This manuscript, therefore, presents a detailed step-by-step protocol for endotracheal intubation through tracheotomy in non-survival rat models when guaranteed intubation success, constant oxygenation levels, high ventilation pressures, or open thoracotomy are required.
The protocol emphasizes the importance of meticulous surgical technique to ensure consistent and reliable outcomes, especially for researchers who are inexperienced or lack routine in the technique of endoorotracheal intubation via direct laryngoscopy. This procedure is, therefore, expected to minimize animal suffering and unnecessary animal losses.

Here we introduce a standardized procedure for endotracheal intubation via tracheotomy followed by thoracotomy in rats, aimed at enhancing the precision and reproducibility of non-survival applications requiring invasive ventilation and exposure of thoracic organs in in vivo rat models.

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Summary

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