These studies were supported by the National Institutes of Health COBRE grant P20GM125504-01 Sub-Project 8246.

The procedures described here were reviewed and approved by the University of Louisville Institutional Biosafety Committee (protocol # 14-038) and Institutional Animal Care and Use Committee (protocol # 19567).
1. Assembly of the sampling chamber
<ol>
	<li>Create two holes using a 3/4&#34; diamond drill bit on a drill press on the flat surfaces of a 600 mL square glass wide mouth Mason canning jar with 95 mm gasket and airtight lids, with bail and trigger clamp lids (Figure 1). 
	NOTE: A sampling chamber is not commercially available and must be constructed.</li>
	<li>Assemble a brass bulkhead (1/4&#34; NPT internal thread, 3/4-16 UNF external thread) through both the holes in the Mason jar using rubber washers (3/4&#34; inner diameter, 1&#34; outer diameter) on both faces of contact between the bulkhead and glass to ensure an airtight seal.</li>
	<li>Use one bulkhead assembly for a pressure sensor while attaching the other bulkhead by a Luer-connected syringe for calibration purposes.
	<ol>
		<li>For the pressure sensor, wrap the 1/4&#34; NPT threads of a high-performance gauge pressure transducer with Teflon tape and thread them into the bulkhead. Use a soldering iron to connect the pressure sensor wiring to an 8-pin male DIN connector, using the manufacturer wiring instructions for interface with a commercially available high-quality data acquisition device (see Table of Materials). 
		NOTE: This requires the use of a 150 K ohm 1/8 watt 1% metal film resistor within the DIN connector wiring.</li>
		<li>For the calibration port, use a 1/4&#34; male NPT to 1/8&#34; female NPT adaptor to connect a 1/8&#34; male NPT to a female Luer lock nickel-plated connector to the brass bulkhead wrapping threaded connections with Teflon tape. Use a polypropylene threaded male Luer cap to seal the Luer connector when not in use. 
		&#8203;NOTE: Do not overtighten the bulkhead connectors onto the glass jar, as this will develop cracks. If desired, silicone can be added to the rubber gaskets to ensure an airtight seal of the bulkheads to the glass jar.</li>
	</ol>
	</li>
</ol>
2. System setup
<ol>
	<li>Connect the sampling chamber to a bridge amplifier using an 8-pin DIN connector and the bridge amplifier to the data acquisition device, following the manufacturer&#39;s instructions.</li>
	<li>Connect the data acquisition device to a power supply and a computer running physiological data analysis software using the manufacturer&#39;s cables. 
	NOTE: Ensure that the data acquisition device is powered on and warmed up for at least 5 min before use to ensure that the sensor stabilizes its measurements.</li>
	<li>Initiate the software to interface with the data acquisition system.</li>
	<li>Download the optional Spirometry Module within the software, and modify the default unit settings from L/s to &#181;L/s in the Spirometry &#62; Settings window.</li>
</ol>
3. System calibration
<ol>
	<li>Within the software, create a 4-Channel window with the following data windows: Channel 1: Source data at 4 k/s sample rate and 1 mV range; Channel 2: Digital filter of Channel 1 using a high pass 1 Hz auto adjust filter; Channel 3: Smoothing of Channel 2 data by averaging of 100 samples; Channel 4: Spirometry Flow of Channel 3 data (Custom Flowhead, calibrated to Formula (&#181;L/s) = 120,000 x Voltage). 
	NOTE: 120,000 is a placeholder correlation coefficient that will be modified during calibration.</li>
	<li>Set up DataPad analysis of Channel 4 with the following Columns: Column 1: Channel 4 Data, Comments &#62; Full Comment Text; Column 2: Channel 4 data, Cyclic Measurements &#62; Average Cyclic Frequency; Column 3: Channel 4 data, Cyclic Measurements &#62; Average Cyclic Height.</li>
	<li>Set the frame rate to 100:1 on the bottom-right corner of the chart display. Save this window configuration as a template for all future studies.</li>
	<li>Close the Sample Chamber lid and attach a 25 &#181;L Gas Tight syringe to the Luer bulkhead connector. Fit the syringe with a Chaney adaptor set to repeatedly deliver a volume of 20 &#181;L. 
	NOTE: An optional short piece of 1/16&#34; tubing and Luer/barb connectors may be used to connect the syringe to Sample Chamber. However, long tubing should be avoided to avoid significant changes to the total air volume of the Sample Chamber.</li>
	<li>Draw 20 &#181;L of air into the syringe using the depth stop of the Chaney adaptor.</li>
	<li>Zero the pleth in the software (Setup &#62; Zero All Inputs (Alt-Z)) and start a recording.</li>
	<li>While recording, and with a stable baseline, rapidly depress/withdraw the Syringe plunger for approximately 10 repetitions to replicate subject breathing with a measured 20 &#181;L breath. Stop the recording. 
	NOTE: The frequency of the artificial breaths should exceed 2 Hz to maximize the reproducibility of the calibration.</li>
	<li>Label the identity of the measured sample by right-clicking the beginning of numbered pleth recording and click on Add Comment.</li>
	<li>Reset the Syringe, Zero the input, and repeat recording measurements of 20 &#181;L pulses two additional times (three total recording sessions).</li>
	<li>After completing all the measurements, use the computer mouse to select a portion of breathing pleth that accurately represents the artificial 20 &#181;L breaths. 
	NOTE: Within the DataPad module, data will appear in the preview header providing a temporary readout of the breath rate (Average Cyclic Frequency, Hz) and breath depth (Average Cyclic Height, &#181;L). The data preview can be recorded into DataPad using the Add to DataPad icon.</li>
	<li>Review the Column 3 data (Average Cyclic Height), and calculate the average measured breath volume from the three recordings. Perform the following calculation of the average measured breath volume: Calibration coefficient = delivered volume / measured volume x 120,000. 
	&#8203;NOTE: 120,000 was the placeholder Flow Head Calibration coefficient used in step 3.1 now modified from measured data. The system is now calibrated to typical mouse breathing using current environmental temperature and humidity. The system can now monitor the subject breathing, and calibration can be re-performed daily to account for any fluctuation in temperature/humidity.</li>
</ol>
4. Subject monitoring
<ol>
	<li>Open a master template as outlined in step 3.4, or complete steps 4.2 to 4.3.</li>
	<li>Within the software, create a 4-Channel window with the following data processing: Channel 1: Source data at 4 k/s sample rate and 1 mV range; Channel 2: Digital filter of Channel 1 using a High Pass 1 Hz Auto Adjust filter; Channel 3: Smoothing of Channel 2 data by Averaging of 100 Samples; Channel 4: Spirometry Flow of Channel 3 data (Custom Flowhead, calibrated to Formula (&#181;L/s) = 120,000 x Voltage). 
	NOTE: 120,000 is the correlation coefficient calculated for the current pressure sensor; however, the user should perform the system calibration described in step 3 and use this user-defined correlation coefficient instead.</li>
	<li>Set up DataPad analysis of Channel 4 with the following Columns: Column 1: Channel 4 Data, Comments &#62; Full Comment Text; Column 2: Channel 4 data, Cyclic Measurements &#62; Average Cyclic Frequency; Column 3: Channel 4 data, Cyclic Measurements &#62; Average Cyclic Height.</li>
	<li>Place subject in sampling chamber and close lid. For this experiment a conscious 4-12 week female albino C57BL/6J mouse (B6(Cg)-Tyrc-2J/J) was used.</li>
	<li>Equalize the atmospheric pressure in the chamber (from sealing the lid) by briefly loosening the Luer bulkhead cap and retighten.</li>
	<li>Observe that the subject is not actively moving within the sampling chamber before Zeroing All Inputs (Alt-Z shortcut) and beginning a recording. 
	NOTE: If the subject begins to move in the sampling chamber, the baseline may move off the scale, which may be addressed by re-zeroing all inputs in the middle of a recording, which will create a fresh recording on the scale. Suppose the subject engages in exploration or grooming during the recording; note which portion of the plethysmography recording most accurately reflects normal breathing.</li>
	<li>Label the subject&#39;s identity by right-clicking the beginning of numbered pleth recording and click on Add Comment.</li>
	<li>Return the subject to its cage. Limit the time spent in the sealed sampling chamber to 5 min to avoid asphyxiation and stress. 
	NOTE: The risk for asphyxiation is low given that the 600 mL air volume of the Sample Chamber will not be rapidly spent by a healthy mouse breathing at &#60;15 mL/min.</li>
	<li>Use the computer mouse to select a portion of breathing pleth that accurately represents the subject breathing. 
	NOTE: Within the DataPad module, data will appear in the preview header providing a temporary readout of the breath rate (Average Cyclic Frequency, Hz) and breath volume (Average Cyclic Height, &#181;L). The data preview can be recorded into DataPad using the Add to DataPad icon.</li>
	<li>Continue measuring subject mice one at a time and recording representative sections of breathing pleth to DataPad.</li>
	<li>After data recording, export DataPad data to Excel. Calculate the minute volume as follows: Minute Volume (mL/min) = Breath Rate (Hz) x Breath Volume (&#181;l) x 0.06.</li>
</ol>

<ol>
	<li>Warawa, J. M., Long, D., Rosenke, R., Gardner, D., Gherardini, F. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+for+the+Burkholderia+pseudomallei+capsular+polysaccharide+encoded+by+the+wcb+operon+in+acute+disseminated+melioidosis.">Role for the Burkholderia pseudomallei capsular polysaccharide encoded by the wcb operon in acute disseminated melioidosis.</a> Infection and Immunity. 77 (12), 5252-5261 (2009).</li><li>West, T. E., Myers, N. D., Liggitt, H. D., Skerrett, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+pulmonary+infection+and+inflammation+induced+by+inhalation+of+Burkholderia+pseudomallei.">Murine pulmonary infection and inflammation induced by inhalation of Burkholderia pseudomallei.</a> International Journal of Experimental Pathology. 93 (6), 421-428 (2012).</li><li>Lawrenz, M. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+evaluation+of+murine+lung-specific+disease+models+for+Pseudomonas+aeruginosa+applicable+to+therapeutic+testing.">Development and evaluation of murine lung-specific disease models for Pseudomonas aeruginosa applicable to therapeutic testing.</a> Pathogens and Disease. 73 (5), (2015).</li><li>Lim, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+respiratory+function+in+mice+using+unrestrained+whole-body+plethysmography.">Measuring respiratory function in mice using unrestrained whole-body plethysmography.</a> Journal of Visualized Experiments: JoVE. (90), e51755 (2014).</li><li>Drorbaugh, J. E., Fenn, W. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+barometric+method+for+measuring+ventilation+in+newborn+infants.">A barometric method for measuring ventilation in newborn infants.</a> Pediatrics. 16 (1), 81-87 (1955).</li><li>Simon, G., Pride, N. B., Jones, N. L., Raimondi, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relation+between+abnormalities+in+the+chest+radiograph+and+changes+in+pulmonary+function+in+chronic+bronchitis+and+emphysema.">Relation between abnormalities in the chest radiograph and changes in pulmonary function in chronic bronchitis and emphysema.</a> Thorax. 28 (1), 15-23 (1973).</li><li>Gassiep, I., Armstrong, M., Norton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+melioidosis.">Human melioidosis.</a> Clinical Microbiology Reviews. 33 (2), 06-19 (2020).</li><li>Gutierrez, M. G., Warawa, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Attenuation+of+a+select+agent-excluded+Burkholderia+pseudomallei+capsule+mutant+in+hamsters.">Attenuation of a select agent-excluded Burkholderia pseudomallei capsule mutant in hamsters.</a> Acta Tropica. 157, 68-72 (2016).</li><li>Gutierrez, M. G., Pfeffer, T. L., Warawa, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Type+3+secretion+system+cluster+3+is+a+critical+virulence+determinant+for+lung-specific+melioidosis.">Type 3 secretion system cluster 3 is a critical virulence determinant for lung-specific melioidosis.</a> PLoS Neglected Tropical Diseases. 9 (1), 3441 (2015).</li><li>Rocco, P. R. M., Zin, W. A., Gullo, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Anaesthesia, Pain, Intensive Care and Emergency Medicine. , (2002).</li><li>Fodah, R. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlation+of+Klebsiella+pneumoniae+comparative+genetic+analyses+with+virulence+profiles+in+a+murine+respiratory+disease+model.">Correlation of Klebsiella pneumoniae comparative genetic analyses with virulence profiles in a murine respiratory disease model.</a> PLoS One. 9 (9), 107394 (2014).</li><li>Gotts, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinically+relevant+model+of+pneumococcal+pneumonia,+ARDS,+and+nonpulmonary+organ+dysfunction+in+mice.">Clinically relevant model of pneumococcal pneumonia, ARDS, and nonpulmonary organ dysfunction in mice.</a> American Journal of Physiology-Lung Cellular and Molecular Physiology. 317 (5), 717-736 (2019).</li><li>Galan, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Common+themes+in+the+design+and+function+of+bacterial+effectors.">Common themes in the design and function of bacterial effectors.</a> Cell Host &amp; Microbe. 5 (6), 571-579 (2009).</li></ol>

The authors have no conflicts of interest.

sWBP is an attractive approach for enhancing the understanding of respiratory infection in small animal models. Importantly, it is a non-invasive approach, and as such, it does not pose a significant risk of causing undue stress to research animals during an infection challenge. Indeed, the procedure of monitoring subject breathing is a rapid test requiring several minutes and minimal subject handling. The scientific benefit is the high-resolution understanding of how microbial pathogens influence lung function during disease. This approach will provide benefit to basic research, facilitating the understanding of how a pathogen causes disease, as well as providing a translational utility to understand how a novel therapeutic restores a research subject to a state of respiratory health.
In this manuscript, representative results are provided for the pathogen B. pseudomallei, which causes an early lethargic response. Not all bacterial lung infections present in the same manner in mouse infection models. Previous experience with other infection models has demonstrated that the bacterial pathogen Klebsiella pneumoniae presents as an asymptomatic infection up to the point at which mice succumb to infection, also at approximately Day 3 post-infection11. It is hypothesized that the host demand for inspired air (i.e., minute volume) may closely relate to the degree of lethargy with which a given disease presents. Future studies will be required to examine how different bacterial pathogens affect lung function during respiratory disease. It is understood that different pathogens have unique approaches to evade host defense, including differences in, (1) propensity to be intracellular or extracellular pathogens, (2) the ability to cause early/late hypothermic response, and (3) use of different repertoires of virulence determinants3,12,13. Therefore, it is likely that different disease strategies will result in unique effects on lung function and breathing during infection.
The recommended settings described in this protocol can be modified to accommodate unique challenges present during sWBP. One of the common issues experienced during an sWBP recording session is the subject's movement within the Sample Chamber. As mentioned, this movement modifies the baseline and can affect the accuracy of breathing measurements. A Digital filter was used to normalize the shifting baseline, allowing viable breath measurements despite small movements. Excessive movement can push a baseline measurement out of the range of a zeroed input. Recordings are recommended at 1 mV range (Channel 1 setting), which provides a compromise of still observing the peaks of the plethysmography while avoiding loss of data outside of range. For exceptionally active subjects, it may be necessary to extend the recording range &gt;1 mV to avoid persistent out of range signals.
The recommended procedure calls for daily calibration (or at each session) to accommodate environmental humidity/temperature fluctuations. Traditional WBP uses complex calculations that factor in the temperature/humidity of both environment and subject5,6. It has been demonstrated that in the present sWBP apparatus, the effects of the host temperature/humidity do not significantly alter the measured breath volume of a calibration source. Therefore, this approach in sWBP fundamentally differs from the &gt;50-year-old approach of Drorbaugh and Fenn. Here, sWBP directly relates pressure changes to a measured breath volume with no further correction from the host.
It is essential to contrast research animal WBP to that of clinical WBP. The types of biometric data that were attempted to collect by sWBP are breath volume and frequency. Such measurements are collected clinically using simple spirometry equipment in which a patient holds a breath monitor to their mouth and breaths normally into a device monitoring airflow. Similar spirometry in research animals requires restraint, thus contributing to stress and an inherent disruption in breathing. Therefore, simple spirometry is functional clinically but not for research animals. WBP serves an essential purpose in the clinic to collect advanced data, including such measurements as residual lung volume. Such data can only be contained in the context of a subject being able to follow instructions on how they breathe, including Forced Expiration (emptying of their lung by a deep exhale). Research animals cannot be relied upon to follow breathing instructions from a researcher. Many of the advanced measurements collected clinically during WBP cannot be reproduced in research animals. WBP in research animals is fundamentally different from clinical WBP. Animal WBP seeks to collect simple ventilation data (breath rate and volume) in a non-restrained fashion to avoid animal stress and breathing perturbation. Thus far, the use of WBP in research animals appears to replicate the techniques used in clinical WBP, including complex calculations based on environmental and subject temperature and humidity, but without the ability to collect the advanced data from a subject that can follow instructions on how to perform a forced expiration. With this in mind, it was sought to demonstrate whether a simplified version of WBP would suffice to collect the pertinent breathing frequency and volume relevant to respiratory disease studies. A calibration session was employed, which compensated for any variation in environmental temperature and humidity. Further, it was demonstrated with an artificial mouse that subject temperature, and humidity to a measured breath volume have no significant effect on accurately measuring breath volume. It was concluded that sWBP has excellent application to research animal studies, without the requirement of the user to employ cumbersome mathematical treatment of data.

Respiratory bacterial pathogens are often associated with an inflammatory response in the lung leading to lung pathology1,2. In the clinical setting, diagnosis of pneumonia typically includes culture techniques from sputum, blood-oxygen saturation analysis, and chest X-ray. These techniques can be translated for small animal infection models, but only oxygen saturation analysis represents a rapid, real-time analysis in mice for disease severity. The blood oxygen saturation (SpO2) was previously investigated as a method to track disease progression in respiratory disease studies; however, moribund mice have unexpectedly high SpO2 readings both in a Pseudomonas aeruginosa model3, which are not the predictive or moribund disease, likely because mice can modulate their physiological activity. To this end, diagnostic levels of SpO2 were not found for bacterial respiratory disease in mice thus far.
Therefore, this work investigated the use of other clinically relevant methods of detecting the effects of lung disease on lung function as a rapid physiologic measurement. Simplified Whole Body Plethysmography (sWBP) offers the opportunity to investigate breath rate and depth as a rapid, non-invasive biometric analysis. Previous studies have demonstrated how to assemble WBP apparatus in a laboratory4; however, several of the components shown in such studies are not currently commercially available. Further, traditional WBP requires complex data collection and data processing based on humidity and temperature5,6. Hence, it was decided to develop a simplified WBP apparatus that is calibrated daily to room temperature/humidity and assess whether or not the temperature/humidity contribution of the subject itself has any effect on the measured breath volume. Thus, a modified sWBP apparatus has been created which sources the currently available materials. Further, it has been investigated whether this lab-sourced apparatus can detect changes in breathing associated with disease progression during the model of lethal respiratory melioidosis in mice.
The sWBP apparatus constructed for this work used commercially available equipment and software to process analog pressure sensor data into a digital readout. The pressure sensor was fitted to an airtight glass jar with bulkhead connectors. The benefit of a glass jar is the structural rigidity of the material, which will resist changes to the internal pressure of the jar, affecting measurements of volume changes during the monitoring of breathing. The sampling chamber has been engineered to have two ports on the two flat surfaces of the square jar, one to access the chamber by a Luer connector for calibration and the other to house the pressure sensor. The pressure sensor selected has a highly sensitive gauge pressure transducer with a range for small changes in pressure (25 mbar range).
This protocol is demonstrated using a murine model of respiratory melioidosis. Burkholderia pseudomallei (Bp) is the bacterial agent of melioidosis - a disease associated with tropical regions of the world7. Bp is found in the environment, specifically in wet environments of standing water and moist soil, from which it typically causes subcutaneous infections of cuts/scratches of susceptible hosts. However, Bp is also infectious when inhaled and is a potential threat for use in bioterrorism by aerosol dispersal. While fully virulent Bp requires handling in a BSL-3 laboratory, an acapsular mutant strain was previously engineered, which can be safely handled at BSL-2 and excluded from the select agent criteria8. Further, an intubation-mediated intratracheal (IMIT) infection model of respiratory melioidosis has been developed to study respiratory disease progression of Bp5,9. We have used this infection model to characterize the change in breathing that occurs during disease progression through the moribund endpoint.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1/8&#34; NPT Luer adaptor</td>
			<td>Amazon</td>
			<td>B07DH9MY8W</td>
			<td>Calibration port</td>
		</tr>
		<tr>
			<td>1/8&#34; NPT to 1/4&#34; NPT adaptor</td>
			<td>Amazon</td>
			<td>B07T6CR6FS</td>
			<td>Bulkhead to luer adaptor</td>
		</tr>
		<tr>
			<td>150 kohm resistor</td>
			<td>Amazon</td>
			<td>B07GPRYL81</td>
			<td>Pressure transducer excitation voltage selection</td>
		</tr>
		<tr>
			<td>3/4&#34; diamond drill bit</td>
			<td>Drilax</td>
			<td>DRILAX100425</td>
			<td>To drill bulkhead mounts in glass jar</td>
		</tr>
		<tr>
			<td>Bridge Amp</td>
			<td>AD Instruments</td>
			<td>FE221</td>
			<td>One channel option</td>
		</tr>
		<tr>
			<td>Bulkhead fitting</td>
			<td>Legines</td>
			<td>3000L-B</td>
			<td>1/4&#34; NPT, 3/4-16 UNF brass bulkhead coupling</td>
		</tr>
		<tr>
			<td>Chaney adaptor</td>
			<td>Hamilton</td>
			<td>14725</td>
			<td>Gas tight syringe adaptor for set volume</td>
		</tr>
		<tr>
			<td>DIN connector</td>
			<td>AD Instruments</td>
			<td>SP0104</td>
			<td>To connect pressure sensor to Bridge Amp</td>
		</tr>
		<tr>
			<td>Gastight syringe, 25 uL</td>
			<td>Hamilton</td>
			<td>80201</td>
			<td>Calibration syringe</td>
		</tr>
		<tr>
			<td>LabChart</td>
			<td>AD Instruments</td>
			<td></td>
			<td>Life Science Data Acquisition Software</td>
		</tr>
		<tr>
			<td>Luer plug</td>
			<td>Cole Parmer</td>
			<td>45513-56</td>
			<td>Calibration port closure</td>
		</tr>
		<tr>
			<td>PowerLab 4/26</td>
			<td>AD Instruments</td>
			<td>PL2604</td>
			<td>Digital interface to computer</td>
		</tr>
		<tr>
			<td>Pressure transducer</td>
			<td>Omega Engineering</td>
			<td>PX409-10WGV</td>
			<td>High accuracy oil filed gage pressure sensor</td>
		</tr>
		<tr>
			<td>Rubber gasket</td>
			<td>Amazon</td>
			<td>B07LH4C8LS</td>
			<td>To mount bulkheads (4 required per chamber)</td>
		</tr>
		<tr>
			<td>Square glass jar</td>
			<td>Amazon</td>
			<td>B07VNSPR8P</td>
			<td>600 ml with 95 mm silicone gasket</td>
		</tr>
	</tbody>
</table>

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.

System calibration 
The data analysis software allows for direct calibration of a custom flowhead, such as the one described herein. This is performed when setting the Spirometry Flow. As described in step 3.1, an option exists to input the known calibration air volume, which calculates the voltage to volume correlation coefficient within the system. This, however, generates a correlation coefficient based on a single read, and it has been observed that the inherent variation of calibration from an n = 1 standard has poor utility. The current approach can address this shortcoming and allows a user to perform a daily calibration using multiple readings averaged to calculate a calibration coefficient. Calibration with 20 &#181;L of injected air was demonstrated herein, representing a typical high-end breath volume in a typical mouse. The software assumes an origin intercept (0,0) and is thus calibrated from 0-20 &#181;L using this approach.
The methodology proposed here for sWBP calibrates daily, thus accounting for any fluctuations in environmental humidity/temperature. The original methods used for specific WBP, date back to the methodology of Drorbaugh and Fenn from 1955, who developed WBP for measuring ventilation in human infants5. The Drorbaugh and Fenn calculations account for variations in temperature and humidity of the environment and the subject. The current approach corrects for environmental fluctuations by calibrating each sWBP session. Still, it was decided to address whether the heating and humidification of breathing across the nasal cavity/lung of a mouse affect the measurement of a known volume of air. Thus, an artificial apparatus was created to mimic the subject&#39;s effect on heating and humidifying calibrated air measurements. Luer connectors were attached to a 15 mL conical tube and placed this sealed conical in-line between the sample chamber and the gas-tight calibration syringe. A 20 &#181;L calibration was performed using an empty conical tube held at room temperature (23 &#176;C). The conical tube was then partially filled with distilled water to just below the Luer connectors, allowing time to equilibrate the headspace of the conical; the calibration volume was then re-measured to investigate the effect of humidity. The conical tube was placed in a heating block and equilibrated at 37 &#176;C in a humid environment, and finally equilibrated to 37 &#176;C with no water to assess the effect of subject heating and with no additional contribution of humidity. Figure 2 demonstrates that all conditions tested did not significantly impact the calibrated 20 &#181;L measurement delivered by the gas-tight syringe. From this finding, it was concluded that sWBP offers an accessible approach to monitor breathing in research animals without the need for complex calculations predicated on the temperature and humidity of the animal subject, as these make no significant impact on measured breath volume.
Subject monitoring 
sWBP was used to monitor breathing during the disease of lethal respiratory infections with the bacterial pathogen B. pseudomallei. One challenge of monitoring breathing in conscious animals is the curiosity of normal healthy animals moving within the sample chamber. The movement of the mouse creates a constantly moving baseline that can be mitigated in part by pre-conditioning subjects to the chamber over a period of several days prior to measurement.&#160;This issue primarily affects the baseline measurement in healthy mice, as the subjects become lethargic during infection, making sWBP much more manageable with reduced subject activity. It may be tempting to attempt to use some form of restraint, whether physical or anesthesia. The use of physical restraint may affect natural breathing by causing stress. Further, the use of anesthetics is known to have pronounced effects on breath rate and depth10; thus, it was decided to investigate the impact of anesthesia with the in-house sWBP apparatus. Isoflurane is commonly used to perform in vivo diagnostic imaging during the infection models, and therefore, a C57BL/6 mouse was anesthetized and monitored progression until recovery out of anesthesia using sWBP. This trial was conducted with a juvenile 4-week old albino C57BL/6J mouse to prolong the window of recovery from anesthesia. Figure 3 demonstrates that the preferred anesthetic causes mice to exhibit a slow breath rate with a large tidal volume of air. As mice begin to recover from sedation, their breath rate increases and breath volume decreases, with the net effect that total inspired air slowly increases. In this trial, it was found that the breath volume is restored to pre-anesthesia levels within the first 30 s of recovery. The breath rate steadily increases until the baseline breathing is restored to 2-2.5 min post-removal from anesthesia. The minute volume closely followed the effects of breath rate, reaching baseline minute volume by 2.5 min post-removal from anesthesia. This finding supports that anesthesia should not be employed in the sWBP approach. It dramatically affects baseline breathing, not surprisingly, as anesthesia will slow the host metabolism, creating a reduced demand for inspired oxygen.&#160;Sanitation of the sample chamber should also be considered between subjects to address study-specific infection control as well as the impact of pheromones from urine or feces which could impact the stress between subjects.
WBP is an attractive strategy for monitoring lung function in respiratory disease models in a non-invasive manner. sWBP was used to study how breathing changes during lethal respiratory melioidosis infections (Figure 4), with time points mirroring bioluminescence monitoring in the lung. It was observed that this model is associated with an early onset of lethargy, which persists in a slowly progressive manner until the development of the moribund disease at approximately 3 days post-infection. It was also observed that the breath rate and total inspired air (minute volume) of the mice decrease rapidly during the first day of infection and remain low for the remainder of the course of infection (Figure 4A,C). This pattern is consistent with the early onset lethargy, which persists for the next 2 days of the infection. In contrast, the breath volume does not steeply drop during the first 24 h and instead has a slight and steady decrease, which approaches a linear decline over the 3-day course of the disease (Figure 4B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62722/62722fig01.jpg" /> 
Figure 1: sWBP apparatus. A custom Sample Chamber was constructed from a sealable square glass jar with bulkhead connectors on two flat faces. One bulkhead was used to mount a gauge pressure sensor connected to a bridge amplifier and data acquisition device digitizer via an 8-pin DIN connection. The second bulkhead was fitted with a Luer connector for calibration by a gas-tight syringe. The device was connected to a PC running the software. <a href="https://www.jove.com/files/ftp_upload/62722/62722fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62722/62722fig02.jpg" /> 
Figure 2: Effect of subject temperature and humidity on breath volume. A 15 mL conical tube with Luer connectors was installed in-line between the 20 &#181;L calibration syringe and the Sample Chamber. The system was calibrated to 20 &#181;L with no additional temperature/humidity contribution from the conical tube. Other measurements were collected post-equilibration with saturated humidity from distilled water and/or warming of the conical tube from room temperature (23 &#176;C) to body temperature (37 &#176;C). No significant difference was detected from n = 5 measurements of each condition by One-way ANOVA with Tukey&#39;s Multiple Comparison post-test. <a href="https://www.jove.com/files/ftp_upload/62722/62722fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62722/62722fig03.jpg" /> 
Figure 3: Effect of gas anesthesia on breathing in mice. Representative data from a 4-week old female albino C57BL/6J mouse (8.6 g) was sedated for 5 min with 3% isoflurane in oxygen and transferred to an sWBP Sample Chamber. Pleth data was collected for 150 s following removal from anesthesia. The subject began initial ambulation by 100 s following removal from anesthesia. (A) Baseline breathing before anesthesia, measuring a 4.97 Hz breath rate, 9.74 &#181;L breath volume, and 2.91 mL minute volume. (B) The first 60 s of changes to breathing during recovery from anesthesia. (A-B) Vertical axis measuring &#181;L per breath and horizontal axis in seconds. (C-E) Ventilation data was collected during the 150 s of recovery from anesthesia, averaged from &#8805;3 breath cycles per time point for (C) Breath Rate, (D) Breath Volume, and the (E) calculated Minute Volume. The pre-anesthesia baseline values are indicated with a horizontal dotted line in each respective graph. <a href="https://www.jove.com/files/ftp_upload/62722/62722fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62722/62722fig04.jpg" /> 
Figure 4: Effect of respiratory melioidosis on host breathing. Five 8-week female C57BL/6 mice were infected with 4.9 log CFU of bioluminescent B. pseudomallei strain JW270. sWBP was conducted throughout the 3-day course of infection, measuring Breath Rate (A) and Breath Volume (B). The total inspired air was calculated as the Minute Volume (C). Data for each of the five subjects are independently plotted with third-order polynomial regression. <a href="https://www.jove.com/files/ftp_upload/62722/62722fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Simplified Whole Body Plethysmography to Characterize Lung Function During Respiratory Melioidosis at JoVE.com

Simplified Whole Body Plethysmography to Characterize Lung Function During 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 ...

simplified-whole-body-plethysmography-to-characterize-lung-function

Research

JoVE Journal

Immunology and Infection

3.0K Views.  University of Louisville. This protocol presents the construction and use of a Simplified Whole Body Plethysmography apparatus to monitor bacterial respiratory disease progression non-invasively.

Video: Simplified Whole Body Plethysmography to Characterize Lung Function During Respiratory Melioidosis

Establishing a Liquid-covered Culture of Polarized Human Airway Epithelial Calu-3 Cells to Study Host Cell Response to Respiratory Pathogens In vitro

The apical and basolateral surfaces of airway epithelial cells demonstrate directional responses to pathogen exposure in vivo. Thus, ideal in vitro models for examining cellular responses to respiratory pathogens polarize, forming apical and basolateral surfaces. One such model is differentiated normal human bronchial epithelial cells (NHBE). However, this system requires lung tissue samples, expertise isolating and culturing epithelial cells from tissue, and time to generate an air-liquid interface culture.
Calu-3 cells, derived from a human bronchial adenocarcinoma, are an alternative model for examining the response of proximal airway epithelial cells to respiratory insult1, pharmacological compounds2-6, and bacterial7-9 and viral pathogens, including influenza virus, rhinovirus and severe acute respiratory syndrome - associated coronavirus10-14. Recently, we demonstrated that Calu-3 cells are susceptible to respiratory syncytial virus (RSV) infection in a manner consistent with NHBE15,16 . Here, we detail the establishment of a polarized, liquid-covered culture (LCC) of Calu-3 cells, focusing on the technical details of growing and culturing Calu-3 cells, maintaining cells that have been cultured into LCC, and we present the method for performing respiratory virus infection of polarized Calu-3 cells.
To consistently obtain polarized Calu-3 LCC, Calu-3 cells must be carefully subcultured before culturing in Transwell inserts. Calu-3 monolayer cultures should remain below 90% confluence, should be subcultured fewer than 10 times from frozen stock, and should regularly be supplied with fresh medium. Once cultured in Transwells, Calu-3 LCC must be handled with care. Irregular media changes and mechanical or physical disruption of the cell layers or plates negatively impact polarization for several hours or days. Polarization is monitored by evaluating trans-epithelial electrical resistance (TEER) and is verified by evaluating the passive equilibration of sodium fluorescein between the apical and basolateral compartments17,18 . Once TEER plateaus at or above 1,000 &Omega;&times;cm2, Calu-3 LCC are ready to use to examine cellular responses to respiratory pathogens.

Establishing a Liquid-covered Culture of Polarized Human Airway Epithelial Calu-3 Cells to Study Host Cell Response to Respiratory Pathogens In vitro

The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the Centers for Disease Control and Prevention.

The  apical and basolateral surfaces of airway epithelial cells demonstrate  directional responses to pathogen exposure in  vivo. Thus, ideal in vitro ...

Simplified Human Neutrophil Extracellular Traps (NETs) Isolation and Handling

Neutrophil Extracellular Traps (NETs) have been recently identified as part of the neutrophil&#8217;s antimicrobial armamentarium. Apart from their role in fighting infections, recent research has demonstrated that they may be involved in many other disease processes, including cancer progression. Isolating purified NETs is a crucial element to allow the study of these functions.
In this video, we demonstrate a simplified method of cell free NET isolation from human whole blood using readily available reagents. Isolated NETs can then be used for immunofluorescence staining, blotting or various functional assays. This enables an assessment of their biologic properties in the absence of the potential confounding effects of neutrophils themselves.
A density gradient separation technique is employed to isolate neutrophils from healthy donor whole blood. Isolated neutrophils are then stimulated by phorbol 12-myristate 13-acetate (PMA) to induce NETosis. Activated neutrophils are then discarded, and a cell-free NET stock is obtained.
We then demonstrate how isolated NETs can be used in an adhesion assay with A549 human lung cancer cells. The NET stock is used to coat the wells of a 96 well cell culture plate O/N, and after ensuring an adequate NET monolayer formation on the bottom of the wells, CFSE labeled A549 cells are added. Adherent cells are quantified using a Nikon TE300 fluorescent microscope. In some wells, 1000U DNAse1 is added 10 min before counting to degrade NETs

Simplified Human Neutrophil Extracellular Traps NETs Isolation and Handling

In the following protocol, we describe a very simple way to isolate Neutrophil Extracellular traps (NETs) from human whole blood using readily available reagents. We then demonstrate how the isolated NETs can be used in an in vitro adhesion assay with cancer cells.

Neutrophil Extracellular Traps (NETs) have been recently identified as part of the neutrophil&#8217;s antimicrobial armamentarium. Apart from their role ...

Phage Phenomics: Physiological Approaches to Characterize Novel Viral Proteins

Current investigations into phage-host interactions are dependent on extrapolating knowledge from (meta)genomes. Interestingly, 60 - 95% of all phage sequences share no homology to current annotated proteins. As a result, a large proportion of phage genes are annotated as hypothetical. This reality heavily affects the annotation of both structural and auxiliary metabolic genes. Here we present phenomic methods designed to capture the physiological response(s) of a selected host during expression of one of these unknown phage genes. Multi-phenotype Assay Plates (MAPs) are used to monitor the diversity of host substrate utilization and subsequent biomass formation, while metabolomics provides bi-product analysis by monitoring metabolite abundance and diversity. Both tools are used simultaneously to provide a phenotypic profile associated with expression of a single putative phage open reading frame (ORF). Representative results for both methods are compared, highlighting the phenotypic profile differences of a host carrying either putative structural or metabolic phage genes. In addition, the visualization techniques and high throughput computational pipelines that facilitated experimental analysis are presented.

Here, we present phenomic approaches for the functional characterization of putative phage genes. Techniques include a developed assay capable of monitoring host anabolic metabolism, the Multi-phenotype Assay Plates (MAPs), in addition to the established method of metabolomics, capable of measuring effects to catabolic metabolism.

Current investigations into phage-host interactions are dependent on extrapolating knowledge from (meta)genomes. Interestingly, 60 - 95% of all phage ...

Biomimetic Materials to Characterize Bacteria-host Interactions

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The biochemical and functional characterization of adhesins mediating these initial bacteria-host interactions is often compromised by the presence of other bacterial factors, such as cell wall components or secreted molecules, which interfere with the analysis. This protocol describes the production and use of biomimetic materials, consisting of pure recombinant adhesins chemically coupled to commercially available, functionalized polystyrene beads, which have been used successfully to dissect the biochemical and functional interactions between individual bacterial adhesins and host cell receptors. Protocols for different coupling chemistries, allowing directional immobilization of recombinant adhesins on polymer scaffolds, and for assessment of the coupling efficiency of the resulting &#8220;bacteriomimetic&#8221; materials are also discussed. We further describe how these materials can be used as a tool to inhibit pathogen mediated cytotoxicity and discuss scope, limitations and further applications of this approach in studying bacterial - host interactions.

Bacterial attachment to host cells is a key step during host colonization and infection. This protocol describes the generation of polymer-coupled recombinant adhesins as biomimetic materials which allow analysis of the contribution of individual adhesins to these processes, independent of other bacterial factors.

Bacterial attachment to host cells is one of the earliest events during bacterial colonization of host tissues and thus a key step during infection. The ...

Legionella pneumophila Outer Membrane Vesicles: Isolation and Analysis of Their Pro-inflammatory Potential on Macrophages

Bacteria are able to secrete a variety of molecules via various secretory systems. Besides the secretion of molecules into the extracellular space or directly into another cell, Gram-negative bacteria can also form outer membrane vesicles (OMVs). These membrane vesicles can deliver their cargo over long distances, and the cargo is protected from degradation by proteases and nucleases. 
Legionella pneumophila (L. pneumophila) is an intracellular, Gram-negative pathogen that causes a severe form of pneumonia. In humans, it infects alveolar macrophages, where it blocks lysosomal degradation and forms a specialized replication vacuole. Moreover, L. pneumophila produces OMVs under various growth conditions. To understand the role of OMVs in the infection process of human macrophages, we set up a protocol to purify bacterial membrane vesicles from liquid culture. The method is based on differential ultracentrifugation. The enriched OMVs were subsequently analyzed with regard to their protein and lipopolysaccharide (LPS) amount and were then used for the treatment of a human monocytic cell line or murine bone marrow-derived macrophages. The pro-inflammatory responses of those cells were analyzed by enzyme-linked immunosorbent assay. Furthermore, alterations in a subsequent infection were analyzed. To this end, the bacterial replication of L. pneumophila in macrophages was studied by colony-forming unit assays. 
Here, we describe a detailed protocol for the purification of L. pneumophila OMVs from liquid culture by ultracentrifugation and for the downstream analysis of their pro-inflammatory potential on macrophages.

Legionella pneumophila Outer Membrane Vesicles: Isolation and Analysis of Their Pro-inflammatory Potential on Macrophages

Here, we describe the purification of Legionella pneumophila (L. pneumophila) outer membrane vesicles (OMVs) from liquid cultures. These purified vesicles are then used for the treatment of macrophages to analyze their pro-inflammatory potential.

Bacteria are able to secrete a variety of molecules via various secretory systems. Besides the secretion of molecules into the extracellular space or ...

A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment

Assessment of antibiotic action with new drug development directed towards anaerobic bacteria is difficult and technically demanding. To gain insight into possible MOA, morphologic changes associated with antibiotic exposure can be visualized using scanning electron microscopy (SEM). Integrating SEM imaging with traditional kill curves may improve our insight into drug action and advance the drug development process. To test this premise, kill curves and SEM studies were conducted using drugs with known but different MOA (vancomycin and metronidazole). C. difficile cells (R20291) were grown with or without the presence of antibiotic for up to 48 h. Throughout the 48 h interval, cells were collected at multiple time points to determine antibiotic efficacy and for imaging on the SEM. Consistent with previous reports, vancomycin and metronidazole had significant bactericidal activity following 24 h of treatment as measured by colony-forming unit (CFU) counting. Using SEM imaging we determined that metronidazole had significant effects on cell length (&#62; 50% reduction in cell length for each antibiotic; P&#60; 0.05) compared to controls and vancomycin. While the phenotypic response to drug treatment has not been documented previously in this manner, they are consistent with the drug&#39;s MOA demonstrating the versatility and reliability of the imaging and measurements and the application of this technique for other experimental compounds.

A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment

Antibiotic efficacy is most commonly determined by conducting killing kinetic studies and measuring colony forming units (CFUs). By integrating scanning electron microscopy (SEM) with these standard methods, we can distinguish the pharmacological effects of treatment between different antibiotics.

Assessment of antibiotic action with new drug development directed towards anaerobic bacteria is difficult and technically demanding. To gain insight into ...

Screening Assays to Characterize Novel Endothelial Regulators Involved in the Inflammatory Response

The endothelial layer is essential for maintaining homeostasis in the body by controlling many different functions. Regulation of the inflammatory response by the endothelial layer is crucial to efficiently fight against harmful inputs and aid in the recovery of damaged areas. When the endothelial cells are exposed to an inflammatory environment, such as the outer component of gram-negative bacteria membrane, lipopolysaccharide (LPS), they express soluble pro-inflammatory cytokines, such as Ccl5, Cxcl1 and Cxcl10, and trigger the activation of circulating leukocytes. In addition, the expression of adhesion molecules E-selectin, VCAM-1 and ICAM-1 on the endothelial surface enables the interaction and adhesion of the activated leukocytes to the endothelial layer, and eventually the extravasation towards the inflamed tissue. In this scenario, the endothelial function must be tightly regulated because excessive or defective activation in the leukocyte recruitment could lead to inflammatory-related disorders. Since many of these disorders do not have an effective treatment, novel strategies with a focus on the vascular layer must be investigated. We propose comprehensive assays that are useful to the search of novel endothelial regulators that modify leukocyte function. We analyze endothelial activation by using specific expression targets involved in leukocyte recruitment (such as, cytokines, chemokines, and adhesion molecules) with several techniques, including: real-time quantitative polymerase chain reaction (RT-qPCR), western-blot, flow cytometry and adhesion assays. These approaches determine endothelial function in the inflammatory context and are very useful to perform screening assays to characterize novel endothelial inflammatory regulators that are potentially valuable for designing new therapeutic strategies.

Vascular endothelium tightly controls leukocyte recruitment. Inadequate leukocyte extravasation contributes to human inflammatory diseases. Therefore, searching for novel regulatory elements of endothelial activation is necessary to design improved therapies for inflammatory disorders. Here, we describe a comprehensive methodology to characterize novel endothelial regulators that can modify leukocyte trafficking during inflammation.

The endothelial layer is essential for maintaining homeostasis in the body by controlling many different functions. Regulation of the inflammatory ...

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 ...

Evaluation of Extracellular Vesicle Function During Malaria Infection

Malaria is a life-threatening disease caused by Plasmodium parasites, with P. falciparum being the most prevalent on the African continent and responsible for most malaria-related deaths globally. Several factors including parasite sequestration in tissues, vascular dysfunction, and inflammatory responses influence the evolution of the disease in malaria-infected people. P. falciparum-infected red blood cells (iRBCs) release small extracellular vesicles (EVs) containing different kinds of cargo molecules that mediate pathogenesis and cellular communication between parasites and host. EVs are efficiently taken up by cells in which they modulate their function. Here we discuss strategies to address the role of EVs in parasite-host interactions. First, we describe a straightforward method for labeling and tracking EV internalization by endothelial cells, using a green cell linker dye. Second, we report a simple way to measure permeability across an endothelial cell monolayer by using a fluorescently labeled dextran. Finally, we show how to investigate the role of small non-coding RNA molecules in endothelial cell function.

In this work, we describe protocols to investigate the role of extracellular vesicles (EVs) released by Plasmodium falciparum infected erythrocytes. In particular, we focus on the interactions of EVs with endothelial cells.

Malaria is a life-threatening disease caused by Plasmodium parasites, with P. falciparum being the most prevalent on the African continent and responsible ...

Mouse Body Temperature Measurement Using Infrared Thermometer During Passive Systemic Anaphylaxis and Food Allergy Evaluation

Mouse body temperature measurement is of paramount importance for investigating allergies and anaphylactic symptoms. Rectal probes for temperature readings is common, and they have been proven to be accurate and invaluable in this regard. However, this method of temperature measurement requires the mice to be anesthetized in order to insert the probe without injury to the animal. This limits the ability to observe other phenotypes of the mouse simultaneously. In order to investigate other phenotypes while measuring temperatures, rectal probes are not ideal, and another method is desired. Here, we introduce a noninvasive method of temperature measurement that foregoes the requirement for mouse anesthesia while maintaining equal reliability to rectal probes in measuring body temperature. We use an infrared thermometer that detects body surface temperatures at ranges between 2 and 150 mm. This method of body temperature measurement is successful in reliably replicating temperature change trends during passive system anaphylaxis experiments in mice. We show that body surface temperatures are about 2.0 &#176;C lower than rectal probe measurements, but the degree of temperature drop follows the same trend. Furthermore, we use the same technique to observe mice in a food allergy model to evaluate temperature and activity levels simultaneously.

Here we present a new method to accurately measure body temperature differences in passive systemic anaphylaxis (PSA) and food allergy mouse models using an infrared thermometer. This procedure has been accurately duplicated in previous PSA results.