This work has been supported by NIH HL-1034.

The Johns Hopkins University Animal Care and Use Committee approved all animal protocols.
1. Equipment
The composite system set up, ready to measure the PV curve is shown in Figure 1.
<ol>
	<li>Volume measurement:
	<ol>
		<li>Generate a constant rate of inflation and deflation by using a syringe pump with a switch that allows the user to quickly reverse the pump after reaching the pressure limits. For mouse PV curves, use a very lightly greased 5 ml glass syringe with the initial volume (prior to inflation) set at 3 ml of air. 3 ml is sufficiently large to measure volumes in nearly all mouse PV curves.</li>
		<li>Measure the volume delivered by the pump by attaching a linear differential transformer to the pump housing, with a small sensor rod connected to the moving syringe plunger. 
		Note: An empirical means to correct for gas compression in the system is described under the PV curve recording section.</li>
	</ol>
	</li>
	<li>Pressure measurement:
	<ol>
		<li>Use a standard inexpensive pressure gauge with a range of 0-60 cm H2O (0-1 PSI).</li>
	</ol>
	</li>
	<li>Recording measurement:
	<ol>
		<li>To record the PV curve use any digital recorder with XY capabilities (e.g., PowerLab). Set one channel to record the corrected volume signal and another channel to record the transpulmonary pressure (Ptp), in order to graph the PV curve. Use a bridge preamplifier that connects to the main Powerlab to measure the pressure. Calibrate the pressure channel from 0-40 cm H2O, and calibrate the volume channel from 0-3 ml.</li>
	</ol>
	</li>
</ol>
2. Correction for Gas Compression
Note: This is a critical initial step in the set up, since as the pressure increases, the gas volume decreases, and thus the volume of air delivered to the mouse will be increasingly less than the displacement of the syringe barrel.
<ol>
	<li>Close the stopcock that will connect the PV system to the lungs, so no gas can leave the system. Start the infusion and observe if the corrected Volume channel on the recorder shows any measurable changes as the pressure increases to about 40 cm H2O. If so, then correct as in the next steps.
	<ol>
		<li>Correct for gas compression empirically by subtracting from the plunger displacement measurement (i.e., the uncorrected volume) a term proportional to the inflation pressure. Do this on a Powerlab channel (called Vc) to show the volume signal minus a coefficient times the pressure.</li>
		<li>Determine the coefficient in the equation. First, make an initial guess, turn the chart recording on, and start the pump. Since the inflation tubing is sealed, adjust the pressure coefficient multiplier to make the Vc channel read zero as the pressure rises from 0-40 cm H2O. If it goes up or down, simply adjust the correction factor until it stays flat over this pressure range. This correction factor will always be the same, if the same 3 ml starting volume in the syringe is not changed.</li>
	</ol>
	</li>
</ol>
3. Experimental Tests in Mice
<ol>
	<li>Procedure for measurement of the PV curve in mice. All animal protocols were approved by the Johns Hopkins University Animal Care and Use Committee.
	<ol>
		<li>Anesthetize mice (C57BL/6 mice at 6-12 weeks of age) with ketamine (90 mg/kg) and xylazine (15 mg/kg), and confirm anesthesia by the absence of reflex motion. 
		Note: The PV curve can be completed in anesthetized mice in less than 10 min and is a terminal procedure.&#160;</li>
		<li>Tracheostomize the mice with an 18 G stub needle cannula. Do this by making a small incision in the skin overlying the trachea, locating the trachea, then making a small slit in the trachea, where the stub needle can be inserted. Secure the cannula by tying with thread.</li>
		<li>Allow the mice to breath 100% oxygen for at least 4 min. This can be via spontaneous breathing from a bag or with a ventilator nominally set with a tidal volume of 0.2 ml at 150 breaths/min.</li>
		<li>Close the tracheal cannula and allow 3-4 min for the mouse to absorb all the oxygen. This oxygen absorption procedure results the death of the animals and in a nearly complete degassing of the lung11. Confirm death of the mouse by measuring the cessation of the heartbeat with ECG electrodes or direct observation.</li>
		<li>Once the degassing of the lung is complete and the lung volume is zero, begin inflating the lung with room air in the syringe pump&#160;at a rate of 3 ml/min. Monitor the pressure trace on the digital recorder, and when it reaches 35 cm H2O, reverse the pump.</li>
		<li>Follow the deflation curve until pressure reaches negative 10 cm H2O, by which time the airways have collapsed, trapping air in alveoli preventing further volume reduction. Immediately reverse the pump again, allowing the lung to reinflate as the collapsed airways open. This heterogeneous opening is normally apparent by the noisy looking inflation limb at the initial part of this 2nd inflation.</li>
		<li>When the pressure again reaches 35 cm H2O, reverse the pump direction, and continue to deflate the lung until this 2nd deflation limb reaches 0 cm H2O. Then stop the pump.</li>
		<li>View the PowerLab chart record of pressure and flow and the PV curve. Then analyze the PV curve to detect phenotypic changes in lung parenchyma that occur with different lung pathologies.</li>
	</ol>
	</li>
</ol>

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None of the authors have any financial interests that would be in conflict with the material presented in this paper.

In this paper a straightforward reproducible method has been described to measure in mice a classical method of phenotyping lung elasticity, the total lung PV curve. Such curves were instrumental in the discovery of pulmonary surfactant and its importance in providing lung stability. Here it is shown how the PV curve is also useful in providing a means to measure several variables related to lung elasticity in adult mouse lungs. There were highly significant changes in all variables in two commonly used mouse models to g...

The mouse is now the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measuring the changes in lung elasticity. Although there are many ways to measure elasticity, the classical method is that of the total pressure-volume (PV) curve measured from residual volume (RV) to total lung capacity (TLC). This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years1-3. Such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development4-7. Despite the PV curve&#8217;s importance as a measurement of the lung&#8217;s phenotype, there has been no standardized way to perform this measurement. It has been done simply by inflating and deflating the lung with discrete steps (waiting a variable time for equilibration after each) or with pumps that can continuously inflate and deflate the lung. The PV curve is often done over a volume range between zero and some user-define lung capacity, but the time duration of each pressure volume loop reported by different labs has been extremely variable, varying from a few sec8 to hr2. Some investigators refer to this total lung PV curve as static or quasistatic, but these are qualitative terms that offer little insight, and they are not used here. In addition, the PV curve has not been widely reported in the mouse, despite the fact that it can provide useful information on the macroscopic effects of structural changes in the lung.
Several issues have resulted in variability in PV curve acquisition including: 1) the rate of inflation and deflation; 2) the pressure excursions for inflation and deflation; and 3) the means to determine an absolute lung volume measurement. In the method present here, a rate of 3 ml/min was chosen as a compromise, being not too short as to reflect the dynamic elasticity associated with normal ventilation and not too slow as to make the measurement impractical, particularly when studying large cohorts. Since a nominal total lung capacity in a C57BL/6 healthy mouse is on the order of 1.2 ml9, this rate typically allows two complete closed PV loops to be done in about 1.5 min.
In the extended literature where PV curves have been reported, the peak inflation pressure used has been extremely variable, varying from as low as 20 to over 40 cm H2O. Part of this variability may be related to species, but a primary goal of setting the upper pressure limit for PV curves is to inflate the lung to total lung capacity (TLC), or maximal lung volume. The TLC in humans is defined by the maximal voluntary effort an individual can make, but unfortunately this can never be duplicated in any animal model. Thus, the maximal volume in experimental PV curves is determined by a maximal pressure arbitrarily set by the investigator. The goal is to set a pressure where the PV curve is flat, but unfortunately the inflation limb of a mammalian lung PV curve is never flat. So most investigators set a pressure where the inflation curve begins to flatten substantially, typically 30 cm H2O. In the mouse, however, the PV curve is even more complex with a double hump on the inflation limb, and where this inflation limb is often still rising steeply at 30 cm H2O10, so 30 is not a good end point for the PV curve. For this reason, we use 35 cm H2O as the pressure limit for the mouse PV curve, which is a pressure at which the inflation limbs of all strains we have examined begin to flatten.
Since the PV curve itself is very nonlinear, the appearance of a PV loop will depend on the volume from where the curve starts. Some commercial ventilators allow users to do large PV loops, starting from FRC, but if the FRC volume is unknown then it is impossible to interpret changes in such PV curve with any pathology, since these changes could simply result from a change in starting volume, and not structural alterations in the lung. Thus without an absolute volume measurement, PV curves are almost impossible to interpret and thus have little utility. Although, there are several ways to measure lung volumes, these&#160;are often cumbersome and require special equipment. In the simple approach described here, the PV curve starts at zero volume after an in vivo degassing procedure.
In summary, this paper demonstrates a straightforward method to standardize lung PV curve measurement in the mouse lung, and defines several metrics that can be calculated from this curve that are linked to lung structure. The PV curve thus provides a pulmonary function test that has direct application in being able to detect phenotypic structural changes in mice with common lung pathologies such as emphysema and fibrosis.

<table><tbody><tr><td>Syringe pump</td><td>Harvard Apparatus</td><td>55-2226</td><td>Infuse/withdraw syringe pump</td></tr><tr><td>Pump 22 reversing switch</td><td>Harvard Apparatus</td><td>552217</td><td>Included with pump</td></tr><tr><td>Linear displacement transformer</td><td>Trans-Tek, Inc.</td><td>0244-0000</td><td></td></tr><tr><td>5 ml glass syringe</td><td>Becton Dickenson</td><td></td><td>Several other possible vendors</td></tr><tr><td>Digital recorder</td><td>ADInstruments</td><td>PL3504</td><td>Several other possible vendors</td></tr><tr><td>Bridge amp signal conditioner</td><td>ADInstruments</td><td>FE221</td><td></td></tr><tr><td>Gas tank, 100% oxygen</td><td>Airgas, Inc</td><td></td><td>Any supplier or hospital source will work</td></tr><tr><td>Pressure transducer: 0 - 1 psi&amp;nbsp;mV output</td><td>Omega Engineering</td><td>PX-137</td><td>Range &amp;asymp; 0 - 60 cm H&lt;sub&gt;2&lt;/sub&gt;O</td></tr></tbody></table>

measurement of the pressure-volume curve in mouse lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Although the procedure for the PV curves is demonstrated in the video only for control healthy mice, we examined the ability of the PV curve to detect functional and pathologic changes in mice with two different common pathologies, emphysema and fibrosis. Details of these traditional models described elsewhere12,13. Very briefly, after anesthesia with 3% isoflurane the emphysema was caused by 3 or 6 U porcine pancreatic elastase instilled into the trachea and studied 3 weeks later, and the fibrosis was caused ...

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Measurement of the Pressure-volume Curve in Mouse Lungs

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

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18.5K Views.  Johns Hopkins University. Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung’s structural changes with developing pathology.

Video: Measurement of the Pressure-volume Curve in Mouse Lungs

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

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

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

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

Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is a common modifier of heart and lung function, by elaborating cellular infiltration, production of cytokines and growth factors, and by initiating remodeling processes 1.
Compared to the left ventricle, the right ventricle is a low-pressure pump that operates in a relatively narrow zone of pressure changes. Increased pulmonary artery pressures are associated with increased pressure in the lung vascular bed and pulmonary hypertension 2. Pulmonary hypertension is often associated with inflammatory lung diseases, for example chronic obstructive pulmonary disease, or autoimmune diseases 3. Because pulmonary hypertension confers a bad prognosis for quality of life and life expectancy, much research is directed towards understanding the mechanisms that might be targets for pharmaceutical intervention 4. The main challenge for the development of effective management tools for pulmonary hypertension remains the complexity of the simultaneous understanding of molecular and cellular changes in the right heart, the lungs and the immune system.
Here, we present a procedural workflow for the rapid and precise measurement of pressure changes in the right heart of mice and the simultaneous harvest of samples from heart, lungs and immune tissues. The method is based on the direct catheterization of the right ventricle via the jugular vein in close-chested mice, first developed in the late 1990s as surrogate measure of pressures in the pulmonary artery5-13. The organized team-approach facilitates a very rapid right heart catheterization technique. This makes it possible to perform the measurements in mice that spontaneously breathe room air. The organization of the work-flow in distinct work-areas reduces time delay and opens the possibility to simultaneously perform physiology experiments and harvest immune, heart and lung tissues.
The procedural workflow outlined here can be adapted for a wide variety of laboratory settings and study designs, from small, targeted experiments, to large drug screening assays. The simultaneous acquisition of cardiac physiology data that can be expanded to include echocardiography5,14-17 and harvest of heart, lung and immune tissues reduces the number of animals needed to obtain data that move the scientific knowledge basis forward. The procedural workflow presented here also provides an ideal basis for gaining knowledge of the networks that link immune, lung and heart function. The same principles outlined here can be adapted to study other or additional organs as needed.

A specific and rapid protocol to simultaneously investigate right heart function, lung inflammation, and the immune response is described as a learning tool. Video and figures describe physiology and microdissection techniques in an organized team-approach that is adaptable to be used for small to large sized studies.

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is ...

Immunology and Infection

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), representing the gas-exchange capacity of the lung, is a critical criterion to assess respiratory function. However, observer bias can significantly influence measured values for lung morphological parameters. The protocol that we describe here minimizes variations during measurements of two morphological parameters used for ISA calculation: internal lung volume (ILV) and mean linear intercept (MLI). Using ISA as a morphometric and functional parameter to determine the outcome of alveolar regeneration in both pneumonectomy (PNX) and prosthesis implantation mouse models, we found that the increased ISA following PNX treatment was significantly blocked by implantation of a prosthesis into the thoracic cavity1. The ability to accurately quantify ISA is not only expected to improve the reliability and reproducibility of lung function studies in injured-induced alveolar regeneration models, but also to promote mechanistic discoveries of multiple pulmonary diseases.

Internal lung surface area (ISA) is a critical criterion for assessing lung morphology and physiology in lung diseases and injury-induced alveolar regeneration. We describe here a standardized method that can minimize the measurement bias for ISA in both lung pneumonectomy and prosthesis implantation mouse models.

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), ...

Developmental Biology

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  ...

Videomorphometric Analysis of Hypoxic Pulmonary Vasoconstriction of Intra-pulmonary Arteries Using Murine Precision Cut Lung Slices

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ventilation. Up to now, the underlying mechanisms are not fully understood. The major vascular segment contributing to HPV is the intra-acinar artery. This vessel section is responsible for the blood supply of an individual acinus, which is defined as the portion of lung distal to a terminal bronchiole. Intra-acinar arteries are mostly located in that part of the lung that cannot be selectively reached by a number of commonly used techniques such as measurement of the pulmonary artery pressure in isolated perfused lungs or force recordings from dissected proximal pulmonary artery segments1,2. The analysis of subpleural vessels by real-time confocal laser scanning luminescence microscopy is limited to vessels with up to 50 &#181;m in diameter3.
We provide a technique to study HPV of murine intra-pulmonary arteries in the range of 20-100 &#181;m inner diameters. It is based on the videomorphometric analysis of cross-sectioned arteries in precision cut lung slices (PCLS). This method allows the quantitative measurement of vasoreactivity of small intra-acinar arteries with inner diameter between 20-40 &#181;m which are located at gussets of alveolar septa next to alveolar ducts and of larger pre-acinar arteries with inner diameters between 40-100 &#181;m which run adjacent to bronchi and bronchioles. In contrast to real-time imaging of subpleural vessels in anesthetized and ventilated mice, videomorphometric analysis of PCLS occurs under conditions free of shear stress. In our experimental model both arterial segments exhibit a monophasic HPV when exposed to medium gassed with 1% O2 and the response fades after 30-40 min at hypoxia.

Hypoxic pulmonary vasoconstriction (HPV) is an important physiological phenomenon by which at alveolar hypoxia lung perfusion is matched to ventilation. The major vascular segment contributing to HPV is the intra-acinar artery. Here, we describe our protocol for the analysis of HPV of murine pulmonary vessels with diameters of 20-100 &mu;m.

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ...

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 ...

Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured with independent control over pulmonary arterial flow rate, flow rate waveform, airway pressure and left atrial pressure. Pulmonary vascular resistance is calculated based on multi-point, steady pressure-flow curves; pulmonary vascular impedance is calculated from pulsatile pressure-flow curves obtained at a range of frequencies. As now recognized clinically, impedance is a superior measure of right ventricular afterload than resistance because it includes the effects of vascular compliance, which are not negligible, especially in the pulmonary circulation. Three important metrics of impedance - the zero hertz impedance Z0, the characteristic impedance ZC, and the index of wave reflection RW - provide insight into distal arterial cross-sectional area available for flow, proximal arterial stiffness and the upstream-downstream impedance mismatch, respectively. All results obtained in isolated, ventilated and perfused lungs are independent of sympathetic nervous system tone, volume status and the effects of anesthesia. We have used this technique to quantify the impact of pulmonary emboli and chronic hypoxia on resistance and impedance, and to differentiate between sites of action (i.e., proximal vs. distal) of vasoactive agents and disease using the pressure dependency of ZC. Furthermore, when these techniques are used with the lungs of genetically engineered strains of mice, the effects of molecular-level defects on pulmonary vascular structure and function can be determined.

The following protocol outlines the process of isolating, ventilating and instrumenting mouse lungs to measure steady or pulsatile pulmonary vascular pressure-flow relationships in order to quantify the effects of blood flow, airflow, airway changes and vascular changes on right ventricular afterload.

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured ...

Biology

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 ...

Lung Fixation under Constant Pressure for Evaluation of Emphysema in Mice

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung fixation to produce reliable histological specimens of the lung. Due to the nature of the lung&rsquo;s structural composition, which consists largely of air and tissue, there is a risk that it collapses or deflates during the fixation process. Various lung fixation methods exist, each of which has its own advantages and disadvantages. The lung fixation method presented here utilizes constant pressure to enable optimal tissue evaluation for studies using an emphysematous mouse lung model. The main advantage is that it can fix many lungs with the same condition at one time. Lung specimens are obtained from chronic cigarette smoke-exposed mice. Lung fixation is performed using specialized equipment that enables the production of constant pressure. This constant pressure maintains the lung in a reasonably inflated state. Thus, this method generates a histological specimen of the lung that is suitable to evaluate cigarette smoke-induced mild emphysema.

Presented here is a useful protocol for lung fixation that creates a stable condition for histological evaluate of lung specimens from a mouse model of emphysema. The main advantage of this model is that it can fix many lungs with the same constant pressure without lung collapse or deflation.

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung ...

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 ...