Funding and support were acquired through NIH grant K22 AI153678-01 and Rutgers School of Graduate Studies. We thank Fawad Yousufzai and Luke Fritzky from the Rutgers Biomedical Health Sciences Cellular Imaging and Histology Core for their work and expertise in obtaining immunofluorescent images.

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
<ol>
	<li>Stain 1 x 106 spores in carboxyfluorescein succinimidyl ester (CFSE), Cell Tracker Orange, or Cell Tracker Red (or intracellular dye of choice) at 5 &#181;M for 30 min at 30 &#176;C, then wash with PBS, and spin the spores at 12,000 x g. Repeat this wash once.</li>
	<li>Prepare the spore inoculum at the appropriate concentration so that 25 &#181;L contains the appropriate spore dose for one mouse.</li>
</ol>
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.
<ol>
	<li>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.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67556/67556fig01.jpg" /> 
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. <a href="https://www.jove.com/files/ftp_upload/67556/67556fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>Prior to exposing the mouse to anesthesia, prepare a pipet with 25 &#181;L 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.</li>
	<li>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).</li>
	<li>Confirm&#160;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.</li>
	<li>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.</li>
</ol>
3. Euthanasia of mice
<ol>
	<li>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.</li>
	<li>Wait 10 min post-injection until the mouse enters a surgical plane of anesthesia, confirmed by lack of response to toe pinch.</li>
</ol>
4. Perfusion of lungs with PBS and formalin
<ol>
	<li>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&#39;s superior half over its head, pulling out the arms from the skin.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
5. Air-inflating the perfused lung
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
6. Immersion fixation and dehydration
<ol>
	<li>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 &#176;C for 24-48 h.</li>
	<li>Rinse the lungs in PBS, then place in a 30% sucrose-PBS (w/v) solution for 72-96 h at 4 &#176;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 &#176;C.</li>
</ol>
7. Cryosectioning
<ol>
	<li>Equilibrate the specimens in OCT cryoblocks to the -20 &#176;C temperature of the cryostat for 1 h prior to sectioning. Section the blocks at thicknesses of 20-100 &#181;m. 
	NOTE: Air inflation can lead to increased fragility of the specimen, and cutting at increased thickness can prevent tissue breakage during sectioning.</li>
	<li>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 &#176;C at this stage.</li>
</ol>
8. Slide preparation and blocking
<ol>
	<li>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 &#181;m) 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.</li>
	<li>Dry the slides after OCT has dissolved away from the tissue. Use a wipe to dry excess droplets from the slide&#39;s perimeter around the tissue specimen. </li>
	<li>Use a hydrophobic Pap pen to draw a perimeter around the specimen, and let it dry for 5 min.</li>
	<li>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 &#181;L 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.</li>
	<li>Leave the blocking solution on the slides for 1 h at RT.</li>
</ol>
9. Immunostaining
<ol>
	<li>Wash the slide by pipetting off the fluid and adding 300-500 &#181;L of PBS.&#160;Repeat the wash once.</li>
	<li>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 &#181;L per slide of this solution and stain overnight at 4 &#176;C. 
	NOTE: Antibodies should be tested empirically for staining concentration, time, and temperature. Thin sections (8-20 &#181;m) are stained for 30 min at 37 &#176;C and thicker sections (20-100 &#181;m) at 4 &#176;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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
10. Imaging via fluorescent microscopy
<ol>
	<li>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.</li>
	<li>Collect sufficient image tiles to capture the entire lung lobe (e.g., ~200 tiles of size 400 &#956;m x 400 &#956;m). Use the microscope software program (e.g., Zeiss ZEN 3.7 or similar) to export the merged tile image for downstream processing.</li>
</ol>
11. Spatial analysis via QuPath
<ol>
	<li>In QuPath, a free open-source software14, create a project file and add the fluorescent images to the file.</li>
	<li>Classify the EpCAM+ pixels as epithelium by clicking the top menu option Classify, then Pixel classification &#62; 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>To conduct spatial analysis of the spores, select the top menu option Analyze &#62; Spatial Analysis &#62; 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 &#62; Export Measurements.</li>
</ol>

Preserving Fungal Spore Localization in Lungs for Fluorescent Microscopy Analysis

<ol><li>American Society for Microbiology. One Health: Fungal Pathogens of Humans, Animals, and Plants. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=%22Report+on+an+American+Academy+of+Microbiology+Colloquium%22">Report on an American Academy of Microbiology Colloquium</a>. , Washington, DC. (2019).</li>
<li>Crossen, A. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Human+airway+epithelium+responses+to+invasive+fungal+infections%3A+A+critical+partner+in+innate+immunity%5BTitle%5D)+AND+%22J+Fungi+%28Basel%29%22%5BJournal%5D)">Human airway epithelium responses to invasive fungal infections: A critical partner in innate immunity.</a> J Fungi (Basel). 9 (1), 40(2022).</li>
<li>Yamamoto, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Particle-size+distributions+and+seasonal+diversity+of+allergenic+and+pathogenic+fungi+in+outdoor+air%5BTitle%5D)+AND+%22ISME+J%22%5BJournal%5D)">Particle-size distributions and seasonal diversity of allergenic and pathogenic fungi in outdoor air.</a> ISME J. 6 (10), 1801-1811 (2012).</li>
<li>Thakur, A. K., Kaundle, B., Singh, I. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aThakur%2C+AK+%22Mucoadhesive+Drug+Delivery+Systems+in+Respiratory+Diseases.Targeting+Chronic+Inflammatory+Lung+Diseases+Using+Advanced+Drug+Delivery+Systems%22">Mucoadhesive Drug Delivery Systems in Respiratory Diseases.Targeting Chronic Inflammatory Lung Diseases Using Advanced Drug Delivery Systems</a>. , Elsevier, Academic Press. (2020).</li>
<li>Darquenne, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Aerosol+deposition+in+health+and+disease%5BTitle%5D)+AND+%22J+Aerosol+Med+Pulm+Drug+Deliv%22%5BJournal%5D)">Aerosol deposition in health and disease.</a> J Aerosol Med Pulm Drug Deliv. 25 (3), 140-147 (2012).</li>
<li>Kradin, R. L., Mark, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+pathology+of+pulmonary+disorders+due+to+Aspergillus+spp%5BTitle%5D)+AND+%22Arch+Pathol+Lab+Med%22%5BJournal%5D)">The pathology of pulmonary disorders due to Aspergillus spp.</a> Arch Pathol Lab Med. 132 (4), 606-614 (2008).</li>
<li>Karnak, D., Avery, R. K., Gildea, T. R., Sahoo, D., Mehta, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Endobronchial+fungal+disease%3A+An+under-recognized+entity%5BTitle%5D)+AND+%22Respiration%22%5BJournal%5D)">Endobronchial fungal disease: An under-recognized entity.</a> Respiration. 74 (1), 88-104 (2006).</li>
<li>Amich, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Three-dimensional+light+sheet+fluorescence+microscopy+of+lungs+to+dissect+local+host+immune-Aspergillus+fumigatus+interactions%5BTitle%5D)+AND+%22mBio%22%5BJournal%5D)">Three-dimensional light sheet fluorescence microscopy of lungs to dissect local host immune-Aspergillus fumigatus interactions.</a> mBio. 11 (1), e02752-e02819 (2020).</li>
<li>Wiesner, D. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Club+cell+TRPV4+serves+as+a+damage+sensor+driving+lung+allergic+inflammation%5BTitle%5D)+AND+%22Cell+Host+Microbe%22%5BJournal%5D)">Club cell TRPV4 serves as a damage sensor driving lung allergic inflammation.</a> Cell Host Microbe. 27 (4), 614-628.e6 (2020).</li>
<li>Evans, S. E., Hahn, P. Y., McCann, F., Kottom, T. J., Pavlovic', Z. V., Limper, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pneumocystis+cell+wall+%CE%B2-glucans+stimulate+alveolar+epithelial+cell+chemokine+generation+through+nuclear+factor-%CE%BAB-dependent+mechanisms%5BTitle%5D)+AND+%22Am+J+Respir+Cell+Mol+Biol%22%5BJournal%5D)">Pneumocystis cell wall β-glucans stimulate alveolar epithelial cell chemokine generation through nuclear factor-κB-dependent mechanisms.</a> Am J Respir Cell Mol Biol. 32 (6), 490-497 (2005).</li>
<li>Bauer, C., Krueger, M., Lamm, W. J. E., Glenny, R. W., Beichel, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((lapdMouse%3A+associating+lung+anatomy+with+local+particle+deposition+in+mice%5BTitle%5D)+AND+%22J+Appl+Physiol%22%5BJournal%5D)">lapdMouse: associating lung anatomy with local particle deposition in mice.</a> J Appl Physiol. 128 (2), 309-323 (2020).</li>
<li>Srirama, P. K., Wallis, C. D., Lee, D., Wexler, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Imaging+extra-thoracic+airways+and+deposited+particles+in+laboratory+animals%5BTitle%5D)+AND+%22J+Aerosol+Sci%22%5BJournal%5D)">Imaging extra-thoracic airways and deposited particles in laboratory animals.</a> J Aerosol Sci. 45, 40-49 (2012).</li>
<li>Thomas, S. M., Bednarek, J., Janssen, W. J., Hume, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Air-inflation+of+murine+lungs+with+vascular+perfusion-fixation%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Air-inflation of murine lungs with vascular perfusion-fixation.</a> J Vis Exp. 168, e62215(2021).</li>
<li>Bankhead, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((QuPath%3A+Open+source+software+for+digital+pathology+image+analysis%5BTitle%5D)+AND+%22Sci+Rep%22%5BJournal%5D)">QuPath: Open source software for digital pathology image analysis.</a> Sci Rep. 7 (1), 16878(2017).</li>
<li>Davenport, M. L., Sherrill, T. P., Blackwell, T. S., Edmonds, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Perfusion+and+inflation+of+the+mouse+lung+for+tumor+histology%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Perfusion and inflation of the mouse lung for tumor histology.</a> J Vis Exp. 162, e60605(2020).</li>
<li>Hsia, C. C. W., Hyde, D. M., Ochs, M., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+official+research+policy+statement+of+the+American+Thoracic+Society%2FEuropean+Respiratory+Society%3A+Standards+for+quantitative+assessment+of+lung+structure%5BTitle%5D)+AND+%22Am+J+Respir+Crit+Care+Med%22%5BJournal%5D)">An official research policy statement of the American Thoracic Society/European Respiratory Society: Standards for quantitative assessment of lung structure.</a> Am J Respir Crit Care Med. 181 (4), 394-418 (2010).</li>
<li>Gil, J., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Extracellular+lining+of+bronchioles+after+perfusion-fixation+of+rat+lungs+for+electron+microscopy%5BTitle%5D)+AND+%22Anat+Rec%22%5BJournal%5D)">Extracellular lining of bronchioles after perfusion-fixation of rat lungs for electron microscopy.</a> Anat Rec. 169 (2), 185-199 (1971).</li>
<li>Bachofen, H., Ammann, A., Wangensteen, D., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Perfusion+fixation+of+lungs+for+structure-function+analysis%3A+credits+and+limitations%5BTitle%5D)+AND+%22J+Appl+Physiol+Respir+Environ+Exerc+Physiol%22%5BJournal%5D)">Perfusion fixation of lungs for structure-function analysis: credits and limitations.</a> J Appl Physiol Respir Environ Exerc Physiol. 53 (2), 528-533 (1982).</li>
<li>Evans, C. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+polymeric+mucin+Muc5ac+is+required+for+allergic+airway+hyperreactivity%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">The polymeric mucin Muc5ac is required for allergic airway hyperreactivity.</a> Nat Commun. 6, 6281(2015).</li>
<li>Galati, D. F., Asai, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Immunofluorescence+microscopy%5BTitle%5D)+AND+%22Curr+Protoc%22%5BJournal%5D)">Immunofluorescence microscopy.</a> Curr Protoc. 3 (8), e842(2023).</li>
<li>Hickey, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fluorescence+microscopy-An+outline+of+hardware%2C+biological+handling%2C+and+fluorophore+considerations%5BTitle%5D)+AND+%22Cells%22%5BJournal%5D)">Fluorescence microscopy-An outline of hardware, biological handling, and fluorophore considerations.</a> Cells. 11 (1), 35(2021).</li>
<li>Risling, T. E., Caulkett, N. A., Florence, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Open-drop+anesthesia+for+small+laboratory+animals%5BTitle%5D)+AND+%22Can+Vet+J%22%5BJournal%5D)">Open-drop anesthesia for small laboratory animals.</a> Can Vet J. 53 (3), 299-302 (2012).</li>
<li>Bodnar, M. J., Ratuski, A. S., Weary, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mouse+isoflurane+anesthesia+using+the+drop+method%5BTitle%5D)+AND+%22Lab+Anim%22%5BJournal%5D)">Mouse isoflurane anesthesia using the drop method.</a> Lab Anim. 57 (6), 623-630 (2023).</li>
</ol>

The authors have nothing to disclose.

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/cts3&#916; 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 alve...

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 &#181;m and 10 &#181;m 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 (&#62; 6 &#181;m) deposit in the upper airway, medium-sized particles (2-6 &#181;m) can deposit in smaller airways, and small particles (&#60;2 &#181;m) reach the alveolar region5. Aspergillus spores (2-3 &#181;m) 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 work2,9,10.
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/cts3&#916; arthroconidia (3-5 &#181;m in size) accumulate more proximally than previously shown, namely in distal bronchioles and broncho-alveolar junctions rather than terminal alveoli. This information can focus&#160;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.

<table><tbody><tr><td>18 G, 1 1/2 needle (305185)</td><td>Fisher</td><td>305185</td><td></td></tr><tr><td>1 L&amp;nbsp; Screwtop Jar&amp;nbsp; (Nalgene)</td><td>Fisher Scientific</td><td>11-823-33</td><td></td></tr><tr><td>Air-Tite Bulk Unsterile Syringes 10 mL Luer Lock</td><td>Fisher</td><td>14-817-175</td><td></td></tr><tr><td>AnaSed Injection (xylazine sterile solution)</td><td>Akorn</td><td>59399-110-20</td><td></td></tr><tr><td>Animal-Free Blocker and Diluent, R.T.U.</td><td>Vector</td><td>SP-5035-100</td><td></td></tr><tr><td>BD Insyte Autoguard Winged Shielded IV Catheter with BD Vialon Catheter Material 18 G x 1.88 in</td><td>BD</td><td>381547</td><td></td></tr><tr><td>BD Pharmingen Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block&amp;trade;)</td><td>BD</td><td>553142</td><td></td></tr><tr><td>CellTracker Orange CMRA Dye</td><td>Fisher Scientific</td><td>NC0873640</td><td></td></tr><tr><td>CFSE</td><td>Labviva</td><td>75003</td><td></td></tr><tr><td>Coccidioides posadasii cts2/ard1/cts3&amp;Delta;&amp;nbsp;</td><td>BEI Resources</td><td>NR-166</td><td></td></tr><tr><td>Corning 70 micron strainers</td><td>VWR</td><td>10054-456</td><td></td></tr><tr><td>EpCAM AlexaFluor647 monoclonal antibody</td><td>Biolegend</td><td>118211</td><td></td></tr><tr><td>Exel International HYpodermic Needles 30 G x 1/2&quot;</td><td>Labviva</td><td>EN3012</td><td></td></tr><tr><td>Fisherbrand Sterile Syringes for Single Use (1mL, Leur Slip)</td><td>Fisher</td><td>14-955-462</td><td></td></tr><tr><td>Glucose Monohydrate</td><td>Azer Scientific&amp;nbsp;</td><td>ES17530-500G</td><td></td></tr><tr><td>High Vacuum Grease</td><td>VWR</td><td>59344-055</td><td></td></tr><tr><td>Hoechst 33342 Solution 20 mM (5 mL)</td><td>ThermoFischer</td><td>62249</td><td></td></tr><tr><td>Isoflurane USP</td><td>Covetrus</td><td>29405</td><td></td></tr><tr><td>Ketamine Hydrochloride</td><td>Dechra</td><td>1000001250</td><td></td></tr><tr><td>KIMWIPES Delicate Task Wipers (4.4&#039;&#039; x 8.4&quot;)</td><td>VWR</td><td>21905-026</td><td></td></tr><tr><td>Lexer Baby Scissors</td><td>FST</td><td>14078-10</td><td></td></tr><tr><td>Micro-Adson Forceps</td><td>FST</td><td>11018-12</td><td></td></tr><tr><td>Neutral Buffered Formalin (10%) (Azer Scientific)</td><td>Fisher</td><td>22-026-350</td><td></td></tr><tr><td>Nunc EasYFlask tissue culture flasks, T75, filter caps</td><td>VWR</td><td>15708-134</td><td></td></tr><tr><td>PBS</td><td>VWR</td><td>45000-446</td><td></td></tr><tr><td>Peel-A-Way embedding molds</td><td>Sigma</td><td>E6032-1CS</td><td></td></tr><tr><td>QuPath 0.5.1 Software</td><td>Open-source</td><td>https://qupath.github.io/</td><td></td></tr><tr><td>Silk Suture thread size 3/0</td><td>FST</td><td>18020-30</td><td></td></tr><tr><td>SlidesMicro Slides Superfrost Plus&amp;nbsp;</td><td>VWR</td><td>48311-703</td><td></td></tr><tr><td>SlowFade Glass Soft-set Antifade Mountant (2 mL)</td><td>ThermoFischer</td><td>S36917</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S0389-500G</td><td></td></tr><tr><td>Tissue-Tek O.C.T. Compound, Sakura Finetek</td><td>VWR</td><td>25608-930</td><td></td></tr><tr><td>Tween 20</td><td>ThermoFischer</td><td>J20605.AP</td><td></td></tr><tr><td>Vector Laboratories ImmEDGE Hydrophobic Barrier Pen Set Of 2</td><td>Fisher Scientific</td><td>NC9545623</td><td></td></tr><tr><td>VWR Micro Cover Glasses, Rectangular (24 mm x 40 mm #1.5)</td><td>VWR</td><td>48393-230</td><td></td></tr><tr><td>White Plastic Wire Mesh</td><td>MAPORCH</td><td>789862904922</td><td></td></tr><tr><td>Yeast Extract</td><td>Fisher</td><td>BP9727-500</td><td></td></tr><tr><td>Zeiss AxioScan 7&amp;nbsp;</td><td>Carl Zeiss Microscopy GmbH</td><td>https://www.zeiss.com/microscopy/us/products/imaging-systems/axioscan-for-biology.html</td><td>multichannel fluorescent microscope</td></tr><tr><td>ZEN 3.7 Software</td><td>Carl Zeiss Microscopy GmbH</td><td>https://www.zeiss.com/microscopy/us/products/software/zeiss-zen.html</td><td>microscopy software</td></tr></tbody></table>

efficient method for imaging murine lungs that preserves spatial dynamics of fungal spores in the airways

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/cts3&#916; 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.

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 &#34;crackles&#34; 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 ...

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

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.

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and ...

efficient-method-for-imaging-murine-lungs-that-preserves-spatial

Research

JoVE Journal

Immunology and Infection

309 Views.  Rutgers - The State University of New Jersey. Dit onderzoek heeft tot doel een efficiënt protocol op te stellen voor het behoud van de luchtweginhoud van muizen in longinfectiemodellen. We ontwikkelen het vermogen om de vraag te beantwoorden waar ingeademde schimmelsporen in de longen terechtkomen. Onlangs hebben veel groepen de rol geïdentificeerd van verschillende luchtwegepitheelcellen bij het detecteren van infecties en het initiëren van beschermende immuunresponsen.Deze epitheelcellen vari...

Video: Efficiënte methode voor het in beeld brengen van muizenlongen die de ruimtelijke dynamiek van schimmelsporen in de luchtwegen behoudt

Long-term High-Resolution Intravital Microscopy in the Lung with a Vacuum Stabilized Imaging Window

Metastasis to secondary sites such as the lung, liver and bone is a traumatic event with a mortality rate of approximately 90% 1. Of these sites, the lung is the most difficult to assess using intravital optical imaging due to its enclosed position within the body, delicate nature and vital role in sustaining proper physiology. While clinical modalities (positron emission tomography (PET), magnetic resonance imaging (MRI) and computed tomography (CT)) are capable of providing noninvasive images of this tissue, they lack the resolution necessary to visualize the earliest seeding events, with a single pixel consisting of nearly a thousand cells. Current models of metastatic lung seeding postulate that events just after a tumor cell&#39;s arrival are deterministic for survival and subsequent growth. This means that real-time intravital imaging tools with single cell resolution 2 are required in order to define the phenotypes of the seeding cells and test these models. While high resolution optical imaging of the lung has been performed using various ex vivo preparations, these experiments are typically single time-point assays and are susceptible to artifacts and possible erroneous conclusions due to the dramatically altered environment (temperature, profusion, cytokines, etc.) resulting from removal from the chest cavity and circulatory system 3. Recent work has shown that time-lapse intravital optical imaging of the intact lung is possible using a vacuum stabilized imaging window 2,4,5 however, typical imaging times have been limited to approximately 6 hr. Here we describe a protocol for performing long-term intravital time-lapse imaging of the lung utilizing such a window over a period of 12 hr. The time-lapse image sequences obtained using this method enable visualization and quantitation of cell-cell interactions, membrane dynamics and vascular perfusion in the lung. We further describe an image processing technique that gives an unprecedentedly clear view of the lung microvasculature.

This protocol describes the use of multiphoton microscopy to perform long-term high-resolution, single cell imaging of the intact lung in real time using a vacuum stabilized imaging window.

Metastasis to secondary sites such as the lung, liver and bone is a traumatic event with a mortality rate of approximately 90% 1. Of these sites, the lung ...

Cancer Research

Direct Observation of Phagocytosis and NET-formation by Neutrophils in Infected Lungs using 2-photon Microscopy

After the gastrointestinal tract, the lung is the second largest surface for interaction between the vertebrate body and the 
environment. Here, an effective gas exchange must be maintained, while at the same time avoiding infection by the multiple pathogens that are inhaled 
during normal breathing. To achieve this, a superb set of defense strategies combining humoral and cellular immune mechanisms exists. One of the most 
effective measures for acute defense of the lung is the recruitment of neutrophils, which either phagocytose the inhaled pathogens or kill them by 
releasing cytotoxic chemicals. A recent addition to the arsenal of neutrophils is their explosive release of extracellular DNA-NETs by which bacteria 
or fungi can be caught or inactivated even after the NET releasing cells have died. We present here a method that allows one to directly observe neutrophils, 
migrating within a recently infected lung, phagocytosing fungal pathogens as well as visualize the extensive NETs that they have produced throughout the 
infected tissue. The method describes the preparation of thick viable lung slices 7 hours after intratracheal infection of mice with conidia of the mold 
Aspergillus fumigatus and their examination by multicolor time-lapse 2-photon microscopy. This approach allows one to directly investigate antifungal 
defense in native lung tissue and thus opens a new avenue for the detailed investigation of pulmonary immunity.

We show, how to use 2-photon microscopy for the observation of the dynamics of neutrophil granulocytes in infected lungs while they phagocytose pathogens or produce neutrophil extracellular traps (NETs).

After the gastrointestinal tract, the lung is the second largest surface for interaction between the vertebrate body and the 
environment. Here, an ...

Ex vivo Live Imaging of Lung Metastasis and Their Microenvironment

Metastasis is a major cause for cancer-related morbidity and mortality. Metastasis is a multistep process and due to its complexity, the exact cellular and molecular processes that govern metastatic dissemination and growth are still elusive. Live imaging allows visualization of the dynamic and spatial interactions of cells and their microenvironment. Solid tumors commonly metastasize to the lungs. However, the anatomical location of the lungs poses a challenge to intravital imaging. This protocol provides a relatively simple and quick method for ex vivo live imaging of the dynamic interactions between tumor cells and their surrounding stroma within lung metastasis. Using this method, the motility of cancer cells as well as interactions between cancer cells and stromal cells in their microenvironment can be visualized in real time for several hours. By using transgenic fluorescent reporter mice, a fluorescent cell line, injectable fluorescently labeled molecules and/or antibodies, multiple components of the lung microenvironment can be visualized, such as blood vessels and immune cells. To image the different cell types, a spinning disk confocal microscope that allows long-term continuous imaging with rapid, four-color image acquisition has been used. Time-lapse movies compiled from images collected over multiple positions and focal planes show interactions between live metastatic and immune cells for at least 4 hr. This technique can be further used to test chemotherapy or targeted therapy. Moreover, this method could be adapted for the study of other lung-related pathologies that may affect the lung microenvironment.

Ex vivo Live Imaging of Lung Metastasis and Their Microenvironment

We describe a relatively simple method for ex vivo live imaging of the tumor cell-stroma interactions within lung metastasis, utilizing fluorescent reporters in mice. Using spinning-disk confocal microscopy, this technique enables visualization of live cells for at least 4 hr and could be adapted to study other inflammatory lung conditions.

Metastasis is a major cause for cancer-related morbidity and mortality. Metastasis is a multistep process and due to its complexity, the exact cellular ...

Medicine

A Multimodal Imaging Approach Based on Micro-CT and Fluorescence Molecular Tomography for Longitudinal Assessment of Bleomycin-Induced Lung Fibrosis in Mice

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which causes significant deterioration in lung function and subsequent death from respiratory failure.
The pathogenesis of IPF in experimental animal models has been induced by bleomycin administration. In this study, we investigate an IPF-like mouse model induced by a double intratracheal bleomycin instillation. Standard histological assessments used for studying lung fibrosis are invasive terminal procedures. The goal of this work is to monitor lung fibrosis through noninvasive imaging techniques such as Fluorescent Molecular Tomography (FMT) and Micro-CT. These two technologies validated with histology findings could represent a revolutionary functional approach for real time non-invasive monitoring of IPF disease severity and progression. The fusion of different approaches represents a step further for understanding the IPF disease, where the molecular events occurring in a pathological condition can be observed with FMT and the subsequent anatomical changes can be monitored by Micro-CT.

We describe a non-invasive multimodal imaging approach based on Micro-CT and fluorescence molecular tomography for longitudinal assessment of the mouse lung fibrosis model induced by double intratracheal instillation of bleomycin.

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which ...

Imaging Mycobacterium tuberculosis in Mice with Reporter Enzyme Fluorescence

Reporter enzyme fluorescence (REF) utilizes substrates that are specific for enzymes present in target organisms of interest for imaging or detection by fluorescence or bioluminescence. We utilize BlaC, an enzyme expressed constitutively by all M. tuberculosis strains. REF allows rapid quantification of bacteria in lungs of infected mice. The same group of mice can be imaged at many time points, greatly reducing costs, enumerating bacteria more quickly, allowing novel observations in host-pathogen interactions, and increasing statistical power, since more animals per group are readily maintained. REF is extremely sensitive due to the catalytic nature of the BlaC enzymatic reporter and specific due to the custom flourescence resonance energy transfer (FRET) or fluorogenic substrates used. REF does not require recombinant strains, ensuring normal host-pathogen interactions. We describe the imaging of M. tuberculosis infection using a FRET substrate with maximal emission at 800 nm. The wavelength of the substrate allows sensitive deep tissue imaging in mammals. We will outline aerosol infection of mice with M. tuberculosis, anesthesia of mice, administration of the REF substrate, and optical imaging. This method has been successfully applied to evaluating host-pathogen interactions and efficacy of antibiotics targeting M. tuberculosis.

Imaging Mycobacterium tuberculosis in Mice with Reporter Enzyme Fluorescence

We describe the optical imaging of mice infected with Mycobacterium tuberculosis (M. tuberculosis) using reporter enzyme fluorescence (REF). This protocol facilitates the sensitive and specific detection of M. tuberculosis in pre-clinical animal models for pathogenesis, therapeutics and vaccine research.

Reporter enzyme fluorescence (REF) utilizes substrates that are specific for enzymes present in target organisms of interest for imaging or detection by ...

Histological Quantification to Determine Lung Fungal Burden in Experimental Aspergillosis

The quantification of lung fungal burden is critical for the determination of the relative levels of immune protection and fungal virulence in mouse models of pulmonary fungal infection. Although multiple methods are used to assess fungal burden, quantitative polymerase chain reaction (qPCR) of fungal DNA has emerged as a technique with several advantages over previous culture-based methods. Currently, a comprehensive assessment of lung pathology, leukocyte recruitment, fungal burden, and gene expression in mice with invasive aspergillosis (IA) necessitates the use of a significant number of experimental and control animals. Here the quantification of lung histological staining to determine fungal burden using a reduced number of animals was examined in detail. Lung sections were stained to identify fungal structures with Gomori&#39;s modified methanamine silver (GMS) staining. Images were taken from the GMS-stained sections from 4 discrete fields of each formalin-fixed paraffin-embedded lung. The GMS stained areas within each image were quantified using an image analysis program, and from this quantification, the mean percentage of stained area was determined for each sample. Using this strategy, eosinophil-deficient mice exhibited decreased fungal burden and disease with caspofungin therapy, while wild-type mice with IA did not improve with caspofungin. Similarly, fungal burden in mice lacking &#947;&#948; T cells were also improved by caspofungin, as measured by qPCR and GMS quantification. GMS quantification is therefore introduced as a method for the determination of relative lung fungal burden that may ultimately reduce the quantity of experimental animals required for comprehensive studies of invasive aspergillosis.

Here we describe a protocol to determine pulmonary fungal burden in mice with invasive aspergillosis by quantification of Gomori&#39;s modified methanamine silver staining in histological sections. Use of this method resulted in comparable results with less animals compared to assessment of fungal burden by quantitative PCR of lung fungal DNA.

The quantification of lung fungal burden is critical for the determination of the relative levels of immune protection and fungal virulence in mouse ...

Imaging Cell Interaction in Tracheal Mucosa During Influenza Virus Infection Using Two-photon Intravital Microscopy

The analysis of cell-cell or cell-pathogen interaction in vivo is an important tool to understand the dynamics of the immune response to infection. Two-photon intravital microscopy (2P-IVM) allows the observation of cell interactions in deep tissue in living animals, while minimizing the photobleaching generated during image acquisition. To date, different models for 2P-IVM of lymphoid and non-lymphoid organs have been described. However, imaging of respiratory organs remains a challenge due to the movement associated with the breathing cycle of the animal.
Here, we describe a protocol to visualize in vivo immune cell interactions in the trachea of mice infected with influenza virus using 2P-IVM. To this purpose, we developed a custom imaging platform, which included the surgical exposure and intubation of the trachea, followed by the acquisition of dynamic images of neutrophils and dendritic cells (DC) in the mucosal epithelium. Additionally, we detailed the steps needed to perform influenza intranasal infection and flow cytometric analysis of immune cells in the trachea. Finally, we analyzed neutrophil and DC motility as well as their interactions during the course of a movie. This protocol allows for the generation of stable and bright 4D images necessary for the assessment of cell-cell interactions in the trachea.

In this study, we present a protocol to perform two-photon intravital imaging and cell interaction analysis in the murine tracheal mucosa after infection with influenza virus. This protocol will be relevant for researchers studying immune cell dynamics during respiratory infections.

The analysis of cell-cell or cell-pathogen interaction in vivo is an important tool to understand the dynamics of the immune response to infection. ...

Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung

Aspergillus fumigatus conidia are airborne pathogens that can penetrate human airways. Immunocompetent people without allergies exhibit resistance and immunological tolerance, while in immunocompromised patients, conidia can colonize airways and cause severe invasive respiratory disorders. Various cells in different airway compartments are involved in the immune response that prevents fungal invasion; however, the spatio-temporal aspects of pathogen elimination are still not completely understood. Three-dimensional (3D) imaging of optically cleared whole-mount organs, particularly the lungs of experimental mice, permits detection of fluorescently labeled pathogens in the airways at different time points after infection. In the present study, we describe an experimental setup to perform a quantitative analysis of A. fumigatus conidia distribution in the airways. Using fluorescent confocal laser scanning microscopy (CLSM), we traced the location of fluorescently labeled&#160;conidia in the bronchial branches and the alveolar compartment 6 hours after oropharyngeal application to mice. The approach described here was previously used for detection of the precise pathogen location and identification of the pathogen-interacting cells at different phases of the immune response. The experimental setup can be used to estimate the kinetics of the pathogen elimination in different pathological conditions.

Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung

We describe the method for quantitative analysis of the distribution of Aspergillus fumigatus conidia (3 &#181;m in size) in the airways of mice. The method also can be used for the analysis of microparticles and nanoparticle agglomerate distribution in the airways in various pathological condition models.

Aspergillus fumigatus conidia are airborne pathogens that can penetrate human airways. Immunocompetent people without allergies exhibit resistance and ...

Advanced Imaging of Lung Homing Human Lymphocytes in an Experimental In Vivo Model of Allergic Inflammation Based on Light-sheet Microscopy

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an attractive therapeutic target in the clinical management of affected patients. In order to further optimize strategies aiming at therapeutic regulation of pathologically imbalanced tissue infiltration of pro-inflammatory immune cells, it will be of particular importance to achieve improved insights into disease- and organ-specific homing properties of peripheral lymphocytes. The here described experimental protocol allows to monitor lung accumulation of fluorescently labeled and adoptively transferred human lymphocytes in the context of papain-induced pulmonary inflammation. In contrast to standard in vitro assays frequently used for the analysis of immune cell migration and chemotaxis, the now introduced in vivo setting takes into account lung-specific aspects of tissue organization and the influence of the complex inflammatory scenario taking place in the living murine organism. Moreover, three-dimensional cross-sectional light-sheet fluorescence microscopic imaging does not only provide quantitative data on infiltrating immune cells, but also depicts the pattern of immune cell localization within the inflamed lung. Overall, we are able to introduce an innovative technique of high value for immunological research in the field of chronic inflammatory lung diseases, which can be easily applied by following the provided step-by-step protocol.

The here introduced protocol allows characterization of the lung homing capacity of primary human lymphocytes under in vivo inflammatory conditions. Pulmonary infiltration of adoptively transferred human immune cells in a mouse model of allergic inflammation can be imaged and quantified by light-sheet fluorescence microscopy of chemically cleared lung tissue.

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an ...

In vivo Evaluation of Mucociliary Clearance in Mice

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a metachronal, synchronal fashion, these multiple, motile, actin-based organelles generate a cephalad fluid flow clearing the respiratory tract of inhaled pollutants and pathogens. With increasing environmental pollution, novel viral pathogens and emerging multi-drug resistant bacteria, cilia generated mucociliary clearance (MCC) is essential for maintaining lung health. MCC is also depressed in multiple congenital disorders like primary ciliary dyskinesia, cystic fibrosis as well as acquired disorders like chronic obstructive pulmonary disease. All these disorders have established, in some case multiple, mouse models. In this publication, we detail a method using a small amount of radioactivity and dual-modality SPECT/CT imaging to accurately and reproducibly measure MCC in mice in vivo. The method allows for recovery of mice after imaging, making serial measurements possible, and testing potential therapeutics longitudinally over time. The data in wild-type mice demonstrates the reproducibility of the MCC measurement as long as adequate attention to detail is paid, and the protocol strictly adhered to.

In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a ...

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

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