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A Leak-Free Head-Out Plethysmography System to Accurately Assess Lung Function in Mice

Mice are a valuable model for elegant studies of complex, systems-dependent diseases, including pulmonary diseases. Current tools to assess lung function in mice are either terminal or lack accuracy. We set out to develop a low-cost, accurate, head-out variable-pressure plethysmography system to allow for repeated, non-terminal measurements of lung function in mice. Current head-out plethysmography systems are limited by air leaks that prevent accurate measures of volume and flow. We designed an inflatable cuff that encompasses the mouse’s neck preventing air leak. We wrote corresponding software to collect and analyze the data, remove movement artifacts, and automatically calibrate each dataset. This software calculates inspiratory/expiratory volume, inspiratory/expiratory time, breaths per minute, enhanced pause, mid-expiratory flow, and end-inspiratory pause. To validate the use, we established our plethysmography system accurately measured tidal breathing, the bronchoconstrictive response to methacholine, sex and age associated changes in breathing, and breathing changes associated with house dust mite sensitization. Our estimates of volume, flow, and timing of breaths are in line with published estimates, we observed dose-dependent decreases in volume and flow in response to methacholine (P < 0.05), increased lung volume and decreased breathing rate with aging (P < 0.05), and that house dust mite sensitization decreased tidal volume and flow (P <0.05) while exacerbating the methacholine induced increases in inspiratory and expiratory time (P < 0.05). We describe an accurate, sensitive, low-cost, head-out plethysmography system that allows for longitudinal studies of pulmonary disease in mice. New & Noteworthy We describe a variable-pressure head-out plethysmography system that can be used to assess lung function in mice. A balloon cuff that inflates around the mouse’s neck prevents air leak, allowing for accurate measurements of lung volume and air flow. Custom software facilitates system calibration, removes movement artifacts, and eases data analysis. The system was validated by measuring tidal breathing, responses to methacholine, and changes associated with house dust mite sensitization, sex, and aging. Contributions to Study Stephanie Bruggink: development of head-out plethysmography chamber, measurement of breathing, data analysis, prepared manuscript Kyle Kentch: development of head-out plethysmography chamber, programmed software to collect and analyze data, prepared manuscript Jason Kronenfeld: development of tools to analyze data, analysis of data Benjamin Renquist: development of head-out plethysmography chamber, statistical analysis, prepared manuscript


Introduction
pressure change resulting from expansion of the ribcage is not negated by a simultaneous removal 111 of air from the chamber (17). Unlike the forced oscillation technique, head-out plethysmography 112 is non-terminal and performed in awake, conscious mice. In turn, head-out plethysmography 113 allows for repeated measurements of physiologically relevant breathing. All published and 114 commercial plethysmography chambers use a piece of latex (or other flexible plastic) to create an 115 airtight seal around the mouse's neck (17). In preliminary studies we found this set-up did not 116 create an airtight seal. Without an airtight seal, pressure-volume relationships to determine tidal 117 volume and flow are inaccurate. Accordingly, as described, head-out plethysmography can provide 118 accurate measures of the timing of breaths but cannot be applied to accurately assess either volume 119 or flow. 120 Double-chamber plethysmography utilizes a similar set-up to head-out plethysmography with 121 an additional chamber fit around the head or nose of the mouse. Pressure changes resulting from 122 nasal flow and thoracic movements are measured separately (18). Importantly, this allows for 123 measurement of specific airway resistance, assessed by the time delay between the nasal and 124 thoracic flows. The change in lung volume proceeds the change in nasal flow, creating a pressure 125 gradient to pull air into the lung. The delay between thoracic and nasal flows is short in healthy with holes that allow air to pass from the outside to inside of the ring to inflate the balloon cuff. balloon ends are wrapped over the outside of the ring and held in place with O-rings (Figure 1.6). 161 To allow inflation of the balloon, we use a needle (any size and brand) to perforate the balloon 162 where the balloon ends overlap and occlude the holes along the midline of this ring (Figure 1.6). 163 We created a small air port (hole created with needle) in the inner syringe barrel to inflate 164 the balloon cuff (Figure 1.2). Next, we placed the outer chamber over the inner chamber making 165 sure not to move the 3D printed spacers, a process facilitated with a small amount of petroleum 166 jelly. A female luer ( Figure 1H thread style to 500 series barb 1/16 inch ID tubing Cole-Parmer 167 Instrument #SK-45508-01 connected to tubing Picture 1B Silastic Laboratory Tubing #508-005) 168 was briefly heated to melt the plastic and this melted leur was placed over the top of the hole and 169 allowed to cool and adhere creating an air port onto which tubing could be attached. The tubing, 170 cut to ~22 cm, is then connected to another female luer and a stopcock ( Figure 1M Cole-Parmer 171 #30600-00). This allows for the balloon cuff on the inside of the inner chamber to be inflated by a 172 3 ml syringe from the stopcock. Since each mouse displaces different chamber volumes, the user must calibrate the volume-202 pressure relationship prior to each breathing measurement. We calibrate by rapidly injecting 203 known volumes of air that range from 0.05-0.4 ml (0.05, 0.1, 0.2, 0.3, and 0.4 ml), as the inspiratory 204 and expiratory volume is expected to be < 0.4 ml. We then quantify the pressure change associated 205 with injection of a given volume. The chamber pressure is reset to atmospheric pressure by 206 opening the 3-way stopcock, or simply removing the syringe from the stopcock, allowing room air 207 to flow into the air port between each injection. To ensure precise calibration, each volume is 208 injected 4 times. The software has a built-in annotation marker, allowing us to log the injection 209 volume and timing (Figure 2A, C). Automated calibration software (use described below) allows 210 the user to enumerate the pressure-volume relationship by regressing the change in pressure on 211 volume injected and fitting an equation that will be used to extrapolate all changes in chamber 212 pressure to changes in chamber volume associated with breathing ( Figure 1.10).

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After calibration, breathing measurements are initiated. We waited at least 5 minutes for 215 the mice to become accustomed to the chamber before collecting data, which was about the time 216 it took to perform the calibration. When collecting only baseline measurements, we collected 10 217 minutes of tidal breathing. In studies to understand the response to methacholine, we collected a 218 5-minute baseline measurement followed by exposure to nebulized methacholine (MP 219 Biomedicals, LLC). As multiple measurements were completed at different concentrations of 220 methacholine, we averaged the 5-minute baseline measurements for each mouse to report tidal 221 breathing. We exposed the mice to methacholine by placing the end of the neubulizer (Aeroneb 222 lab nebulizer small unit, Kent Scientific) directly over the mouse's head to ensure delivery into the airway. We delivered methacholine (dissolved in PBS) for 30 seconds at 0.191 ml/30 seconds at 224 concentrations of 0, 25, 50, and 100 mg/ml of methacholine. Breathing was then measured for 10 225 minutes after the conclusion of methacholine administration. When completed, the balloon cuff 226 was deflated then the syringe plunger was removed allowing the mouse to back out of the chamber.

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Mice were allowed to fully recover after each dose of methacholine by allowing at least 1 hour 228 between measurements. Time course data is presented as 1-minute rolling averages taken every 30 229 seconds, while baseline data is presented as a 5-minute average. When applicable, we normalized 230 to baseline values and present the data as change from baseline.

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The user interface for data collection is simple (Figure 2A). Upon opening the program, 248 the identifier for the mouse and measurement identifier was entered into CH1 or CH2 Mouse ID 249 and measurement initiated by selecting Start CH1/CH2. For calibration, the volume injected was 250 written into CH1/CH2 Calibration Volume, the volume was injected, and LOG was selected.

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Selecting LOG marked that timepoint with the entered volume (shown in Figure 2C). Of note, this

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To the smoothed pressure signal, we then applied a moving step fit algorithm in each region 281 of interest (20). The published method was modified to optimize volume-pressure correlation and 282 use the most datapoints to develop this correlation. To account for differences in signal-to-noise 283 ratio, the half-window size, w, varies with calibration injection volume, v, such that = 200 ⁄ .

284
Instead of a first-degree polynomial, we use a zero-degree polynomial for the fitting functions, 285 under the assumption that the slope before and after a calibration step is zero. After obtaining 286 (the indicator for the probability of datapoint, i, to be a potential step position) a step height 287 threshold is selected at 0.25 standard deviations from = 0. Datapoints outside this threshold 288 represent step regions and datapoints within the threshold represent plateau regions ( Figure 2C).

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To be used in the analysis, a region of interest must consist of an initial baseline plateau followed by an upward step to a calibration plateau containing the calibration injection timepoint.

291
Furthermore, each plateau must exceed a step width threshold of 0.5 seconds. The change in 292 pressure for a calibration injection is calculated as the difference between the mean unsmoothed 293 pressure signal values of the calibration and baseline plateaus.

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The pressure changes of the calibration injections were matched with known injection 295 volumes and a linear regression was used to determine the relationship between pressure and 296 volume ( Figure 1.10). Where Vinjection is the injection volume, and m is the slope. Given that there 297 is no change in pressure without a change in volume, we set the ΔP-intercept (y-intercept) to 0.
To ensure accuracy of the calibration, all injection volumes were repeated in quadruplicate 300 and any points with a residual value exceeding one standard deviation away from the mean residual 301 value were excluded.

302
A calibration injection causes an equivalent decrease in chamber volume (Vinjection= -303 ΔVchamber). Injection volume is substituted by change in chamber volume to calculate the mouse's 304 respiration signal: For our analyses, we report change in plethysmography chamber volume as a proxy for changes 307 lung volume.

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We used a Gaussian filter (sigma of 5 datapoints) to reduce noise present in the volume 309 signal. This signal was then detrended to correct for any global drift in chamber pressure by fitting

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To determine the presence of pauses in breathing between inspiratory and expiratory 327 phases (end-inspiratory pause) we again adopted a previously described algorithm (21). Since we 328 were primarily interested in pauses occurring between the inhale and exhale phases, and not those 329 occurring midway through one of these phases, we limited the search for pauses to those that 330 occurred within the first 25% of volume increase (expiration). For each breath, the airway flow 331 signal between the breath's peak and trough was used to identify the end of inhalation, the pause 332 after inhalation, and the onset of exhalation.  based on residuals from a known and calculated volume and our known volume (as measured 374 using a 1 ml syringe) likely had error greater than 10 µl, this is likely an underestimate of our 375 sensitivity using this system.

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The software for calibration and analysis are a single program 377 (https://github.com/bjrenquist/plethysmography; Figure 2B). We stopped the recording after was assessed using a two-way ANOVA. Differences between means were corrected for multiple 405 comparisons using a Tukey's adjustment.

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Chamber validation 408 To validate our system, we measured tidal breathing in lean adult male mice. We report  Table 2).

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We applied our head-out plethysmography system to measure breathing changes caused

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We further validated the use of our system to measure tidal breathing changes associated 432 with sex and aging. Inspiratory (Male P = 0.0002; Female P = 0.0016; Figure 6A) and expiratory 433 (Male P = 0.0026; Female P = 0.0102; Figure 6B) volume, inspiratory time (Male P = 0.0071;

434
Female P = 0.0007; Figure 6C), and Penh (Male P < 0.0001; Female P = 0.0565; Figure 6F) 435 increased with age in both male and female mice. Mid-expiratory flow increased with age in 436 male mice (P = 0.0003; Figure 6G) and tended to increase with age in female mice (P = 0.11; 437 Figure 6G). End-inspiratory pause (P = 0.016; Figure 6H) and end-inspiratory pause and  to ensure a more accurate time-dependent model. 499 We report greater amounts of bronchoconstriction (both percent change and 500 minimum/maximum percent change) at lower methacholine doses than previously reported in 501 studies using whole-body plethysmography (gray Supplementary better software capturing data for each breath, and/or enhanced system sensitivity. We aerosolized 508 methacholine directly onto the face of the mouse instead of into a whole-body plethysmography 509 chamber. In addition, we utilized a nebulizer that produces small particle size (4.0-6.0 µm) which 510 are more easily delivered further into the lung. Finally, our airtight chamber allows for accurate 511 determination of changes in volume and flow.  Construction of a leak proof, variable-pressure head-out plethysmography chamber that can be calibrated. 1) Head-out plethysmography chamber materials. A. Plunger fitting in 60 ml syringe (L) to prevent mouse from backing out of chamber B. Tubing to create balloon cuff air port C. Tubing to create calibration air port D. 300 ml outer syringe E. Two O-rings to secure balloon to 3-D printed ring for balloon cuff (J) F. Four inner O-rings to secure 3D printed spacer (into I) G. Two outer O-rings to secure 3D printed spacer (into I) H. Female luers to create air port and attach tubing to stopcocks (M) I. Two 3D printed spacers J. 3D printed ring for balloon cuff K. Two options for balloons to create balloon cuff L. Inner chamber 60 ml syringe M. Stopcocks to allow air flow into balloon cuff and out chamber 2) Inner chamber with 3D printed spacers and O-rings fitted into grooves, picturing creation of hole for balloon port 3) Preparation of balloon cuff to be placed directly under hole (created in 2) 4) Pressure transducer with tip removed from one end and small piece of plastic tubing to place over the tip 5) Inner chamber complete with holes for adequate assessment of pressure changes by pressure transducer 6) Sample of how to perforate balloon cuff to allow inflation 7) Complete set-up with inner and outer chamber, balloon cuff, pressure transducer, and ports to allow for airflow into the chamber or into the balloon 8) mouse in plethysmography chamber 9) Sample pressure trace with breaths signified in black and the red line indicating average pressure which shifts after injection of a calibrating quantity of air at 63 seconds. 10) Sample of calibration regression lines created for 3 mice (pink, marine, and black) with automatically excluded points in corresponding lighter color.

Figure 5.
Minimal or maximal 1 minute rolling average over the 10 minutes after expose to methacholine (0, 25, 50, or 100 mg/ml exposure for 30 seconds nebulized at flow rate of 0.191 ml/30 sec) before and after sensitization with house dust mite. *Indicates significant differences (P < 0.05) before (white) and after (gray) house dust mite exposure. Data are presented as mean + SEM (n = 8-9). . Tidal breathing in male and female mice aged 6 months (mo.) and 18 mo. Data are presented as mean ± SEM . a,b,c bars that do not share a common letter differ significantly (P < 0.05; n =14-30).