SML is supported by a studentship from the Canadian Institutes of Health Research, MG is supported by a bursary from the Respiratory Health Network of the FRQS (Fonds de recherche du Qu&#233;bec &#8211; Sant&#233;) and YB is a research scholar from FRQS.
AUTHORS&#8217; CONTRIBUTION 
All authors contributed to the conception of the manuscript and/or the video. SML and LD collected the data. SML, LD, YB, DM, DB and AR contributed to data analysis, the generation of figures and manuscript writing. YB, AR, KL and MG were involved in preparing the video script. The play was performed by YB, KL and MG.

The following procedures were approved by the Quebec Heart and Lung Institute Animal Care Committee in accordance with the guidelines of the Canadian Council on Animal Care (CCAC).
1. Preparation
<ol>
	<li>Study
	<ol>
		<li>(Critical) Prior to performing any experiment, obtain the appropriate approvals (e.g., IACUC) and trainings (e.g., animal handling).</li>
		<li>Familiarize yourself with the equipment and the operating software. Read the user manuals and, if needed, create a configuration file to define the number of sites, input signals, analyzers, and parameters. 
		Note: Make sure to select a high sampling rate (2 KHz).</li>
		<li>Configure the analyzers settings for the parameters of interest.
		<ol>
			<li>Select Tuning from the tool bar and then&#160;Analyzers.</li>
			<li>Adjust the flow threshold to a value that correctly separates the breaths (mouse: 0.5 mL/s) and opt for Ti + Te to compute the breathing rate.</li>
			<li>Set a value for the barometric pressure (760 mm Hg) and specify the maximal deviation of inspiratory/expiratory volume for a breath to be considered valid (mouse: 20%).</li>
			<li>Click on Setting in the Calculated parameters field to adjust the limits of acceptance. 
			Note: The following settings were used for the described mouse experiments: Inspiration time, 50 to 170 ms; expiration time, 40 to 180 ms; frequency of breathing, 30 to 450 bpm; specific airway resistance, 0 to 15 cmH2O&#183;s; midexpiratory flow with a precision of at least 3 decimals.</li>
			<li>Once completed, select Apply and Close to exit the dialog window.</li>
		</ol>
		</li>
		<li>From the tool bar menu, go to Tuning again and then Storage to set a desired data storage rate. Click on Apply and Close to exit the window. 
		Note: An average at every 10 s is typically used.</li>
		<li>Create a protocol within the operating software to define a sequence of commands and the desired timing for each one.&#160;An example is displayed in Figure 1.</li>
		<li>If the experiment involves the administration of a substance by aerosol, prepare the appropriate solutions and dilutions according to the concentrations to be tested.</li>
	</ol>
	</li>
	<li>Animal
	<ol>
		<li>Work in a quiet area remote from the housing room. Allow the animals to adjust to the change of environment.</li>
		<li>Weigh the animals and select the appropriate restraint size.</li>
		<li>(Critical) Acclimate the animals to the restrainer and procedures prior to the start of the experiment. Depending on the experimental design, multiple acclimation sessions of increasing duration (e.g., 5-30 min) may be needed. 
		Note:&#160;Animals that do not acclimate should be removed from the study.
		<ol>
			<li>At each acclimation session, insert the animal within the restrainer, proceeding from the back opening; holding the device vertically can be helpful.</li>
			<li>Once the animal is in position, insert the back plunger and gently lock it in place without applying an excessive force.</li>
			<li>(Critical) Visually check that the animal breathes normally. If needed, adjust its position by moving the locking mechanism. Make sure&#160;that the animal&#39;s nares are protruding outside of the nose-cone with its snout resting against the inner walls of the restrainer.</li>
			<li>Detach the back panel of the thoracic chamber, insert the restrainer containing the animal through the rubber opening in the thoracic chamber, and close the chamber.</li>
			<li>Attach the head chamber and provide a bias flow. Use a flow of 0.5 L/min for a mouse.</li>
			<li>Allow the animal to relax for 5 min.</li>
			<li>Once the animal is calm, start the recording of the nasal and thoracic flow signals. Verify on the computer screen that the traces are smooth and that they display a regular breathing pattern; see example in Figure 2. 
			Note: If the protocol involves the aerosol administration of a substance, a saline challenge can be included in the acclimation procedure.</li>
			<li>At the end of each session, remove the animal from the thoracic chamber and the restrainer and return it to its housing cage and room.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Equipment
	<ol>
		<li>On the day of the experiment, start an experimental session and load the appropriate configuration file. 
		Note: Verify that it contains the desired protocol for the experiment.</li>
		<li>Go to Run in the tool bar menu. Enter the experiment and animal&#39;s information. Once done, click on the Run button at the bottom of the window.</li>
		<li>Proceed with the calibration of the system. Calibrate each site and input signal separately.
		<ol>
			<li>Turn on the bias flow generator, connect it to the head chamber via a piece of tubing, and adjust the flow rate.</li>
			<li>Close the top opening of the head chamber with a cap.</li>
			<li>Detach the back panel of thoracic chamber, firmly insert the calibration tool inside the rubber opening between the head and the body chamber to create a hermetic seal. Then close and re-attach the back panel of the thoracic chamber.</li>
			<li>Verify that the side port of the thoracic chamber is capped.</li>
			<li>From the software tool bar menu, go to Tuning and then Calibrate.</li>
			<li>Go to Input 1 (thoracic) and select Calibrate to launch the calibration dialog box for the thoracic flow signal.</li>
			<li>(Critical) Verify that the listed parameters in the calibration dialog window display the appropriate settings, i.e., Physical stress applied low value: 0; Physical stress applied high value: -20 mL/s; Samples: Integrate. Once done, click on Low in the Samples window.</li>
			<li>Verify that the signal generated is constant across the display window and then click on Close.</li>
			<li>Connect a 20 mL-syringe through the side port of the thoracic chamber using a plastic connector and a piece of tubing.</li>
			<li>(Critical) Select High in the Samples window and immediately inject 20 mL of air into the chamber over a 2 s period at a flow rate as constant as possible.</li>
			<li>Verify that the signal generated appears completely inside the display window. Use the arrow icon to verify whether the signal is centered and symmetric around the zero line. Then, click on Close. Remove any offset from zero by clicking on Remove AC offset in the Samples window 
			Note: The high value calibration can be redone, if needed.</li>
			<li>Verify that the resulting Scaled Input Range is&#160;within the recommended range (mouse: &#177;280 to &#177;420 mL/s). (Critical) Repeat the calibration steps if the values are outside the acceptable range.</li>
			<li>Calibrate the head chamber in a similar manner as the thoracic chamber (step 1.3.3.6). This time, select Input 2 (nasal). 
			Note: (Important) The value for Physical stress applied high value must be set to +20 mL/s. This will change the polarity of the flow in the front chamber in relation to the rear chamber. Hence, when the animal is breathing, the two flow signals will almost be in phase, aside from the delay used to calculate sRaw.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Lung Function Measurements
<ol>
	<li>Weigh the animals and note their body weights.</li>
	<li>Insert the animal within the restrainer and place it inside the thoracic plethysmograph chamber (Steps 1.2.3.1 to 1.2.3.5).</li>
	<li>Allow the animal to relax for at least 5 min.</li>
	<li>Start the protocol of commands by selecting the first step and then, click on Execute.</li>
	<li>Check on the computer screen that the animal&#39;s breathing signals are regular and smooth (Figure 2). The software automatically displays the calculated parameters on a breath-by-breath basis.&#160;Verify that the animal&#39;s parameters are stable.</li>
	<li>Record the breathing pattern under baseline conditions for up to 10 minutes.</li>
	<li>For protocols involving the administration of a test substance by aerosol, do as follows:
	<ol>
		<li>Adjust the nebulizer on-time and duty cycle, as required. 
		Note: In the examples demonstrated in this article, the nebulizer was operated at a 5% duty cycle for 10 s.</li>
		<li>Perform a vehicle challenge (e.g., saline) and record the response.</li>
		<li>If needed, expose the animal to increasing concentrations of the test substance by changing the concentration in the nebulizer in escalating steps (e.g., doubling concentrations). Record the response after each administration.</li>
		<li>At the end of the experimental session, if not done automatically, stop the recording and return the animal to its housing cage and room.</li>
	</ol>
	</li>
	<li>If needed, select Run from the tool bar menu to run another experimental session.
	<ol>
		<li>Between sessions, clean the plethysmograph chambers and rinse the nebulizer with water. 
		Note: The use of alcohol may cause irreversible damage to the plethysmograph.</li>
	</ol>
	</li>
	<li>If the study involves repeated evaluations over time, repeat the whole measurement sequence at each chosen timepoint. 
	Note: It is wisely recommended to complement the study with some precise measurements of respiratory mechanics and/or lung volumes1,15.</li>
</ol>
3. Data Analysis
Note: The software automatically saves the experimental file and exports the recorded parameters when the experimental session is closed.
<ol>
	<li>Calculate a baseline average for the parameters of interest for each animal and experimental group. 
	Note: Table 1 lists a number of typical parameters, categorized based on the type of information they provide.</li>
	<li>When relevant, evaluate the effect of the aerosolized substance studied on the parameters of interest at each concentration using a specific point (e.g., maximal or minimal value), an average, or the full time-course; a normalization to baseline can also be considered.</li>
	<li>Report the results using the group means and errors in a table or graphic format. Analyze the results statistically. 
	Note:&#160;In the present study, two-way ANOVAs with repeated measures were used to assess the effect of methacholine, the allergen &#8211; house-dust mite (HDM) &#8211;&#160; and their interaction on different DCP readouts (sRaw and EF50), as well as on different FOT readouts (RN, G and H), in both strains of mice. Sidak&#8217;s multiple comparison tests were then used to determine the concentrations of methacholine at which the allergic mice differ from the control mice. The same tests were used to assess the effect of days, HDM and their interaction at baseline (i.e., before methacholine) on readouts of airway obstruction (sRaw and EF50) and of ventilation pattern (BF, TV, MV and EIP). Pearson&#8217;s correlations were used to evaluate the correlations between sRaw obtained with the DCP and RN obtained with the FOT. All statistical analyses and graphs were performed using alternative standard statistical software (e.g., GraphPad Prism). p &#8804; 0.05 was considered sufficient to reject the null hypothesis.</li>
</ol>

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiologic+assessment+of+allergic+rhinitis+in+mice:+role+of+the+high-affinity+IgE+receptor+(FcepsilonRI).">Physiologic assessment of allergic rhinitis in mice: role of the high-affinity IgE receptor (FcepsilonRI).</a> J Allergy Clin Immunol. 116 (5), 1020-1027 (2005).</li></ol>

DB and AR are employed by SCIREQ Scientific Respiratory Equipment Inc, a commercial entity involved in topics related to the content of this article. DB also owns stock. SCIREQ Inc. is an emka TECHNOLOGIES company.

The ability to measure lung function in conscious animals is clearly warranted in respiratory research. In general, the DCP is an interesting approach to evaluate the ventilation function of the respiratory system in conscious and spontaneously breathing animals26. More specifically, the DCP, or its head-out variant, often strikes a right balance between the quality of the information provided and the desired level of invasiveness3 (Table 2). The technique can be adapted to various species (e.g., mouse, rat, guinea pig) or animal sizes and can be used in many research applications. It is particularly useful to assess numerous animals at once in a parallel study design, to monitor the respiratory function in repeated manner, and to capture the kinetics of a response over time. In addition, the technique is straightforward and can be learned in a relatively timely manner. In the present paper, a protocol employing DCP measurements in mice was used as an example to describe the hands-on aspects of this restrained plethysmography technique as well as to discuss the critical steps and related outcomes.
When working with conscious animals, it is essential to control the conditions of the surrounding environment (e.g., quiet room with a limited number of people or activity) in order to generate reproducible results. Since the restrainers come in various dimensions, it is important to start with the appropriate size so that the breathing movements are unperturbed. It is also helpful and often required to acclimate the animals to the experimental set-up and procedures, as it is well established in mice that restraining affects the breathing frequency12. Depending on the experimental design or conditions, multiple sessions of incremental durations may be needed. Finally, allowing time at the start of an experiment for the animals to adjust to the room change and necessary handling is a simple consideration that proved effective to ensure that the breathing pattern is consistently regular and relaxed at baseline. Working under conditions where the animals are comfortable, well-adapted, and calm will also be beneficial in terms of result variability and quality. It also limits any stress-induced release of catecholamine, which may increase airway caliber and attenuate an induced bronchoconstriction.
It is important to understand that there is a need to separate&#160;as hermetically as possible&#160;the nasal and thoracic flows. Depending on the system or species studied, the sealing mechanism can vary in shape as well as in efficacy. In the DCP we described herein, the seal is created between the animal&#39;s snout and the restraining device. When assessing the respiratory function by DCP, it is also essential to provide a sufficient and continuous bias flow, as a decrease in the level of oxygen available to the animal will result in significant effects. Taking into account the well-being of the animal in the restrainer limits the propensity for air leaks created by agitation and thereby maximizes the quality of the data. Contrastingly, a breaking in the seal will result in either rejected datasets or an underestimation of some parameters.
In addition to enable the separate recording of the nasal flow signal, the head chamber is typically used to expose the animal to aerosolized substances. As illustrated in this article, this can be utilized to perform a bronchoprovocation test to demonstrate differing degrees of responsiveness. In such experiments, adjusting the range of tested concentrations may be needed depending on the species, strain, or sex of the animals studied. As previously demonstrated8,9,10,27, the present results show that the methacholine-induced changes in sRaw correlated well with invasive FOT measurements of airway resistance. The results also demonstrate that the DCP technique is not as sensitive as the FOT for&#160;its ability to detect respiratory dysfunction and to identify an altered response localized within the lower compartments of the lung (lung tissue and/or small peripheral airways). Since the animal&#39;s airways are intact, the presence of the upper airways, which accounts for the largest part of the total respiratory resistance to airflow28, can affect the aerosol distribution and deposition in addition to dampen the contribution from the lower airways to a measurement. Table 3 summarizes other differences between the DCP technique and the FOT. Finally, while it would be theoretically possible to estimate the animal&#39;s total airway resistance (including upper airways) from a measurement of sRaw, it is generally recommended to complement the DCP evaluation with an invasive measurement technique such as the FOT29 to obtain direct measurements of detailed respiratory mechanics. Depending on the objectives of the study, measurements of upper airway resistance can also be considered30,31,32.
Conclusion&#160; 
Owing to its limited degree of invasiveness, the DCP is a technique that can fulfill an important need in respiratory research. It is able to provide accurate readouts of ventilation pattern in conscious animals concurrently with some undisputed indexes of airflow obstruction. The information obtained also truly complements that from more invasive approaches.

The increasing use of small animals to model human respiratory diseases has urged the development of techniques to quantitatively assess the functions of the respiratory system in those animals. Currently, the forced oscillation technique (FOT) is recognized as the most precise approach to assess respiratory mechanics in small animals1,2. However, as stated by the phenotyping uncertainty principle, what is gained in measurement precision with the FOT is traded off against a loss in noninvasiveness3. Indeed, FOT measurements are acquired under highly controlled experimental conditions that necessitate anesthesia, tracheotomy or oral intubation, as well as mechanical ventilation; a scenario far from real-life.
In situations where the experimental requirements proscribe the use of anesthetic agents or call for little or no deviation from the animal&#39;s natural physiological state, double-chamber plethysmography (DCP) can be considered. As its name indicates, a DCP setup consists of two connected rigid chambers built to isolate as hermetically as possible the animal&#39;s head (or nose), in the front chamber, from its thorax, in the rear chamber. Within the setup, the animal is conscious and breathes spontaneously while being restrained. Because the walls of the chambers cannot expand or retract, the motion of air in-and-out of the animal generates a corresponding but opposite waveform inside the rear&#160;chamber, as a result of the compression/decompression of surrounding air. The waveform due to nasal flow in the front chamber and the one related to the thoracic motion in the rear chamber can thus be separated and captured simultaneously. Depending on the design of the DCP setup, these waveforms can be acquired using&#160;a set of either pressure transducers or pneumotachographs to respectively record the changes in chamber pressure or airflow in-and-out of the chambers as a function of time. The latter approach is more common nowadays.
While the animal&#39;s breathing frequency can be accurately determined by any kind of plethysmography techniques, the situation is not the same for the determination of tidal volume and its related ventilation parameters (e.g., minute ventilation, expiratory volume, etc.). As opposed to the whole-body plethysmograph (WBP) technique, where the animal&#39;s tidal volume is estimated from the box signal4,5, the DCP technique provides accurate assessments of tidal volume. This is related to the direct acquisition of the animal&#39;s thoracic movement in the rear chamber, which are proportional to the changes in lung volume during breathing.
In addition to these accurate ventilatory parameters (e.g., tidal volume, breathing frequency, and minute ventilation), some disturbances in the shape of the respiratory cycle can also be used to investigate neuronal aspects that govern the respiratory drive or respiratory reflexes. A specific example of that application would be the evaluation of the irritation potential of inhaled substances on the upper airway sensory neurons6. Here, the duration of a pause at the onset of expiration is determined using a parameter called end-inspiratory pause (EIP), also referred to as duration of braking6. The prolongation of this pause by an irritant substance is associated with the closure of the animal&#39;s glottis, causing a measurable period of braking in the first part of the expiration6,7.
Another important advantage of the DCP is that it provides two validated and undisputed parameters that are sensitive to airflow obstruction. One is called the flow at mid-tidal expiratory volume and is abbreviated EF508,9,10. It is the airflow at midway volume of each tidal breath during expiration. EF50 is extracted from the thoracic flow trace and can thus be measured without the front chamber (i.e., in a head-out configuration). The other one is called specific airway resistance and is abbreviated sRaw11,12,13. The determination of sRaw requires the simultaneous recording of the animal&#39;s nasal and thoracic flows as it is computed from the time delay between these separate respiratory traces at the point of zero flow at the end of inspiration. The rationale that describes the basis by which this delay relates to sRaw was expatiated previously11. Put simply, the changes in lung volume precede the air movement since a pressure gradient needs to develop in order to drive airflow. In a healthy animal breathing quietly, this delay is typically very small. However, the pressure gradient that is required to accommodate a given flow (e.g., a flow sufficient to provide adequate ventilation) is influenced by the degree of airway resistance. During bronchoconstriction, for example, the gradient of pressure needed to accommodate a given flow is greater, which implies that the animal has to work harder for breathing. A greater gradient of pressure in the animal&#39;s thorax also implies that a greater portion of the flow in-and-out of the rear chamber is due to decompression/compression of air within the thorax, which is the portion of the total thoracic expansion/retraction that is out of phase with the nasal flow. The increased resistance due to bronchoconstriction will thus increase the delay between the rear and the front chambers and thereby increases sRaw. The gradient of pressure that drives airflow in-and-out of the lung is also influenced by the initial thoracic gas volume (TGV). At a greater TGV for example, the expansion/retraction of the thorax needed to generate a given gradient of pressure is greater (simply because the volume displacement that is required to generate a given gradient of pressure is greater), which also implies that the animal has to work harder for breathing. Again, these extra thoracic displacements are the ones required to decompress/compress air in the thorax and are thus out of phase with the nasal flow. So, an increased TGV will also increase the delay between the chambers and thereby increases sRaw. As can be seen, both bronchoconstriction and increased TGV result in a more important effort to draw air in-and-out of the lung. This is, in essence, the physiological meaning of sRaw. It represents the work required for breathing5,14.
It is thus important to understand that two distinct factors influence sRaw: airway resistance and TGV. In fact, sRaw can be expressed as the product of airway resistance and TGV11. Conscious animals can modify their TGV at will, as to adapt their ventilation to a given environment. Under such conditions, where the animal&#39;s natural physiological state is unaltered, it is thus impossible to discern whether a change in sRaw stems from an alteration in airway resistance, from a change in TGV, or from a mix of the two. Hence, it is recommended to complement the DCP evaluation with more invasive measurements of respiratory mechanics and/or lung volumes, such as those provided by the FOT1,15.
To date, the DCP has been used in various research applications. The technique can be used with or without the head chamber to quantitatively and accurately evaluate the effect of various substances, such as pharmaceutical agents, allergens, irritants or other mediators, on respiratory function in conscious small animals16,17,18. The front chamber can also be used as an exposing chamber to aerosolized substances or varying gas concentrations (hypoxia, hypercapnia, etc.)19. Conveniently, it allows one to concomitantly measure the acute effects of these exposures. In fact, one of the common uses of the DCP is to assess the degree of responsiveness to aerosolized methacholine in different models of respiratory diseases20,21,22,23,24,25.
Although the DCP technique is seemingly straightforward, some practical challenges could potentially discourage inexperienced users or impair the accuracy and reproducibility of the results. The present paper provides a detailed description of the recommended procedures to successfully record respiratory function by DCP in conscious, restrained, spontaneously breathing mice. The description is specific to the stated equipment (please refer to the Table of Materials). The utility and value of the DCP is also demonstrated in a common model of pulmonary allergic inflammation in two strains of mice tested at baseline and in response to aerosolized methacholine.

<table><tbody><tr><td>Acetyl-&amp;beta;-methylcholine chloride&amp;nbsp;</td><td>Sigma-Aldrich</td><td>A-2251</td><td>Methacholine</td></tr><tr><td>Phosphate buffered saline</td><td>Multicell</td><td>311-506-CL</td><td>PBS 10X</td></tr><tr><td>House dust mite extract</td><td>GREER</td><td>290902</td><td>HDM&amp;nbsp;</td></tr><tr><td>DCP complete system</td><td>&amp;nbsp;SCIREQ Inc. /emka TECHNOLOGIES</td><td></td><td></td></tr><tr><td>iox software</td><td>&amp;nbsp;SCIREQ Inc. /emka TECHNOLOGIES</td><td></td><td></td></tr><tr><td>Aerogen Aeroneb nebulizer</td><td>&amp;nbsp;SCIREQ Inc. /emka TECHNOLOGIES</td><td></td><td></td></tr><tr><td>flexiVent FX complete system</td><td>&amp;nbsp;SCIREQ Inc. /emka TECHNOLOGIES</td><td></td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography at JoVE.com

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

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 ...

assessment-respiratory-function-conscious-mice-double-chamber

Research

JoVE Journal

Medicine

42.8K Views.  Université Laval. Het doel van dit artikel is bedoeld als een gedetailleerde beschrijving van de aanbevolen procedures voor het evalueren van de ademhalingsfunctie in bewuste muizen door dubbele-kamer plethysmography.

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

Hyperinsulinemic-euglycemic Clamps in Conscious, Unrestrained Mice

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold standard" method for assessing insulin action in vivo. During an insulin clamp, hyperinsulinemia is achieved by a constant insulin infusion. Euglycemia is maintained via a concomitant glucose infusion at a variable rate. This variable glucose infusion rate (GIR) is determined by measuring blood glucose at brief intervals throughout the experiment and adjusting the GIR accordingly. The GIR is indicative of whole-body insulin action, as mice with enhanced insulin action require a greater GIR. The insulin clamp can incorporate administration of isotopic 2[14C]deoxyglucose to assess tissue-specific glucose uptake and [3-3H]glucose to assess the ability of insulin to suppress the rate of endogenous glucose appearance (endoRa), a marker of hepatic glucose production, and to stimulate the rate of whole-body glucose disappearance (Rd).
The miniaturization of the insulin clamp for use in genetic mouse models of metabolic disease has led to significant advances in diabetes research. Methods for performing insulin clamps vary between laboratories. It is important to note that the manner in which an insulin clamp is performed can significantly affect the results obtained. We have published a comprehensive assessment of different approaches to performing insulin clamps in conscious mice1 as well as an evaluation of the metabolic response of four commonly used inbred mouse strains using various clamp techniques2. Here we present a protocol for performing insulin clamps on conscious, unrestrained mice developed by the Vanderbilt Mouse Metabolic Phenotyping Center (MMPC; URL: www.mc.vanderbilt.edu/mmpc). This includes a description of the method for implanting catheters used during the insulin clamp. The protocol employed by the Vanderbilt MMPC utilizes a unique two-catheter system3. One catheter is inserted into the jugular vein for infusions. A second catheter is inserted into the carotid artery, which allows for blood sampling without the need to restrain or handle the mouse. This technique provides a significant advantage to the most common method for obtaining blood samples during insulin clamps which is to sample from the severed tip of the tail. Unlike this latter method, sampling from an arterial catheter is not stressful to the mouse1. We also describe methods for using isotopic tracer infusions to assess tissue-specific insulin action. We also provide guidelines for the appropriate presentation of results obtained from insulin clamps.

The hyperinsulinemic-euglycemic clamp, or insulin clamp, is the gold standard for assessing insulin action in vivo. A method for performing insulin clamps in mice is described. This includes a method for arterial catheterization that permits experiments to be performed in conscious, unrestrained mice with minimal stress.

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold ...

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

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment (RMCA) in Individuals with Chronic Spinal Cord Injury

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination is disrupted by spinal cord injury (SCI), control of respiratory muscles innervated below the injury level is compromised1,2 leading to respiratory muscle dysfunction and pulmonary complications. These conditions are among the leading causes of death in patients with SCI3. Standard pulmonary function tests that assess respiratory motor function include spirometrical and maximum airway pressure outcomes: Forced Vital Capacity (FVC), Forced Expiratory Volume in one second (FEV1), Maximal Inspiratory Pressure (PImax) and Maximal Expiratory Pressure (PEmax)4,5. These values provide indirect measurements of respiratory muscle performance6. In clinical practice and research, a surface electromyography (sEMG) recorded from respiratory muscles can be used to assess respiratory motor function and help to diagnose neuromuscular pathology. However, variability in the sEMG amplitude inhibits efforts to develop objective and direct measures of respiratory motor function6. Based on a multi-muscle sEMG approach to characterize motor control of limb muscles7, known as the voluntary response index (VRI)8, we developed an analytical tool to characterize respiratory motor control directly from sEMG data recorded from multiple respiratory muscles during the voluntary respiratory tasks. We have termed this the Respiratory Motor Control Assessment (RMCA)9. This vector analysis method quantifies the amount and distribution of activity across muscles and presents it in the form of an index that relates the degree to which sEMG output within a test-subject resembles that from a group of healthy (non-injured) controls. The resulting index value has been shown to have high face validity, sensitivity and specificity9-11. We showed previously9 that the RMCA outcomes significantly correlate with levels of SCI and pulmonary function measures. We are presenting here the method to quantitatively compare post-spinal cord injury respiratory multi-muscle activation patterns to those of healthy individuals.

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment RMCA in Individuals with Chronic Spinal Cord Injury

The purpose of this publication is to present our original work on a multi-muscle surface electromyographic approach to quantitatively characterize respiratory muscle activation patterns in individuals with chronic spinal cord injury using vector-based analysis.

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination ...

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

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese ...

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 ...

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 ...

Biology

Simplified Whole Body Plethysmography to Characterize Lung Function During Respiratory Melioidosis

Surrogate animal models of disease are subject to the 3Rs of Responsible Research. There is a frequent revisiting of refinements to animal models to ensure that both animal welfare and scientific insights advance with the availability of new technologies. This article demonstrates the use of Simplified Whole Body Plethysmography (sWBP) to non-invasively study respiratory failure in a model of lethal respiratory melioidosis. sWBP has the sensitivity to detect breathing in mice through the entirety of the course of the disease, allowing for the moribund-associated symptoms (bradypnea and hypopnea) to be measured and potentially used to develop humane endpoint criteria.
Some of the benefits of sWBP in the context of respiratory disease are that host breath monitoring comes the closest of any physiologic measurement at assessing dysfunction of the primary infected tissue, namely, the lung. In addition to biological significance, the use of sWBP is rapid and non-invasive, minimizing stress in research animals. This work demonstrates the use of in-house sWBP apparatus to monitor disease throughout the course of respiratory failure in the murine model of respiratory melioidosis.

This protocol presents the construction and use of a Simplified Whole Body Plethysmography apparatus to monitor bacterial respiratory disease progression non-invasively.

Surrogate animal models of disease are subject to the 3Rs of Responsible Research. There is a frequent revisiting of refinements to animal models 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 ...

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

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