Efficient Method for Imaging Murine Lungs that Preserves Spatial Dynamics of Fungal Spores in the Airways

Summary

We present a method for a fungal pathogenesis model that preserves the natural positioning of fungal spores in the lung airways for analysis via fluorescent microscopy.

Abstract

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and bronchioles, branch until terminating in the alveolar airspace where gas exchange occurs. Infections originating in the bronchioles or alveoli elicit distinct host responses and disease manifestations. Therefore, understanding precisely where spores naturally localize in the lungs, particularly soon after infection, expands opportunities for investigation of host-pathogen interactions. Herein, we detail an in-situ analysis of lungs from mice infected with Coccidioides posadasii cts2/ard1/cts3Δ arthroconidia. Conventional methods for histological preservation involve liquid inflation of the airways with a fixative solution, which displaces the natural location of aspirated fungal particles, pushing spores from proximal bronchioles to terminal airspaces.

Conversely, this method of air-inflation with blood vasculature perfusion-fixation preserves the physiologic position of fungal spores within the bronchioles. Moreover, we describe a simple approach to cryopreserving, embedding, and imaging lung specimens. We also share high-throughput computational techniques via the open-source QuPath program to analyze the spatial distribution of fungal spores within the lung. The method presented here is simple and quick, requires minimal equipment to perform, and can be easily adapted for use with many respiratory fungal infection models.

Introduction

Humans can inhale up to billions of spores per day from a variety of environmental fungi¹. To understand our barrier defenses against these inhaled spores, we must appreciate the precise microanatomical environments where these spores land within the airways and lung parenchyma. The cellular composition of the airways (i.e., epithelial cells) significantly transforms along the trachea, bronchi, bronchioles, and alveoli. Each of these distinct regions is composed of different cell types with discrete functions which avails an arsenal of defenses to prevent pathologic infection.

The precise location of pulmonary fungal spore deposition can vary between the airway lumens of columnar epithelial-lined bronchioles, alveolar ducts, or alveoli². Most clinically relevant fungal species produce spores between 1 µm and 10 µm in diameter³. The deposition of these spore particles in the lung depends on several factors, such as aerodynamics, density, electrical charge, and phoretic forces, which can influence the mechanism of sedimentation after inhalation⁴. Generally, large particles (> 6 µm) deposit in the upper airway, medium-sized particles (2-6 µm) can deposit in smaller airways, and small particles (<2 µm) reach the alveolar region⁵. Aspergillus spores (2-3 µm) have been reported to reach alveolar spaces, but clinical pathology reports also indicate a significant burden of bronchial and bronchiolar disease⁶. There is also increasing recognition of endobronchial fungal infections of Aspergillus fumigatus, Coccidioides immitis, Candida species, Cryptococcus neoformans, Histoplasma capsulatum, and Zygomycetes due to the increasing popularity of flexible bronchoscopy⁷. Recent advances in microscopic imaging of Aspergillus infections in mice have also revealed that more proximal airspaces such as bronchi and bronchioles may bear the highest burden for pathologic fungal proliferation⁸. Research on the host response to pulmonary fungal infections has shown that both bronchiolar epithelial cells and alveolar epithelial cells play important roles as immune sentinels, so elucidating the exact sites of spore deposition and epithelial interaction will be vital for future work^2,9,10.

Studying these proximal airway pathogenesis dynamics is difficult because standard lung fixation and sectioning preparation techniques can displace spores from these proximal positions in epithelial-lined airways and push them toward distal terminal alveolar regions. Commonly, 10% formalin or 4% paraformaldehyde is used to inflate the lungs and rapidly expose the entire lung to fixative. When preparing lungs for cryosectioning, some groups administer optimal cutting temperature (OCT) compound into the lungs to improve cryosectioning performance¹¹. These practices are useful in the right context but have been found by our lab and other groups to displace spores and particles from proximal locations, thereby interfering with interpretations about the spore-exposed cell types and the subsequent host response¹².

To accurately establish the microanatomical localization of inhaled fungal spores, we have developed a quick, low-resource method for the preservation of the location of fungal spores in the airways of mice. We adapted a murine air-inflation vascular perfusion-fixation method from Thomas et al. (2021), where we reduced the amount of equipment, time, and technical skill required to achieve a satisfactory result¹³. From the method described here, we have observed that Coccidioides posadasii cts2/ard1/cts3Δ arthroconidia (3-5 µm in size) accumulate more proximally than previously shown, namely in distal bronchioles and broncho-alveolar junctions rather than terminal alveoli. This information can focus biological questions concerning the critical cell types associated with these regions of the lung and their influence on the early responses to inhaled fungal spores.

Protocol

All methods described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of Rutgers Biomedical Health Sciences.

1. Label spores with intracellular fluorescence

1. Stain 1 x 10⁶ spores in carboxyfluorescein succinimidyl ester (CFSE), Cell Tracker Orange, or Cell Tracker Red (or intracellular dye of choice) at 5 µM for 30 min at 30 °C, then wash with PBS, and spin the spores at 12,000 x g. Repeat this wash once.
2. Prepare the spore inoculum at the appropriate concentration so that 25 µL contains the appropriate spore dose for one mouse.

2. Inoculating mouse with spores through aspiration inhalation using isoflurane anesthesia

CAUTION: Isoflurane is a volatile anesthetic agent and must be used within a ducted biosafety cabinet or fume hood.

1. To prepare the isoflurane exposure jar, place a folded napkin into a 1 L screw-top Nalgene jar and place a plastic mesh circle over the napkin as a platform for the mice. Add 2 mL of isoflurane to the napkin, close the jar, and wait 1-2 min for the gas to equilibrate in the container.

Figure 1: Apparatus for isoflurane open drop method of spore inoculation. A folded napkin is placed in the bottom of a 1 L screw top jar, and then a circular plastic mesh is inserted to act as a platform for the mice. Please click here to view a larger version of this figure.

2. Prior to exposing the mouse to anesthesia, prepare a pipet with 25 µL of the inoculum. Place the mouse in the isoflurane container and tilt the container to monitor for loss of the righting reflex, where the mouse has lost consciousness and is upside down. This usually takes about 10 s.
3. Monitor the respiration rate (RR) until it slows to a consistent 50-80 breaths per minute (bpm), about 50% of resting RR. This usually takes another 10 s. At this time, remove the mouse from the isoflurane jar.
   NOTE: This is a high concentration of isoflurane, which will produce rapid induction of anesthesia, and the mouse must be very closely monitored. Lethal isoflurane overdose is a risk, as the concentration of isoflurane can be variable, and the dose-response curve to isoflurane is very steep. If the breathing rate dips below 50 bpm and/or is irregular, remove the mouse from isoflurane, allow it to recover for 3-5 min, and try again (a maximum of two repeated attempts).
4. Confirm the anesthetic depth by lack of response to toe pinch. Then, position the mouse in a supine position in one hand, and allow the mouse to take 3-5 regular gasping breaths with increasing rate (beginning at 50-60 bpm) and vigor. As this begins, place the pipet tip at the posterior of the mouse oropharynx and release the inoculum during an inhalation.
   NOTE: These gasps through the mouth jolt the head inferiorly due to the recruitment of accessory muscles for breathing. When the pipet is inserted during these gasps, the mouth should open with each gasp. If the mouth does not open, the mouse is insufficiently anesthetized and will likely not inhale contents from the oropharynx. The pipet should depress the tongue, pushing it towards the bottom and front of the mouth to prevent swallowing of the inoculum. When the pipet is in the mouth and the mouse is adequately anesthetized, the mouse will open its mouth with each gasp.
5. With the hand and thumb holding the mouse, feel for crackles on the posterior and anterior aspects of the mouse thorax to confirm inhalation of the inoculum.

3. Euthanasia of mice

1. Inject the mouse intraperitoneally with a cocktail of 200 mg/kg ketamine and 24 mg/kg xylazine 10 min prior to the sacrifice time point.
2. Wait 10 min post-injection until the mouse enters a surgical plane of anesthesia, confirmed by lack of response to toe pinch.

4. Perfusion of lungs with PBS and formalin

1. Spray the anesthetized mouse with 70% ethanol. Use scissors to snip back the skin of the mouse and pull apart the skin to expose the peritoneum on all sides. Pull the skin from the mouse's superior half over its head, pulling out the arms from the skin.
2. Cut the peritoneal membrane at the sternum and along the inferior aspect of the ribcage, exposing the peritoneum. Displace the liver inferiorly, revealing the diaphragm. Use scissors to carefully puncture the diaphragm away from the lung parenchyma to avoid puncturing a hole in the lungs themselves.
   NOTE: Inadvertently puncturing the lungs will make subsequent air inflation impossible.
3. After the diaphragm is punctured, air enters the thorax, collapsing the lungs. Cut the ribcage on the left side to reveal the heart. Inject 5-10 mL of PBS into the right ventricle with a 30 G needle at a rate of 1 mL per 5 s (to avoid damaging vessels), rapidly whitening the lungs.
   CAUTION: Formalin is a hazardous volatile substance that should be handled within a ducted biosafety cabinet or fume hood.
4. When finished, remove the needle and use a new needle to inject 5 mL of 10% neutral-buffered formalin (NBF) into the right ventricle at the same rate.

5. Air-inflating the perfused lung

1. Remove the anterior half of the ribcage. To expose the trachea, cut the superior ribs and collarbones to the right and left of the neck. Take care to avoid cutting near the midline where the trachea will lie. With forceps, clasp the remaining top ribs and collarbones on the neck midline and pull superiorly until the trachea is exposed.
2. Prepare a suture thread of 10 cm in length, pull it under the trachea, and pre-tie a loose knot at the inferior end of the exposed trachea. Prepare a 1 mL syringe of air with an 18 G catheter. Use an 18 G needle to make a hole in the trachea at the superior end of the exposed region.
3. Place the catheter into that hole, ensuring a relatively tight fit to prevent air from escaping. Slowly inject the 1 mL of air into the lungs over the course of 10 s, watching for lung inflation of all lobes. Full inflation results in the lungs wrapping slightly around the heart and filling the volume they occupy in the unpunctured diaphragm.
4. Pull the knot tight around the trachea, and remove the catheter. Hold the trachea and use blunt scissors to remove the lungs from the mouse, being careful not to puncture the lung.

6. Immersion fixation and dehydration

1. Place the lungs on the top edge of a 50 mL conical tube filled with 20 mL of 10% NBF. Keep the suture threads outside of the conical, screw down the cap, and invert the conical so that the lungs are suspended upside down in 20 mL of NBF. Place it at 4 °C for 24-48 h.
2. Rinse the lungs in PBS, then place in a 30% sucrose-PBS (w/v) solution for 72-96 h at 4 °C to dehydrate the lungs in preparation for cryopreservation. Remove the lungs from sucrose, place the lungs in a cryomold with optimal cutting temperature (OCT) medium, and freeze the specimen at -80 °C.

7. Cryosectioning

1. Equilibrate the specimens in OCT cryoblocks to the -20 °C temperature of the cryostat for 1 h prior to sectioning. Section the blocks at thicknesses of 20-100 µm.
   NOTE: Air inflation can lead to increased fragility of the specimen, and cutting at increased thickness can prevent tissue breakage during sectioning.
2. Collect sections onto glass slides and let dry for 30 min to 1 h to ensure tissue adherence to the slide.
   NOTE: Slides may be held at -80 °C at this stage.

8. Slide preparation and blocking

1. Place slides in PBS bath (in a dish or Coplin jar) for 30 min at room temperature (RT) to remove OCT from the slides.
   NOTE: Thicker sections (40-100 µm) may not adhere to glass slides as well as thinner sections, so it is best to keep them flat and upright in a dish rather than placed on their side in a Coplin jar.
2. Dry the slides after OCT has dissolved away from the tissue. Use a wipe to dry excess droplets from the slide's perimeter around the tissue specimen.
3. Use a hydrophobic Pap pen to draw a perimeter around the specimen, and let it dry for 5 min.
4. Prepare a blocking buffer of animal-free blocking solution with 0.3% Tween-20 and 1:100 Fc Block in 300 mL per slide.
   NOTE: For intracellular targets, 1% Tween-20 may be used. A volume of 300 µL is usually sufficient, but the necessary volume to cover the tissue fully depends on the perimeter size of the hydrophobic marker. The tissue should not dry from this point onwards.
5. Leave the blocking solution on the slides for 1 h at RT.

9. Immunostaining

1. Wash the slide by pipetting off the fluid and adding 300-500 µL of PBS. Repeat the wash once.
2. Prepare primary AlexaFluor-647 conjugated rat anti-mouse EpCAM (clone G8.8) antibody (airway epithelial marker) in animal-free blocker solution with 0.33% Tween-20 (or 1% for intracellular targets). Add 300 µL per slide of this solution and stain overnight at 4 °C.
   NOTE: Antibodies should be tested empirically for staining concentration, time, and temperature. Thin sections (8-20 µm) are stained for 30 min at 37 °C and thicker sections (20-100 µm) at 4 °C overnight using 1:100 dilutions for most antibodies. If using unconjugated antibodies, wash 2x (as above) and apply a secondary fluorescent antibody for an empirically determined time and temperature. Generally, secondary antibody staining is done at RT for 1 h at 1:1000 dilution.
3. Remove the antibody solution from the specimen, and wash the slide with 300 mL of PBS for 5 min. Repeat this wash once. Dry the slides, removing all excess fluid from the specimen and surrounding glass.
4. Add one drop of RT soft set mounting media (e.g., SlowFade Glass) to each piece of tissue. Take care to avoid bubbles by letting the bottle settle while upside down and slowly dropping each drop onto the tissues. Place an appropriately sized cover slip to extend beyond the sample and hydrophobic marker perimeter.

10. Imaging via fluorescent microscopy

1. Use a multichannel fluorescent microscope to scan whole lung sections in an unbiased manner using an objective with sufficient resolution to resolve individual spores and host cells (e.g., Zeiss Axioscan 7 using 20x oil objective with NA = 0.8 or similar).
   NOTE: Ensure that laser power and PMT voltage gain are set to maximize the signal-to-noise ratios of the target over the background determined with a fluorescence minus one (FMO) control.
2. Collect sufficient image tiles to capture the entire lung lobe (e.g., ~200 tiles of size 400 μm x 400 μm). Use the microscope software program (e.g., Zeiss ZEN 3.7 or similar) to export the merged tile image for downstream processing.

11. Spatial analysis via QuPath

1. In QuPath, a free open-source software¹⁴, create a project file and add the fluorescent images to the file.
2. Classify the EpCAM+ pixels as epithelium by clicking the top menu option Classify, then Pixel classification > Create Thresholder. Select the resolution, channel, smoothing sigma, and threshold value that best identifies the target region. Save the classifier under a unique name, and select Create Objects.
3. In the new Create objects window, select New object type as Annotation. For lung epithelium, set the minimum object size and minimum hole size to 100 mm². There is no need to select Split Objects here. After selecting OK, the annotations will be created, and they can be modified by eye using the Brush tool in the upper left-hand area.
4. To classify spores, repeat the steps from 11.3-11.4 using the channel in which the spores were captured and make the new object type Detection rather than Annotation. Set minimum object size and minimum hole size to 0 mm², and select Split Objects.
5. To conduct spatial analysis of the spores, select the top menu option Analyze > Spatial Analysis > Calculate signed distance to annotations 2D. The distances to EpCAM+ epithelium will be calculated for each detected spore. Export these via the top menu option Measure > Export Measurements.

Results

This method ultimately produces immunofluorescent images of mouse lungs using physiologic air inflation to leave the airways undisturbed. Importantly, multiple checkpoints along the way will confirm that components of the protocol have been performed successfully. During the inoculation, it is important to confirm that the inoculum was aspirated by feeling for "crackles" on the posterior chest wall of the mouse that indicate the liquid has entered the airways. If there is no sensation of crackles, it is possible that the mouse swallowed the inoculum. The perfusion of mouse lungs with PBS should result in white mottling of the lungs followed by complete whitening of the entire lung (Figure 2A). If there is incomplete whitening, the subsequent formalin perfusion will not reach all areas of the lung parenchyma (Figure 2B). When inflating the lung with air, the lungs should slowly distend until they reach physiologic size. After tying the tracheal suture to secure the air inside the lungs, the lungs should not contract in size. If they do, the lung was likely punctured during the procedure.

Here, we show representative images of sections with labeled Coccidioides posadasii cts2/ard1/cts3Δ spores (green) and EpCAM+ columnar bronchiolar epithelium (magenta) in both air-inflated and formalin liquid-inflated lungs (Figure 3A,B respectively). These spores primarily accumulate within distal bronchioles and in the alveolar spaces immediately adjacent to those distal bronchioles, likely in alveolar ducts. The air-inflated fixative-perfused lungs contain spores that appear more frequently within bronchioles and cluster closer to bronchioles (Figure 3A), rather than the liquid fixative where spores appear slightly more dispersed from the bronchiolar epithelium (Figure 3B). Through the QuPath spatial analysis, we measure the distance of individual spores to the nearest EpCAM+ bronchiolar epithelium to indicate the level of dispersion of the spores away from the bronchiole and into more distal alveolar spaces. We show that liquid inflation disperses spores further away from bronchioles compared with physiologic air inflation (Figure 4A,B). The air-inflation vascular perfusion preserves the natural positioning of these spores in distal bronchioles and prevents the artificial dispersion of these spores into more distal alveolar spaces by intratracheal fixative instillation.

Figure 2: Lung perfusion success indication. (A) Successfully perfused lungs will be completely white. (B) Incompletely perfused lungs will appear pink or mottled in appearance. Please click here to view a larger version of this figure.

Figure 3: Representative Images of spore distribution in air-inflated formalin-perfused lungs vs. formalin-inflated lungs. Intracellular fluorescent spores (green) are seen to be distributed in closer proximity to EpCAM+ bronchiolar epithelium (magenta) in (A) air-inflated lungs than in (B) formalin-inflated lungs. Images were obtained from sections with a depth of 1.2 mm from the posterior surface of the left lobes of mice in each treatment group on a multichannel fluorescent microscope at 20x magnification, and tiles were stitched using microscopy software. Scale bar = 800 µm. Please click here to view a larger version of this figure.

Figure 4: Comparison of spore distance (mm) to bronchiolar epithelium between air-inflated and formalin-inflated spore-treated lungs. Lungs from 3 mice per treatment group were harvested, processed, and imaged for 1.5 h following inoculation with 1 x 10⁶ Coccidioides posadasii cts2/ard1/cts3Δ arthroconidia. Four sections per mouse were sampled at 0.8 mm, 1.2 mm, 1.6 mm, and 2.0 mm from the posterior surface of the left lobe, stained, and imaged on a multichannel fluorescent microscope. All spores from each group were detected and spatially analyzed in QuPath to determine the (A) distance to the nearest EpCAM+ columnar bronchiolar epithelium, and (B) the median and 95% confidence intervals are displayed in units of micrometers. Air-inflation spores, n = 11,673. Liquid-inflation spores, n = 9,837. Mann-Whitney t-test was performed due to lack of normality, and p < 0.0001. Please click here to view a larger version of this figure.

Discussion

We have established a pipeline for spore inhalation and analysis of the spatial deposition of the inhaled spores. This pipeline provides valuable information to determine the relevant stromal regions of the lung affected by inhalation of Coccidioides posadasii cts2/ard1/cts3Δ arthroconidia. We have observed that Coccidioides spores and similarly sized inert particles (data not shown) accumulate in distal bronchiolar regions and broncho-alveolar junctions rather than fully dispersed throughout alveolar airway lumens. This model requires a relatively low level of equipment and technical skill to produce cryosections for immunofluorescence (or other downstream pipelines such as immunohistochemistry [IHC]) that maintain the physiological integrity of the airway contents while maintaining natural lung morphology. Without air inflation, the collapse of alveoli distorts the natural architecture of lung parenchyma surrounding bronchioles^13,15. Loss of the air volume of the lung during histological processing was indicated as a "serious problem" by a joint official research policy statement of the American Thoracic Society and European Respiratory Society in defining standards for quantitative assessment of lung structure¹⁶. Furthermore, intratracheal instillation of fixative can disrupt the nature and location of proximal airway molecular and cellular components such as mucins, surfactant, and airway cilia^17,18,19. The method described here sufficiently preserves both the air volume of the lung along with the molecular and cellular components along airways that interact with inhaled fungal spores.

Immunofluorescence microscopy provides a powerful tool to interrogate spatial dynamics of lung components, and it is well-reviewed^20,21. There are no limitations to the targets that can be stained via immunofluorescence of these sections, except that antibodies with epitopes sensitive to formalin fixation may not bind their targets satisfactorily. When using a new antibody, it is best to prepare a fluorescence minus one (FMO) control to determine whether the antibody generates any signal above the background. If using a secondary, use a treatment group without the primary antibody to control for off-target binding of the secondary antibody. Primary conjugated monoclonal antibodies are very specific, so staining at higher temperatures (37 °C) for less time (30 min) is common and generally binds specifically to their target. On the other hand, unconjugated polyclonal antibodies are more sensitive to detect antigens but less specific, so longer incubations (~16 h) at colder temperatures (4 °C) optimize the signal-to-noise ratio. All antibodies will require empirical optimization.

The spatial analysis model used in this study for spore localization measures the distance of individual spores to the nearest bronchiolar epithelium. This requires appropriate identification of the bronchiolar epithelium using its high EpCAM signal and morphologically distinct appearance as a columnar epithelium lining tube-like structures. EpCAM can also be expressed at lower levels by alveolar epithelium, so it is important to threshold the EpCAM signal in QuPath to limit the defined annotations to morphologically confirmed bronchiolar regions. As bronchioles have an anatomical minimum size, we can limit the QuPath definition of these regions to the minimum area of 100 mm², as stated in step 11.3. This will exclude isolated high EpCAM-expressing cells or artifacts throughout alveolar spaces, such as alveolar type II epithelial cells, from the QuPath annotations. These steps allow us to define exclusively bronchiolar epithelium as QuPath annotations and determine the proximity of spores to this defined bronchiolar epithelium.

The infection model described here uses isoflurane anesthesia to induce an oral gasp response in mice for aspiration of the fungal spores. The isoflurane open drop method has been developed as a quick, low-resource technique to induce short-term anesthesia in mice^9,22,23. This method takes some practice to perform consistently. The time under anesthesia must be long enough to sufficiently anesthetize the mouse to aspirate pharyngeal contents. If the mouse is insufficiently anesthetized, it can swallow the inoculum instead of aspirating it. The sign of sufficient anesthetization is reached when the mouse produces a regular oral gasp of increasing vigor at a rate of about 50-60 gasps/min. The supine mouse's head should move forward with each gasp. Insufficient anesthetization can be determined by a lack of gasp or any movement of whiskers, tongue, or limbs. It is best practice to depress the tongue with the pipet tip while administering the inoculum to prevent swallowing. Overdose of isoflurane can result in the death of the mouse and must be prevented. A good rule of thumb to prevent this is to remove the mouse from isoflurane after the same amount of time taken for the mouse to lose the righting reflex. For example, if the mouse takes 10 s to lose the righting reflex, remove the mouse after an additional 10 s. As the isoflurane gas escapes over the course of multiple mice treatments, this time frame may increase. In our hands, we can treat 6-8 mice without adding more isoflurane.

There are some limitations to the components of this protocol. The inoculation procedure uses spores suspended in 25 µL of fluid PBS medium, which is less physiologic than an aerosolized inoculation of spores. An open question remains whether the use of aspirated fluid alters the localization of spores/particles compared with other inoculation methods, such as inhalation of airborne spores. However, aerosolized inhalation requires extensive resource investment and has other limitations (limited inoculum size, variable exposure, occupational risk) that may not be optimal depending on the nature of the study and lab environment. Another limitation of this protocol is that tissue fragility may occur during cryosectioning, which could be due to either the presence of air throughout the tissue or the reduced effectiveness of the vascular fixation compared with intratracheal fixation. This cryosectioning fragility can be overcome by cutting thicker sections of 60-100 µm and using confocal microscopy to image two-dimensional (2D) planes within the tissue.

We believe the simplicity and adaptability of this method will enable other groups to find utility with their chosen inhaled pathogens and host responses of interest.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
18 G, 1 1/2 needle (305185)	Fisher	305185
1 L  Screwtop Jar  (Nalgene)	Fisher Scientific	11-823-33
Air-Tite Bulk Unsterile Syringes 10 mL Luer Lock	Fisher	14-817-175
AnaSed Injection (xylazine sterile solution)	Akorn	59399-110-20
Animal-Free Blocker and Diluent, R.T.U.	Vector	SP-5035-100
BD Insyte Autoguard Winged Shielded IV Catheter with BD Vialon Catheter Material 18 G x 1.88 in	BD	381547
BD Pharmingen Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block™)	BD	553142
CellTracker Orange CMRA Dye	Fisher Scientific	NC0873640
CFSE	Labviva	75003
Coccidioides posadasii cts2/ard1/cts3Δ	BEI Resources	NR-166
Corning 70 micron strainers	VWR	10054-456
EpCAM AlexaFluor647 monoclonal antibody	Biolegend	118211
Exel International HYpodermic Needles 30 G x 1/2"	Labviva	EN3012
Fisherbrand Sterile Syringes for Single Use (1mL, Leur Slip)	Fisher	14-955-462
Glucose Monohydrate	Azer Scientific	ES17530-500G
High Vacuum Grease	VWR	59344-055
Hoechst 33342 Solution 20 mM (5 mL)	ThermoFischer	62249
Isoflurane USP	Covetrus	29405
Ketamine Hydrochloride	Dechra	1000001250
KIMWIPES Delicate Task Wipers (4.4'' x 8.4")	VWR	21905-026
Lexer Baby Scissors	FST	14078-10
Micro-Adson Forceps	FST	11018-12
Neutral Buffered Formalin (10%) (Azer Scientific)	Fisher	22-026-350
Nunc EasYFlask tissue culture flasks, T75, filter caps	VWR	15708-134
PBS	VWR	45000-446
Peel-A-Way embedding molds	Sigma	E6032-1CS
QuPath 0.5.1 Software	Open-source	https://qupath.github.io/
Silk Suture thread size 3/0	FST	18020-30
SlidesMicro Slides Superfrost Plus	VWR	48311-703
SlowFade Glass Soft-set Antifade Mountant (2 mL)	ThermoFischer	S36917
Sucrose	Sigma	S0389-500G
Tissue-Tek O.C.T. Compound, Sakura Finetek	VWR	25608-930
Tween 20	ThermoFischer	J20605.AP
Vector Laboratories ImmEDGE Hydrophobic Barrier Pen Set Of 2	Fisher Scientific	NC9545623
VWR Micro Cover Glasses, Rectangular (24 mm x 40 mm #1.5)	VWR	48393-230
White Plastic Wire Mesh	MAPORCH	789862904922
Yeast Extract	Fisher	BP9727-500
Zeiss AxioScan 7	Carl Zeiss Microscopy GmbH	https://www.zeiss.com/microscopy/us/products/imaging-systems/axioscan-for-biology.html	multichannel fluorescent microscope
ZEN 3.7 Software	Carl Zeiss Microscopy GmbH	https://www.zeiss.com/microscopy/us/products/software/zeiss-zen.html	microscopy software