This work was supported by the Guangzhou Science and Technology Planning Project (202002030151), the Major Project of Guangzhou National Laboratory (GZNL2024A02001), and the grant of State Key Laboratory of Respiratory Disease (SKLRD-Z-202202).

All the procedures were approved by the Animal Care and Use Committee of Guangzhou Medical University (20240248) and were performed in strict accordance with approved guidelines. Male specific pathogen-free C57BL/6 mice weighing 20-25 g were used in this study. All mice were housed under controlled temperature (22 &#177; 2 &#176;C), humidity (50% &#177; 20%), and lighting (6:30 AM to 6:30 PM) in solid bottom cages with food and water available ad libitum. The time line of the protocol is shown in Figure 1.
1. Establishment of a mouse model of cough
<ol>
	<li>Use influenza A/California/7/2009 (H1N1) virus. Determine the lethal dose of the H1N1 virus in mice.
	<ol>
		<li>Briefly, anesthetize mice (n = 10 per group) with 2.5% isoflurane and then intranasally infect with a 10-fold serially diluted virus.</li>
		<li>Monitor the death over a period of 15 days. The survival rates and body weights of mice were recorded daily. Calculate the median lethal dose (LD50) by the Reed-Muench method26. 
		NOTE: Mice were painlessly sacrificed with an overdose of pentobarbital sodium when they reached or exceeded 30% body weight loss.</li>
	</ol>
	</li>
	<li>Infect the mouse with the H1N1 virus.
	<ol>
		<li>Dissolve 0.8 x LD50 of H1N1 virus in 50 &#181;L of phosphate-buffered saline (PBS).</li>
		<li>Anesthetize the mice with 2.5% isoflurane.</li>
		<li>When the mice were deeply anesthetized, place the mouse in a supine position with the nostrils facing upward.</li>
		<li>Instill 5-10 &#181;L of H1N1 virus solution or PBS solution (as a control) intranasally into one nostril of mice using a pipette (Figure 2A).</li>
		<li>Then, hold the mouse&#39;s mouth closed with the thumb to make its nose inhale hard so that the H1N1 virus solution in the nasal cavity is completely inhaled into the lungs (Figure 2B).</li>
		<li>Repeat steps 1.2.4 and 1.2.5 in the other nasal cavity of the mice.</li>
		<li>When all H1N1 virus prepared in step 1.2.1 is intranasally instilled into the mouse&#39;s lungs, place it in the supine position for rest (Figure 2C).</li>
	</ol>
	</li>
	<li>Measure the cough sensitivity of the mouse after model establishment using the Buxco small animal whole-body plethysmography system. On day 21, intraperitoneally anesthetize the mouse with pentobarbital sodium (150 mg&#183;kg-1). Collect and process blood, spleen, BALF, lung, and trachea tissues (Figure 1).</li>
</ol>
2. The cough sensitivity measurement
<ol>
	<li>Prepare citric acid (0.4 M): Place 0.1537 mg of citric acid into a 5 mL centrifuge tube and add normal saline to a volume of 2 mL.</li>
	<li>Instrument inspection
	<ol>
		<li>Check the channels: Connect the whole-body plethysmograph chambers according to the manufacturer&#39;s instructions and click the calibration button-the calibration button changes from orange to green, indicating that calibration is successful (Figure 3A).</li>
		<li>Calibrate the nebulizer:
		<ol>
			<li>After connecting the nebulizer, add 500 &#181;L of normal saline into the nebulizer and click the nebulization button.</li>
			<li>When the liquid in the nebulizer is completely aerosolized, click the nebulization button again to see the power of the nebulizer, which is generally about 0.3 mL/min. Click the nebulization button again to accept the current nebulized power.</li>
		</ol>
		</li>
		<li>After checking the channels, ensure the error value is less than 0.5%.</li>
	</ol>
	</li>
	<li>Setting parameters
	<ol>
		<li>Click Create New Study, select the Cough option, select Mouse in Species, and click Next.</li>
		<li>Select the CCnt parameters: Set the duration of the acclimation period to 1 min, the response time to 10 min, the aerosol volume to 1 mL, and the delivery duration to 10 min.</li>
		<li>Place conscious unrestrained mouse in individual plastic transparent whole-body plethysmograph chambers. Input the Weight and Subject ID of the mouse and click Next. 
		NOTE: Mice are thigmotaxic and do not need to habituate to the small chambers; hence, habituation is not performed before the experiment.</li>
		<li>Add 1 mL of citric acid (0.4 M) solution to the nebulizer and observe real-time changes in CCnt (Figure 3B).</li>
		<li>After citric acid is completely used, click File and End Session to complete the experiment.</li>
	</ol>
	</li>
</ol>
3. Harvesting of blood, spleen, BALF, lung, and trachea tissues of mouse (Figure 4)
<ol>
	<li>Collect blood.
	<ol>
		<li>After cough detection, anesthetize the mouse with pentobarbital sodium (150 mg&#183;kg-1) by intraperitoneal injection. Collect blood from the orbits of the anesthetized mouse. Mix 1 mL of blood in 0.1 mL of anticoagulant buffer (9.9 mg/mL heparin sodium dissolved in PBS) at 4 &#176;C (Figure 4A). 
		NOTE: The method of retroorbital blood harvesting is simple and can get a large volume of blood.</li>
		<li>Shake the blood collection tube to fully mix the blood and anticoagulant to prevent blood coagulation.</li>
		<li>Centrifuge the collected blood at 800 x g for 5 min at 4 &#176;C. Collect the supernatant, store it at -80 &#176;C (used for cytokine measurement), and resuspend the pellet in 1 mL of D-Hank&#39;s solution. Spread 10 &#181;L of the blood cell suspension on the glass slide to determine cell profiles.</li>
	</ol>
	</li>
	<li>Harvest the spleen.
	<ol>
		<li>Open the mouse&#39;s chest, collect the blood, and exsanguinate via the artery (Figure 4B).</li>
		<li>Remove the left auricle, and then perfuse the pulmonary and systemic circulation with 5 mL of normal saline (Figure 4C, D). Remove the whole spleen from the mouse using surgical forceps (Figure 4E).</li>
		<li>After measuring the weight of the spleen, cut it into two halves. Fix the first half with 4% paraformaldehyde at room temperature (RT) for histopathological analysis. Store the second half at -80 &#176;C for cytokine measurement.</li>
	</ol>
	</li>
	<li>Bronchoalveolar lavage
	<ol>
		<li>Make the bronchoalveolar lavage tube using a Pasteur pipet. Heat the Pasteur pipette using an alcohol lamp. When it becomes soft, lengthen it to form a thin tube. Glue the catheter portion of a slip-tip catheter to the thin tube. The length of the bronchoalveolar lavage tube is 5 cm. The upper diameter is 5 mm, and the lower diameter is 1 mm (Figure 5). 
		NOTE: This bronchoalveolar lavage tube is a viable tool for BALF collection.</li>
		<li>For BALF collection, separate the right lung by ligating at the right mainstem bronchus. Collect the BALF from the left lungs by lavaging three times with 0.5 mL of PBS pre-cooled on ice. The recovery ratio of BALF is more than 80% (Figure 4F).</li>
		<li>Centrifuge the collected BALF at 800 x g for 5 min at 4 &#176;C. Collect the supernatant and measure the total protein concentration, uric acid concentration, LDH activity, and cytokine concentrations. 
		NOTE: The inflammatory markers, including total protein concentration, the uric acid concentration, and the LDH activity in the BALF supernatant, are detected using the assay kit according to the manufacturer&#39;s guidelines24.</li>
		<li>Resuspend the pellet in 200 &#181;L of PBS, and count the leukocytes in BALF with the help of a counting slide.</li>
		<li>To determine cell profiles by differential counts, smear 50 &#181;L of the cell suspension onto the glass slides and allow it to dry.</li>
		<li>Fix the slide with 4% paraformaldehyde overnight, and then stain with the hematoxylin-eosin (HE). Classify at least 400 cells into neutrophils, macrophages, lymphocytes, or eosinophils per slide.</li>
	</ol>
	</li>
	<li>Harvesting the lung and trachea tissues for histopathological analysis
	<ol>
		<li>After BALF collection, remove and fix half of the right lung (Figure 4G) and trachea (Figure 4H) with 4% paraformaldehyde at RT for histopathological analysis. Store the other half at -80 &#176;C for western blotting, Real-time quantitative polymerase chain reaction (qPCR), and Enzyme-linked immunosorbent assay (ELISA).</li>
	</ol>
	</li>
</ol>

Noninvasive Real-Time Whole-Body Plethysmography for Cough Sensitivity in Mice

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The authors have nothing to disclose.

Some chronic refractory and postinfectious coughs are common conditions associated with respiratory virus infection27. In order to better evaluate cough sensitivity and airway inflammation, a mouse model of cough was established using the H1N1 virus in this study. Appropriate mouse cough models should be selected for other studies according to the purpose of the study. Most previous studies used the guinea pig as an animal model in mechanistic studies or new drug trials for cough...

Cough is an important defense behavior to maintain airway patency and protect the lungs from potentially harmful substances. However, when dysregulated, cough becomes a pathological condition1. Chronic cough, usually defined as lasting eight or more weeks, is one of the most frequent symptoms requiring medical attention2. Because chronic cough frequently persists for years, patients suffer from a huge socioeconomic burden and a marked decrement in quality of life3,4,5. Chronic cough is widely considered a cough hypersensitivity syndrome and is characterized by troublesome coughing often triggered by low levels of thermal, mechanical, or chemical exposure6. The occurrence of cough hypersensitivity is closely related to airway inflammation7. However, the pathophysiological mechanisms underlying the modulation of cough sensitivity need to be further elucidated.
Animal models can mimic the complex pathophysiology of the cough and are important tools for cough research8,9. Previous studies have found that viral infection, intrapulmonary Interferon-&#947; (IFN-&#947;) instillation, esophageal perfusion of hydrochloric acid, pollutant exposure, cigarette smoke, and citric acid can induce cough in animals10,11,12,13,14,15,16,17. In order to better evaluate cough and airway inflammation, a mouse model of cough was established using a nonlethal dose of the H1N1 virus in this study. For the detection of cough, some cough measurement tools have been established clinically to measure cough, including subjective and objective methods18. The subjective evaluation tools to assess the cough severity primarily include a visual analog scale, the cough score, and quality of life questionnaires, etc19,20. However, they are unlikely to be used to assess cough in animals. In addition, cough can be objectively assessed through a cough challenge test and cough frequency monitoring. The cough challenge test using a whole-body plethysmography (WBP) system is an objective method widely used in animal studies to measure cough sensitivity and reveal the underlying mechanisms of cough13,16. Based on the neuroanatomical characteristics of the cough reflex, citric acid, capsaicin, adenosine 5&#39;-triphosphate (ATP), allyl isothiocyanate (AITC), and inflammatory mediator bradykinin are commonly used as the tussive agents to induce cough21,22. Citric acid is one of the earliest and the most widely used tussive agents triggering cough re&#64258;exes, which has been validated for measuring cough sensitivity. Besides, the citric acid challenge has good safety, feasibility, and tolerability and is suggested to assess cough reflex sensitivity in response to cough therapies23. Therefore, this article will describe the method for measuring cough sensitivity in response to citric acid in mice using a noninvasive and real-time WBP system.
The studies of the pathophysiology of cough require test samples, including samples from the blood, bronchoalveolar lavage fluid (BALF), and lung and trachea tissues to verify changes in the levels of key factors24. Currently, there is a lack of normative procedures for harvesting tissue samples of mice, and related studies employ different approaches that complicate the assessment of airway inflammation. Bronchoalveolar lavage is an important method to evaluate airway inflammation in respiratory diseases25. Different methods of bronchoalveolar lavage will lead to a lack of comparability between related studies. Moreover, different bronchoalveolar lavage methods have an impact on inflammatory cells and inflammatory cytokines in BALF. Therefore, this article will describe the establishment of a mouse model of cough with a nonlethal dose of the H1N1 virus, the measurement of cough using a WBP system, and a reliable, safe, and highly successful bronchoalveolar lavage method of mice.

<table><tbody><tr><td>4% paraformaldehyde&amp;nbsp;</td><td>Biosharp</td><td>BL539A</td><td></td></tr><tr><td>Buxco Small Animal Whole Body Plethysmography System&amp;nbsp;</td><td>DSI</td><td>&amp;mdash;</td><td></td></tr><tr><td>Calcium-free and magnesium-free Hank&amp;rsquo;s Balanced Salt Solution</td><td>Beyotime</td><td>C0219</td><td></td></tr><tr><td>Citric acid</td><td>Sigma-Aldrich</td><td>C2404</td><td></td></tr><tr><td>Hematoxylin-Eosin</td><td>BASO Biotechnology</td><td>BA-4098</td><td></td></tr><tr><td>Heparin sodium&amp;nbsp;</td><td>Alfa Aesar</td><td>A16198</td><td></td></tr><tr><td>Influenza A/California/7/2009 (H1N1) virus</td><td>ATCC</td><td>VR-1894</td><td></td></tr><tr><td>Isoflurane</td><td>RWD</td><td>R510-22</td><td></td></tr><tr><td>Lactate dehydrogenase assay kit</td><td>Nanjing Jiancheng Bioengineering Institute</td><td>A020-2-2</td><td></td></tr><tr><td>Normal saline</td><td>Guangzhou Zhongbo Biotechnology</td><td>1234-1</td><td></td></tr><tr><td>Pasteur pipet</td><td>NEST</td><td>318415</td><td></td></tr><tr><td>Pentobarbital sodium</td><td>Merck</td><td>P3761</td><td></td></tr><tr><td>Phosphate buffered saline&amp;nbsp;</td><td>Meilunbio</td><td>MA0015</td><td></td></tr><tr><td>Total protein assay kit</td><td>Nanjing Jiancheng Bioengineering Institute</td><td>A045-3</td><td></td></tr><tr><td>Uric acid assay kit</td><td>Thermo Fisher Scientific</td><td>A22181</td><td></td></tr></tbody></table>

methods for detecting cough and airway inflammation in mice

Chronic cough, which lasts for more than 8 weeks, is one of the most common complaints requiring medical attention, and patients suffer from a huge socioeconomic burden and a marked decrement in quality of life. Animal models can mimic the complex pathophysiology of the cough and are important tools for cough research. The detection of cough sensitivity and airway inflammation is of great significance for studying the complex pathological mechanism of cough. This article describes the measurement of cough using a noninvasive and real-time whole-body plethysmography (WBP) system and the normative procedures for harvesting tissue samples (including blood, lung, spleen, and trachea) of mice. It introduces some methods to assess airway inflammation, including pathological changes in hematoxylin and eosin (HE)-stained lung and trachea sections, the total protein concentration, the uric acid concentration, and the lactate dehydrogenase (LDH) activity in the supernatant of bronchoalveolar lavage fluid (BALF), and the leukocytes and differential cell counts of BALF. These methods are reproducible and serve as valuable tools to study the complex pathophysiology of cough.

Figure 6 shows representative images of pathological changes in HE-stained lung (Figure 6A,B), trachea (Figure 6C,D), and spleen (Figure 6E,F). H1N1 virus infection resulted in inflammatory changes in mouse lungs, including edema and many lymphocytes and neutrophil infiltration. H1N1 virus infection also induced inflammatory changes in mouse tracheae, i...

Methods for Detecting Cough and Airway Inflammation in Mice

Here, we describe the measurement of cough using a noninvasive and real-time whole-body plethysmography (WBP) system and the normative procedures for harvesting tissue samples of mice and introduce some methods to assess airway inflammation.

Chronic cough, which lasts for more than 8 weeks, is one of the most common complaints requiring medical attention, and patients suffer from a huge ...

methods-for-detecting-cough-and-airway-inflammation-in-mice

Research

JoVE Journal

Medicine

551 Views.  The First Affiliated Hospital of Guangzhou Medical University. Mijn onderzoek richt zich op chronische hoest. Om de pathofysiologie van hoest te begrijpen, hadden mijn studies testmonsters nodig om veranderingen in sleutelfactorniveaus te verifiëren. Ik streef ernaar gestandaardiseerde procedures op te stellen voor het oogsten van weefselmonsters van muizen.Overgevoeligheid voor hoesten wordt vaak veroorzaakt door een virale infectie. Voor het eerst ontdekten we dat virale infectie overgevoeligheid voor hoest kan...

Video: Auteur in de schijnwerpers: onderzoek naar de pathofysiologie van chronische hoest

Using Visual and Narrative Methods to Achieve Fair Process in Clinical Care

The Institute of Medicine has targeted patient-centeredness as an important area of quality improvement. A major dimension of patient-centeredness is respect for patient's values, preferences, and expressed needs. Yet specific approaches to gaining this understanding and translating it to quality care in the clinical setting are lacking. From a patient perspective quality is not a simple concept but is best understood in terms of five dimensions: technical outcomes; decision-making efficiency; amenities and convenience; information and emotional support; and overall patient satisfaction. Failure to consider quality from this five-pronged perspective results in a focus on medical outcomes, without considering the processes central to quality from the patient's perspective and vital to achieving good outcomes. In this paper, we argue for applying the concept of fair process in clinical settings. Fair process involves using a collaborative approach to exploring diagnostic issues and treatments with patients, explaining the rationale for decisions, setting expectations about roles and responsibilities, and implementing a core plan and ongoing evaluation. Fair process opens the door to bringing patient expertise into the clinical setting and the work of developing health care goals and strategies. This paper provides a step by step illustration of an innovative visual approach, called photovoice or photo-elicitation, to achieve fair process in clinical work with acquired brain injury survivors and others living with chronic health conditions. Applying this visual tool and methodology in the clinical setting will enhance patient-provider communication; engage patients as partners in identifying challenges, strengths, goals, and strategies; and support evaluation of progress over time. Asking patients to bring visuals of their lives into the clinical interaction can help to illuminate gaps in clinical knowledge, forge better therapeutic relationships with patients living with chronic conditions such as brain injury, and identify patient-centered goals and possibilities for healing. The process illustrated here can be used by clinicians, (primary care physicians, rehabilitation therapists, neurologists, neuropsychologists, psychologists, and others) working with people living with chronic conditions such as acquired brain injury, mental illness, physical disabilities, HIV/AIDS, substance abuse, or post-traumatic stress, and by leaders of support groups for the types of patients described above and their family members or caregivers.

This paper illustrates an innovative visual approach (photovoice or photo-elicitation) to achieve fair process in clinical care for patients living with chronic health conditions, illuminate gaps in clinical knowledge, forge better therapeutic relationships, and identify patient-centered goals and possibilities for healing.

The Institute of Medicine has targeted patient-centeredness as an important area of quality improvement. A major dimension of patient-centeredness is ...

Murine Model of Allergen Induced Asthma

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, with the prevalence increasing.2 As with other diseases, animal models of allergic airway disease greatly facilitate understanding of the underlying pathophysiology, help identify potential therapeutic targets, and allow preclinical testing of possible new therapies. Models of allergic airway disease have been developed in several animal species, but murine models are particularly attractive due to the low cost, ready availability, and well-characterized immune systems of these animals.3 Availability of a variety of transgenic strains further increases the attractiveness of these models.4 Here we describe two murine models of allergic airway disease, both employing ovalbumin as the antigen. Following initial sensitization by intraperitoneal injection, one model delivers the antigen challenge by nebulization, the other by intratracheal delivery. These two models offer complementary advantages, with each mimicking the major features of human asthma.5
The major features of acute asthma include an exaggerated airway response to stimuli such as methacholine (airway hyperresponsiveness; AHR) and eosinophil-rich airway inflammation. These are also prominent effects of allergen challenge in our murine models,5,6 and we describe techniques for measuring them and thus evaluating the effects of experimental manipulation. Specifically, we describe both invasive7 and non-invasive8 techniques for measuring airway hyperresponsiveness as well as methods for assessing infiltration of inflammatory cells into the airways and the lung. Airway inflammatory cells are collected by bronchoalveolar lavage while lung histopathology is used to assess markers of inflammation throughout the organ. These techniques provide powerful tools for studying asthma in ways that would not be possible in humans.

Experimental mouse models of allergic asthma offer new possibilities for studying disease pathogenesis and developing new therapeutics. These models are well suited to measuring factors governing the allergic immune response, airway inflammation, and pulmonary pathophysiology.

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, ...

Immunology and Infection

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 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

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 ...

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

The forced oscillation technique (FOT) is a powerful, integrative and translational tool permitting the experimental assessment of lung function in mice in a comprehensive, detailed, precise and reproducible manner. It provides measurements of respiratory system mechanics through the analysis of pressure and volume signals acquired in reaction to predefined, small amplitude, oscillatory airflow waveforms, which are typically applied at the subject's airway opening. The present protocol details the steps required to adequately execute forced oscillation measurements in mice using a computer-controlled piston ventilator (flexiVent; SCIREQ Inc, Montreal, Qc, Canada). The description is divided into four parts: preparatory steps, mechanical ventilation, lung function measurements, and data analysis. It also includes details of how to assess airway responsiveness to inhaled methacholine in anesthetized mice, a common application of this technique which also extends to other outcomes and various lung pathologies. Measurements obtained in na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage are presented to illustrate how this tool can contribute to a better characterization and understanding of studied physiological changes or disease models as well as to applications in new research areas.

The present protocol provides a detailed step-by-step description of the procedures required to execute measurements of respiratory system mechanics as well as the assessment of airway responsiveness to inhaled methacholine in mice using the forced oscillation technique (flexiVent; SCIREQ Inc, Montreal, Qc, Canada).

The  forced oscillation technique (FOT) is a powerful, integrative and translational  tool permitting the experimental assessment of lung function in mice ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Preserving Fungal Spore Localization in Lungs for Fluorescent Microscopy Analysis

Efficient Method for Imaging Murine Lungs that Preserves Spatial Dynamics of Fungal Spores in the Airways

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and bronchioles, branch until terminating in the alveolar airspace where gas exchange occurs. Infections originating in the bronchioles or alveoli elicit distinct host responses and disease manifestations. Therefore, understanding precisely where spores naturally localize in the lungs, particularly soon after infection, expands opportunities for investigation of host-pathogen interactions. Herein, we detail an in-situ analysis of lungs from mice infected with Coccidioides posadasii cts2/ard1/cts3&#916; arthroconidia. Conventional methods for histological preservation involve liquid inflation of the airways with a fixative solution, which displaces the natural location of aspirated fungal particles, pushing spores from proximal bronchioles to terminal airspaces.
Conversely, this method of air-inflation with blood vasculature perfusion-fixation preserves the physiologic position of fungal spores within the bronchioles. Moreover, we describe a simple approach to cryopreserving, embedding, and imaging lung specimens. We also share high-throughput computational techniques via the open-source QuPath program to analyze the spatial distribution of fungal spores within the lung. The method presented here is simple and quick, requires minimal equipment to perform, and can be easily adapted for use with many respiratory fungal infection models.

We present a method for a fungal pathogenesis model that preserves the natural positioning of fungal spores in the lung airways for analysis via fluorescent microscopy.

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and ...

In vivo Evaluation of Mucociliary Clearance in Mice

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a metachronal, synchronal fashion, these multiple, motile, actin-based organelles generate a cephalad fluid flow clearing the respiratory tract of inhaled pollutants and pathogens. With increasing environmental pollution, novel viral pathogens and emerging multi-drug resistant bacteria, cilia generated mucociliary clearance (MCC) is essential for maintaining lung health. MCC is also depressed in multiple congenital disorders like primary ciliary dyskinesia, cystic fibrosis as well as acquired disorders like chronic obstructive pulmonary disease. All these disorders have established, in some case multiple, mouse models. In this publication, we detail a method using a small amount of radioactivity and dual-modality SPECT/CT imaging to accurately and reproducibly measure MCC in mice in vivo. The method allows for recovery of mice after imaging, making serial measurements possible, and testing potential therapeutics longitudinally over time. The data in wild-type mice demonstrates the reproducibility of the MCC measurement as long as adequate attention to detail is paid, and the protocol strictly adhered to.

In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a ...

Citric Acid-Induced Mouse Model for Cough Hypersensitivity

Establishment of a Mouse Model with Cough Hypersensitivity via Inhalation of Citric Acid

Cough is one of the most common symptoms of many respiratory diseases. Chronic cough significantly impacts quality of life and imposes a considerable economic burden. Increased cough sensitivity is a pathophysiological hallmark of chronic cough. It has been observed that cough hypersensitivity is related to airway inflammation, remodeling of airway sensory nerves, and alterations in the central nervous system. However, the precise molecular mechanisms remain unclear and require further elucidation using suitable animal models. Previous studies have utilized guinea pigs as models for studying cough, but these models present several experimental limitations, including high costs, a lack of transgenic tools, and a scarcity of commercial reagents. In addition, guinea pigs typically exhibit poor environmental tolerance and high mortality when exposed to stimuli. In contrast, mice are smaller, easier to maintain, more cost-effective, and amenable to genetic manipulation, making them more suitable for mechanistic investigations. In this study, we established a mouse model with cough hypersensitivity&#160;via&#160;continuous inhalation of citric acid (CA). This model is straightforward to operate and yields reproducible results, making it a valuable tool for further studies on the mechanisms and potential novel treatments for chronic cough.

This protocol describes the development of a mouse model with cough hypersensitivity, which can serve as an ideal model for studying the mechanisms of chronic cough.

Cough is one of the most common symptoms of many respiratory diseases. Chronic cough significantly impacts quality of life and imposes a considerable ...

Methods for Detecting Cough and Airway Inflammation in Mice

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