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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>00117XF-Stopcock 1 way 100/PK M Luer</td>
			<td>Cole-Parmer</td>
			<td>Mfr # VPB1000050N &#8211; 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

6.6K Views.  National Jewish Health. 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.

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

Whole Animal Perfusion Fixation for Rodents

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small pieces of tissue, larger specimens like the intact brain pose a problem for immersion fixation because the fixative does not reach all regions of the tissue at the same rate 5,7. Often, changes in response to hypoxia begin before the tissue can be preserved 12. The advantage of directly perfusing fixative through the circulatory system is that the chemical can quickly reach every corner of the organism using the natural vascular network. In order to utilize the circulatory system most effectively, care must be taken to match physiological pressures 3. It is important to note that physiological pressures are dependent on the species used. Techniques for perfusion fixation vary depending on the tissue to be fixed and how the tissue will be processed following fixation. In this video, we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain for immunohistochemistry. The main advantage of this technique (vs. gravity-fed systems) is that the circulatory system is utilized most effectively.

Here we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain.

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

The Corneal Micropocket Assay: A Model of Angiogenesis in the Mouse Eye

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets containing specific growth factors that trigger blood vessel growth throughout the naturally avascular cornea, angiogenesis can be measured and quantified. In this assay the angiogenic response is generated over the course of several days, depending on the type and dose of growth factor used. The induction of neovascularization is commonly triggered by either basic fibroblast growth factor (bFGF) or vascular endothelial growth factor (VEGF). By combining these growth factors with sucralfate and hydron (poly-HEMA (poly(2-hydroxyethyl methacrylate))) and casting the mixture into pellets, they can be surgically implanted in the mouse eye. These uniform pellets slowly-release the growth factors over five or six days (bFGF or VEGF respectively) enabling sufficient angiogenic response required for vessel area quantification using a slit lamp. This assay can be used for different applications, including the evaluation of angiogenic modulator drugs or treatments as well as comparison between different genetic backgrounds affecting angiogenesis. A skilled investigator after practicing this assay can implant a pellet in less than 5 min per eye.

The protocol describes the corneal micropocket assay as developed in mice.

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets ...

Non-invasive Parenchymal, Vascular and Metabolic High-frequency Ultrasound and Photoacoustic Rat Deep Brain Imaging

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central nervous system of small animals. However, transdermal and transcranial neuroimaging is frequently affected by low sensitivity, image aberrations and loss of space resolution, requiring scalp or even skull removal before imaging. To overcome this challenge, a new protocol is presented to gain significant insights in brain hemodynamics by photoacoustic and high-frequency ultrasounds imaging with the animal skin and skull intact. The procedure relies on the passage of ultrasound (US) waves and laser directly through the fissures that are naturally present on the animal cranium. By juxtaposing the imaging transducer device exactly in correspondence to these selected areas where the skull has a reduced thickness or is totally absent, one can acquire high quality deep images and explore internal brain regions that are usually difficult to anatomically or functionally describe without an invasive approach. By applying this experimental procedure, significant data can be collected in both sonic and optoacoustic modalities, enabling to image the parenchymal and the vascular anatomy far below the head surface. Deep brain features such as parenchymal convolutions and fissures separating the lobes were clearly visible. Moreover, the configuration of large and small blood vessels was imaged at several millimeters of depth, and precise information were collected about blood fluxes, vascular stream velocities and the hemoglobin chemical state. This repertoire of data could be crucial in several research contests, ranging from brain vascular disease studies to experimental techniques involving the systemic administration of exogenous chemicals or other objects endowed with imaging contrast enhancement properties. In conclusion, thanks to the presented protocol, the US and PA techniques become an attractive noninvasive performance-competitive means for cortical and internal brain imaging, retaining a significant potential in many neurologic fields.

The present work describes a new protocol to perform non-invasive high-frequency ultrasound and photoacoustic based imaging on rat brain, to efficiently visualize deep subcortical regions and their vascular patterns by directing signals on skull foramina naturally present on animal cranium.

Photoacoustics and high frequency ultrasound stands out as powerful tools for neurobiological applications enabling high-resolution imaging on the central ...

Isolation of Primary Murine Brain Microvascular Endothelial Cells

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional complex of tight and adherens junctions. Together with other cell-types such as astrocytes or pericytes, they form the neurovascular unit (NVU), which specifically regulates the interchange of fluids, molecules and cells between the peripheral blood and the CNS. Through this complex and dynamic system BMECs are involved in various processes maintaining the homeostasis of the CNS. A dysfunction of the BBB is observed as an essential step in the pathogenesis of many severe CNS diseases. However, specific and targeted therapies are very limited, as the underlying mechanisms are still far from being understood. 
Animal and in vitro models have been extensively used to gain in-depth understanding of complex physiological and pathophysiological processes. By reduction and simplification it is possible to focus the investigation on the subject of interest and to exclude a variety of confounding factors. However, comparability and transferability are also reduced in model systems, which have to be taken into account for evaluation. The most common animal models are based on mice, among other reasons, mainly due to the constantly increasing possibilities of methodology. In vitro studies of isolated murine BMECs might enable an in-depth analysis of their properties and of the blood-brain-barrier under physiological and pathophysiological conditions. Further insights into the complex mechanisms at the BBB potentially provide the basis for new therapeutic strategies.
This protocol describes a method to isolate primary murine microvascular endothelial cells by a sequence of physical and chemical purification steps. Special considerations for purity and cultivation of MBMECs as well as quality control, potential applications and limitations are discussed.

Brain microvascular endothelial cells (BMEC) are interconnected by specific junctional proteins forming a highly regulated barrier separating blood and the central nervous system (CNS), the so-called blood-brain-barrier (BBB). The isolation of primary murine brain microvascular endothelial cells, as discussed in this protocol, enables detailed in vitro studies of the BBB.

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional ...

Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic neuroinflammatory state within the central nervous system (CNS), thought to be driven principally by virally-mediated activation of microglia and brain resident macrophages. HIV-1 infection is also accompanied by changes in cerebrovascular blood flow (CBF), raising the possibility that HIV-associated chronic neuroinflammation may lead to changes in CBF and/or in cerebral vascular architecture. To address this question, we have used a mouse model for HIV-induced neuroinflammation, and we have tested whether long-term exposure to this inflammatory environment may damage brain vasculature and result in rarefaction of capillary networks. In this paper we describe a method to quantify changes in cortical capillary density in a mouse model of neuroinflammatory disease (HIV-1 Tat transgenic mice). This generalizable approach employs in vivo two-photon imaging of cortical capillaries through a thin-skull cortical window, as well as ex vivo two-photon imaging of cortical capillaries in mouse brain sections. These procedures produce images and z-stack files of capillary networks, respectively, which can be then subjected to quantitative analysis in order to assess changes in cerebral vascular architecture.

This paper describes a method by which the vascular architecture in the brain can be quantified using in vivo and ex vivo two-photon microscopy.

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic ...

Evaluation of Bioenergetic Function in Cerebral Vascular Endothelial Cells

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types that comprise the BBB; these cells have a very high-energy demand, which requires optimal mitochondrial function. In the case of disease or injury, the mitochondrial function in these cells can be altered, resulting in disease or&#160;the opening of the BBB. In this manuscript, we introduce a method to measure mitochondrial function in CVE cells by using whole, intact cells and a bioanalyzer. A mito-stress assay is used to challenge the cells that have been perturbed, either physically or chemically, and evaluate their bioenergetic function. Additionally, this method also provides a useful way to screen new therapeutics that have direct effects on mitochondrial function. We have optimized the cell density necessary to yield oxygen consumption rates that allow for the calculation of a variety of mitochondrial parameters, including ATP production, maximal respiration, and spare capacity. We also show the sensitivity of the assay by demonstrating that the introduction of the&#160;microRNA, miR-34a, leads to a pronounced and detectable decrease in mitochondrial activity. While the data shown in this paper is optimized for the bEnd.3 cell line, we have also optimized the protocol for primary CVE cells, further suggesting the utility in preclinical and clinical models.

Endothelial cell mitochondria are critical to maintain blood-brain-barrier integrity. We introduce a protocol to measure bioenergetic function in cerebral vascular endothelial cells.

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types ...

Inducement and Evaluation of a Murine Model of Experimental Myopia

Murine model of myopia can be a powerful tool for myopia research because of the comparatively easy genetic manipulation. One way to induce myopia in animals is to put clear minus lenses in front of eyes for weeks (lens-induced myopia, LIM). However, extant protocols for inducement and evaluation vary from laboratory to laboratory. Here, we described a highly practical and reproducible method to induce LIM in mice using newly designed eyeglasses. The method fixes the lens stably in front of the mouse eye while allows the lens to be taken off for cleaning or topical drug administration. The phenotype is robust and efficient, and the variance is small. The method described here can be applied to mice right after weaning which extends the possible duration for experiments. We also gave technical advises for achieving reproducible results in refraction and axial length measurements. We hope the step-by-step protocol described here and the detailed article can help researchers perform myopia experiments with myopia more smoothly and make the data comparable across laboratories.

In this protocol, we describe the full process of experimental myopia inducement in mice using newly designed eyeglasses and the technic needed for achieving stable and reproducible results in ocular parameter measurements.

Murine model of myopia can be a powerful tool for myopia research because of the comparatively easy genetic manipulation. One way to induce myopia in ...

Retinal Vascular Reactivity as Assessed by Optical Coherence Tomography Angiography

The vascular supply to the retina has been shown to dynamically adapt through vasoconstriction and vasodilation to accommodate the metabolic demands of the retina. This process, referred to as retinal vascular reactivity (RVR), is mediated by neurovascular coupling, which is impaired very early in retinal vascular diseases such as diabetic retinopathy. Therefore, a clinically feasible method of assessing vascular function may be of significant interest in both research and clinical settings. Recently, in vivo imaging of the retinal vasculature at the capillary level has been made possible by the FDA approval of optical coherence tomography angiography (OCTA), a noninvasive, minimal risk and dyeless angiography method with capillary level resolution. Concurrently, physiological and pathological changes in RVR have been shown by several investigators. The method shown in this manuscript is designed to investigate RVR using OCTA with no need for alterations to the clinical imaging procedures or device. It demonstrates real time imaging of the retina and retinal vasculature during exposure to hypercapnic or hyperoxic conditions. The exam is easily performed with two personnel in under 30 min with minimal subject discomfort or risk. This method is adaptable to other ophthalmic imaging devices and the applications may vary based on the composition of the gas mixture and patient population. A strength of this method is that it allows for an investigation of retinal vascular function at the capillary level in human subjects in vivo. Limitations of this method are largely those of OCTA and other retinal imaging methods including imaging artifacts and a restricted dynamic range. The results obtained from the method are OCT and OCTA images of the retina. These images are amenable to any analysis that is possible on commercially available OCT or OCTA devices. The general method, however, can be adapted to any form of ophthalmic imaging.

This article describes a method for measuring retinal vasculature reactivity in vivo with human subjects using a gas breathing provocation technique to deliver vasoactive stimuli while acquiring retinal images.

The vascular supply to the retina has been shown to dynamically adapt through vasoconstriction and vasodilation to accommodate the metabolic demands of ...

Preparation of Acute Human Hippocampal Slices for Electrophysiological Recordings

Epilepsy affects about 1% of the world population and leads to a severe decrease in quality of life due to ongoing seizures as well as high risk for sudden death. Despite an abundance of available treatment options, about 30% of patients are drug-resistant. Several novel therapeutics have been developed using animal models, though the rate of drug-resistant patients remains unaltered. One of probable reasons is the lack of translation between rodent models and humans, such as a weak representation of human pharmacoresistance in animal models. Resected human brain tissue as a preclinical evaluation tool has the advantage to bridge this translational gap. Described here is a method for high quality preparation of human hippocampal brain slices and subsequent stable induction of epileptiform activity. The protocol describes the induction of burst activity during application of 8 mM KCl and 4-aminopyridin. This activity is sensitive to established AED lacosamide or novel antiepileptic candidates, such as dimethylethanolamine (DMEA). In addition, the method describes induction of seizure-like events in CA1 of human hippocampal brain slices by reduction of extracellular Mg2+ and application of bicuculline, a GABAA receptor blocker. The experimental set-up can be used to screen potential antiepileptic substances for their effects on epileptiform activity. Furthermore, mechanisms of action postulated for specific compounds can be validated using this approach in human tissue (e.g., using patch-clamp recordings). To conclude, investigation of vital human brain tissue ex vivo (here, resected hippocampus from patients suffering from temporal lobe epilepsy) will improve the current knowledge of physiological and pathological mechanisms in the human brain.

The presented protocol describes the transport and preparation of resected human hippocampal tissue with the ultimate goal to use vital brain slices as a preclinical evaluation tool for potential antiepileptic substances.

Epilepsy affects about 1% of the world population and leads to a severe decrease in quality of life due to ongoing seizures as well as high risk for ...