The authors thank Lucia Rosas, Lauren Doolittle, Lisa Joseph and Lindsey Ferguson for their technical assistance and the University Laboratory Animal Resources for their animal care support. This work is funded by NIH T35OD010977 and R01-HL102469.

All animal activities described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of The Ohio State University and were conducted in AAALAC-accredited facilities.
1. Procedure Preparation
<ol>
	<li>Construct the intubation platform. To achieve the appropriate platform slope, use a three-inch (7.6 cm) 3-ring binder. Fold a 15&#8722;20 cm length of 3-0 silk or other thread material in half and adhere the ends of the thread to the top of the inclined platform with tape to create a suspension loop (Figure 1).</li>
	<li>Select a cannula of the appropriate size and length. 
	NOTE: For a 20&#8722;30 g mouse, a 1&#8722;1.5 inch (2.5&#8722;3.8 cm) long catheter up to 18 G can be used. For this study, 18 week-old female BALB/c and 10-week-old C57BL/6 mice (n = 3 of each strain) were used. An opaque white catheter sheath provides the best transcutaneous visualization.</li>
	<li>Cut a bevel at the distal tip of the catheter and smooth the cut surface with abrasive paper to create a rounded bevel tip. Gently create a slight bend in the cannula approximately 1 cm from the bevel (Figure 2). 
	NOTE: A new catheter should be used for each mouse.</li>
	<li>Anesthetize the mouse with ketamine (5.4 mg/g body weight) and xylazine (16 &#181;g/g body weight) administered intraperitoneally.&#160; Apply sterile ophthalimic ointment to the eyes. 
	NOTE: Proper anesthetic depth is achieved by lack of response of the mouse to a firm toe pinch.</li>
	<li>Suspend the mouse in a supine position on the intubation platform by hooking the upper incisors around the silk thread at the top of the angled surface (Figure 3). Once the mouse is squarely positioned in dorsal recumbency, gently grasp the base of the tail and retract the tail towards the table. Place a piece of tape over the base of tail to secure the mouse.</li>
	<li>Apply depilatory cream (Table of Materials) to the ventral cervical region for 30&#8722;45 s then remove all depilatory cream from the cervical region using a dry gauze. Repeat application process if needed. Thoroughly rinse the skin with saline or distilled water to remove any residue then wipe dry.</li>
</ol>
2. Intubation Procedure
<ol>
	<li>Use straight, flat forceps in the nondominant hand to gently retract the tongue in a manner that sufficiently opens the mouth for introduction of the cannula. 
	NOTE: Rat tooth forceps should not be used as this will damage the tongue.</li>
	<li>With the dominant hand, advance the cannula into the mouth such that the end that is distal to the slight bend is against the roof of the subject&#8217;s mouth.</li>
	<li>Release the tongue and slide the flat edge of the closed forceps caudally along the ventral neck until the manubrium is reached. This motion laterally displaces the salivary glands and flattens the muscle covering the trachea. The trachea appears transcutaneously as a white line (Figure 3A). If necessary, rotate the forceps in a craniodorsal direction while maintaining tension on the skin in a caudal direction to cause the laterally displaced salivary glands to peak. This maneuver creates more contrast around the trachea (Figure 3B). 
	NOTE: Avoid excessive force on the ventral neck as it can collapse the trachea and impair breathing.</li>
	<li>Advance the cannula while simultaneously angling the distal tip of the cannula ventrally by supination of the dominant hand with simultaneous flexion of the wrist.</li>
	<li>The proper placement of the cannula is indicated by visualization of the opaque cannula in the trachea (Figure 4B,D). If the cannula has been advanced past the level of the origin of the masseter muscle and visualization of the cannula in the trachea has not been confirmed, retract the cannula and reattempt the maneuver.</li>
	<li>Confirm proper cannula placement by connecting a lung inflation bulb to the cannula and observing thoracic expansion with concurrent depression of the device.</li>
	<li>Without displacing the cannula, carefully unhook the incisors of the mouse from the intubation platform. Move the mouse to a horizontal platform (Table of Materials) and insert the cannula to the adaptor on the ventilator. Following the deep inflation, ventilate the mouse for 60 s then measure respiratory resistance.</li>
</ol>
3. Recovery
<ol>
	<li>Once the procedure is complete, move the mouse to a warmed platform. Provide constant stimulation via light toe or tail pinches to encourage spontaneous respiration.</li>
	<li>Extubation can occur when the mouse just begins to chew. Grasp the cannula at the level of the hub and gently pull the tube cranially and away from the mouse until the cannula is completely removed from the subject&#8217;s mouth. 
	NOTE: It is preferable to provide airway support with the rigid cannula for as long as possible during the recovery process.</li>
	<li>Once extubated, transfer the mouse to a clean recovery cage with heat support. Continuously monitor the mouse until it is fully ambulatory, and recovery is complete.</li>
</ol>

<ol><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+%22Laboratory+Animals%22%5BJournal%5D)">A novel and simple method for endotracheal intubation of mice.</a> Laboratory Animals. 41 (1), 128-135 (2007).</li>
<li>Rivera, B., Miller, S. R., Brown, E. M., Price, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Novel+Method+for+Endotracheal+Intubation+of+Mice+and+Rats+Used+in+Imaging+Studies%5BTitle%5D)+AND+%22Contemporary+Topics+in+Laboratory+Animal+Science%22%5BJournal%5D)">A Novel Method for Endotracheal Intubation of Mice and Rats Used in Imaging Studies.</a> Contemporary Topics in Laboratory Animal Science. 44 (2), 52-55 (2005).</li>
<li>Sparrowe, J., Jimenez, M., Rullas, J., Martinez, A. E., Ferrer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Refined+Intratracheal+Intubation+Technique+in+the+Mouse%2C+Complete+Protocol+Description+for+Lower+Airway+Models%5BTitle%5D)+AND+%22Global+Journal+of+Animal+Scientific+Research%22%5BJournal%5D)">Refined Intratracheal Intubation Technique in the Mouse, Complete Protocol Description for Lower Airway Models.</a> Global Journal of Animal Scientific Research. 3 (2), 363-369 (2015).</li>
<li>Deyo, D. J., Wei, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Novel+Method+of+Intubation+and+Ventilation+in+Mice%5BTitle%5D)+AND+%22Anesthesia+%26+Analgesia%22%5BJournal%5D)">A Novel Method of Intubation and Ventilation in Mice.</a> Anesthesia & Analgesia. 88 (2), 179(1999).</li>
<li>Vergari, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+new+method+of+orotracheal+intubation+in+mice%5BTitle%5D)+AND+%22European+Review+for+Medical+and+Pharmacological+Sciences%22%5BJournal%5D)">A new method of orotracheal intubation in mice.</a> European Review for Medical and Pharmacological Sciences. 8 (3), 103-106 (2004).</li>
<li>Brown, R. H., Walters, D. M., Greenberg, R. S., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+method+of+endotracheal+intubation+and+pulmonary+functional+assessment+for+repeated+studies+in+mice%5BTitle%5D)+AND+%22Journal+of+Applied+Physiology%22%5BJournal%5D)">A method of endotracheal intubation and pulmonary functional assessment for repeated studies in mice.</a> Journal of Applied Physiology. 87 (6), 2362-2365 (1999).</li>
<li>Das, S., MacDonald, K., Chang, H. S., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Simple+Method+of+Mouse+Lung+Intubation%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">A Simple Method of Mouse Lung Intubation.</a> Journal of Visualized Experiments. (73), e50318(2013).</li>
<li>Limjunyawong, N., Mock, J., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Instillation+and+Fixation+Methods+Useful+in+Mouse+Lung+Cancer+Research%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Instillation and Fixation Methods Useful in Mouse Lung Cancer Research.</a> Journal of Visualized Experiments. (102), e52964(2015).</li>
<li>Qin, W., Baran, U., Wang, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphatic+response+to+depilation-induced+inflammation+assessed+with+label-free+optical+lymphangiography%5BTitle%5D)+AND+%22Lasers+in+Surgery+and+Medicine%22%5BJournal%5D)">Lymphatic response to depilation-induced inflammation assessed with label-free optical lymphangiography.</a> Lasers in Surgery and Medicine. 47 (8), 669-676 (2015).</li>
<li>McGovern, T. K., Robichaud, A., Fereydoonzad, L., Schuessler, T. F., Martin, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Evaluation+of+Respiratory+System+Mechanics+in+Mice+using+the+Forced+Oscillation+Technique%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique.</a> Journal of Visualized Experiments. (75), e50107(2013).</li>
</ol>

The authors have nothing to disclose.

Intubation using the transcutaneous tracheal visualization technique offers a refined approach to the standard skin incision method. With special attention to several key steps, intubation can be easily and quickly achieved. The animal must be placed squarely in dorsal recumbency on the intubation platform with the mouse secured in gentle retraction. This will extend the animal into vertical alignment and proper positioning for intubation. In addition, the depilatory cream should not remain in contact with the animal&#82...

The laboratory mouse is a common animal model for human respiratory disease. Thus, there are several published methods for mouse intubation for the purpose of both instillation of treatments and measurement of respiratory mechanics. Most of the described procedures require visualization of the glottis through the oral cavity with specialized equipment such as a laryngoscope or fiber-optic light source1,2,3,4,5,6,7. However, this can be difficult when a relatively large cannula is required, as it can obscure the view of the researcher. Limjunyawong et al.8 have addressed this concern with a method of intubation in which a small cutaneous incision is made along the midline of the ventral neck allowing for visualization of the trachea. Following the procedure, the incision is closed with tissue adhesive.
For studies requiring frequent repeated intubations, successive incising and closure of this site requires debridement of the skin margins and tissue trauma to the ventral neck. The purpose of the transcutaneous tracheal visualization approach to oral intubation is to provide a refined, noninvasive technique specifically suitable for repeated intubation studies as well as single intubation events in mice.

<table><tbody><tr><td>18 G x 1 1/4&quot; intravenous catheter, Safelet</td><td>Fisher Scientific</td><td>#14-841-14</td><td>Cannula for intubation</td></tr><tr><td>70% ethanol, 10 L</td><td>Fisher Scientific</td><td>25467025</td><td>Cleaning cannula</td></tr><tr><td>Abrasive paper (sandpaper)</td><td>Porter-Cable</td><td>74001201</td><td>Cannula preparation</td></tr><tr><td>AnaSed (xylazine sterile solution) injection (100 mg/mL)</td><td>Akorn Animal Health</td><td>NDC# 59399-111-50</td><td>Anesthesia</td></tr><tr><td>Blue labeling tape (0.5 in x 14 yds)</td><td>Fisher Scientific</td><td>15966</td><td>Restraint on intubation platform</td></tr><tr><td>Braided silk suture without needle, nonsterile, (3-0)</td><td>Henry Schein</td><td>Item #1007842</td><td>Intubation platform</td></tr><tr><td>Deltaphase Isothermal Pad</td><td>Braintree Scientific</td><td>39DP</td><td>Mouse thermoregulation and recovery</td></tr><tr><td>Deltaphase Operating Board</td><td>Braintree Scientific</td><td>39OP</td><td>Mouse recovery (prior to extubation)</td></tr><tr><td>Distilled water</td><td>ThermoFisher</td><td>15230253</td><td>Cleaning mouse following depilation</td></tr><tr><td>Eye Scissors, angled, sharp/sharp</td><td>Harvard Apparatus</td><td>72-8437</td><td>Cannula preparation</td></tr><tr><td>FlexiVent (FX Module 2)</td><td>Scireq</td><td>N/A</td><td>Record lung function data (not required to perform procedure, used in this study to validate procedure)</td></tr><tr><td>Gauze sponges</td><td>Fisher scientific</td><td>13-761-52</td><td>Hair removal</td></tr><tr><td>Heavy-Duty 3&quot; 3-Ring View Binders</td><td>Staples</td><td>24690CT</td><td>Intubation platform</td></tr><tr><td>Instat Software</td><td>Graphpad</td><td>N/A</td><td>Statistical analysis software</td></tr><tr><td>Insulin syringe (0.5 cc, U100)</td><td>Fisher Scientific</td><td>329461</td><td>Anesthesia administration</td></tr><tr><td>Ketamine HCl Injection, USP (100 mg/mL)</td><td>Llyod Laoratories</td><td>List No. 4871</td><td>Anesthesia</td></tr><tr><td>Lung inflation bulb</td><td>Harvard Apparatus</td><td>72-9083</td><td>Confirm cannula placement</td></tr><tr><td>Micro Forceps, Curved, Smooth</td><td>Harvard Apparatus</td><td>72-0445</td><td>Retract tongue and create tension on neck for cannula visualization</td></tr><tr><td>Nair (hair removal lotion), 9 oz bottle</td><td>Church &amp;amp; Dwight</td><td>42010440</td><td>Hair removal</td></tr><tr><td>Sterile saline (0.9%), 10 mL</td><td>Fisher Scientific</td><td>NC9054335</td><td>Anesthesia, cleaning skin following hair removal</td></tr></tbody></table>

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.

Serial monitoring of baseline pulmonary function 
Eighteen-week-old female BALB/c and 10-week-old C57BL/6 mice (n = 3 of each strain) were intubated using the described method on day 0, 3, 10, and 17. Following intubation on each day, the subject was connected to a mechanical ventilator supplied with 100% oxygen (Table of Materials). Respiratory resistance (Rrs) was measured using the forced oscillation technique for 60 s following a deep inflation to 25 cm H2O held for 5...

Watch this Scientific Journal Video about Repeated Orotracheal Intubation in Mice at JoVE.com

Repeated Orotracheal Intubation in Mice

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 ...

Research

JoVE Journal

Medicine

15.9K Views.  The Ohio State University. Dit protocol stelt onderzoekers in staat om herhaalde metingen van de longfunctie in individuele muizen uit te voeren, waardoor de experimentele variabiliteit afneemt en het aantal muizen dat nodig is voor elk experiment wordt verminderd. Het belangrijkste voordeel van dit protocol is dat het geen visualisatie van de luchtpijp door de mondholte of traumatische dissectie van de nek voor intubatie vereist. Deze methode zal van bijzondere waarde zijn voor onderzoekers die chronische en progressi...

Video: Repeated Orotracheal Intubation in 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  ...

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

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 ...

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

Experimental Model to Evaluate Resolution of Pneumonia

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to high morbidity and mortality. Currently, there are no pre-clinical models that recapitulate the complexity of human ARDS. However, infectious models of pneumonia (PNA) can replicate the main pathophysiological features of ARDS. Here, we describe a model of PNA induced by the intratracheal instillation of live Streptococcus pneumoniae and Klebsiella pneumoniae in C57BL6 mice. In order to evaluate and characterize the model, after inducing injury, we carried out serial measurements of body weight and bronchoalveolar lavage (BAL) for measuring markers of lung injury. Additionally, we harvested lungs for cell count and differentials, BAL protein quantification, cytospin, bacterial colony-forming unit counts, and histology. Lastly, high dimensional flow cytometry was performed. We propose this model as a tool to understand the immune landscape during the early and late resolution phases of lung injury.

This manuscript describes the establishment of an infectious model of pneumonia in mice and the respective characterization of injury resolution along with methods for growing bacteria and intratracheal instillation. A novel approach using high-dimensional flow cytometry to evaluate the immune landscape is also described.

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to ...

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 ...

Creating a Murine Model of RV Hypertrophy and Failure by Pulmonary Trunk Banding

A Murine Model of Pressure Overload-Induced Right Ventricular Hypertrophy and Failure by Pulmonary Trunk Banding

Right ventricular (RV) failure caused by pressure overload is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary diseases. The pathogenesis of RV failure is complex and remains inadequately understood. To identify new therapeutic strategies for the treatment of RV failure, robust and reproducible animal models are essential. Models of pulmonary trunk banding (PTB) have gained popularity, as RV function can be assessed independently of changes in the pulmonary vasculature.
In this paper, we present a murine model of RV pressure overload induced by PTB in 5-week-old mice. The model can be used to induce different degrees of RV pathology, ranging from mild RV hypertrophy to decompensated RV failure. Detailed protocols for intubation, PTB surgery, and phenotyping by echocardiography are included in the paper. Furthermore, instructions for customizing instruments for intubation and PTB surgery are given, enabling fast and inexpensive reproduction of the PTB model.
Titanium ligating clips were used to constrict the pulmonary trunk, ensuring a highly reproducible and operator-independent degree of pulmonary trunk constriction. The severity of PTB was graded by using different inner ligating clip diameters (mild: 450 &#181;m and severe: 250 &#181;m). This resulted in RV pathology ranging from hypertrophy with preserved RV function to decompensated RV failure with reduced cardiac output and extracardiac manifestations. RV function was assessed by echocardiography at 1 week and 3 weeks after surgery. Examples of echocardiographic images and results are presented here. Furthermore, results from right heart catheterization and histological analyses of cardiac tissue are shown.

Author Spotlight: Investigating the Underlying Mechanisms of Right Ventricular Failure in Pulmonary Hypertension

We describe a murine model of right ventricular pressure overload-induced by pulmonary trunk banding. Detailed protocols for intubation, surgery, and phenotyping by echocardiography are included in the paper. Custom-made instruments are used for intubation and surgery, allowing for fast and inexpensive reproduction of the model.

Right ventricular (RV) failure caused by pressure overload is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary ...

Repeated Orotracheal Intubation in Mice

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