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 border="0" cellpadding="0" cellspacing="0" width="1031">
	<tbody>
		<tr height="17">
			<td height="17" style="height:17px;width:263px;">LabChart 7 software (for Macintosh)</td>
			<td style="width:139px;">ADINSTRUMENTS</td>
			<td style="width:155px;">MLU60/7</td>
			<td style="width:475px;">used in protocol step 4</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">PowerLab 8/30 (model ML870)</td>
			<td>ADINSTRUMENTS</td>
			<td>PL3508</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Octal Bridge Amp (model ML228)</td>
			<td>ADINSTRUMENTS</td>
			<td>FE228</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Black BNC to BNC cable (1m)</td>
			<td>ADINSTRUMENTS</td>
			<td style="width:155px;">MLAC01&#160;</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Macintosh OS&#160;</td>
			<td>Apple Inc.&#160;</td>
			<td>-</td>
			<td style="width:475px;">Mac OS X 10.4 or later</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Surgipack Digital Rectal Thermometer&#160;</td>
			<td>Vega Technologies</td>
			<td>MT-918</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;">Grass volumeteric pressure transducer PT5A</td>
			<td>Grass Instruments Co.</td>
			<td style="width:155px;">Model number PT5A; serial No. L302P4.</td>
			<td style="width:475px;"></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">1ml Syringe</td>
			<td>Becton Dickinson (BD)</td>
			<td>309628</td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;width:263px;">5ml serological syringe pipettes</td>
			<td>Greiner Bio One</td>
			<td>606160</td>
			<td style="width:475px;">Connected via plastic tubing</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Balance/Scales</td>
			<td>VWR International, Pty Ltd</td>
			<td>SHIMAUW220D</td>
			<td style="width:475px;">Any weighing balance with of 0.1 gram resolution</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">HM40 Humidity &#38; temperature meter&#160;</td>
			<td>Vaisala&#160;</td>
			<td>HM40A1AB</td>
			<td style="width:475px;"></td>
		</tr>
		<tr height="53">
			<td height="53" style="height:53px;">Barometer</td>
			<td>Barometer World</td>
			<td>1586</td>
			<td></td>
		</tr>
		<tr height="52">
			<td height="52" style="height:52px;">Laboratory tubing</td>
			<td>Dow Corning&#160;</td>
			<td>508-101</td>
			<td style="width:475px;">Used to connect water column to the syringe and pressure transducer</td>
		</tr>
		<tr height="136">
			<td height="136" style="height:136px;width:263px;">Cylindrical Perspex Chamber</td>
			<td>Dynalab Corp.</td>
			<td></td>
			<td style="width:475px;">Custom built cylindrical chamber with internal dimensions as follows: 50mm(w) x 1500mm(l). There are two lids for each side, with dimensions 80mm(l) x 80mm(w). Each lid has a 60mm wide circular hole cut on the face of the lid 50mm 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 5mm in diameter. For attachment of humidity probe, the openings are 25mm in diameter.&#160;</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">80% Ethanol (4L)</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...

Watch this Scientific Journal Video about Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography at JoVE.com

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 ...

measuring-respiratory-function-mice-using-unrestrained-whole-body

Research

JoVE Journal

Biology

30.1K Views.  Monash Institute of Medical Research.  Оценка физиологией дыхания традиционно полагались на методы, которые требуют сдержанности или торможение животного. Безудержная плетизмография всего тела, однако, обеспечивает точное, неинвазивный, количественный анализ физиологией дыхания в животных мод...

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 ...

Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume. A specially designed differential pressure transducer and an occlusion tail-cuff measure the total blood volume in the tail without the need to obtain the individual pulse signal. Special attention is afforded to the length of the occlusion cuff in order to derive the most accurate blood pressure readings. VPR can easily obtain readings on dark-skinned rodents, such as C57BL6 mice and is MRI compatible. The CODA system provides you with measurements of six (6) different blood pressure parameters; systolic and diastolic blood pressure, heart rate, mean blood pressure, tail blood flow, and tail blood volume. Measurements can be made on either awake or anesthetized mice or rats. The CODA system includes a controller, laptop computer, software, cuffs, animal holders, infrared warming pads, and an infrared thermometer. There are seven different holder sizes for mice as small as 8 grams to rats as large as 900 grams.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. This tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA ...

Visualization of Mitochondrial Respiratory Function using Cytochrome C Oxidase / Succinate Dehydrogenase (COX/SDH) Double-labeling Histochemistry

Mitochondrial DNA (mtDNA) defects are an important cause of disease and may underlie aging and aging-related alterations 1,2. The mitochondrial theory of aging suggests a role for mtDNA mutations, which can alter bioenergetics homeostasis and cellular function, in the aging process 3. A wealth of evidence has been compiled in support of this theory 1,4, an example being the mtDNA mutator mouse 5; however, the precise role of mtDNA damage in aging is not entirely understood 6,7.
Observing the activity of respiratory enzymes is a straightforward approach for investigating mitochondrial dysfunction. Complex IV, or cytochrome c oxidase (COX), is essential for mitochondrial function. The catalytic subunits of COX are encoded by mtDNA and are essential for assembly of the complex (Figure 1). Thus, proper synthesis and function are largely based on mtDNA integrity 2. Although other respiratory complexes could be investigated, Complexes IV and II are the most amenable to histochemical examination 8,9. Complex II, or succinate dehydrogenase (SDH), is entirely encoded by nuclear DNA (Figure 1), and its activity is typically not affected by impaired mtDNA, although an increase might indicate mitochondrial biogenesis 10-12. The impaired mtDNA observed in mitochondrial diseases, aging, and age-related diseases often leads to the presence of cells with low or absent COX activity 2,12-14. Although COX and SDH activities can be investigated individually, the sequential double-labeling method 15,16 has proved to be advantageous in locating cells with mitochondrial dysfunction 12,17-21.
Many of the optimal constitutions of the assay have been determined, such as substrate concentration, electron acceptors/donors, intermediate electron carriers, influence of pH, and reaction time 9,22,23. 3,3'-diaminobenzidine (DAB) is an effective and reliable electron donor 22. In cells with functioning COX, the brown indamine polymer product will localize in mitochondrial cristae and saturate cells 22. Those cells with dysfunctional COX will therefore not be saturated by the DAB product, allowing for the visualization of SDH activity by reduction of nitroblue tetrazolium (NBT), an electron acceptor, to a blue formazan end product 9,24. Cytochrome c and sodium succinate substrates are added to normalize endogenous levels between control and diseased/mutant tissues 9. Catalase is added as a precaution to avoid possible contaminating reactions from peroxidase activity 9,22. Phenazine methosulfate (PMS), an intermediate electron carrier, is used in conjunction with sodium azide, a respiratory chain inhibitor, to increase the formation of the final reaction products 9,25. Despite this information, some critical details affecting the result of this seemly straightforward assay, in addition to specificity controls and advances in the technique, have not yet been presented.

Visualization of Mitochondrial Respiratory Function using Cytochrome C Oxidase / Succinate Dehydrogenase COX/SDH Double-labeling Histochemistry

The cytochrome c oxidase/sodium dehydrogenase (COX/SDH) double-labeling method allows for direct visualization of mitochondrial respiratory enzyme deficiencies in fresh-frozen tissue sections. This is a straightforward histochemical technique and is useful in investigating mitochondrial diseases, aging, and aging-related disorders.

Mitochondrial DNA (mtDNA) defects are an important cause of disease and may underlie aging and aging-related alterations 1,2. The mitochondrial theory of ...

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 ...

Whole-Body Nanoparticle Aerosol Inhalation Exposures

Inhalation is the most likely exposure route for individuals working with aerosolizable engineered nano-materials (ENM). To properly perform nanoparticle inhalation toxicology studies, the aerosols in a chamber housing the experimental animals must have: 1) a steady concentration maintained at a desired level for the entire exposure period; 2) a homogenous composition free of contaminants; and 3) a stable size distribution with a geometric mean diameter &lt; 200 nm and a geometric standard deviation &sigma;g &lt; 2.5 5. The generation of aerosols containing nanoparticles is quite challenging because nanoparticles easily agglomerate. This is largely due to very strong inter-particle forces and the formation of large fractal structures in tens or hundreds of microns in size 6, which are difficult to be broken up. Several common aerosol generators, including nebulizers, fluidized beds, Venturi aspirators and the Wright dust feed, were tested; however, none were able to produce nanoparticle aerosols which satisfy all criteria 5.
A whole-body nanoparticle aerosol inhalation exposure system was fabricated, validated and utilized for nano-TiO2 inhalation toxicology studies. Critical components: 1) novel nano-TiO2 aerosol generator; 2) 0.5 m3 whole-body inhalation exposure chamber; and 3) monitor and control system. Nano-TiO2 aerosols generated from bulk dry nano-TiO2 powders (primary diameter of 21 nm, bulk density of 3.8 g/cm3) were delivered into the exposure chamber at a flow rate of 90 LPM (10.8 air changes/hr). Particle size distribution and mass concentration profiles were measured continuously with a scanning mobility particle sizer (SMPS), and an electric low pressure impactor (ELPI). The aerosol mass concentration (C) was verified gravimetrically (mg/m3). The mass (M) of the collected particles was determined as M = (Mpost-Mpre), where Mpre and Mpost are masses of the filter before and after sampling (mg). The mass concentration was calculated as C = M/(Q*t), where Q is sampling flowrate (m3/min), and t is the sampling time (minute). The chamber pressure, temperature, relative humidity (RH), O2 and CO2 concentrations were monitored and controlled continuously. Nano-TiO2 aerosols collected on Nuclepore filters were analyzed with a scanning electron microscope (SEM) and energy dispersive X-ray (EDX) analysis.
In summary, we report that the nano-particle aerosols generated and delivered to our exposure chamber have: 1) steady mass concentration; 2) homogenous composition free of contaminants; 3) stable particle size distributions with a count-median aerodynamic diameter of 157 nm during aerosol generation. This system reliably and repeatedly creates test atmospheres that simulate occupational, environmental or domestic ENM aerosol exposures.

A whole-body nanoparticle aerosol inhalation exposure facility was constructed for nano-sized titanium dioxide (TiO2) inhalation toxicology studies. This system provides nano-TiO2 aerosol test atmospheres that have: 1) a steady mass concentration; 2) a homogenous composition free of contaminants; and 3) a stable particle size distribution during aerosol generation.

Inhalation is the most likely exposure route for individuals working with aerosolizable engineered nano-materials (ENM). To properly perform nanoparticle ...

Assessing Murine Resistance Artery Function Using Pressure Myography

Pressure myograph systems are exquisitely useful in the functional assessment of small arteries, pressurized to a suitable transmural pressure. The near physiological condition achieved in pressure myography permits in-depth characterization of intrinsic responses to pharmacological and physiological stimuli, which can be extrapolated to the in vivo behavior of the vascular bed. Pressure myograph has several advantages over conventional wire myographs. For example, smaller resistance vessels can be studied at tightly controlled and physiologically relevant intraluminal pressures. Here, we study the ability of 3rd order mesenteric arteries (3-4 mm long), preconstricted with phenylephrine, to vaso-relax in response to acetylcholine. Mesenteric arteries are mounted on two cannulas connected to a pressurized and sealed system that is maintained at constant pressure of 60 mmHg. The lumen and outer diameter of the vessel are continuously recorded using a video camera, allowing real time quantification of the vasoconstriction and vasorelaxation in response to phenylephrine and acetylcholine, respectively.
	To demonstrate the applicability of pressure myography to study the etiology of cardiovascular disease, we assessed endothelium-dependent vascular function in a murine model of systemic hypertension. Mice deficient in the &alpha;1 subunit of soluble guanylate cyclase (sGC&alpha;1-/-) are hypertensive when on a 129S6 (S6) background (sGC&alpha;1-/-S6) but not when on a C57BL/6 (B6) background (sGC&alpha;1-/-B6). Using pressure myography, we demonstrate that sGC&alpha;1-deficiency results in impaired endothelium-dependent vasorelaxation. The vascular dysfunction is more pronounced in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice, likely contributing to the higher blood pressure in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice. 
	Pressure myography is a relatively simple, but sensitive and mechanistically useful technique that can be used to assess the effect of various stimuli on vascular contraction and relaxation, thereby augmenting our insight into the mechanisms underlying cardiovascular disease.

In pressure myography, an intact small segment of a vessel is mounted onto two small cannulas and pressurized to a suitable luminal pressure. Here, we describe the method to measure vasorelaxation response of the mouse 3rd order mesenteric arteries in c57 and sGC&alpha;1-/- mice using pressure myography.

Pressure myograph systems are  exquisitely useful in the functional assessment of small arteries, pressurized  to a suitable transmural pressure. The  ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Measuring RAN Peptide Toxicity in C. elegans

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis (ALS) and Huntington&#8217;s disease. Recently, repeat expansion-containing RNA was shown to be the substrate for a novel type of protein translation called repeat-associated non-AUG-dependent (RAN) translation. Unlike canonical translation, RAN translation does not require a start codon and only occurs when repeats exceed a threshold length. Because there is no start codon to determine the reading frame, RAN translation occurs in all reading frames from both sense and antisense RNA templates that contain a repeat expansion sequence. Therefore, RAN translation expands the number of possible disease-associated toxic peptides from one to six. Thus far, RAN translation has been documented in eight different repeat expansion-based neurodegenerative and neuromuscular diseases. In each case, deciphering which RAN products are toxic, as well as their mechanisms of toxicity, is a critical step towards understanding how these peptides contribute to disease pathophysiology. In this paper, we present strategies to measure the toxicity of RAN peptides in the model system C. elegans. First, we describe procedures for measuring RAN peptide toxicity on the growth and motility of developing C. elegans. Second, we detail an assay for measuring postdevelopmental, age-dependent effects of RAN peptides on motility. Finally, we describe a neurotoxicity assay for evaluating the effects of RAN peptides on neuron morphology. These assays provide a broad assessment of RAN peptide toxicity and may be useful for performing large-scale genetic or small molecule screens to identify disease mechanisms or therapies.

Measuring RAN Peptide Toxicity in C. elegans

Repeat-associated non-ATG-dependent translational products are emerging pathogenic features of several repeat expansion-based diseases. The goal of the protocol described is to evaluate toxicity caused by these peptides using behavioral and cellular assays in the model system C. elegans.

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis ...