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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Hyperpolarized xenon MRI can quantify regional lung microstructure (air-space dimensions) and physiology (ventilation and gas exchange) in translational research and clinical care. Although challenging, it can provide comparable pulmonary insights in preclinical studies. This protocol describes the infrastructure and procedures needed to perform routine xenon lung MRI in mice.

Abstract

Hyperpolarized (HP) xenon-129 (129Xe) is an inhaled magnetic resonance imaging (MRI) contrast agent with unique spectral and physical properties that can be exploited to quantify pulmonary physiology, including ventilation, restricted diffusion (alveolar-airspace size), and gas exchange. In humans, it has been used to evaluate disease severity and progression in a variety of pulmonary disorders and is approved for clinical use in the United States and United Kingdom. Beyond its clinical applications, the ability of 129Xe MRI to noninvasively assess pulmonary pathophysiology and provide spatially resolved information is valuable for preclinical research. Among animal models, mice are the most widely used due to the accessibility of genetically modified disease models. Here, 129Xe MRI is promising as a minimally invasive, radiation-free, and sensitive technique to longitudinally monitor lung disease progression and therapy response (e.g., in drug discovery). This technique can extend to preclinical applications by incorporating an MRI-triggered, free-breathing apparatus or mechanical ventilator to deliver gas. Here, we describe the steps and provide checklists to ensure robust data collection and analysis, including creating a thermally polarized xenon gas phantom for quality control, optimizing polarization, animal handling (sedation, intubation, ventilation, and care for mice), and protocols for ventilation, restricted diffusion, and gas exchange data. While preclinical 129Xe MRI can be applied in various animal models (e.g., rats, pigs, sheep), this protocol focuses on mice due to the challenges posed by their small anatomy, which are balanced by their affordability and the availability of many disease models.

Introduction

While pulmonary disorders remain the leading causes of global morbidity and mortality1, the last decade has seen dramatic improvements in patient outcomes. These improvements are driven in part by two factors. Firstly, Phase III clinical trials now prioritize changes in lung function as endpoints rather than mortality, accelerating drug trials2,3,4,5. Secondly, advancements in improved animal models have provided insights into disease mechanisms and aided therapy development6,7. Mouse models are often favored for translational research because they offer physiological parallels to humans, affordability, and rapid disease development. Genetic engineering has expanded the range and quality of available models, with the International Mouse Strain Resource now boasting over 32,000 mouse strains8, compared to only 4,218 rat strains (Rat Genome Database9). These models have opened new avenues for investigating mechanistic drivers and therapy responses for a range of lung diseases, including chronic obstructive pulmonary disease (COPD)10, cystic fibrosis (CF)11, pulmonary fibrosis12,13, pulmonary hypertension14,15, and asthma16.

Unfortunately, lung research involving mice is limited by the techniques available to quantify disease burden. Studies often rely on terminal procedures that 1) provide whole-lung information (biochemical assays) or localized information (histology) and 2) demand cross-sectional designs and large sample sizes. Thus, they capture neither spatial nor temporal disease dynamics. In contrast, non-invasive, three-dimensional imaging can assess structure, molecular processes, and function in the lungs over time.

Lung structure (e.g., airway abnormalities and interstitial fibrosis) can be visualized with ultra-short echo-time (UTE) MRI and microcomputed tomography (Β΅CT) at high resolution. Functional and mechanistic information (e.g., ventilation, perfusion, tumor metabolism, and inflammatory processes) can be obtained with exogenous contrast agents (e.g., xenon-enhanced CT and oxygen-enhanced UTE) and ionizing nuclear medicine approaches (i.e., positron emission tomography [PET], and single-photon emission computed tomography [SPECT]). However, functional imaging is challenging due to the modest contrast-to-noise (particularly for oxygen-enhanced UTE at the high magnetic field strengths used for preclinical MRI, where T1 is lengthened) available without employing ionizing modalities with higher than normal levels of radiation. While imaging with these modalities is well tolerated in animal models using conventional doses, cumulative radiation may confound results in studies on immunology, inflammation, and lung cancer17. However, hyperpolarized (HP) xenon-129 (129Xe) magnetic resonance imaging (MRI) provides minimally invasive, non-irradiating, and highly sensitive structural and functional information. While this technique has been employed in preclinical research to characterize conditions including emphysema18,19, fibrosis20, lung cancer21, COPD22, and radiation-induced lung injury23 at single or multiple time points, it remains underutilized in the preclinical setting.

To enable routine, preclinical 129Xe MRI, several prerequisites are required, including institutional regulatory support, a hyperpolarization device, aΒ 129Xe-tuned radiofrequency (RF) coil, and a multi-nuclear-capable scanner. Although advanced applications24,25,26,27,28,29,30,31,32,33 require vendor-specific pulse programming that is outside of the scope of this protocol, basic applications can be achieved with modest software modifications. Therefore, we focus on quality control, magnetization handling, data collection, and animal handling procedures β€” including mechanical ventilation β€” that are unique toΒ preclinicalΒ 129Xe MRI (Figure 1).

To date, small animal 129Xe imaging has employed three MR-safe gas delivery approaches, each with advantages and disadvantages: free-breathing, piston-driven, and pressure-drop. Free-breathing allows spontaneous inhalation without risk of injury from intubation or tracheostomy but consumes significantly more HP gas and can introduce motion artifacts34,35. Commercial piston-driven devices are self-calibrating and easy to use out-of-the-box but may be prohibitively expensive36. The pressure-drop-based approach used here is well described in the literature, modular, customizable, and run by open-source code37,38,39,40. Furthermore, it is cost-effective, typically totaling less than $10k and a few weeks of dedicated build time. The pressure-drop ventilator delivers 129Xe from a dose bag within a pressurized cannister while monitoring the airway pressure of an intubated mouse.

figure-introduction-6202
Figure 1: Overview of the protocol to collect routine xenon-129 (129Xe) magnetic resonance imaging (MRI) in mice. (A) Steps for initial setup. (Note: scanner programming is unique to each vendor and not described in this protocol). (B) Steps to collect daily quality assurance (QA) and animal data. (C)Β Steps for successful experiment conclusion and data analysis. Please click here to view a larger version of this figure.

Here, we collect and analyze the three common classes of 129Xe MRI data: ventilation, diffusion-weighted imaging (alveolar-airspace size), and gas exchange. Ventilation images depict the distribution of inhaled 129Xe gas. Regions of the lungs with reduced airflow appear dark in HP gas images, and pathology is quantified by the volume of defective ventilation. In humans, the ventilation defect percentage (VDP) has shown strong repeatability41,42 and high sensitivity to lung obstruction in diseases like COPD43,44,45 and asthma46,47.

The restricted diffusion of the 129Xe atoms in the airspace can be measured via the apparent diffusion coefficient (ADC) and serves as a surrogate for air-space size. The ADC is calculated by acquiring a baseline image (b0) without diffusion weighting and one or more images acquired in the presence of bipolar gradient-induced diffusion weighting (bN). An elevated ADC reflects an increaseΒ in airspace size due to aging or emphysematous remodeling18,48. Further, using multiple b-value images (β‰₯4) allows more detailed morphometric information (e.g., mean linear intercept) to be calculated49,50.

Gas exchange can be characterized due to 1) the solubility of 129Xe in the capillary membrane tissue, plasma, and RBCs (red blood cells) and 2) the >200 ppm downfield chemical shift of 129Xe when dissolved in these compartments. Both spectroscopic and imaging data provide insight into cardiopulmonary diseases (e.g., pulmonary hypertension and left heart failure51,52,53). While many species (humans, canines, and rats) display unique spectral peaks originating from each compartment, mice lack a unique RBC signal due to differences in hemoglobin-xenon binding site interactions. Instead, all dissolved components are combined into a single signal in mice54. However, it is possible to observe a distinct RBC resonance in transgenic mice expressing human hemoglobin, such as those used in models of sickle cell disease54. Overall, dissolved 129Xe spectroscopy and imaging provide unique insights into cardiopulmonary pathophysiology in mice55,56.

Before attempting this protocol, it is necessary to understand background information about the MRI scanner, mechanical ventilation, and mouse handling techniques required for mouse studies. Prior to initiating animal studies, all procedures must be approved by the local Institutional Animal Care and Use Committee (IACUC)57. Because the total magnetic moment available in the mouse lung is intrinsically low (i.e., tidal volume ~250 Β΅L), voxel size must be 1000-fold smaller than in humans to achieve anatomically equivalent resolution. The murine breathing rate is also exceedingly rapid (>100 breaths/minute). As such, the single-breath-hold procedures typically used for human imaging are not feasible. Instead, only a few RF excitations can be applied within each breath, so 129Xe images must be encoded over tens to hundreds of breaths. Pulse programming may be required to permit external triggering of acquisitions and to properly loop slices, phase encodes, and/or diffusion-weighted images while balancing signal-to-noise ratio (SNR), resolution, and scan duration.Β Here, the ventilator outputs a transistor-transistor logic (TTL) pulse once per breath to trigger data acquisition (Figure 2).

figure-introduction-11108
Figure 2: Representative mechanical ventilation and data acquisition timing. (A) User-controlled ventilation can trigger data acquisition at end-inspiration, during the breath hold, or at end-expiration. (B) For this 3D radial ventilation sequence, the user defines the total number of projections acquired and the number of projections per breath. (C) For a slice-selective, 2D diffusion-weighted image, the user defines the order of slices, b-value images, and phase encodes. Please click here to view a larger version of this figure.

To enable reliable ventilation andΒ 129Xe delivery, robust sedation and intubation procedures are required. For each study, the downstream effects of each anesthetic must be considered - including changes to minute ventilation, heart rate (HR), and blood pressure58,59,60,61,62,63,64,65,66. While a variety of sedatives have been used for preclinical HP gas MRI, we employ a mixture of ketamine, xylazine, and acepromazine, due to its availability, cost-effectiveness, reliability, and duration67,68. Once sedated, animals must be intubated for effective mechanical ventilation. Intubation of mice is difficult due to the small size of their anatomy, and thus, it is important to train thoroughly in this technique. We encourage investigators to review published video protocols69,70. Because most commercial intubation cannulas contain stainless steel, we introduce a technique to craft metal-free (i.e., MRI and HP-gas compatible), wedge-shaped cannulas that can be customized to match airway diameter to create an airtight seal with the mouse tracheal wall.

Because 129Xe images are collected over many breaths, ventilator settings are critical. Protective ventilation strategies must be carefully considered to prevent lung injury71,72,73,74. In particular, the use of low tidal volume (TV), moderate positive end-expiratory pressure (PEEP), and alveolar recruitment maneuvers (RMs) reduce the risk of ventilator-induced lung injury in human patients and animal models75,76,77,78,79,80,81. Here, we recommend a simple technique that is compatible with pressure-drop 129Xe mechanical ventilation that is protective and provides sufficient 129Xe image SNR. Specifically, we apply PEEP by adding a commercial PEEP valve to the exhale line of the ventilator. To perform RMs, the exhalation line must be closed so that the animal receives multiple inhalations without exhalation until a target pressure and duration have been reached.

Throughout, we provide general ventilation settings, but it is advised to review the literature to address specific study goals82,83. In addition to monitoring the peak inspiratory pressure during mechanical ventilation,Β it is important to monitor the animal's temperature, which can be done using standard mouse temperature monitoring methods. While not required for imaging, monitoring heart rate via electrocardiogram (ECG) can be advantageous; ECG can indicate if an animal is waking from sedation, overdosed, or distressed, allowing the researcher to intervene.

The protocol we describe is designed to collect 129Xe 3D radial ventilation data61, 2D GRE diffusion-weighted data76, and dynamic pulse-acquire spectroscopy gas exchange data. This protocol aims to bridge the gap between preclinical research in small animal models and the potential for 129Xe MRI to advance our understanding of pulmonary disorders.

Protocol

All methods described here were approved by the Institutional Animal Care and Use Committee (IACUC) of Cincinnati Children's Hospital Medical Center.

1. Initial site preparation

  1. Create and test a thermally polarized 129Xe gas phantom (Figure 3).
    1. Obtain a borosilicate glass vessel (~60 mL), a plunger valve with a front-sealing O-ring, and a ground borosilicate glass stem, all rated up to 150 psig. Ensure there are no magnetic parts. Attach a compression fitting to the glass stem. Tighten to produce a gas-tight seal.
    2. Connect the vessel to a vacuum pump and oxygen reservoir according to Figure 3A. Vacuum vessel to less than 100 mTorr absolute pressure.
    3. Fill the vessel with oxygen to the pressure of 1.5 atm to reduce the 129Xe T1 from > 30 min to ~2 s (at 7 T magnetic field strength; For a 9.4 T scanner, use 1.6 atm oxygen). Seal vessel.
      NOTE: For higher field strengths, somewhat higher oxygen partial pressure will be required to achieve T1≀ 2 s84.
    4. Fill a gas-impermeable reservoir with 400 mL of isotopically enriched xenon (85% 129Xe).
      NOTE: Natural abundance xenon (26% 129Xe) can also be used, but signal taveraging will need to be increased to mitigate the ~3-fold reduction in SNR.
    5. Connect the vessel to the reservoir of 129Xe. Vacuum tubing to less than 100 mTorr absolute pressure.
    6. Fill an open-mouth, benchtop liquid N2 Dewar to ~90%. Submerge the bottom of the vessel (~5 cm) in liquid nitrogen to condense O2 and create a vacuum (Figure 3B). While submerged, open the valve to allow 129Xe from the reservoir to flow into the vessel (Figure 3C).
    7. Seal the plunger valve by slowly pulling the stem until the inflow hole in the plunger is pulled past the O-ring. Immediately after the hole has passed the O-ring, hand tighten to seal the vessel. Remove the vessel from liquid nitrogen and allow it to thaw.
      NOTE: Once thawed, the vessel will pressurize to ~4.5 atm (2 atm O2 + 2.5 atm 129Xe).
    8. Protect glassware (e.g., insert vessel in padded polyvinyl chloride (PVC) pipe container, Figure 3D).
      NOTE: If properly maintained, the phantom can maintain pressure for a decade or longer.
    9. Measure the T1 of the phantom (e.g., using a spectroscopic inversion recovery sequence). Confirm T1 < 2 s for 7 T scanners. Track signal and T1 over time for quality assurance (QA).

figure-protocol-2945
Figure 3: Creation of a thermally polarized 129Xe gas phantom guided by the protocol detailed in StepΒ 1.1. The O2 and 129Xe partial pressures can be altered to customize the T1 to yield appropriate 129Xe T1 times and signal strength at a given field strength84. Please click here to view a larger version of this figure.

  1. Perform quality assurance with the thermally polarized gas phantom (Table 1Β andΒ Figure 4).
    1. Place the 129Xe coil at the isocenter of the magnet and center the 129Xe gas phantom within the coil. In a single pulse-and-acquire sequence ("single pulse"), set the working frequency to match the approximate 129Xe gas frequency in the phantom (~83.07 MHz at 7T).
    2. Center the acquisition and excitation frequencies to the 129Xe Larmor frequency, and use this frequency for all phantom calibration and quality control 129Xe scans. See Table 1 for experimental parameters for all QA scans. Confirm that the phantom is centered with a phantom localizer (Figure 4A).
    3. Perform nutation experiment to calibrate flip angle: Assuming SNR is sufficient, use single RF pulses with repetition time (TR) spacing > 5 x T1. For each acquisition, incrementally increase RF power until the signal nulls and begins to invert. The standard used here is: number of pulses = 65; TR = 10 s; pulse duration = 125 Β΅s; RF power = 1-13.8 W, incremented by 0.2 W
    4. Fourier transform and phase the first spectrum (i.e., the spectrum acquired with the lowest RF power). Apply the same phasing for all spectra. Plot the real spectra as a function of the RF pulse power (Figure 4B).
    5. The power that produces a null peak (i.e., minimum peak height) corresponds to the 180° flip angle. Achieve a 90˚ flip angle by using the same power at half the pulse length needed to produce the 180˚ flip angle. Assuming the scanner software allows, set this 90° reference power and pulse length for subsequent flip angle scaling.
    6. Use a single pulse to shim by minimizing the full width at half maximum of the 129Xe spectrum (TR ~ 1 s). If necessary, recenter the frequency after shimming. Record the full-width half max.
    7. Run the 129Xe QA scan (Table 1 and Figure 4C). Record QA data: SNR, mean phantom signal, and standard deviation of the noise.

figure-protocol-5999
Figure 4: Pre-scan quality assurance. (A) A low-resolution 2D GRE coronal phantom localizer ensures the phantom is centered in the magnet. (B) A nutation experiment to set a 90˚ pulse shows a null peak at the 180° pulse. (C) After localizing and calibrating the flip angle, acquire a higher resolution 2D GRE QA image. Please click here to view a larger version of this figure.

Protocol Short-Hand NameSequence DescriptionTR (ms)TE (ms)Averages / RepetitionsFlip Angle (˚)Matrix Size or NptsFOV (mm2)RF BW (kHz)Slice / Slab Thickness (mm)Scan Duration
Single pulsePulse acquire10001 / 1602048101 s
Phantom localizer2D GRE2003.720 / 14860 Γ— 32120 Γ— 483602 min
Flip angle calibrationPulse acquire70001 / 652020485.127.5 min
129Xe QAΒ 2D GRE50003.38 / 19032232234021 min

Table 1: Phantom calibration quality assurance sequence parameters. TR = repetition time, TE = echo time, Npts = number of points, FOV = field of view, BW = bandwidth. Please click here to download this Table.

  1. Plan polarization (Figure 5A, B).
    1. Select polarized 129Xe volume and accumulation time: 400 mL in 15 min is optimal for this protocol (Figure 5) but can easily be adjusted for other applications and equipment.
    2. Assuming a known dead volume within the hyperpolarizer (e.g., Vdead vol = 80 mL), calculate the flow rate (FXe in SLM) for a dispensed volume Vdesired, 129Xe gas fraction f, and accumulation time tacc:
      figure-protocol-12148Β  Β  Β (1)
      NOTE: While longer production times will typically yield higher polarization, they may not be practical for in vivo imaging. Use an appropriate model for the hyperpolarizer85,86,87,88 to determine a flow rate that will balance production time and polarization. Here, the model of J. W. PlummerΒ et al.89 (Figure 5A) was used. This applies to continuous-flow polarizers and is not applicable to stopped-flow hyperpolarizers90.
    3. Polarize gas according to these parameters, measure the polarization with a commercial or homebuilt device and compare it to the predicted polarization for QA.
    4. Measure the polarization loss during transportation. If polarization decreases by a sufficiently large amount (e.g., >10%), build a magnetic carrying case to protect polarization during transportation. See Supplementary File 1: Managing polarization during transportation and Supplementary Figure 1.

figure-protocol-13533
Figure 5:Β Polarization management. (A)Β Polarization and produced volume are a function of accumulation time and flow rate. A 400 mL bag of gas provides high initial polarization (~35%) over 20 min. While using 1 L of gas might seem attractive, it will have a lower initial polarization (~20%). (B) After ~15 minutes of ventilation, a 1 liter batch of HP 129Xe would deplete to <10% polarization while 600 mL of gas remained116. Thus, using multiple 400 mL bags of 129Xe maintains higher average delivered polarization. C) Locations where the primary field and active shielding field intersect (red box at position (N,N,N)) can cause rapid relaxation of HP 129Xe. Characterizing the magnet's fringe field helps identify safe zones where reservoirs of HP 129Xe can be placed without rapid relaxation (green box at position (0,0,n)). Please click here to view a larger version of this figure.

Supplementary File 1: Managing polarization during transportation. Please click here to download this File.

  1. Measure the position-dependent T1 of HP 129Xe within the fringe field (Figure 5C).
    1. Create reference points with known distances and positions relative to the magnet isocenter along the X, Y, and Z dimensions. Label isocenter and label the other positions n through N. The number of positions to investigate will depend on the space available.
    2. Hyperpolarize a small volume of 129Xe (~250 mL) and transport it to the MRI control room. Fill a large syringe (50-100 mL) with 129Xe and place it at the isocenter within the magnet (Position 1 in Figure 5C). Play out a ~1˚ flip angle pulse to measure the signal.
    3. Leave the syringe in position for ~10 min, then acquire another spectrum. Remeasure the signal every 10 min until the signal has decayed at least 1 T1 (i.e., the signal has decayed to ~1/3rd its initial value).
    4. Start a new T1 experiment with a new syringe of 129Xe by repeating StepΒ 1.4.2.
    5. Move the syringe to a new position (e.g., Position n in Figure 5C) and leave it there for 10 min. Return the syringe to the isocenter to acquire an additional ~1˚ flip angle spectrum.
    6. Repeat this process: move the syringe to Position n, wait for 10 min, put it back to the isocenter, and remeasure the signal until it has decayed at least 1 T1.
    7. Repeat StepsΒ 1.4.4 - 1.4.6 for the remaining reference positions covering the X, Y, and Z directions.
    8. The initial signal (S0) will decay monoexponentially over n RF pulses with aΒ ΞΈ flip angle. Fit the signal (S) as a function of time (t) in the fringe field to calculate the T1 in each position:
      figure-protocol-16988Β  Β Β  (2)
    9. Identify a position with sufficient T1 (>20 min) for 129Xe reservoir placement.
  2. Create metal-free intubation cannulas (Figure 6).
    1. Obtain two veinous indwelling polytetrafluoroethylene (PTFE) catheters with Luer connectors. Discard needles into a sharps disposal container.
      ​NOTE: For mice >25 g, use 18 G and 20 G catheters. For smaller animals, use 20 G and 22 G catheters.
    2. Cut the Luer connector off the catheters. Feed the smaller catheter into the upper end of the larger catheter to create a more sharply tapered and longer cannula. Cut the composite cannula to ~4.6 cm with a beveled end,Β not including the Luer base (Figure 6A,Β B).
    3. Cut the wider end off of a 200 Β΅L pipette tip to a length of ~2.6 cm (Figure 6C).
    4. Coat the inside of the pipette tip with general mold release lubricant. Use another pipette tip inserted inside to thinly distribute the lubricant. Fill the pipette tip with acetoxy silicone vulcanizing paste epoxy (Figure 6D, E).
    5. Plug the cannula with 22 G wire extending out of both ends.Β Feed the cannula tube through silicone in the pipette tip. Extend the tube ~7 mm past the end of the pipette tip (Figure 6F, G).
    6. Slide the cannula tube on the wider side of the pipette tip through a plastic male slip Luer connector, gluing the pieces together with the epoxy. Trim tubing that extends past the Luer connector (Figure 6H).
    7. Wait for the epoxy to dry (>24 h), then carefully remove the silicone intubation cannula from the pipette tip mold. Remove the wire from the cannula, ensuring the tube has not been occluded (Figure 6I).
    8. To make a handle for easy intubation, connect tubing (1/16" or 1/8") to a female Luer connector. When ready for intubation, connect this female Luer connector to the male Luer intubation cannula. This piece can be easily detached post-intubation (Figure 6J).
    9. Sanitize before each use on animals: wipe the outside of the cannula with 70% alcohol. Wipe 20 G wire with sanitizer then feed the wire through the cannula to sanitize the inside and ensure no blockages.

figure-protocol-19712
Figure 6:Β Creating MRI and HP 129Xe compatible mouse intubation cannulas. These cannulas are constructed of venous catheters, pipette tips, and silicon sealant, as described in StepΒ 1.5. Please click here to view a larger version of this figure.

2. Daily data collection

NOTE: See Supplementary File 2: Preclinical scan QA checklist.

Supplementary File 2: Preclinical scan QA checklist. Please click here to download this File.

  1. Complete daily scanner quality control and setup ventilator (Figure 4).
    1. With phantoms, run QA protocols on the scanner (see Table 1 for QA scan parameters and StepΒ 1.2 for daily quality control steps).
    2. Calibrate Ventilator according to the method of J. Nouls et al.38. See Supplementary File 3: Ventilator calibration, Supplementary Figure 2, and Supplementary Figure 3.
    3. Set the ventilator settings for imaging at end-inspiration (Table 2). Place the animal bed on the scanner rack and the life support module (i.e., mechanical ventilator parts) on the table next to the scanner.
    4. Activate the site-specific animal heating system. Set a heater to 35.5 - 40 Β°C, turn on circulating air, and place the air hose within ~5" of where the animal's head will rest to pre-warm the bore of the scanner.
Ventilation SettingRecommendation for HP 129Xe MRINotes
Tidal volume (TV)8–10 mL/kg of ideal body weightModerate TV; low TV requires higher BR which may cause motion artifacts in images
Positive end-expiratory pressure (PEEP)2–6 cmH2O
Breath rate (BR)80–120 br/min
Recruitment maneuvers (RMs)~35 cmH2O for 6 s every 5 min
Ventilation duration; Position< 6 h; supineSupine to better see chest motion
Fraction of inspired oxygen (FIO2)0.3–0.5Prevent hypoxia in anesthetized mice
Inspiratory to expiratory ratio (I:E)1:2–1:4
Inspiratory to total cycle duration0.2–0.4
Minute ventilationβ‰₯0.57 mLΒ·g-1Β·min-1
Our standards:
BR = 80 br/min, inspiration duration = 200 ms, FIO2 = 0.3
Imaging at end-inspiration: breath hold = 200 ms, trigger delay = 200 ms after start of inspiration
Imaging during breath hold: breath hold = 250 ms, trigger delay = 250 ms after start of inspiration
Imaging at end-expiration: breath hold = 200 ms, trigger delay = 650 ms after start of inspiration

Table 2: Recommended ventilator settings for 129Xe imaging. Parameters can be fine-tuned for specific study objectives and experimental conditions117,118,119,120,121,122,123,124. Please click here to download this Table.

Supplementary File 3: Ventilator calibration. Please click here to download this File.

  1. Sedate and intubate animal.
    1. Turn on the incubator to 27.7 Β°C and/or the electric heating pad to 37.7 Β°C. Measure and record the body mass of the animal. Calculate sedative dosing based on mass. See Table 3 for a typical dosing regimen.
    2. Inject the sedative intraperitoneally. Note the time of injection and set the timer for the next dose of sedative.
      NOTE: Complete the remaining steps in SectionΒ 2 (Daily data collection) as quickly as possible to minimize the time under sedation and the risk of overdose.
    3. Apply eye lubricant to theΒ animal's eyes and place the animal in a cage on the heating pad or within an incubator to prevent hypothermia.
    4. Confirm the animal is fully sedated by performing a toe pinch test 10-15 min after sedative injection68. Intubate following procedures outlined in Das et al.69.
      NOTE: The article by Das et al.69 is accompanied by a comprehensive video demonstration of the technique. The steps are as follows:
    5. Hang the animal supine by the teeth on a slanted board. Use a rodent tongue depressor to pull out the tongue.
    6. To ensure vocal cords are visible, supply white light via a fiber optic cable within the intubation cannula or a bright light placed on the outside of the throat. Insert the cannula less than 5 mm past the vocal cords.
    7. Ensure the cannula is in the trachea, not the esophagus, by connecting it to a piece of tubing with a small droplet of water inside. If the water droplet moves in time with the animal's breaths, the positioning is likely correct.

AgentDoseRouteDurationComments
Inhaled Agents
IsofluraneInduction: 4%–5%
Maintenance: 1%–
3% or to effect		InhaledDuring continuous flowβ€’ Requires use of calibrated vaporizer
Injectable Agents
Recommended: Ketamine + xylazine + acepromazine90 + 9 + 3 mg/kgIntraperitoneal20–60 minβ€’ Creates susceptibility to hypothermia
β€’ For repeated dosing, it is recommended to switch to a ketamine + xylazine mixture to prevent overdose
β€’ Causes shaking as it wears off. For imaging, strictly adhere to dosing schedule
β€’ May cause bradycardia
Ketamine + xylazine90 + 9 mg/kgIntraperitoneal20–40 minβ€’ See above (Ketamine + xylazine + acepromazine)
Pentobarbital50 - 70 mg/kgIntraperitoneal20–60 minβ€’ Depresses respiratory rate and motion
β€’ Expense may be cost prohibitive
β€’ Pharmaceutical grade may not be available
Disclaimer: These are general guidelines. Consult a veterinarian for more information before implementation.

Table 3: Common anesthetic formulary for mice. Please click here to download this Table.

  1. Ventilate animal (Table 2).
    1. Attach the animal to ventilator via Luer connector on the intubation cannula. Monitor diaphragmatic motion and peak inspiratory pressure (~10-12 cm H2O for a tidal volume of 10 mL/kg of body weight). If the pressure or breathing motion is abnormal, carefully adjust the neck angle and depth of the cannula as needed.
    2. Ensure the intubation cannula is airtight by performing a recruitment maneuver: prevent exhalation (e.g., with a finger, block the exhale port) such that the animal inhales multiple times without exhaling.
      NOTE: If the airway pressure reaches 35 cmH2O peak inspiratory pressure over ~6 s, the airway seal is sufficiently tight. If not, see the discussion for troubleshooting.
    3. Allow normal exhalation to resume. Perform recruitment maneuvers between scans and every ~5 min when not scanning to maintain lung compliance and prevent atelectasis. Connect the PEEP valve to the exhale line. Set PEEP to 4 cmH2O. Watch the peak inspiratory pressure increase by that amount.
    4. After successful intubation, plan and initialize the production of HP 129Xe around the sedative redose schedule to prevent the animal from waking during a scan. Monitor body temperature throughout the experiment.
  2. Data acquisition: Collect ventilation images.
    NOTE: Data Acquisition StepsΒ 2.4, 2.5, and 2.6 can be done in any order
    1. Set the ventilator according to Table 2 for imaging at end-inspiration.
    2. Load the following setup protocols: proton animal localizer, single pulse to center on gas frequency in the mouse lungs, and 129Xe animal localizer. See Table 4 for scan parameters.
    3. Position the animal at the isocenter and confirm that the thoracic cavity is in the center of the field of view with proton localization. If using single-frequency coils, replace the proton coil with a 129Xe-tuned RF coil.
    4. Record 129Xe polarization and transport it to the MRI scanner. See Supplementary File 4: Xenon polarization QA checklist.
    5. Place a bag of 129Xe within the ventilator cannister and seal. Connect the canister in line with the ventilator and allow the cannister to pressurize (3 - 6 psig).
    6. Initiate 129Xe mechanical ventilation. Each time 129Xe ventilation is activated, allow the animal to complete ~5 breaths prior to starting a scan to turn over the functional residual capacity of the lungs.
      NOTE: Switch to the N2/O2 mixture between 129Xe scans to conserve hyperpolarized gas.
    7. Using a single pulse, adjust the working frequency to match the in vivo resonance frequency of gaseous 129Xe (~83.07 MHz at 7 T). Copy the frequency to all subsequent gas phase 129Xe scans. Perform 129Xe localization to confirm lungs are at isocenter.
    8. Load and run the 129Xe radial ventilation sequence. Monitor peak inspiratory pressure.
      NOTE: If the 129Xe gas runs out before the protocol is finished, the peak inspiratory pressure will decline rapidly. A 400 mL bag of 129Xe can ventilate a 30 g mouse for ~24 min whenΒ ventilated with 70% 129XeΒ at 80 breaths per minute with a tidal volume of 10 mL/kg of ideal body weight.
    9. When the scan is finished, switch to ventilating with the N2/O2 mixture and remove the empty bag of 129Xe.
    10. For imaging at end-expiration, change the breath hold duration and trigger delay according to Table 2 and repeat Steps 2.4.2 - 2.4.9.
    11. Export the raw data from the scanner.
Protocol Short-Hand NameSequence DescriptionTriggerTR (ms)TE (ms)RepetitionsFlip Angle (˚) Matrix Size or  NptsFOV (mm2)RF BW (kHz)Slice/Slab Thickness (mm)Scan Duration
Single pulsePulse acquire (gas phase)Optional10001602048101 s
Animal localizer2D GREYes501.716064232232560 s
Radial Ventilation3D Multi-echo radialYes20See caption13061322332.053016 min
Dissolved phase single pulsePulse acquire (dissolved phase)No8019051210.3580 ms
Dissolved phase dynamic spec.Pulse acquire (dissolved phase)No5010009051210.550 s
Diffusion weighted2D GREYes12.28.144564232231.518 min


Table 4: In vivo sequence parameters. The 3D multi-echo radial ventilation sequence described previously39 acquires images at 6 echo times. Results are shown for the first echo image (TE = 1.12 ms, Figure 7). Please click here to download this Table.

Supplementary File 4: Xenon polarization QA checklist. Please click here to download this File.

  1. Data acquisition: Run dynamic dissolved phase spectroscopy.
    1. Set the ventilator according to Table 2 for imaging during breath hold. Set the BR to 100 breaths/min. Prepare for a new bag of 129Xe as in StepsΒ 2.4.2Β - 2.4.5.
    2. Initiate 129Xe mechanical ventilation. Load and run a single pulse to adjust the working frequency to match the dissolved frequency (~83.084 MHz at 7 T). Copy the working frequency to the dissolved phase dynamic spectroscopy.
      NOTE: This will be a single peak in mice with wild-type hemoglobin54.
    3. Load and run the dynamic spectroscopy sequence centered on the dissolved frequency in animal lungs.Β When the scan is finished, switch to ventilating with the N2/O2 mixture and remove the empty bag of 129Xe. Export the raw data from the scanner.
  2. Data acquisition: Collect diffusion-weighted images.
    1. Set the ventilator according to Table 2 for imaging during breath hold. Prepare for a new bag of 129Xe as in StepsΒ 2.4.2 -Β 2.4.7.
    2. Load and run the diffusion-weighted sequence. When the scan is finished, switch to ventilating with the N2/O2 mixture and remove the empty bag of 129Xe. Export the raw data from the scanner.

3. Concluding the experiment

  1. Recover animal.
    1. Pull the intubation cannula straight out of the mouth of the animal. If the animal does not immediately begin breathing spontaneously, administer light chest compressions. If available, administer a light flow of medical-grade oxygen near the animal's face to support sedation recovery.
    2. Once the animal is steadily breathing on its own and if the animal was sedated for >2 h, administer 0.5 - 1 mL of normal saline subcutaneously to prevent dehydration.
    3. Return the animal to a cage by itself. Place the cage in an incubator or on a heating pad.
      NOTE: Sedated animals are vulnerable to cannibalism and cannot be placed in a cage with cage mates until fully recovered (i.e., ambulating independently). Sedated animals cannot regulate their body temperature. Use the back of the hand to feel the temperature of the animal every few minutes.
    4. Monitor the animal closely until their righting reflex has returned (i.e., it can independently flip from a supine to a prone position).
    5. Remove the animal from heat support once it can ambulate independently. Return the animal to a cage with its cage mates.
      NOTE: if the animal has no cage mates, it is more susceptible to sedation-induced hypothermia. Provide the animal with extra bedding, and if available, leave the animal in an incubator overnight.
    6. Record the animal's weight once per week for 2 weeks to monitor its health.
      ​NOTE: If the animal sustained an injury to the mouth or esophagus from intubation, it may stop eating. If the animal loses >20% of its initial body weight, consult a veterinarian about euthanasia.
  2. Analyze ventilation images (Figure 7).
    1. Load raw data into a programming platform. Download the open-source reconstruction framework for non-Cartesian Images91.
    2. Reconstruct images according to open-source framework instructions. Normalize the first point on each radial projection39.
    3. Segment the lung parenchyma in the images, including voxels with low or no 129Xe signal. Do not include large airways. Segment the background noise in the image, excluding lungs, airways, and imaging artifacts.
      NOTE: The proton localizer image can aid in determining parenchymal boundaries.
    4. Calculate the SNR using the formula:
      figure-protocol-45911Β  Β  Β (3)
    5. Quantify defective ventilation.
      NOTE: Various methods have been proposed to quantify impaired ventilation in small animal models. Analysis methods remain an open area of inquiry, but easily implemented approaches include: (i) Semi-quantitative manual segmentation92, (ii) Histogram approach using the signal from the trachea to normalize parenchymal signal47, and (iii) Segmenting the total lung volume (TLV) and defining a signal threshold (e.g., <60% of the whole lung mean) to divide the lungs into defective volume and ventilated lung volume (VV). Quantify VDP93,94Β according to:
      figure-protocol-46779Β  Β  Β (4)
  3. Analyze dissolved phase dynamic spectroscopy (Figure 8).
    1. Load raw data into a programming platform. Perform a fast Fourier transform and phase the spectra (simultaneous manual, zero-order phasing of spectra is sufficient for this application.)
    2. Calculate standard nuclear magnetic resonance (NMR) data95,96: SNR, full width half max, integrated area, chemical shift, and phase of both peaks.
    3. From the magnitude data, divide the signal amplitude of the dissolved spectrum by that of the gas spectrum for each repetition to find the dissolved-to-gas ratio over time.
  4. Analyze diffusion-weighted images (Figure 9).
    1. Load raw data into a programming platform. Segment the lung parenchyma of the b0 image as in StepΒ 3.2.3. Calculate the SNR for each b-value image.
    2. Calculate the signal-value-to-noise ratio, SVNR0, by dividing the signal in each voxel of the b0 image by the standard deviation of the image noise. Exclude voxels with SVNR0 < 2.5 times the image noise97.
      NOTE: The SVNR0 is an individual voxel metric, while the parenchymal SNR is a whole-lung metric.
    3. Calculate ADC by fitting the signal decay over the b-values (bi)Β according to Equation 598,99:
      figure-protocol-48480Β  Β  Β (5)

Results

Ventilation Images

If animal preparation and ventilation procedures are implemented properly, 3D radial imaging can successfully capture ventilation patterns when data acquisition is performed at either inspiration or expiration (Figure 7). While these images are collected over many breaths, the method described here is similar to the single-breath imaging method used in humans. This is because all lines of k-space are collected at a specific time...

Discussion

Hyperpolarized 129Xe MRI is emerging as a sophisticated and powerful technique to study lung microstructure and function in small animal models. This protocol is intended to guide initial site preparation and describe experimental procedures needed to quantify ventilation, diffusion, and gas exchange in mouse lungs with HP 129Xe. Key prerequisites for experiments include establishing a 129Xe gas phantom and ensuring high gas polarization is delivered to the animal. The latter requires car...

Disclosures

Peter Niedbalski is a consultant for Polarean Imaging, Plc.

Acknowledgements

The authors extend their heartfelt gratitude to Jerry Dalke for being a guiding light in ventilator construction. We'd like to thank Carter McMaster for brewing HP 129Xe gas. We'd also like to thank Dr. Matthew Willmering and Dr. Juan Parra-Robles for their thought-provoking scientific discussions. Figures created with BioRender.com. This work was funded by the National Institutes of Health (Grant Nos: NHLBI R01HL143011, R01HL151588)

Materials

NameCompanyCatalog NumberComments
1 mL syringefisher scientificCatalog No.14-955-464https://www.fishersci.com/shop/products/sterile-syringes-single-use-12/14955464
10 mL graduated cylinderCole-ParmerΒ UX-34502-69https://www.coleparmer.com/i/cole-parmer-essentials-graduated-cylinder-glass-hexagonal-base-10-ml-2-pk/3450269?PubID=UX&persist=true&ip=no&
gad_source=1&gclid=CjwKCAi
A6KWvBhAREiwAFPZM7h3do
-ssjascARuVviKd7V7kC5ztdIB6
_70DnMr-K3qk9RKeJ7-IrhoCeT
0QAvD_BwE
18 G - veinous PFTE catheters (nonsterile)Terumo SurfloSROX1832CAhttps://www.shopmedvet.com/product/iv-catheter-18-x-1-25inch?r=GSS17&p=GSS17&utm_source=
google&utm_medium=google_
shopping&gad_source=1&gclid=
CjwKCAiA0bWvBhBjEiwAtEsoW
4oTvZkAgWQCda6ocVtQlulVrG
2536FNbu5soMVSFN8xK_g1Uh
pXIRoCGwoQAvD_BwE
20 G - veinous PFTE catheters (nonsterile)Terumo SurfloSROX2051CAhttps://www.shopmedvet.com/product/iv-catheter-20-x-2inch?r=GSS17&p=GSS17&utm_source
=google&utm_medium=google_
shopping&gad_source=1&gclid=
CjwKCAiA0bWvBhBjEiwAtEsoW
87ggCkgToD_XF_UgpQBTpmN
dgSNfCml6TkDKlW8k27Dq_daR
itPuhoCnBQQAvD_BwE
22 G - veinous PFTE catheters (nonsterile)Terumo SurfloSROX2225CAhttps://www.shopmedvet.com/product/iv-catheter-22-x-1inch?r=GSS17&p=GSS17&utm_source=
google&utm_medium=google_
shopping&gad_source=1&gclid
=CjwKCAiA0bWvBhBjEiwAtEso
W9IM6mpee6m7e-lBfR8dZhSN
KYbMUs7qgEU4gYCRTW_rJAs
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400 mL tedlar bagsJensen Inert ProductsGST-001S-3507TJCNA
60 mL syringefisher scientificCatalog No.14-955-461https://www.fishersci.com/shop/products/sterile-syringes-single-use-12/14955461
70% alcoholCole-ParmerUX-80024-34https://www.coleparmer.com/i/labchem-isopropyl-alcohol-70-v-v-500-ml/8002434?PubID=UX&persist=true&ip=
no&gad_source=1&gclid=CjwKC
AiA6KWvBhAREiwAFPZM7gGh
p8g7MBHBBKadaRCAwfEMgV
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Dewar for liquid nitrogenTerra Universal4LDBhttps://www.laboratory-equipment.com/tw-4ldb-liquid-nitrogen-dewar-ic-biomedical.html?srsltid=AfmBOooxwMtOA1Z2TweR
P8V5Iy5EvYT3alZuzoiY
3UF3Ib9RgFnDxVTfWP0
Eye lubricantRefreshΒ REFRESH P.M.https://www.refreshbrand.com/Products/refresh-pm
Fiber optic lightAmScopeHL250-AYhttps://amscope.com/products/hl250-ay?tw_source=google
&tw_adid=&tw_campaign=
16705014684&gad_source=
1&gclid=CjwKCAiA6KWvBhA
REiwAFPZM7p-DpyvHJaGxR
pAD1385hzGf1oPdKHHLFDR
Sp8yrtxry11SNJeJnKxoCtAoQ
AvD_BwE
GaussmeterApex MagnetsGMHT201https://www.apexmagnets.com/magnets/accessories/ht-digital-gaussmeter-with-peak-hold-can-display-gauss-or-tesla
Glass vessel (phantom)Ace Glass8648-24https://aceglass.com/results.php?t=8648-24&t=8648-24
Heating padOffice Depot9206211 Β Β Β Pure Enrichment PureRelief Express Designer Series Heating Pad 12 x 15 Palm Aqua - Office Depot
HyperpolarizerPolarean9820https://polarean.com/xenon-mri-platform/
Intubation boardHallowell EMC000A3467https://hallowell.com/product/rodent-tilting-workstand/
Intubation suppliesParts list published elsewhereNAhttps://app.jove.com/t/50318/a-simple-method-of-mouse-lung-intubation
Isotopically enriched xenon cylinderLinde IsotopesΒ XE-129(1%)N2(10%)HE CGMP 302SZNA
Liquid nitrogenLindeNI LC160-22https://www.lindedirect.com/store/product-detail/nitrogen_n2_nitrogen_liquid
_lc160_22_psi_ni_lc160_22
/ni-lc160-22?cat_id=shop&node=b89
Male slip luerCole-ParmerUX-21943-27https://www.coleparmer.com/i/diba-omnifit-t-series-solvent-waste-cap-adapter-polypropylene-male-luer-slip-x-1-16-id-hose-barb-5-pk/2194327
ManometerGrainger3T294https://www.grainger.com/product/3T294?gucid=N:N:PS:
Paid:GGL:CSM-2295:4P7A1P:
20501231&gad_source=1&gclid
=CjwKCAiAi6uvBhADEiwAWiyR
dltxrPJmmcm0bFiYLuPrB25HV
QFdEfKMBqvgJBNdQUs3DZ7b
TLr8CRoCanAQAvD_BwE&
gclsrc=aw.ds
Minivent ventilatorharvard apparatus73-0044https://www.harvardapparatus.com/minivent-ventilator-for-mice-single-animal-volume-controlled-ventilators.html
Mouse ear puncherfisher scientific13-812-201https://www.fishersci.com/shop/products/fisherbrand-animal-ear-tag-punch/13812201
Mouse tongue depressorMedical ToolsVRI-617https://medical-tools.com/shop/rodent-tongue-depressor.html
Mouse weight scaleCole-ParmerUX-11712-12https://www.coleparmer.com/i/adam-equipment-cqt2000-core-portable-balance-2000g-x-1g-220-v/1171212?PubID=UX&persist=true&ip=no&gad
_source=1&gclid=CjwKCAiA6K
WvBhAREiwAFPZM7iYnAG5Ilc
Z5DZWrdJ6wcLDZSCSfNJHOH
m2PQOpyyWe0TjFa75R3tBoCjB
sQAvD_BwE
MRI scannerΒ Bruker7T Biospec horizontal systemhttps://www.bruker.com/de/products-and-solutions/preclinical-imaging/mri/biospec.html
MultimeterHome Depot1007898529https://www.homedepot.com/p/Klein-Tools-600-Volt-Digital-Multi-Meter-Manual-Ranging-MM325/320822947
Natural abundance xenonLinde IsotopesΒ UN 2036NA
NeedleΒ Β fisher scientific305194https://www.fishersci.com/shop/products/bd-general-use-precisionglide-hypodermic-needles-20/148266C?keyword=true
Needle safe syringe holderfisher scientificNC2703873https://www.fishersci.com/shop/products/ndlsafe-ii-syr-uncap-deca/NC2703873#?keyword=needlesafe
Nitrogen cylinderLindeNI M-Khttps://www.lindedirect.com/store/product-detail/nitrogen_n2_nitrogen_nf_k/ni-m-k?cat_id=shop&node=b89
Oxygen cylinderLindeOX M-Khttps://www.lindedirect.com/store/product-detail/oxygen_o2_oxygen_usp_k/ox-m-k?cat_id=shop&node=b90
Oxygen sensorApogee instrumentsMO-200https://www.apogeeinstruments.com/mo-200-oxygen-sensor-with-handheld-meter/
Oxygen sensor inline flowheadApogee instrumentsAO-002https://www.apogeeinstruments.com/ao-002-oxygen-meter-sensor-flow-through-head/
PEEP valveHallowell EMC000A6556Ahttps://hallowell.com/product/adjustable-peep-valve-with-exhaust-port-range-5-20cm-disposable/
Pipette tipsfisher scientificCatalog No.02-707-108Fisherbrand Stack-Rack Space-Saver Tips: 101-1000 L Standard; Blue; Volume: | Fisher Scientific
Plunger valveΒ Ace glass8648-20https://www.aceglass.com/results.php?t=8648
Preclinical coilDoty scientificcustom builthttps://dotynmr.com/products/bmax-xy-low-e/
Pressure regulatorsCole-ParmerUX-98202-11https://www.coleparmer.com/i/cole-parmer-single-stage-regulator-1500-scfh-capacity-346-cga-fitting/9820211?PubID=UX&persist=true&ip=no&
gad_source=1&gclid=CjwKCAi
A6KWvBhAREiwAFPZM7pruR
xCAiaj52nA_8Y1nveQZRsD6B
f0QO65o2DKFYqRoz0PopSkX
QxoCxqcQAvD_BwE
Pressure-drop ventilatorParts list published elsewhereNAhttps://sites.duke.edu/driehuyslab/resources/
PVC pipe for phantomHome Depot193682https://www.homedepot.com/p/IPEX-1-2-in-x-10-ft-White-PVC-SCH-40-Potable-Pressure-Water-Pipe-30-05010HD/319692959
SAI animal heating systemSAIIModel 1030https://i4sa.com/product/model-1030-monitoring-gating-system/
SalineFarris Laboratories Inc.0409488820-1https://www.farrislabs.com/products/bacteriostatic-sodium-chloride-0-9-30ml-bottle?variant=42807174824167&currency
=USD&utm_medium=product_
sync&utm_source=google&utm_
content=sag_organic&utm_
campaign=sag_organic&utm_
campaign=gs-2021-09-24&utm
_source=google&utm_medium
=smart_campaign&gad_source
=1&gclid=CjwKCAiA6KWvBh
AREiwAFPZM7oS3-hFDETO_2f6OWOoKyBMb
WuDuWqYxdWRYUWEkY
M2Py73VfGzVtRoC2FQQAvD_BwE
Sharps containerfisher scientific22-730-455https://www.fishersci.com/shop/products/sharps-container-47/p-7250579#?keyword=needle%20safe
Silicone epoxyGrainger3KMY7https://www.grainger.com/product/3KMY7?gucid=N:N:PS:Paid:GGL:CSM-
2295:4P7A1P:20501231&gad_
source=1&gclid=CjwKCAiA6KW
vBhAREiwAFPZM7voahkm8tda
t1Euql1A8DFhC6AZVJ0wXzCE
PfE6iUzrIJXV-Hl8o4xoCQLYQA
vD_BwE&gclsrc=aw.ds
Silicone mold release lubricantGrainger19MW95https://www.grainger.com/product/CRC-Mold-Release-Agent-16-oz-19MW95
SpirometerADInstrumentsFE141https://www.adinstruments.com/products/spirometer
Spirometer - mouse flowheadADInstrumentsMLT1Lhttps://www.adinstruments.com/products/respiratory-flow-heads
Tubing - 1/4 ODClippardURH1-0402-CLT-050https://www.clippard.com/part/URH1-0402-CLT-050
Tubing - 1/8 ODClippardURH1-0804-CLT-050https://www.clippard.com/part/URH1-0402-CLT-050
Vacuum pumpCole-ParmerΒ UX-60062-11https://www.coleparmer.com/i/environmental-express-diaphragm-pump-high-volume-120v/6006211?PubID=UX&persist=true&ip=no&gad
_source=1&gclid=CjwKCAiA6K
WvBhAREiwAFPZM7uFGwmW
pRelHNFgZVvJJV09vDUVyfyG
HoKeZTiFNIiVTe-05IpJJPxoCO
PoQAvD_BwE
Wire - 18 gaugeDigikey2328-18H240-NDhttps://www.digikey.com/en/products/detail/remington-industries/18H240/15202027?s=N4
IgjCBcoOwBxVAYygMwIYBsDOB
TANCAPZQDa4YATPAGwgC6h
ADgC5QgDKLATgJYB2AcxAB
fQmAAMAFkqIQKSBhwFiZEA
GZNATi0SGzNpE48BwsSErqw
6uQqV5CJSOQCsMF%2Bq11
GIVuy58QqLmss4gALbogvy4L
AAEAO683LgMIkA
Xenon polarization measurement stationPolareanNAhttps://polarean.com/xenon-mri-platform/

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