Name: measuring respiratory function in mice using unrestrained whole-body plethysmography
Uploaded: 2014-08-12T14:00:00.000+00:00
Duration: 8 min 51 s
Description: Watch this Scientific Journal Video about Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography at JoVE.com

We would like to thank Prof David Walker for his technical advice and provision of equipment in the development of this technique. This work is supported by the Victorian Government&#8217;s Operational Infrastructure Support Program. This work was partly supported by the Victorian Government&#8217;s Operational Infrastructure Support Program.

NOTE: The following experimental procedure is approved by the Animal Ethics Committee at Monash University and conducted in accordance with the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes (2006). Adult female C57BL/6 mice used to generate the representative results were obtained from the Monash Animal Services. The mice were housed in a specific pathogen free, temperature and humidity controlled room with a 12 hr light-dark cycle. These mice had free access to food and water.
1. Initial Setup
<ol>
	<li>Connect the laptop/desktop to the data acquisition machine for recording via a USB cable.</li>
	<li>Connect the Bridge Amplifier from &#8216;output 1&#8217; to &#8216;input 1&#8217; of the data acquisition machine via a BNC cable.</li>
	<li>Insert the pressure transducer into &#8216;channel 1&#8217; of the Octal Bridge Amp. Turn the data acquisition machine on and open the analysis software. The software should automatically detect the equipment setup (see materials and equipment table).</li>
	<li>Open Channel settings found in the setup tool bar of the software. Change the number of channels being recorded to 1.</li>
	<li>Set up the barometer to measure room pressure and the water column apparatus to calibrate the Bridge Amplifier. The water column apparatus includes two 5 ml serological syringe pipettes connected by plastic tubing.</li>
	<li>Fill the columns with water and ensure the water levels are balanced with a ruler. Connect one piece of plastic tubing to the top of each pipette. Figure 2 shows the water column set up.</li>
</ol>
2. Bridge Amplifier Calibration
Note: To calibrate the bridge amplifier an injection of air into the water column is required to create a 1 cm H2O deflection. This will occur under a single set of conditions and is dependent upon the user&#39;s apparatus. For clarification these steps demonstrate how this laboratory would perform the calibration.
<ol>
	<li>Withdraw a 1 ml syringe to 300 &#956;l; attach the syringe to the stopcock at the end of the tubing on the right hand side of the water column. NOTE: Ensure that the stopcock is open to the syringe and the water column, and closed to the room air. If the water levels are not balanced at this point, turn the stopcock so that it is open to room air and the water column, this will rebalance the water. The tubing on the left hand side of the water column should be connected to the pressure transducer to measure the change in pressure induced by plunging the syringe.</li>
	<li>Attach the tubing from the water column on the left hand side to the connector on the pressure transducer (top ring of the transducer).</li>
	<li>Select the scroll down menu found next to channel 1 at the main screen on the right hand side of the software and select &#8220;Bridge amp&#8221; (see materials and equipment table).</li>
	<li>Enter the settings to 5 mV, 20 Hz low pass, tick the &#8216;invert&#8217; box and click &#8216;zero&#8217;. Click &#8216;zero&#8217; to set the trace at ~0 mV. Reduce the window size to 4:1 for easier viewing.</li>
	<li>With everything set up, depress the 1 ml syringe, leaving it for 3 sec. This will show a sudden spike on the software because the pressure has changed. When the 300&#160;&#956;l is depressed the pressure will move the water in the water column by 1 cm. This known value will help calibrate the Bridge Amplifier. 
	NOTE: The pressure increase in the chamber due to the 300&#160;&#956;l depression corresponds to the PK value used for later calculations.</li>
	<li>Select &#8216;input units&#8217; found on the bottom left hand corner of the Bridge Amp window.</li>
	<li>Highlight the &#8220;background trace&#8221; prior to the spike otherwise known as the &#8216;Zero Region&#8217;.
	<ol>
		<li>Click the arrow next to &#8216;point 1&#8217; and this will produce the background signal within the range&#160;of -0.002 mV-0.002 mV (the value will never be exactly at 0 mV).</li>
		<li>Type &#8216;0&#8217; in the window adjacent to the background signal window.</li>
	</ol>
	</li>
	<li>Highlight the &#8220; increased pressure region of graph&#8221; from when the syringe is depressed. Click the arrow next to point 2 and the value should be in range of 0.9-1.2 mV.
	<ol>
		<li>Type &#8216;1&#8217; in the window next to the &#8220;increased pressure&#8221; window. For a visual clarification on steps 2.7 and 2.8 refer to Figure 3. Values found outside the ranges specified may indicate damage to the Octal Bridge Amp.</li>
	</ol>
	</li>
	<li>Go to &#8216;define units&#8217; found at the top right corner of the window and select &#8220;cmH2O&#8221;. If this option is not available, it may be entered manually. Click ok.</li>
	<li>Return to the &#8216;Bridge Amp&#8217; menu (refer to 2.1). Select 1 mV and set amplifier to &#8216;zero&#8217;. This will complete calibration and the water column can safely be removed.</li>
</ol>
3. Recording Lung Function
<ol>
	<li>Weigh the mouse (g). NOTE: One week prior to physiological assessment introduce the mouse to the plethysmography chamber environment. This will aid in acclimatization and reduce stress when carrying out this procedure at a later date. For an overall schematic demonstrating the UWBP setup, please refer to Figure 4.</li>
	<li>Measure the body temperature with a rectal thermometer. Lubricate the thermometer with petroleum jelly prior to insertion. Record the temperature reading and clean the lubricant off with 80% (v/v) ethanol. If using very small animals such as neonatal mouse pups, the mean body temperature value may be determined with an infra-red thermometer instead.</li>
	<li>Place the temperature/relative humidity probe on the one-hole end of the plethysmography chamber. Record the temperature, humidity and barometric pressure inside the plethysmography chamber prior to placing the mouse within.</li>
	<li>Place the mouse in the plethysmography chamber, cover the open end slightly. This allows the mouse to acclimatize. Close the chamber.</li>
	<li>With the temperature/humidity probe inserted in the side of the plethysmography chamber with one hole, now insert the transducer and syringe in the other side with the two holes.</li>
	<li>Press &#8216;Start&#8217; on the software program and record for approximately 15-45 sec. Record 5-10 sec of data where the animal is not moving. Movement will alter the animal&#8217;s basal respiratory physiology and provide poor results. Respiration should oscillate in a linear path on the program. These are useable data. Note: Urination or defecation can lead to an increase in the temperature and humidity inside the plethysmography chamber. This will obscure results during analysis. In the event of urination or defecation, stop recording immediately and clean the plethysmography chamber with 80% (v/v) ethanol. Refer to Figure 6 for a visual representation of suboptimal results, where data should be rejected.</li>
	<li>After recording for 45 sec, press &#8216;Stop&#8217; on the software (see materials and equipment table) program. Remove the mouse from the plethysmography chamber and immediately record the chamber temperature and humidity. Do not continually record for more than 45 sec as this may stress the animal.</li>
	<li>Return the mouse to its cage, spray and wipe the chamber with 80% (v/v) ethanol.</li>
	<li>Allow the chamber to dry and return to baseline temperature and humidity before proceeding on to the next mouse. Repeat steps 3.1 to 3.9 for subsequent animals. Note: If multiple animals are being studied, ensure that the chamber temperature and humidity return to near baseline values before each new animal is put in the chamber.</li>
</ol>
4. Plethysmography Analysis
Note: To calculate respiratory parameters such as tidal volume (VT) and minute volume the following variables need to be measured: respiration rate (breaths/min), total breathing cycle time (sec), inspiration/expiration time (Ti/ Te, seconds) and change in pressure due to each tidal volume (PT). Figure 1 illustrates the variables that can be measured from a trace. The following steps use a software (see materials and equipment table) to measure these variables. When analyzing, avoid regions of the trace containing sniffing or movement. For reproducible results, at least 5 seconds of good breathing trace is required. For an example of different breathing traces refer to Figure 5 and 6.
<ol>
	<li>Open the screen to full screen, set view to 1:1 and select 5 sec of usable data. A representative snapshot of this is shown in Figure 5.</li>
	<li>Open the mini data pad window found at the top of the program in the DataPad tab. Select channel 1 and select &#8216;cycle measurements&#8217; in the left hand column and &#8216;average cyclic height&#8217; in the right hand column.
	<ol>
		<li>Select &#8216;Option&#8217; and set the scale for minimum peak detection to 1 (msec). This will allow detection of every peak value and becomes extremely important when using small animals that produce small oscillations.</li>
		<li>Click &#8216;OK&#8217;. This will present &#8216;Pressure deflection due to each tidal volume&#8217; (PT) measurement.</li>
	</ol>
	</li>
	<li>In the mini data pad, select &#8216;cycle measurements&#8217; followed by &#8216;event count&#8217; and click &#8216;OK&#8217;. This will present the &#8216;frequency&#8217; (f) measurement.
	<ol>
		<li>Frequency needs to be converted to breaths/min. This is done by multiplying the value by 60 sec and dividing the answer by the overall time of recording (min).</li>
	</ol>
	</li>
	<li>In the mini data pad, select &#8216;cycle measurements&#8217; followed by &#8216;period&#8217; and click &#8216;Ok&#8217;. This will present the &#8216;total breathing cycle time&#8217; (Ttot, sec) measurement.</li>
	<li>The next steps are used to create a macroinstruction to generate peak inspiration and expiration time values. Ensure the cursor is directly over the maximum of the peak/trough and add a comment on 9 sequential peaks and troughs. Begin with the peak of the oscillation as shown in Figure 5.</li>
	<li>Subsequently, select window: data pad and column 1. In the window that appears click &#8216;selection information&#8217; in the left hand column, &#8216;duration&#8217; in right hand column and click &#8216;OK&#8217;.</li>
	<li>Select macro found at the top of program and then start recording. Now select commands: &#8216;Find&#8217;, &#8216;Go&#8217;, &#8216;Start of File&#8217; and click &#8216;Find&#8217;.</li>
	<li>Select commands: &#8216;Find&#8217; and &#8216;Find comments&#8217;. Type the same phrase typed for the comment box in the &#8216;containing&#8217; box provided. Choose the &#8216;Select to Previous Point&#8217; tab and &#8216;Find&#8217;.</li>
	<li>Select commands: &#8216;Add to data pad&#8217;. Next, select macro: macro commands and begin repeat. The repeat count window that appears should be set at 9.</li>
	<li>Select command: &#8216;Find next&#8217;. Select command: &#8216;Add to data pad&#8217;. Finally select macro commands and end repeat.
	<ol>
		<li>Now select the macro and stop the recording. Save and name the macro after the animal number. NOTE: Setting up the macro for each animal allows the macro to be used for longitudinal studies and saves time.</li>
	</ol>
	</li>
	<li>The macro can now be run to obtain the Inspiration (Ti) and Expiration (Te) time between each comment. The data will appear under channel 1 of the datapad. Expiration and inspiration occurs consecutively and the data will appear in this order.
	<ol>
		<li>The data need to be manually split into inspiration and expiration values. Average the four data values of each parameter to obtain the mean Ti and Te.</li>
	</ol>
	</li>
	<li>Once the primary values have been derived the tidal volume (VT, ml) can be calculated. To obtain the tidal volume the equation of Drorbaugh and Fenn8 is used: 
	VT (ml)= (PT / PK) x (VK) x ((TCORE(PB - PC)) / (TCORE(PB - PC) - TC(PB - PCORE))) 
	 
	Where 
	VT: Tidal Volume 
	Pk: Pressure deflection due to each injection of 1 ml (Refer to step 2.5) 
	Tcore: Core temperature of each animal 
	PC: Water vapor pressure at chamber temperature X Relative humidity in chamber 
	TC: temperature in animal chamber 
	Pcore: Pressure at body temperature (water vapor pressure at body temperature x 1.0) 
	Pt: Pressure deflection due to each tidal volume 
	Vk: Volume injection for calibration 
	&#160;PB: Barometric pressure 
	&#160;</li>
	<li>Once the tidal volume has been calculated the following parameters can also be determined:
	<ul>
		<li>Minute Volume (ml/min) = VT x f</li>
		<li>Minute Volume (ml/min/kg) = (VT x f) / Body weight (kg)</li>
		<li>VT (ml/kg) = VT (ml) / Body weight (kg)</li>
		<li>Inspiratory Duty Cycle (%) = Ti / Ttot</li>
		<li>Inspiratory Flow Rate (ml/sec) = VT / Ti</li>
		<li>Ratio of inspiration time to expiration time = Ti / Te</li>
		<li>Total cycle time (sec) = Inspiration time (sec) + Expiration time (sec)</li>
	</ul>
	</li>
</ol>

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The authors declare that they have no competing interests. The authors have no conflicts to disclose.

The technique described here is a non-invasive method for assessment of respiratory parameters of unrestrained and unanesthetized mice. The strengths of this protocol include its simplicity and precision to measure lung function longitudinally with minimal artefacts. There are, however, some limitations and critical steps to be noted about the procedure. Firstly and most importantly, the mouse must stay calm within the chamber for a minimum of five seconds. Added stress will disrupt the breathing pattern of the mouse and...

Lung dysfunction is one of the leading causes of morbidity and mortality in the world. The condition is characterized by inadequate oxygen exchange, synonymous with coughing, chest pains and dyspnea. Respiratory disease accounts for ~10% of mortality worldwide1. According to the World Health Organization, mortality rates are set to rise due to persistent smoking, pollution &#38; occupational irritants. UWBP is a useful addition for studying lung physiology, which strongly compliments traditional biochemical and histological analyses2. Other procedures used for lung assessment do not provide the same advantages as UWBP. Invasive plethysmography is a commonly used technique that requires the animal to be anesthetised3,4 and thus, resulting respiratory measurements are not necessarily reflective of a natural state. Further, the requirement for mechanical ventilation and chemical challenges preclude future measurements3,4. Another method of collecting respiratory data is by forced oscillation, which is more sensitive to finer changes in respiratory parameters compared to UWBP5. Forced oscillation is, however, an invasive technique and requires animal termination for data collection5-7.
UWBP involves placing an animal inside a specialized chamber. During inspiration, the tidal air is warmed and humidified within the lungs increasing water vapor pressure and causes thermal expansion of gas8. This effect causes a net change in air volume creating an increase in pressure within the plethysmograph chamber8. The opposite occurs during expiration creating a respiratory waveform from the animal. Waveform analysis is then used to measure from the respiratory trace: respiration rate (breaths/min), total breathing cycle time (sec), inspiration/expiration time (Ti/ Te, sec) and changes in pressure due to each tidal volume (PT). Figure 1 illustrates each measurements origin from a respiratory trace. These measurements are simple to calculate and multiple respiratory parameters may be derived from these measurements. These parameters include: Tidal volume (the volume of air moved between normal inhalation and exhalation), minute volume (volume of gas inhaled from the lungs per minute), inspiratory duty cycle (the percentage of inspiration time to the total breathing cycle duration) and inspiratory flow rate (the amount of air inspired in a given time).
UWBP provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models and can be used for measuring the progression of respiratory disease and lung function6,9. Contrary to other plethysmography techniques, UWBP avoids the use of anesthesia, restraints and invasive manipulations that produce artefacts and experimental variability6,9. Anesthesia can suppress respiration, alter heart rate and can be challenging to regulate10. Restraints induce an increase in respiration due to additional stress via corticosterone and epinephrine release11,13. The key feature of UWBP is repeated physiological assessment making it amenable to longitudinal studies. UWBP is strongly recommended for the longitudinal assessment of lung physiology and offers a valuable skill for future respiratory drug assessment.
Bleomycin, ovalbumin, and hypoxia have been utilized to induce respiratory challenges in several studies and UWBP has successfully measured accurate lung physiological assessment7,9,13-16. The protocol described is designed for standard adult laboratory mice. However, UWBP has been adapted to other animals such as rats, guinea pigs, and non-human primates17-20. UWBP is not restricted only to assessing pulmonary dysfunction but has also been used for the assessment of lung maturation3. The versatility, simplicity and reproducibility of UWBP have established an excellent technique for assessing pulmonary function in animals. Various software (see materials and equipment table) will be required to follow this procedure. An experienced scientist would be able to perform this protocol with a mouse within 1&#160;hr.

<table><tbody><tr><td>LabChart 7 software (for Macintosh)</td><td>ADINSTRUMENTS</td><td>MLU60/7</td><td>used in protocol step 4</td></tr><tr><td>PowerLab 8/30 (model ML870)</td><td>ADINSTRUMENTS</td><td>PL3508</td><td></td></tr><tr><td>Octal Bridge Amp (model ML228)</td><td>ADINSTRUMENTS</td><td>FE228</td><td></td></tr><tr><td>Black BNC to BNC cable (1 m)</td><td>ADINSTRUMENTS</td><td>MLAC01</td><td></td></tr><tr><td>Macintosh OS</td><td>Apple Inc.</td><td></td><td>Mac OS X 10.4 or later</td></tr><tr><td>Surgipack Digital Rectal Thermometer</td><td>Vega Technologies</td><td>MT-918</td><td></td></tr><tr><td>Grass volumeteric pressure transducer PT5A</td><td>Grass Instruments Co.</td><td>Model number PT5A; serial No. L302P4.</td><td></td></tr><tr><td>1 ml Syringe</td><td>Becton Dickinson (BD)</td><td>309628</td><td></td></tr><tr><td>5 ml Serological syringe pipettes</td><td>Greiner Bio One</td><td>606160</td><td>Connected via plastic tubing</td></tr><tr><td>Balance/Scales</td><td>VWR International, Pty Ltd</td><td>SHIMAUW220D</td><td>Any weighing balance with of 0.1 gram resolution</td></tr><tr><td>HM40 Humidity &amp;amp; temperature meter</td><td>Vaisala</td><td>HM40A1AB</td><td></td></tr><tr><td>Barometer</td><td>Barometer World</td><td>1586</td><td></td></tr><tr><td>Laboratory tubing</td><td>Dow Corning</td><td>508-101</td><td>Used to connect water column to the syringe and pressure transducer</td></tr><tr><td>Cylindrical Perspex Chamber</td><td>Dynalab Corp.</td><td></td><td>Custom built cylindrical chamber with internal dimensions as follows: 50 mm(w) x 1,500 mm(l). There are two lids for each side, with dimensions 80 mm(l) x 80 mm(w). Each lid has a 60 mm wide circular hole cut on the face of the lid 50 mm deep. This allows the chamber to fit into the lid. A rubber ring is fitted around each hole of the lid where the chamber will fit. For attachment of syringe and pressure transducer, the openings are 5 mm in diameter. For attachment of humidity probe, the openings are 25 mm in diameter.</td></tr><tr><td>80% Ethanol (4 L)</td><td>VWR International, Pty Ltd</td><td>BDH1162-4LP</td><td></td></tr></tbody></table>

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.

When this procedure has been followed correctly, a consistent oscillating trace is created on the data analysis software. The procedure provides a respiratory trace within a few minutes after setup with simple computing calculations to determine respiratory parameters listed. Figure 5 represents a suitable breathing trace from a control (healthy) mouse. Appropriate oscillating data is produced when the animal is not actively moving.
UWBP is an extremely useful and reliable ass...

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Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

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 ...

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Research

JoVE Journal

Biology

30.3K Views.  Monash Institute of Medical Research.  De beoordeling van respiratoire fysiologie is traditioneel ingeroepen technieken, die beperkingen of sedatie van het dier nodig. Ongebreideld hele lichaam plethysmografie, echter zorgt voor een nauwkeurige, niet-invasieve, kwantitatieve analyse van respiratoire fysiologie in diermodellen. Bovendien kan de techniek herhaalde respiratoire evaluatie van muizen waardoor longitudinale studies. 

Video: Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

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

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 ...

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 ...

Medicine

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

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

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

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

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

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

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

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

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 ...

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

Methods for Detecting Cough and Airway Inflammation in Mice

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

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

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

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

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