This work was funded by the National Heart, Lung, and Blood Institute (NHLBI) grants HL140039 and HL130938. The authors would like to thank Shannon Hott and Jazalle McClendon for their technical expertise.

All methods described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of National Jewish Health.
NOTE: The protocol is organized into three components. The first component details the construction of the air-inflation with perfusion/fixation equipment. A second section describes how to set up the equipment for an experiment. The final section describes how to prepare the animal and perform the experiment.
1. Construction of the water column apparatus (Figure 1B)
<ol>
	<li>Remove the plunger from a 60 mL slip tip syringe.</li>
	<li>Attach tape around the syringe at the 30 mL mark. Set the height of the syringe to this mark for the initial inflation pressure of 25 cmH2O. This is also where the water level in the column should be throughout the entire procedure. Label the tape either as &#34;25 cm&#34; (as depicted in Figure 1A) or &#34;inflation&#34;. 
	NOTE: An inflation pressure of 25 cmH2O is used to ensure recruitment of collapsed airspaces. Once alveoli have been recruited, the pressure is lowered to 20 cmH2O to ensure that the airspaces are not distended.</li>
	<li>Measure 5 cm from the tape towards the plunger end and attach another piece of tape to the syringe. Move the syringe down to this mark to lower the inflation pressure to 20 cmH2O during fixation. Label the tape either as &#34;20 cm&#34; (as depicted in Figure 1A) or &#34;fixation&#34;.</li>
	<li>Attach 180 polyvinyl chloride (PVC) tubing to the slip tip end of the syringe. The length of tubing is dependent upon the distance between the syringe and air inflation chamber (approximately 25-30 cm).</li>
	<li>Place a male Luer (2) thread style tee (1.219 inches length, 0.904 inches height, 0.0904 inches inner diameter) into the other end of the tubing. This male Luer will connect to the stopcock of the air-inflation chamber (Step 2.4).</li>
</ol>
2. Construction of the air-inflation chamber (Figure 1C)
<ol>
	<li>Drill two holes (approximately 4 mm in diameter) in a 500 mL plastic container with a screw-on cap. The holes should be the same size as the female Luers (1.224 inches length, 0.312 inches height, 0.098 inches inner diameter). 
	NOTE: Empty media or buffer containers can be utilized for the air-inflation chamber.</li>
	<li>Coat threads of female Luers with silicone gasket maker and place the smaller side into the pre-drilled holes of the container.</li>
	<li>Add silicone gasket maker around the female Luers where they enter the container to ensure an air-tight seal.</li>
	<li>Screw on a one-way stopcock on the lower female Luer on the air-inflation chamber.</li>
	<li>Cut tubing to the length of roughly 25 cm and attach male Luers to the free ends of the tubing. Connect the male Luer on one end of the tubing to the free female Luer on the air-inflation chamber. The other male Luer will connect to the animal-processing container.</li>
</ol>
3. Construction of the animal-processing container (Figure 1D)
<ol>
	<li>Drill a hole (approximately 4 mm diameter) in the side of a large plastic container. The hole should be the diameter of the female Luer. The plastic container is needed to catch excess fixation solution. 
	NOTE: A 30 cm x 22 cm, 3.8 L plastic storage container was used.</li>
	<li>Coat threads of a female Luer with a silicone gasket maker and place the smaller side into the predrilled hole of the container.</li>
	<li>Add silicone gasket maker around the female Luer where it enters and exits the container to ensure an air-tight seal.</li>
	<li>Screw a one-way stopcock onto the female Luer. Tubing from the air-inflation chamber will attach to this stopcock.</li>
</ol>
4. Preparation of solutions
<ol>
	<li>Heparin solution
	<ol>
		<li>Fill a container with calcium-free PBS and heparin (20 U/mL). Prepare a total of 10 mL heparin solution for each mouse. Heparin is an anti-coagulant that prevents blood clots from forming in the vessels during perfusion-fixation. Heparin solution will be used to flush blood from the lungs prior to perfusion-fixation.</li>
	</ol>
	</li>
	<li>Fixative solution 
	CAUTION: Fixatives can present a health hazard and should be used in a chemical fume hood. All apparatuses are set up in a chemical fume hood to prevent inhalation of fixatives.
	<ol>
		<li>Fill a container with calcium-free PBS and paraformaldehyde (4% final concentration). Prepare a total of 50 mL fixative solution for each mouse. 
		&#8203;NOTE: The type of fixative used may vary and will depend upon downstream histologic studies.</li>
	</ol>
	</li>
</ol>
5.&#160;Preparation of perfusion apparatus
NOTE: A peristaltic pump is suggested for delivery of fluids into vasculature to ensure constant flow rate. The following directions are for setting up the peristaltic pump and may be different for each model. Alternatively, if a peristaltic pump is unavailable, a second water column apparatus may be constructed to perfuse fluids from a height of 35 cm H2O.
<ol>
	<li>First place the tubing around the roller assembly.</li>
	<li>Secure the tubing into the notched posts.</li>
	<li>Latch the levers into place by first placing the left lever around the tubing and securing it in place with the top and then the right lever.</li>
	<li>Place the proximal end of the tubing into the heparin solution and the distal end into the animal processing container.</li>
	<li>Preload heparin solution into the tubing by running the pump to expel air from the tubing.</li>
	<li>Secure a 25G x 5/8&#34; needle onto the end of the left side of the tubing.</li>
</ol>
6. Preparation of air-inflation apparatus
<ol>
	<li>Place the syringe for the water column into a ring holder.</li>
	<li>Measure a vertical height of 25 cm from the animal platform to the &#34;25 cm&#34; tape mark (Step 1.2) on the water column.</li>
	<li>Attach the end of the water column&#39;s tube to the stopcock on the air chamber.</li>
	<li>Attach a tube from the female Luer of the air chamber to the stopcock on the animal processing container. 
	NOTE: If the air-inflation chamber is constructed as depicted in Figure 1C, attaching tubing in reverse order may cause water to leak into the tubing that connects to the tracheal cannula.</li>
	<li>Ensure that the cap to the air chamber is tightly closed.</li>
	<li>Ensure that the stopcock on the outside of the animal processing container is closed and the stopcock on the tubing leading from the water column to the air-inflation chamber is open.</li>
	<li>Fill the syringe with water to the &#34;25 cm&#34; mark. Water will leave the syringe through the tubing into the air chamber. Once pressure is equalized water will stop flowing.
	<ol>
		<li>It is possible that water will slowly leak into the air-inflation chamber as ambient air pressure within the chemical hood fluctuates. Keep an eye on the water level within the syringe and add more if necessary. Maintain the water level at the &#34;25 cm&#34; mark throughout the procedure. 
		NOTE: Usually water levels will stay consistent for the first part of air inflation at 25 cmH2O; however more water will likely need to be added to the syringe during fixation. If the water does not stop flowing, it is likely that an air leak exists within the air-inflation chamber. More silicone gasket maker may need to be applied around the Luers to prevent air leaks.</li>
	</ol>
	</li>
</ol>
7. Preparation of animals (Figure 2)
NOTE: This procedure has been modified from Gage et al13. We have completed this procedure on adult male&#160;and female mice of varying ages and note no&#160;age or sex bias.
<ol>
	<li>Euthanize the animal with sodium pentobarbital (150 mg/kg, intraperitoneally.). Ensure that the animal is dead prior to beginning dissection. 
	NOTE: While this procedure is performed on euthanized animals, this procedure can be performed on live animals to utilize the heart to pump perfusates throughout the animal.</li>
	<li>Make two lateral incisions through the abdominal wall. Make the first incision below the rib cage and the second above the hips. Cut along the midline from the inferior incision towards the superior incision.</li>
	<li>Using blunt scissors, carefully make an incision into the lateral side of the diaphragm. The lungs should collapse as soon as the diaphragm is punctured. 
	NOTE: Care must be taken to avoid puncturing the lungs. A punctured lung is less likely to inflate during later steps.</li>
	<li>Cut transversely along the diaphragm to open the thoracic cavity.</li>
	<li>Cut superiorly along the sternum from the xiphoid process to the jugular notch and laterally above the rib cage to fully expose the heart and lungs. Pin down the sides of the ribcage.</li>
	<li>Make a midline incision in the neck above the trachea. Remove skin, muscle, thyroid gland and connective tissue surrounding the trachea.</li>
	<li>Using curved forceps, slide two pieces of thread or suture under the posterior trachea. Use one piece of suture to hold the inflation Luer-stub adapter in place, and use the other ultimately to tie off the trachea at the conclusion of air-inflation and vascular perfusion-fixation.</li>
	<li>Poke a small hole in the trachea using an 18G x 1&#34; needle or Vannas spring scissors.</li>
	<li>Place a 20G Luer-stub adapter into this hole in the trachea.</li>
	<li>Tie one thread around the trachea immediately distal to where the Luer-stub adapter enters to hold it in place.</li>
	<li>Transfer the animal to the animal processing container.</li>
	<li>Attach the Luer-stub adapter to the female Luer on the inside of the animal processing container.</li>
</ol>
8. Air inflation, perfusion and fixation of the lungs (Figure 2)
<ol>
	<li>Place the 25G x 5/8&#34; needle attached to the perfusion apparatus tubing into the right ventricle of the heart.</li>
	<li>Cut the abdominal aorta to allow blood to drain from the heart and to promote flow of perfusate through the lungs.</li>
	<li>Open the stopcock on the outside of the animal processing container to inflate the lungs. 
	NOTE: It may take time for the lungs to fully inflate. Watch the water level in the syringe, it should not decrease rapidly unless there is a leak in the lungs.</li>
	<li>Inflate the lungs at 25 cmH2O for 5 minutes. Inflation at 25 cmH2O preconditions the lung and assists in recruiting atelectatic lung regions. 
	&#8203;NOTE: A small amount of water may need to be added to the syringe to maintain the 25 cm height. Inflammation and/or experimentally induced lung injury may influence inflation of the lungs. In this case, it may be necessary to increase the inflation pressure up to a maximum of 35 cmH2O to assist in recruiting atelectatic regions.</li>
	<li>During the last minute of the lung inflation, turn on the peristaltic pump to a flow rate of 10 mL/min. Heparin solution should flow from the bottle through the tubing into the animal.
	<ol>
		<li>The goal of heparin infusion is to prevent formation of thrombi in the vessels. Accordingly, infuse heparin until the lungs become white and are devoid of blood. If the lungs do not turn white, adjustment of the right ventricular needle may be necessary.</li>
	</ol>
	</li>
	<li>After inflating for 5 minutes, turn off the peristaltic pump and switch the perfusion tubing from the heparin solution to the fixative.</li>
	<li>Lower the water column syringe to the &#34;20 cm&#34; mark (Step 1.3). It is normal for air bubbles to move within the water column as the pressure changes from 25 to 20 cmH2O.
	<ol>
		<li>Check the water level in the syringe. It should be at the &#34;25 cm&#34; mark. It may be necessary to add more at this time.</li>
	</ol>
	</li>
	<li>Wait for 1 min to allow the lungs to deflate from 25 to 20 cmH2O.</li>
	<li>Restart the perfusion pump at a flow rate of 6.5 mL/min.</li>
	<li>Vascular perfusion-fix for 10 - 15 minutes.</li>
</ol>
9. Extraction of lungs (Figure 3)
<ol>
	<li>Tightly tie the second piece of thread around the trachea distal to the Luer-stub adapter. Remove the Luer-stub adapter from the trachea.</li>
	<li>Remove the needle from the heart.</li>
	<li>Free the lungs and heart from the thoracic cavity by cutting the connective tissue posterior to the mediastinum with blunt scissors. Take care to avoid puncturing the lungs.</li>
	<li>Carefully remove the heart from the lungs.</li>
	<li>Place the lungs in fixative overnight. 
	&#8203;NOTE: Duration of fixative varies depends on downstream histologic studies.
	<ol>
		<li>Place the lungs into a 50 mL conical tube containing 20-25 mL of fixative. Place the thread securing the trachea through the opening of the conical tube and secure by the threads of the cap. Invert the conical tube to ensure that the buoyant, air-inflated lungs remain fully submerged in fixative otherwise they will float to the top of the liquid.</li>
	</ol>
	</li>
	<li>Process the lungs for histologic studies.</li>
</ol>

<ol>
	<li>Hsia, C. C. W., Hyde, D. M., Ochs, M., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+Official+Research+Policy+Statement+of+the+American+Thoracic+Society/European+Respiratory+Society:+Standards+for+Quantitative+Assessment+of+Lung+Structure.">An Official Research Policy Statement of the American Thoracic Society/European Respiratory Society: Standards for Quantitative Assessment of Lung Structure.</a> American Journal of Respiratory and Critical Care Medicine. 181 (4), 394-418 (2010).</li><li>Weibel, E. R., Limacher, W., Bachofen, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscopy+of+rapidly+frozen+lungs:+evaluation+on+the+basis+of+standard+criteria.">Electron microscopy of rapidly frozen lungs: evaluation on the basis of standard criteria.</a> Journal of Applied Physiology: Respiratory, Environmental and Exercise Physiology. 53 (2), 516-527 (1982).</li><li>Bachofen, H., Ammann, A., Wangensteen, D., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+fixation+of+lungs+for+structure-function+analysis:+credits+and+limitations.">Perfusion fixation of lungs for structure-function analysis: credits and limitations.</a> Journal of Applied Physiology: Respiratory, Environmental and Exercise Physiology. 53 (2), 528-533 (1982).</li><li>Gil, J., Bachofen, H., Gehr, P., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alveolar+volume-surface+area+relation+in+air-+and+saline-filled+lungs+fixed+by+vascular+perfusion.">Alveolar volume-surface area relation in air- and saline-filled lungs fixed by vascular perfusion.</a> Journal of Applied Physiology: Respiratory, Environmental and Exercise Physiology. 47 (5), 990-1001 (1979).</li><li>Harris, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pressure-Volume+Curves+of+the+Respiratory+System.">Pressure-Volume Curves of the Respiratory System.</a> Respiratory Care. 50 (1), 78-99 (2005).</li><li>Bachofen, H., Sch&#252;rch, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alveolar+surface+forces+and+lung+architecture.">Alveolar surface forces and lung architecture.</a> Comparative Biochemistry and Physiology Part A: Molecular &amp; Integrative Physiology. 129 (1), 183-193 (2001).</li><li>Mead, J., Takishima, T., Leith, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stress+distribution+in+lungs:+a+model+of+pulmonary+elasticity.">Stress distribution in lungs: a model of pulmonary elasticity.</a> Journal of Applied Physiology. 28 (5), 596-608 (1970).</li><li>Mariano, C. A., Sattari, S., Maghsoudi-Ganjeh, M., Tartibi, M., Lo, D. D., Eskandari, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+Mechanical+Strain+Characterization+of+Ventilated+ex+vivo+Porcine+and+Murine+Lung+using+Digital+Image+Correlation.">Novel Mechanical Strain Characterization of Ventilated ex vivo Porcine and Murine Lung using Digital Image Correlation.</a> Frontiers in Physiology. 11, 600492 (2020).</li><li>Braber, S., Verheijden, K. a. T., Henricks, P. a. J., Kraneveld, A. D., Folkerts, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+fixation+methods+on+lung+morphology+in+a+murine+model+of+emphysema.">A comparison of fixation methods on lung morphology in a murine model of emphysema.</a> American Journal of Physiology - Lung Cellular and Molecular Physiology. 299 (6), 843-851 (2010).</li><li>Brain, J. D., Gehr, P., Kavet, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Airway+Macrophages.">Airway Macrophages.</a> American Review of Respiratory Disease. 129 (5), 823-826 (1984).</li><li>Wheeldon, E. B., Podolin, P. L., Mirabile, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alveolar+Macrophage+Distribution+in+a+Mouse+Model:+The+Importance+of+the+Fixation+Method.">Alveolar Macrophage Distribution in a Mouse Model: The Importance of the Fixation Method.</a> Toxicologic Pathology. 43 (8), 1162-1165 (2015).</li><li>Matulionis, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lung+deformation+and+macrophage+displacement+in+smoke-exposed+and+normal+mice+(Mus+musculus)+following+different+fixation+procedures.">Lung deformation and macrophage displacement in smoke-exposed and normal mice (Mus musculus) following different fixation procedures.</a> Virchows Archiv. A, Pathological Anatomy and Histopathology. 410 (1), 49-56 (1986).</li><li>Gage, G. J., Kipke, D. R., Shain, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+Animal+Perfusion+Fixation+for+Rodents.">Whole Animal Perfusion Fixation for Rodents.</a> JoVE (Journal of Visualized Experiments). (65), e3564 (2012).</li><li>Crosfill, M. L., Widdicombe, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physical+characteristics+of+the+chest+and+lungs+and+the+work+of+breathing+in+different+mammalian+species.">Physical characteristics of the chest and lungs and the work of breathing in different mammalian species.</a> The Journal of Physiology. 158 (1), 1-14 (1961).</li><li>Ramos-Vara, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+and+Methods+of+Immunohistochemistry.">Principles and Methods of Immunohistochemistry.</a> Drug Safety Evaluation: Methods and Protocols. , 115-128 (2017).</li></ol>

The authors have nothing to disclose.

Although commonly used, intratracheal-based fixation methods displace leukocytes from the airways and can alter normal lung architecture. The method of air-inflation with vascular perfusion-fixation that is provided in this protocol overcomes these pitfalls and more accurately preserves lung anatomy. The keys to obtaining high-quality tissue from the vascular perfusion-fixation method include careful monitoring of air-inflation pressures, avoidance of air leaks, and ensuring adequate perfusion of fixative into the vascul...

Lung histology represents the gold standard for assessing lung architecture during health and disease and is one of the most commonly used tools by pulmonary researchers1. One of the most critical aspects of this technique is the proper isolation and preservation of lung tissue, since variability in this step can lead to poor tissue quality and erroneous results1,2,3. In living animals, lung volume is determined by the balance between inward elastic recoil of the lung and outward forces transmitted from the chest wall and diaphragm by surface tension. Accordingly, when the thorax is entered, outward forces are lost and the lung collapses. Histologic sections prepared from collapsed lungs have a crowded appearance and boundaries between anatomic compartments (i.e., airspaces, vasculature, and interstitium) can be difficult to distinguish. To circumvent this challenge, researchers often inflate the lungs during chemical fixation so that airspace size and architecture is maintained.
Lungs can be inflated with air or liquid. The pressure necessary to inflate the lungs to the same volume differs between air- and liquid-inflation due to intermolecular forces at the air-liquid interface. Higher pressure (e.g., 25 cmH2O) is required during air-inflation than liquid inflation (e.g., 12 cmH2O) to overcome surface tension and open the collapsed alveoli4. Once alveoli have been recruited, a lower pressure can keep the alveoli open to the same volume as the pressure-volume curve plateaus, and pressures equalize throughout the lung according to Pascal&#39;s law4,5,6,7,8.
Two main methods of lung inflation and fixation exist to preserve murine lungs for histology. Most commonly, the airspaces are instilled with liquid - often containing a fixative. The main advantage of this approach is that it is relatively easy and requires little training. While intratracheal instillation of fixative may be preferred in studies that focus on the vasculature, liquid that is instilled via the trachea tends to push proximal airway cells and mucins into more distal airspace regions while air inflation does not1,3,4,9,10,11. Moreover, inadvertent detachment of leukocytes from the epithelium during liquid inflation alters their morphology, artifactually giving them a simple, rounded appearance4,10,11,12. Finally, inflation of the lungs with liquid can unintentionally compress the interstitium4,10,11. Together, these factors can distort the normal anatomy and cellular distributions within the preserved lungs, thus limiting the technique.
An alternative method of tissue preservation is vascular perfusion-fixation. In this method, fixative is perfused into the pulmonary vasculature via the vena cava or the right ventricle. This method preserves the location and morphology of cells in the airspace lumen. However, unless the lungs are inflated during perfusion-fixation, the lung tissue is likely to collapse.
Air-inflation with vascular perfusion-fixation harnesses strengths from each of the above fixation techniques. Herein we provide a protocol for this technique. The materials and equipment that are required are relatively inexpensive and can be easily obtained and assembled. The completed setup, shown in Figure 1A, provides constant airway pressure to the lungs by way of an adjustable, fluid-filled column while a peristaltic pump delivers fixative via the right ventricle. Lungs with preserved morphology can then be further processed for structure-function analyses.

<table><tbody><tr><td>00117XF-Stopcock 1 way 100/PK M Luer</td><td>Cole-Parmer</td><td>Mfr # VPB1000050N &amp;ndash; Item # EW-00117-XF</td><td>Stopcock</td></tr><tr><td>BD 60 mL syringe, slip tip</td><td>BD</td><td>309654</td><td>Syringe used to construct the water column</td></tr><tr><td>BD PrecisionGlide Needle 25G x 5/8</td><td>BD Biosciences</td><td>305122</td><td>Needle for vascular perfusion/fixation</td></tr><tr><td>Female Luer Thread Style Panel Mount 1/4-28 UNF to Male Luer</td><td>Nordson Medical</td><td>FTLLBMLRL-1</td><td>Female Luer</td></tr><tr><td>Heparin sodium salt from porcine intestinal mucosa</td><td>Sigma-Aldrich</td><td>H3393</td><td>Heparin solution.</td></tr><tr><td>Luer-Stub Adapter BD Intramedic 20 Gauge</td><td>BD Biosciences</td><td>427564</td><td>Luer-Stub Adapter</td></tr><tr><td>Male Luer (2) to Female Luer Thread Style Tee</td><td>Nordson Medical</td><td>LT787-9</td><td>Male Luer</td></tr><tr><td>Nalgene 180 Clear Plastic PVC Tubing</td><td>ThermoFisher Scientific</td><td>8000-9020</td><td>Tubing</td></tr><tr><td>Paraformaldehyde Aqueous Solution - 32%</td><td>Electron Microscopy Sciences</td><td>15714-S</td><td>Fixative solution. Diluted to 4% with phosphate buffered saline</td></tr><tr><td>Permatex Ultra Blue Multipurpose RTV Silicone Gasket Maker</td><td>Permatex</td><td>81724</td><td>Silicone Gasket Maker for air-tight sealing of chambers</td></tr><tr><td>Phosphate-Buffered Saline, 1x Without Calcium and Magnesium</td><td>Corning</td><td>21-040-CV</td><td>Bottle used to construct the air-inflation chamber, and buffer used for heparin and fixative solutions</td></tr><tr><td>Sterilite Ultra Seal 16.0 cup rectangle food storage container</td><td>Sterilite</td><td>0342</td><td>Animal processing container</td></tr></tbody></table>

air-inflation of murine lungs with vascular perfusion-fixation

Lung histology is often used to investigate the contributions provided by airspace cells during lung homeostasis and disease pathogenesis. However, commonly used instillation-based fixation methods can displace airspace cells and mucus into terminal airways and can alter tissue morphology. In comparison, vascular perfusion-fixation techniques are superior at preserving the location and morphology of cells within airspaces and the mucosal lining. However, if positive airway pressure is not simultaneously applied, regions of the lungs may collapse and capillaries may bulge into the alveolar spaces, leading to distortion of the lung anatomy. Herein, we describe an inexpensive method for air-inflation during vascular perfusion-fixation to preserve the morphology and location of airway and alveolar cells and interstitium in murine lungs for downstream histologic studies. Constant air pressure is delivered to the lungs via the trachea from a sealed, air-filled chamber that maintains pressure via an adjustable liquid column while fixative is perfused through the right ventricle.

In an intact thorax, the lungs are held open by outward forces applied by the chest wall via the pleural space6,14. When the diaphragm is entered during dissection, the integrity of the pleural space is abolished and the lungs should collapse (Figure 2A, 2B). To re-expand the lungs, air inflation is performed. As a first step, 25 cm of water pressure is applied to ensure recruitment of collapsed airspaces. Accordingl...

Watch this Scientific Journal Video about Air-Inflation of Murine Lungs with Vascular Perfusion-Fixation at JoVE.com

Air-Inflation of Murine Lungs with Vascular Perfusion-Fixation

Presented is a method for air-inflation with vascular perfusion-fixation of the lungs that preserves the location of cells within airways, alveoli and interstitium for structure-function analyses. Constant airway pressure is maintained with an air-inflation chamber while fixative is perfused via the right ventricle. Lungs are processed for histologic studies.

Lung histology is often used to investigate the contributions provided by airspace cells during lung homeostasis and disease pathogenesis. However, ...

air-inflation-of-murine-lungs-with-vascular-perfusion-fixation

Research

JoVE Journal

Neuroscience

6.8K Views.  National Jewish Health. Dit is een goedkope methode die de voordelen van luchtinflatie en vasculaire perfusiefixatie combineert voor het behoud van longen voor structurele en functionele analyses. Het belangrijkste voordeel van deze techniek is het behoud van zowel celmorfologie als locatie binnen het luchtruim van de long. Voorzichtigheid is het nodig om de lokstempadapter in de luchtpijp en de perfusiesnaald in de rechterkamer te plaatsen om voldoende inflatie en fixatie te garanderen.Om het luchtinflatieap...

Video: Air-Inflation of Murine Lungs with Vascular Perfusion-Fixation

Perfusion and Inflation of the Mouse Lung for Tumor Histology

The ability to evaluate lung histology is critical for the fields of lung cancer research and cancer metastasis. It is equally important to perform necropsies rapidly and efficiently from studies without sacrificing the quality of the tissues procured. The goal of this protocol is to present a method to rapidly perfuse, inflate, and fix mouse lungs for downstream histological analysis. This method does not standardize lung inflation; thus, it does not require any special procedures or equipment and instead simply instills fixative directly through the trachea following perfusion through the heart. This allows for sufficient estimation of tumor size, histology, and scoring. This also allows for the collection of frozen tissue prior to lung tissue fixation. This method is limited in that it does not allow for later morphometric quantification of the lung; however, it is more than sufficient for lung tumor analysis from genetically engineered mouse models (GEMMs), syngeneic models, as well as xenograft tumor and metastasis studies.

The purpose of this method is to present a simple and efficient method for the perfusion, inflation, and fixation of mouse lungs for the examination of lung tumor pathology and evaluation of metastases to the lung.

The ability to evaluate lung histology is critical for the fields of lung cancer research and cancer metastasis. It is equally important to perform ...

Cancer Research

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), representing the gas-exchange capacity of the lung, is a critical criterion to assess respiratory function. However, observer bias can significantly influence measured values for lung morphological parameters. The protocol that we describe here minimizes variations during measurements of two morphological parameters used for ISA calculation: internal lung volume (ILV) and mean linear intercept (MLI). Using ISA as a morphometric and functional parameter to determine the outcome of alveolar regeneration in both pneumonectomy (PNX) and prosthesis implantation mouse models, we found that the increased ISA following PNX treatment was significantly blocked by implantation of a prosthesis into the thoracic cavity1. The ability to accurately quantify ISA is not only expected to improve the reliability and reproducibility of lung function studies in injured-induced alveolar regeneration models, but also to promote mechanistic discoveries of multiple pulmonary diseases.

Internal lung surface area (ISA) is a critical criterion for assessing lung morphology and physiology in lung diseases and injury-induced alveolar regeneration. We describe here a standardized method that can minimize the measurement bias for ISA in both lung pneumonectomy and prosthesis implantation mouse models.

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), ...

Developmental Biology

Videomorphometric Analysis of Hypoxic Pulmonary Vasoconstriction of Intra-pulmonary Arteries Using Murine Precision Cut Lung Slices

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ventilation. Up to now, the underlying mechanisms are not fully understood. The major vascular segment contributing to HPV is the intra-acinar artery. This vessel section is responsible for the blood supply of an individual acinus, which is defined as the portion of lung distal to a terminal bronchiole. Intra-acinar arteries are mostly located in that part of the lung that cannot be selectively reached by a number of commonly used techniques such as measurement of the pulmonary artery pressure in isolated perfused lungs or force recordings from dissected proximal pulmonary artery segments1,2. The analysis of subpleural vessels by real-time confocal laser scanning luminescence microscopy is limited to vessels with up to 50 &#181;m in diameter3.
We provide a technique to study HPV of murine intra-pulmonary arteries in the range of 20-100 &#181;m inner diameters. It is based on the videomorphometric analysis of cross-sectioned arteries in precision cut lung slices (PCLS). This method allows the quantitative measurement of vasoreactivity of small intra-acinar arteries with inner diameter between 20-40 &#181;m which are located at gussets of alveolar septa next to alveolar ducts and of larger pre-acinar arteries with inner diameters between 40-100 &#181;m which run adjacent to bronchi and bronchioles. In contrast to real-time imaging of subpleural vessels in anesthetized and ventilated mice, videomorphometric analysis of PCLS occurs under conditions free of shear stress. In our experimental model both arterial segments exhibit a monophasic HPV when exposed to medium gassed with 1% O2 and the response fades after 30-40 min at hypoxia.

Hypoxic pulmonary vasoconstriction (HPV) is an important physiological phenomenon by which at alveolar hypoxia lung perfusion is matched to ventilation. The major vascular segment contributing to HPV is the intra-acinar artery. Here, we describe our protocol for the analysis of HPV of murine pulmonary vessels with diameters of 20-100 &mu;m.

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ...

Medicine

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

Instillation and Fixation Methods Useful in Mouse Lung Cancer Research

The ability to instill live agents, cells, or chemicals directly into the lung without injuring or killing the mice is an important tool in lung cancer research. Although there are a number of methods that have been published showing how to intubate mice for pulmonary function measurements, none are without potential problems for rapid tracheal instillation in large cohorts of mice. In the present paper, a simple and quick method is described that enables an investigator to carry out such instillations in an efficient manner. The method does not require any special tools or lighting and can be learned with very little practice. It involves anesthetizing a mouse, making a small incision in the neck to visualize the trachea, and then inserting an intravenous catheter directly. The small incision is quickly closed with tissue adhesive, and the mice are allowed to recover. A skilled student or technician can do instillations at an average rate of 2 min/mouse. Once the cancer is established, there is frequently a need for quantitative histologic analysis of the lungs. Traditionally pathologists usually do not bother to standardize lung inflation during fixation, and analyses are often based on a scoring system that can be quite subjective. While this may sometime be sufficiently adequate for gross estimates of the size of a lung tumor, any proper stereological quantification of lung structure or cells requires a reproducible fixation procedure and subsequent lung volume measurement. Here we describe simple reliable procedures for both fixing the lungs under pressure and then accurately measuring the fixed lung volume. The only requirement is a laboratory balance that is accurate over a range of 1 mg&#8211;300 g. The procedures presented here thus could greatly improve the ability to create, treat, and analyze lung cancers in mice.

The goal of this paper is to describe simple methods that will greatly aid in the setup and analysis of mouse lungs with lung cancer or other pathologies. We present 3 protocols to simply and reliably carry out lung instillations, fixation, and lung volume measurements.

The ability to instill live agents, cells, or chemicals directly into the lung without injuring or killing the mice is an important tool in lung cancer ...

Lung Fixation under Constant Pressure for Evaluation of Emphysema in Mice

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung fixation to produce reliable histological specimens of the lung. Due to the nature of the lung&rsquo;s structural composition, which consists largely of air and tissue, there is a risk that it collapses or deflates during the fixation process. Various lung fixation methods exist, each of which has its own advantages and disadvantages. The lung fixation method presented here utilizes constant pressure to enable optimal tissue evaluation for studies using an emphysematous mouse lung model. The main advantage is that it can fix many lungs with the same condition at one time. Lung specimens are obtained from chronic cigarette smoke-exposed mice. Lung fixation is performed using specialized equipment that enables the production of constant pressure. This constant pressure maintains the lung in a reasonably inflated state. Thus, this method generates a histological specimen of the lung that is suitable to evaluate cigarette smoke-induced mild emphysema.

Presented here is a useful protocol for lung fixation that creates a stable condition for histological evaluate of lung specimens from a mouse model of emphysema. The main advantage of this model is that it can fix many lungs with the same constant pressure without lung collapse or deflation.

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung ...

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or fungal) inflammatory conditions. An overwhelming host response may result in compromised respiration and acute lung injury, which is characterized by lung neutrophil recruitment as a result of the patho-logical host immune, coagulative and tissue remodeling response. Sensitive microscopic methods to visualize and quantify murine lung cellular adaptations, in response to low-dose (0.05 ppm) ozone, a potent environmental pollutant in combination with bacterial lipopolysaccharide, a TLR4 agonist, are crucial in order to understand the host inflammatory and repair mechanisms. We describe a comprehensive fluorescent microscopic analysis of various lung and systemic body compartments, namely the broncho-alveolar lavage fluid, lung vascular perfusate, left lung cryosections, and sternal bone marrow perfusate. We show damage of alveolar macrophages, neutrophils, lung parenchymal tissue, as well as bone marrow cells in correlation with a delayed (up to 36-72 h) immune response that is marked by discrete chemokine gradients in the analyzed compartments. In addition, we present lung extracellular matrix and cellular cytoskeletal interactions (actin, tubulin), mitochondrial and reactive oxygen species, anti-coagulative plasminogen, its anti-angiogenic peptide fragment angiostatin, the mitochondrial ATP synthase complex V subunits, &#945; and &#946;. These surrogate markers, when supplemented with adequate in vitro cell-based assays and in vivo animal imaging techniques such as intravital microscopy, can provide vital information towards understanding the lung response to novel immunomodulatory agents.

Combined ozone and bacterial endotoxin exposed mice show wide-spread cell death, including that of neutrophils. We observed cellular adaptations such as disruption of cytoskeletal lamellipodia, increased cellular expression of complex V ATP synthase subunit &#946; and angiostatin in broncho-alveolar lavage, suppression of the lung immune response and delayed neutrophil recruitment.

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or ...

Immunology and Infection

Quantifying Single Microvessel Permeability in Isolated Blood-perfused Rat Lung Preparation

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with real time imaging, it becomes feasible to determine permeability changes in individual pulmonary microvessels. Herein we describe steps to isolate rat lungs and perfuse them with autologous blood. Then, we outline steps to infuse fluorophores or agents via a microcatheter into a small lung region. Using these procedures described, we determined permeability increases in rat lung microvessels in response to infusions of bacterial lipopolysaccharide. The data revealed that lipopolysaccharide increased fluid leak across both venular and capillary microvessel segments. Thus, this method makes it possible to compare permeability responses among vascular segments and thus, define any heterogeneity in the response. While commonly used methods to define lung permeability require postprocessing of lung tissue samples, the use of real time imaging obviates this requirement as evident from the present method. Thus, the isolated lung preparation combined with real time imaging offers several advantages over traditional methods to determine lung microvascular permeability, yet is a straightforward method to develop and implement.

The isolated blood-perfused lung preparation makes it feasible to visualize microvessel networks on the lung surface. Here we describe an approach to quantify permeability of single microvessels in isolated lungs using real time fluorescence imaging.

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with ...

Biology

Preserving Fungal Spore Localization in Lungs for Fluorescent Microscopy Analysis

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.

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 ...

Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, respectively, hence playing a critical role in the homeostasis of the pulmonary system. Deregulation of SMC contractility contributes to several pulmonary diseases, including asthma and pulmonary hypertension. However, due to limited tissue access and a lack of culture systems to maintain in vivo SMC phenotypes, molecular mechanisms underlying the deregulated SMC contractility in these diseases remain fully identified. The precision-cut lung slice (PCLS) offers an ex vivo model that circumvents these technical difficulties. As a live, thin lung tissue section, the PCLS retains SMC in natural surroundings and allows in situ tracking of SMC contraction and intracellular Ca2+ signaling that regulates SMC contractility. Here, a detailed mouse PCLS preparation protocol is provided, which preserves intact airways and intrapulmonary arteries. This protocol involves two essential steps before subjecting the lung lobe to slicing: inflating the airway with low-melting-point agarose through the trachea and infilling pulmonary vessels with gelatin through the right ventricle. The PCLS prepared using this protocol can be used for bioassays to evaluate Ca2+-mediated contractile regulation of SMC in both the airway and the intrapulmonary arterial compartments. When applied to mouse models of respiratory diseases, this protocol enables the functional investigation of SMC, thereby providing insight into the underlying mechanism of SMC contractility deregulation in diseases.

The present protocol describes preparing and utilizing mouse precision-cut lung slices to assess the airway and intrapulmonary arterial smooth muscle contractility in a nearly in vivo milieu.

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, ...

Air-Inflation of Murine Lungs with Vascular Perfusion-Fixation

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