TKMcG is supported by a studentship from the Canadian Thoracic Society.
AUTHORS' CONTRIBUTION
 All authors took part in the conception of the manuscript. In addition, TKMcG initiated the project, collected experimental results, contributed to the writing of the manuscript and its critical review. AR collected and analyzed experimental results, drafted the manuscript and contributed to its critical review. LF collected experimental results and contributed to the critical review of the manuscript. TFS and JGM contributed to the critical review of the manuscript.

The procedures described below were approved by McGill University Institutional Animal Care Committee in accordance with the guidelines of the Canadian Council on Animal Care (CCAC).
1. Preparatory Steps
<ol>
 <li>Solutions:
 <ol>
 <li>Methacholine: Prepare a stock solution at 50 mg/ml and make serial dilutions (1:1) based on the concentrations to be tested5. Allow the solutions to reach room temperature before nebulizing5.</li>
 <li>Anesthetic agents: Different regimens have been reported in the literature in various strains of mice (Table 1). Note: Regimen 1 was used under the present protocol.</li>
 </ol>
 </li>
 <li>Equipment: The present protocol applies to either of the two flexiVent generations supported by flexiWare 7 software. The software functions are grouped under three modules: Study Definition &amp; Planning, Experimentation Session and Review &amp; Reporting.
 <ol>
 <li>Turn on the system (flexiVent FX only) and/or start the software.</li>
 <li>At the first experimentation session or any time before it, open the Study Definition &amp; Planning module to predefine the study structure.</li>
 <li>Click on the Create a new study button and follow the wizard to create a study, outline the protocol and define the experimental groups and subjects to be studied.</li>
 <li>Initiate an experimentation session by opening the Experimentation Session module and following the start-up sequence for study and template selection.</li>
 <li>Assign a subject to the measurement site and confirm its weight.</li>
 <li>Proceed with the calibration of the system by following the steps described in the operating software. You will be prompted at one point to attach the cannula to be used (step 1.3.3) to the Y-tubing for calibration.</li>
 <li>Critical step. Repeat step 1.2.6 if the calibration values obtained are outside the acceptable range. (Refer to the flexiVent FX or flexiWare 7 user manual for the module specific acceptable ranges of calibration values).</li>
 <li>Cancel prompts to start ventilation and data recording unless ready to start the experiment. These can be initiated at a later point.</li>
 </ol>
 </li>
 <li>Subjects:
 <ol>
 <li>Anesthetize the subject using appropriate doses of anesthetic agents (Table 1).</li>
 <li>Verify that the subject has reached a surgical level of anesthesia. The subject should show no reaction to a toe pinch and its breathing should be regular and not labored.</li>
 <li>Perform a tracheotomy and cannulate the trachea.
 <ol>
 <li>Place the animal on its back and provide a source of heat (e.g. temperature controlled heating blanket or a lamp with a 60 watt bulb situated approximately 45 cm from the mouse to avoid excessive heating).</li>
 <li>Clean the throat area with alcohol and expose the trachea by making a skin incision and gently separating the submaxillary gland and the muscle layer covering it.</li>
 <li>Gently lift the trachea using a pair of micro-forceps and pass suture underneath it.</li>
 <li>Cut between two rings of cartilage nearest the larynx to make a small incision in the trachea without sectioning it.</li>
 <li>Insert the previously calibrated cannula into the incision and advance it gently inside the trachea the length of approximately 5 rings. Note: The present experiments were conducted using a 1.2 cm long metal 18 gauge cannula.</li>
 <li>Critical step. Secure the cannula in place using the suture. The attachment should form an airtight seal around the cannula.</li>
 </ol>
 </li>
 </ol>
 </li>
</ol>
2. Mechanical Ventilation
<ol>
 <li>Bring the animal close to the ventilator.</li>
 <li>Start mechanical ventilation by selecting a predefined or a customized ventilation profile in the ventilation docker.</li>
 <li>Connect the animal to the ventilator via the Y-tubing.</li>
 <li>Critical step. Align the animal to the ventilator and ensure that the tracheal cannula is at the same level as the ventilator to avoid a possible cannula occlusion or tracheal twist.</li>
 <li>Critical Step. Execute a Deep Inflation perturbation by double-clicking on the perturbation name to verify the cannula insertion and attachment. In absence of a leak, the system should be able to hold a pressure of 30 cmH2O over a 3 second period without excessive volume displacement (Figure 1). The recorded volume and pressure traces should also be smooth with no signs of offset or deformation as these could indicate a cannula obstruction or misplacement.</li>
 <li>If needed, connect vital sign transducers for heart rate and body temperature monitoring. Data recording can be initiated either manually or automatically via a script.</li>
</ol>
3. Lung Function Measurements
Measurements or commands (e.g. nebulizer activation, event markers) can be automated using predefined or customized scripts for a highly controlled and repeatable experimental process (Figure 2). Six families of perturbations giving rise to a number of parameters can be used to describe the subject respiratory system mechanics at baseline and following a given challenge (Table 2).
<ol>
 <li>Critical step. When ready to start taking measurements, run a Deep Inflation to recruit closed lung areas and standardize lung volume history.</li>
 <li>Critical step. Verify the absence of spontaneous inspiratory efforts by running a test measurement (e.g. PVs-P or PVs-V). Observe the pressure signal traces in the Selected Dataset view. With stepwise PV curves, pressure plateaus should be well defined with no downwards deflections. A downward swing in pressure would indicate an inspiratory effort from the animal (Figure 3).</li>
 <li>Initiate the selected script by double-clicking on its title. Scripts used in the present study generally included for measurements:
 <ul>
 <li>A sequence of baseline measurements in triplicate.</li>
 <li>Activation of the nebulizer for assessment of airway responsiveness to inhaled methacholine. Note: When prompted by the system, load approximately 100 &mu;l of saline or of a solution of methacholine into the nebulizer. Nebulization will start and stop automatically.</li>
 <li>A sequence of closely spaced measurements (every 10-15 sec) for a period of approximately 3 min following the activation of the nebulizer.</li>
 <li>A prompt to perform another challenge and repeat a sequence of measurements. Note: Drying the inside the nebulizer mount with a swab in between challenges can help prevent droplets or condensation from building-up in the inspiratory line.</li>
 </ul>
 </li>
 <li>At the end of the experiment, stop ventilation and detach the subject.</li>
 <li>Switch to the next subject in the operating software and confirm its weight.</li>
 <li>Critical step. Rinse and dry nebulizer, adaptor, Y-tubing, and cannula between subjects.</li>
 <li>Repeat steps 1.2.6 to 3.6.</li>
 <li>At the end of the day, close the experimental session. Remember to rinse and dry nebulizer, adaptor, Y-tubing, and cannula and to clean the system expiratory valve before leaving the laboratory by following the manufacturer's instructions.</li>
</ol>
4. Data Analysis
The software automatically calculates and displays parameters associated with a perturbation. It also provides a coefficient of determination (COD) which reflects the fit of the mathematical model to the data. Each dataset with an insufficient COD is labelled as excluded by the software. Review of experimental sessions, data re-analysis and creation of export scenarios are done in the Review &amp; Reporting module of the software.
<ol>
 <li>Open the Review &amp; Reporting module and create an export scenario, taking care to only include datasets having a sufficient COD.</li>
 <li>Export as needed parameters, pressure- or flow-volume curve, raw dataset signals or subject information to a spreadsheet application (see Table 3).</li>
 <li>Average baseline measurements for each parameter and plot all measurements as a function of time (see Figure 4). You may then choose to calculate the area under the curve, analyze the general profile of the curves or perform a statistical analysis.</li>
 <li>To express airway responsiveness results as a function of the concentration of methacholine, determine for each subject, parameter and experimental condition either a specific point (e.g. peak) or a specific time after each methacholine challenge. Calculate group averages and report or plot results for each experimental condition (Table 4, Figure 5).</li>
 <li>You may also consider calculating the concentration producing a doubling of a given parameter baseline value (PC200; Figure 5C), applying a normalization (e.g. % baseline) or performing a statistical analysis.</li>
</ol>

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AR, LF, TFS are employed by SCIREQ Scientific Respiratory Equipment Inc. TFS also owns stock. Free access to this article is sponsored by SCIREQ Scientific Respiratory Equipment, Inc.

The continued study of airway dysfunction as it relates to asthma and other lung disorders remains paramount to the understanding of underlying mechanisms of disease and the development of treatment options. The use of mice to model airway disease has been essential in gaining understanding into these disease mechanisms. When considering evaluating airway dysfunction in a subject as small as a mouse, having reliable and accurate tools by which to measure lung function is critical. Furthermore, having tools capable of providing insights on the location of airway dysfunction or therapeutic effect is invaluable. The FOT technique combines all these attributes and provides a powerful, integrative and translational approach to evaluate physiological changes.
In order to succeed with this type of measurement in mice, special attention should be given to a few steps, namely the calibration of the system, the resistance of the endotracheal cannula, the type of nebulizer (as well as its operating settings) the positioning of the animal and the standardization of lung volume history. In addition, it is imperative in order to obtain valid datasets that the subject's respiratory system remains passive during measurements. This can be achieved by the administration of a muscle paralyzing agent, working at a deep plane of anesthesia or by hyperventilating the subject to induce apnea (see Table 1). Investigators can start by mastering the system and its operating software, if desired, with test loads, while acquiring the necessary skills for measurements in mice. It would then be logical to generate reproducible results in na&iuml;ve animals before moving to disease models or treated mice. Since an important proportion of disease models in respiratory research involve exposing animals to agents such as allergens, toxins, pollutants, cigarette smoke or gases, variability in the results obtained with the measurement technique described in this article could therefore be influenced by the exposure procedure used. Standardization of key experimental processes (e.g. using computer-controlled exposure and measurement systems6, 13, 14) could potentially have a significant impact in reducing variability.
Examples presented in this article represent a selection of typical results from na&iuml;ve and chlorine-exposed mice experiments while highlighting the strengths as well as the limitations of the technique. As can be seen for example in Figure 6, the technique is capable of generating reproducible lung function measurements. While similar baseline resistance values were reported between mouse strains, differences in elastance were however observed15. Substantial changes are also to be expected between infant and adult mice16. As for other in vivo physiological assessment, high precision results, such as those generated by the FOT, come with a concession as to the natural state of the subjects. This principle, which is referred to as the phenotyping uncertainty principle1, applies to the present protocol in the sense that measurements need to be made in anesthetized, tracheotomised (or orally intubated) and mechanically ventilated subjects. Another limitation of the technique is observable in Figure 5D-5F where data are not available at the highest concentrations for the chlorine-exposed group because the fit of the Constant Phase Model to the data is poor above moderate levels of bronchoconstriction. However, severely bronchoconstricted animals could be assessed by analyzing Zrs directly15, or by using third party post-analysis software to fit more complex mathematical models, e.g. taking into account the heterogeneity of mechanical function17. Excluded datasets can also be observed if the animal's airways are not sufficiently passive or if the resistance of the cannula is too high. As a rule of thumb, the resistance of the cannula should not exceed the animal's resistance at baseline. Working with a cannula of larger inner diameter and/or shorter length will help reduce the cannula resistance. Finally, the present demonstration of FOT measurements in mice may be perceived as a time-consuming and therefore less efficient methodology or less applicable to longitudinal studies compared to less invasive techniques. However the latter are associated with a great deal of uncertainty as to the basis of their outcomes and are viewed by many as being flawed1. Repeated invasive measurements are possible in orally intubated animals, although technically more challenging17.
From the examples provided, results demonstrated the equivalence of the two generations of the flexiVent system at producing measurements of respiratory mechanics, as well as airway hyperreactivity and hypersensitivity to inhaled methacholine following chlorine-exposure in mice. When used to characterize or understand physiological changes or disease models, the detailed measurement aspect related to the technique can contribute to extend the current state of knowledge.

Adequate characterization of the mechanical properties of the lungs in small animals has become essential since the burgeoning of murine models in respiratory science. When performed using the forced oscillation technique (FOT), a technique also used in human subjects, these measurements provide a powerful, integrative and translational approach to study meaningful physiological changes. FOT measurements are typically obtained by analyzing pressure and volume signals acquired in reaction to a predefined, small amplitude, oscillatory airflow waveform (also referred to as perturbation or input signal) applied at the subject's airway opening1. In its simplest form, a FOT perturbation would be a single sinusoidal waveform at a well-defined frequency. More complex perturbations typically consist of a superposition of a selection of specific (mutually prime) frequency waveforms covering a broad spectrum. The decomposition of the multi-frequency input and output signals into their constituents using the Fourier transform allows the calculation of respiratory system input impedance (Zrs), i.e. the transfer function between the input and output signals, at every frequency included in the perturbation2. Therefore, FOT permits the simultaneous assessment of respiratory mechanics over a range of frequencies in a single maneuver2. Fitting advanced mathematical models (e.g. the Constant Phase Model3) to the impedance data then permit a partitioning of the response into airway (central and peripheral) and parenchymal lung tissue dependent parameters1, 3. Because many factors influencing the physiological response (e.g. breathing frequency, tidal volume, lung volume, upper airways, spontaneous breathing efforts, timing of measurements) are controlled and standardized by the measurement system and experimental procedures,1 the technique is capable of generating precise and reproducible measurements provided that it is performed correctly. The objective of this article is to provide a detailed, chronological description of the procedure needed in order to execute such measurements in mice. The protocol consists of four parts: preparatory steps (reagents, equipment and subjects), mechanical ventilation, lung function measurements, and data analysis. Examples of representative results of respiratory system mechanics generated using a computer-controlled piston ventilator (flexiVent, SCIREQ Inc, Montreal, Qc, Canada) are provided. These were obtained from na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage characterized by airway inflammation, epithelial cell damage and increased airway responsiveness to inhaled aerosolized methacholine4. While this protocol is often used to assess airway responsiveness to inhaled methacholine, it extends to other outcomes and various pathologies including asthma, chronic obstructive pulmonary disease (COPD), emphysema, lung fibrosis, lung injury as well as to transgenic mouse models of pathologies similar to human disease. Research findings using this tool can contribute to a better characterization and understanding of physiological changes or disease models as well as to expansion into new research areas.

<table><tbody><tr><td colspan="4">&lt;strong&gt;REAGENTS&lt;/strong&gt;</td></tr><tr><td>Acetyl-&amp;#946;-methylcholine chloride </td><td>Sigma-Aldrich</td><td>A-2251</td><td>Methacholine</td></tr><tr><td>Micro-Adson forceps, serrated 12 cm</td><td>Fine Science Tools</td><td>11018-12</td><td></td></tr><tr><td>Moria MC31 forceps, serrated-curved</td><td>Fine Science Tools</td><td>11370-31</td><td></td></tr><tr><td>Iris scissors-tough cut, straight 11.5 cm</td><td>Fine Science Tools</td><td>14058-11</td><td></td></tr><tr><td>Spring scissors-2.5 mm blades, straight</td><td>Fine Science Tools</td><td>15000-08</td><td></td></tr><tr><td>Non-sterile blunt needle (18g x &amp;frac12;&quot;)</td><td>Brico Medical Supplies Inc.</td><td>BN1805</td><td>Endotracheal cannula</td></tr><tr><td>Non-sterile 5-0 silk suture</td><td>Seraflex</td><td>IDI58000</td><td></td></tr><tr><td>Phosphate buffered solution</td><td>Gibco</td><td>14190-144</td><td></td></tr><tr><td>15 ml conical tubes</td><td>Starstedt</td><td>SS-4001</td><td></td></tr><tr><td>1 ml TB syringes</td><td>Becton Dickinson</td><td>309626</td><td></td></tr><tr><td>200 &amp;mu;l filter tips</td><td>Biosphere</td><td>70.760.211</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;EQUIPMENT&lt;/strong&gt;</td></tr><tr><td>&lt;em&gt;flexiVent&lt;/em&gt; FX</td><td><a href="http://www.scireq.com/" target="_blank"><img src="/img/new_window.png" /> SCIREQ Inc.</a></td><td>sales@scireq.com</td><td>&lt;a href=&quot;http://www.scireq.com/&quot; target=&quot;_blank&quot;&gt;www.scireq.com&lt;/a&gt;</td></tr><tr><td>Aerogen Aeroneb nebulizer</td><td><a href="http://www.scireq.com/" target="_blank"><img src="/img/new_window.png" /> SCIREQ Inc.</a></td><td>sales@scireq.com</td><td>&lt;a href=&quot;http://www.scireq.com/&quot; target=&quot;_blank&quot;&gt;www.scireq.com&lt;/a&gt;</td></tr></tbody></table>

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.

Respiratory system mechanics measurements. Table 4 shows typical results from na&iuml;ve A/J mice obtained at baseline and following methacholine-induced bronchoconstriction (12.5 mg/ml) using either of the two flexiVent generations supported by flexiWare 7 software. Mechanics of the respiratory system, i.e. under closed chest conditions, were assessed by alternating perturbations of the single and broadband frequency forced oscillation families in a closely spaced manner (SnapShot-150, Quick Prime-3, respectively). Since ventilation is paused during measurements, the Quick Prime-3, which covers a similar frequency range as the Prime-8 but has a shorter duration (3 vs 8 sec), was selected in order to shorten the apneic period, minimize the effect of the perturbation on blood gases and provide a better resolution of the response. Parameters associated with each perturbation were calculated automatically by the operating software. Results illustrate that the two generations of the flexiVent system produced equivalent measurements of respiratory mechanics.
 Site of lung response. Distinguishing the site of lung response allows the investigator to further pinpoint affected regions as well as to identify potential points of pharmacological intervention6. For example, na&iuml;ve A/J mice show an increase of baseline resistance when the end expiratory pressure against which measurements are made is increased from 3 to 9 cmH2O (Figure 6A, SnapShot-150). In the present example, the use of broadband FOT measurements (Quick Prime-3) provided details to clarify the basis for the change in resistance: The change in end expiratory pressure resulted in a decrease in the airway resistance (RN) consistent with the bronchodilating effects of a higher lung volume and the larger inflation pressure (Figure 6D) and an increase in tissue damping (G; Figure 6E), a parameter closely related to tissue resistance that reflects tissue viscoelasticity and possibly the resistance of the small airways7. The latter is known to increase with increasing lung volume.
 Airway hyperresponsiveness. Following chlorine gas exposure, airway responsiveness to inhaled methacholine increases compared to air exposure in Balb/c mice as a result of airway damage4 (Figure 2). Chlorine is known to induce oxidative stress, leading to destruction of structural cells in the airways, in particular epithelial cells, and inducing recruitment of inflammatory cells. As represented in Figure 5, changes in all parameters describing respiratory system mechanics can be seen in response to increasing methacholine challenges. In comparison to air-exposed mice, mice exposed to chlorine gas displayed greater maximal responses at all FOT parameters (Figure 5A, 5B, 5D-5F) as well as a statistically significant leftward shift of the concentration-response curve exemplified by a reduction of the concentration of methacholine required to cause a doubling in resistance and elastance (PC200; Figure 5C). Those results illustrate, respectively, airway hyperresponsiveness and hypersensitivity to inhaled methacholine following exposure to chlorine gas. 
 Other measurements. In addition to FOT, the flexiVent system can also be used to collect other types of lung function8-10 or cardiovascular11 measurements. Figure 7 displays a representative stepwise, pressure-driven pressure-volume curve in na&iuml;ve A/J mice under baseline conditions. The upper portion of the deflation limb of the curve is fit to the Salazar-Knowles equation12 and parameters are automatically calculated by the software.
<img src="/files/ftp_upload/50172/50172table1.jpg" alt="Table 1" fo:src="/files/ftp_upload/50172/50172table1highres.jpg" fo:content-width="6in" /> 
Table 1. Examples of anesthetic regimens used in mice. <a href="https://www.jove.com/files/ftp_upload/50172/50172table1large.jpg" target="_blank"> Click here to view larger table</a>.
 <img src="/files/ftp_upload/50172/50172table2.jpg" alt="Table 2" fo:src="/files/ftp_upload/50172/50172table2highres.jpg" fo:content-width="5in" /> 
Table 2. Perturbations used for lung function measurements in mice. *Extension required to the system. The subject also needs to be in a closed plethysmograph chamber during measurements. <a href="https://www.jove.com/files/ftp_upload/50172/50172table2large.jpg" target="_blank"> Click here to view larger table</a>.
<img src="/files/ftp_upload/50172/50172table3.jpg" alt="Table 3" fo:src="/files/ftp_upload/50172/50172table3highres.jpg" fo:content-width="6in" /> 
Table 3. Example of exported parameters from the single and broadband frequency forced oscillation perturbation families. <a href="https://www.jove.com/files/ftp_upload/50172/50172table3large.jpg" target="_blank"> Click here to view larger table</a>.
 <img src="/files/ftp_upload/50172/50172table4.jpg" alt="Table 4" fo:src="/files/ftp_upload/50172/50172table4highres.jpg" fo:content-width="6in" /> 
Table 4. System Comparison. Comparison of lung mechanics parameters collected using the two generations of the flexiVent system operated by flexiWare 7 software. Results were generated in na&iuml;ve A/J mice (n=5/group) at baseline and following methacholine-induced bronchoconstriction (Mch 12.5 mg/ml). *Groups were compared using a two-way ANOVA for repeated measurements and the log10 of individual responses for homogeneity of variances (GraphPad Prism, version 5.03; GraphPad Software, San Diego, USA).
<img src="/files/ftp_upload/50172/50172fig1.jpg" alt="Figure 1" fo:content-width="4.5in" fo:src="/files/ftp_upload/50172/50172fig1highres.jpg"/> Figure 1. Screenshot of a deep lung inflation. The upper panel shows the volume displaced by the piston of the ventilator (red trace) and the volume delivered to the subject (grey trace). The lower panel displays the cylinder pressure increasing to a set pressure of 30 cmH2O over a period of 3 seconds and held constant for the same time period.
 <img src="/files/ftp_upload/50172/50172fig2.jpg" alt="Figure 2" /> 
Figure 2. Example of a typical script used to assess respiratory system mechanics at baseline.
<img src="/files/ftp_upload/50172/50172fig3.jpg" alt="Figure 3" /> 
Figure 3. Spontaneous inspiratory efforts during the execution of a stepwise pressure-volume curve.
<img src="/files/ftp_upload/50172/50172fig4.jpg" alt="Figure 4" fo:content-width="2.6in" fo:src="/files/ftp_upload/50172/50172fig4highres.jpg"/> 
Figure 4. Time-course response following increasing inhaled methacholine challenges. Results are expressed as the mean (&plusmn; standard deviation) of a group of 5 na&iuml;ve spontaneously hyperresponsive A/J mice. <a href="https://www.jove.com/files/ftp_upload/50172/50172fig4large.jpg" target="_blank">Click here to view larger figure</a>. 
<img src="/files/ftp_upload/50172/50172fig5.jpg" alt="Figure 5" fo:content-width="6in" fo:src="/files/ftp_upload/50172/50172fig5highres.jpg"/> 
Figure 5. Changes in respiratory system mechanics following increasing methacholine challenges in chlorine- and air-exposed Balb/c mice. Peak value was identified for each parameter in every subject and experimental condition. Group averages were then calculated (mean &plusmn; standard deviation; n=4-6). Differences between groups were assessed by analysis of variance using the log10 of individual responses for homogeneity of variances. The concentration of methacholine producing a doubling of baseline (PC200) was obtained by fitting a second order polynomial to individual dose-response curves and interpolation of the fitted curve. Data points are missing in D, E and F in the chlorine-exposed mice at the two highest methacholine concentrations due to insufficiently high coefficients of determination reflecting a poor fit of the mathematical model to the data. <a href="https://www.jove.com/files/ftp_upload/50172/50172fig5large.jpg" target="_blank">Click here to view larger figure</a>.
 <img src="/files/ftp_upload/50172/50172fig6.jpg" alt="Figure 6" fo:content-width="6in" fo:src="/files/ftp_upload/50172/50172fig6highres.jpg"/> 
 Figure 6. Partitioning of the respiratory response into airway and lung tissue mechanics. Experimental trace from a na&iuml;ve A/J mice illustrating single (2.5 Hz) and broadband (1-20.5Hz) frequency forced oscillation measurements of respiratory mechanics in triplicate at two different end expiratory pressures (3 &amp; 9 cmH2O). <a href="https://www.jove.com/files/ftp_upload/50172/50172fig6large.jpg" target="_blank">Click here to view larger figure</a>.
 <img src="/files/ftp_upload/50172/50172fig7.jpg" alt="Figure 7" fo:content-width="4in" fo:src="/files/ftp_upload/50172/50172fig7highres.jpg"/> 
 Figure 7. Pressure-volume curve in na&iuml;ve A/J mice under baseline conditions. Pressure-volume curves were generated using a stepwise pressure-driven perturbation (PVs-P) to ensure that each mouse lungs were inflated to the same pressure, independently of their condition. Salazar-Knowles equation parameters extracted from individual pressure-volume curves were also averaged and reported in a table format. Results are expressed as the mean &plusmn; standard deviation (n=6).

Watch this Scientific Journal Video about Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique at JoVE.com

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

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 ...

evaluation-respiratory-system-mechanics-mice-using-forced-oscillation

Research

JoVE Journal

Biology

56.5K Views.  McGill University. 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).

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

Method to Obtain Pattern of Breathing in Senescent Mice through Unrestrained Barometric Plethysmography

Unrestrained barometric plethysmography (UBP) is a method for quantifying the pattern of breathing in mice, where breathing frequency, tidal volume, and minute ventilation are routinely reported. Moreover, information can be collected regarding the neural output of breathing, including the existence of central apneas and augmented breaths. An important consideration for UBP is obtaining a breathing segment with a minimal impact of anxious or active behaviors, to elucidate the response to breathing challenges. Here, we present a protocol that allows for short, quiet baselines to be obtained in aged mice, comparable to waiting for longer bouts of quiet breathing. The use of shorter time segments is valuable, as some strains of mice may be increasingly excitable or anxious, and longer periods of quiet breathing may not be achieved within a reasonable timeframe. We placed 22 month-old mice in a UBP chamber and compared four 15 s quiet breathing segments between minutes 60&#8211;120 to a longer 10 min quiet breathing period that took&#160;2&#8211;3 h to acquire. We also obtained counts of central apneas and augmented breaths prior to the quiet breathing segments, following a 30 min familiarization period. We show that 10 min of quiet breathing is comparable to using a much shorter 15 s duration. Additionally, the time leading up to these 15 s quiet breathing segments can be used to gather data regarding apneas of central origin. This protocol allows investigators to collect pattern-of-breathing data in a set amount of time and makes quiet baseline measures feasible for mice that may exhibit increased amounts of excitable behavior. The UBP methodology itself provides a useful and noninvasive way to collect pattern-of-breathing data and allows for mice to be tested over several time points.

Unrestrained barometric plethysmography is used to quantify the pattern of breathing in awake mice. We show that 15 s segments under a standardized protocol display similar values to an extended duration of quiet breathing. This methodology also allows for the quantification of apnea and augmented breaths during the first hour in the chamber.

Unrestrained barometric plethysmography (UBP) is a method for quantifying the pattern of breathing in mice, where breathing frequency, tidal volume, and ...

Behavior

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 ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not only ensure faithful genome duplication in the face of endogenous and exogenous DNA damage but also prevent genomic instability, a recognized causative factor in tumor development.
Here, we describe a simple and cost-effective fluorescence microscopy-based method to visualize DNA replication in the avian B-cell line DT40. This cell line provides a powerful tool to investigate protein function in vivo by reverse genetics in vertebrate cells1. DNA fiber fluorography in DT40 cells lacking a specific gene allows one to elucidate the function of this gene product in DNA replication and genome stability. Traditional methods to analyze replication fork dynamics in vertebrate cells rely on measuring the overall rate of DNA synthesis in a population of pulse-labeled cells. This is a quantitative approach and does not allow for qualitative analysis of parameters that influence DNA synthesis. In contrast, the rate of movement of active forks can be followed directly when using the DNA fiber technique2-4. In this approach, nascent DNA is labeled in vivo by incorporation of halogenated nucleotides (Fig 1A). Subsequently, individual fibers are stretched onto a microscope slide, and the labeled DNA replication tracts are stained with specific antibodies and visualized by fluorescence microscopy (Fig 1B). Initiation of replication as well as fork directionality is determined by the consecutive use of two differently modified analogues. Furthermore, the dual-labeling approach allows for quantitative analysis of parameters that influence DNA synthesis during the S-phase, i.e. replication structures such as ongoing and stalled forks, replication origin density as well as fork terminations. Finally, the experimental procedure can be accomplished within a day, and requires only general laboratory equipment and a fluorescence microscope.

DT40, a model vertebrate genetic system, provides a powerful tool to analyze protein function. Here we describe a simple method that allows qualitative analysis of parameters that influence DNA synthesis during the S-phase in DT40 cells at the single molecule level.

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not ...

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...

Medicine

Generation of a Chronic Obstructive Pulmonary Disease Model in Mice by Repeated Ozone Exposure

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high incidence in aging populations. The current conventional therapies for COPD focus mainly on symptom-modifying drugs; thus, the development of new therapies is urgently needed. Qualified animal models of COPD could help to characterize the underlying mechanisms and can be used for new drug screening. Current COPD models, such as lipopolysaccharide (LPS) or the porcine pancreatic elastase (PPE)-induced emphysema model, generate COPD-like lesions in the lungs and airways but do not otherwise resemble the pathogenesis of human COPD. A cigarette smoke (CS)-induced model remains one of the most popular because it not only simulates COPD-like lesions in the respiratory system, but it is also based on one of the main hazardous materials that causes COPD in humans. However, the time-consuming and labor-intensive aspects of the CS-induced model dramatically limit its application in new drug screening. In this study, we successfully generated a new COPD model by exposing mice to high levels of ozone. This model demonstrated the following: 1) decreased forced expiratory volume 25, 50, and 75/forced vital capacity (FEV25/FVC, FEV50/FVC, and FEV75/FVC), indicating the deterioration of lung function; 2) enlarged lung alveoli, with lung parenchymal destruction; 3) reduced fatigue time and distance; and 4) increased inflammation. Taken together, these data demonstrate that the ozone exposure (OE) model is a reliable animal model that is similar to humans because ozone overexposure is one of the etiological factors of COPD. Additionally, it only took 6 - 8 weeks, based on our previous work, to create an OE model, whereas it requires 3 - 12 months to induce the cigarette smoke model, indicating that the OE model might be a good choice for COPD research.

This study describes the successful generation of a new chronic obstructive pulmonary disease (COPD) animal model by repeatedly exposing mice to high concentrations of ozone.

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high ...

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 ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

Employing the Forced Oscillation Technique for the Assessment of Respiratory Mechanics in Adults

There is increasing interest in the use of the forced oscillation technique (FOT) or oscillometry to characterize respiratory mechanics in healthy and diseased individuals. FOT, a complementary method to traditional pulmonary function testing, utilizes a range of oscillatory frequencies superimposed on tidal breathing to measure the functional relationship between airway pressure and flow. This passive assessment provides an estimate of respiratory system resistance (Rrs) and reactance (Xrs) that reflect airway caliber and energy storage and dissipation, respectively. Despite the recent increase in popularity and updated Technical Standards, clinical adoption has been slow which relates, in part, to the lack of standardization regarding the acquisition and reporting of FOT data. The goal of this article is to address the lack of standardization across laboratories by providing a comprehensive written protocol for FOT and an accompanying video. To illustrate that this protocol can be utilized irrespective of a particular device, three separate FOT devices have been employed in the case examples and video demonstration. This effort is intended to standardize the use and interpretation of FOT, provide practical suggestions, as well as highlight future questions that need to be addressed.

As the use of forced oscillation technique (FOT) is increasingly utilized to characterize respiratory mechanics, there is a need to standardize methods with respect to nascent technical guidelines and various manufacturer's recommendations. A detailed protocol is provided including FOT assessment and interpretation for two cases to facilitate the standardization of methods.

There is increasing interest in the use of the forced oscillation technique (FOT) or oscillometry to characterize respiratory mechanics in healthy and ...

Conducting Respiratory Oscillometry in an Outpatient Setting

Respiratory oscillometry is a different modality of pulmonary function testing that is increasingly used in a clinical and research setting to provide information regarding lung mechanics. Respiratory oscillometry is conducted through three acceptable measurements of tidal breathing and can be performed with minimal contraindications. Young children and patients who cannot perform spirometry due to cognitive or physical impairment can usually complete oscillometry. The main advantages of respiratory oscillometry are that it requires minimal patient cooperation and is more sensitive in detecting changes in small airways than conventional pulmonary function tests. Commercial devices are now available. Updated technical guidelines, standard operating protocols, and quality control/assurance guidelines have recently been published. Reference values are also available. 
We conducted oscillometry test audits before and after implementing a formal respiratory oscillometry training program and standard operating protocol. We observed improvement in the quality of tests completed, with a significant increase in the number of acceptable and reproducible measurements. 
The current paper outlines and demonstrates a standard operating protocol to conduct respiratory oscillometry in an outpatient setting. We highlight the key steps to ensuring acceptable and reproducible quality measurements according to the recommended European Respiratory Society (ERS) guidelines, as quality control is critical to measurement accuracies. Potential problems and pitfalls are also discussed with suggestions to resolve technical errors.

We demonstrate a standard operating protocol to conduct respiratory oscillometry, highlighting key quality control and assurance procedures.

Respiratory oscillometry is a different modality of pulmonary function testing that is increasingly used in a clinical and research setting to provide ...