The authors thank the VA-MD Regional College of Veterinary medicine for providing core and technical support for this project. This work is supported by an NIH Career Development Award (K01DK092355).

All studies were conducted under the approval of the Institutional Care and Use Committee (IACUC) for Virginia Tech and in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
1. Intratracheal (i.t.) Inoculation of LPS Using Oropharyngeal Administration
<ol>
	<li>Ensure that each animal is uniquely identified using either an ear punch, ear tag, or other institutionally approved method.</li>
	<li>Record the baseline body weight and body temperature for each animal.</li>
	<li>Prepare the working stock of LPS. Each mouse will receive a 50 &#181;l dose of 1 mg/kg LPS in 1x phosphate buffered saline (PBS). Mock treated animals will receive a 50 &#181;l dose of 1x PBS.</li>
	<li>Prepare an appropriate chamber for the administration of isoflurane using the following drop method. Institutional guidelines vary regarding the use of isoflurane and other anesthetic agents and prior to initiating any studies, persons should contact their institution&#39;s Department of Animal Care/Welfare to insure the all guidelines are adequately met. If drop method isoflurane is not an option, other anesthetic agents are acceptable alternatives.
	<ol>
		<li>In an appropriate safety cabinet, apply approximately 3 ml&#160;of isoflurane to a folded absorbent paper towel and place in the bottom of a 500 ml beaker.</li>
		<li>Place a small piece of aluminum foil on top of the paper towel and cover the beaker with a transparent lid.</li>
	</ol>
	</li>
	<li>Prepare the tools and reagents that will be utilized during the i.t. inoculation. Position the rubber band that will secure the animals head on the intubation stand. This procedure will require a pair of straight forceps, a pair of angled or curved forceps, and a p200 pipette with tips.</li>
	<li>Load 50 &#181;l of LPS into the pipette tip and place in a readily accessible location beside the intubation stand.</li>
	<li>Anesthetize the mouse by placing the animal into the isoflurane chamber and covering the chamber with the transparent lid. Observe the animal&#39;s breathing patterns, which will be rapid and shallow when first placed in the chamber. The animal will be sufficiently anesthetized when breathing rates approach 1 breath/2 sec, which is usually reached within 30 sec of placing in the chamber.&#160;The mouse should be anesthetized as stated in the approved animal protocol using drop method isoflurane.</li>
	<li>Remove the anesthetized mouse from the chamber and suspend the animal on the intubation stand by its front incisors. Insure that the animal is securely restrained and the mouth and tongue are accessible.</li>
	<li>Gently secure the tongue with the straight forceps. Grasp the tongue with the angled forceps and gently pull it out of the mouth until slight resistance is felt. This action is sufficient to block the epiglottis and gain access to the trachea.</li>
	<li>While continuing to hold the tongue in the extended position, administer the 50 &#181;l dose of LPS into the back of the throat and immediately cover the animal&#39;s nostrils with a gloved finger. The LPS will be visible in the back of the throat until the mouse inhales.</li>
	<li>Continue to cover the nostrils and hold the tongue extended for 5-10 additional breaths.</li>
	<li>Remove the mouse from the intubation stand and place the animal back into a new cage until it recovers from the anesthesia.</li>
	<li>Monitor surrogate markers associated with disease progression. Body weight and body temperature should be monitored every 4-8 hr for the duration of the study. Likewise, evaluate behavioral characteristics and clinical symptoms associated with increased morbidity.</li>
	<li>To evaluate disease progression, harvest mice at specific time-points following the LPS exposure. Typical time-points for harvest following LPS exposure include 0, 6, 12, 18, 24, and 48 hr. To evaluate recovery and survival, evaluate the mice for 7 days post-inoculation.</li>
</ol>
2. Serum and Bronchoalveolar Lavage Fluid (BALF) Collection
<ol>
	<li>Sacrifice the mouse using an institutionally approved method.</li>
	<li>Place the animal on its back and secure it to a necropsy board, ideally 0.5 in (1.27 cm) in height.</li>
	<li>Wet the entire mouse with 70% ethanol.</li>
	<li>Using a 1 ml syringe with a 27 G needle, conduct a cardiac puncture prior to making any incisions. It should be possible to withdraw 500-800 &#181;l of whole blood from a single mouse.</li>
	<li>Remove the needle from the syringe and transfer the whole blood to a prelabeled 1.5 ml microcentrifuge tube. For serum collection, allow the whole blood to coagulate for at least 30 min at room temperature. In addition to serum collection, this procedure can also be modified by incorporating an anti-coagulant in the tube and syringe, prior to collecting the whole blood, to study circulating immune cells.</li>
	<li>Make a horizontal incision across the length of the peritoneal cavity and a vertical incision from the peritoneal cavity to the lower jaw of the mouse.</li>
	<li>Using forceps, grasp both sides of the incision and gently pull the skin away from the underlying peritoneal and thoracic cavities.</li>
	<li>Make a large incision along the length of the peritoneal cavity from the genitals to the sternum, taking care to avoid cutting the diaphragm. Gently shift the intestines in the peritoneal cavity to allow access to one of the kidneys.</li>
	<li>Cut the renal vein leading to the kidney to serve as a drainage point for perfusion.</li>
	<li>Nick the diaphragm using scissors, taking care to avoid the lungs and heart, to expose the left side of the heart.</li>
	<li>Manually perfuse the heart using a 10 ml syringe with a 27 G needle and 1x PBS solution. Avoid applying excessive force during the perfusion to ensure that the saline is not forced into the lungs. The remaining blood should drain from the portal vein incision.</li>
	<li>Carefully cut the ribcage along the sternum using blunt/blunt scissors, taking care to avoid cutting the heart and lungs. Accidentally cutting the lungs during this procedure will significantly reduce the amount of BALF collected and will result in an inability to properly inflate the lungs with fixative.</li>
	<li>Gently isolate the heart and lungs away from the ribcage using forceps. Carefully cut each section of the ribcage using blunt/blunt scissors as close to the spine as possible and fully remove both sections.</li>
	<li>Separate the salivary glands using forceps or remove them using scissors, to expose the trachea.</li>
	<li>Carefully cut the muscles that overlay the trachea using scissors. It is essential that the trachea not be cut during this process.</li>
	<li>Using scissors, separate the collarbone overlying the trachea.</li>
	<li>Gently grasp the thymus using forceps and lift the tissue away from the heart. Remove the thymus using scissors, while taking care to avoid cutting the heart or lungs.</li>
	<li>Make a small horizontal incision in the trachea 1-2 rings below the larynx using angled scissors (45-90&#186; angle, sharp/sharp). The incision should be large enough to firmly secure the cannula.</li>
	<li>Insert the cannula so that the tapered end extends 2-3 tracheal rings below the incision and secure the cannula in place using a silk suture. Ensure that the suture passes between the trachea and the esophagus and is pulled tight on the cannula.</li>
	<li>Fill a 1 ml syringe with 1 ml of Hanks Buffered Saline Solution (HBSS) and gently insert its luer into the cannula.</li>
	<li>Using a slow, but constant motion, inject ~900 &#181;l into the trachea, inflating the lungs, and immediately withdraw the HBSS. Place the recovered BALF in a 15 ml conical tube.</li>
	<li>Repeat this lavage 2 additional times to collect approximately 3 ml&#160;of BALF for future analysis. Store the BALF on ice or at 4 &#186;C until ready for use.</li>
</ol>
3. Histopathology Preparation
<ol>
	<li>Insert a 10 ml syringe into the lung inflation stand. Assemble the tubing to the luer, the luer to the stopcock, and the stopcock to the syringe. Ensure the stopcock is in the closed position.&#160;The 10 ml syringe will serve as a gravity flow reservoir. The reservoir should be attached to the inflation stand 4.75 in&#160;above the animal and the top of the reservoir should extend 10.5 in&#160;above the animal.</li>
	<li>Fill the syringe with neutral buffered formalin solution. Ensure that the syringe is completely filled with the fixative.</li>
	<li>Place an absorbent towel under the open end of the tubing and open the stopcock to allow the fixative to fill the tubing. Once the fixative begins flowing from the tubing, immediately close the stopcock and refill the syringe to the top with fixative. It is important that the syringe be completely filled to provide the appropriate amount of pressure for lung inflation.</li>
	<li>Gently pass a second piece of suture thread under the trachea, approximately 1-2 tracheal rings below the end of the cannula.</li>
	<li>Tie a single loose knot in the suture; however, do not pull the knot tight.</li>
	<li>Insert the tubing from the mouse inflation stand into the cannula.</li>
	<li>Open the stopcock and allow the lungs to fill by gravity inflation.</li>
	<li>Once the lungs reach their maximum inflation level, pull the suture tight and tie a second knot.</li>
	<li>Close the stopcock and remove the tubing from the cannula.</li>
	<li>Gently remove the cannula from the trachea by grasping the suture with a pair of forceps and holding the cannula with fingers. Firmly pull the suture down towards the thoracic cavity and the cannula back towards the mouse&#39;s nose.</li>
	<li>Gently grasp the trachea or the sutures with forceps and lift the trachea away from the neck cavity.</li>
	<li>Sever the trachea caudal to the tied suture using blunt scissors.</li>
	<li>Once the trachea is free from the underlying tissues, begin to pull the trachea away from the mouse. Gently lift the lungs out of the thoracic cavity.</li>
	<li>Carefully cut the connective tissue holding the lungs in the thoracic cavity. Continue pulling the trachea away from the mouse&#39;s body and apply steady upward force while cutting the underlying connections. Take care to avoid cutting the lungs. Note that the heart should be left attached to the lungs using this method.</li>
	<li>Once the lungs have been removed, place them in 10 ml&#160;of buffered formalin.</li>
	<li>Remove a section of mouse tail for genotyping.</li>
	<li>Dispose of the mouse carcass following the appropriate institutional guidelines.</li>
</ol>
4. Cytokine Evaluation
<ol>
	<li>Centrifuge the whole blood, collected under step 2.5, at 12,000 x g for 5 min&#160;to isolate the serum. Transfer the serum to a prelabeled 1.5 ml microcentrifuge tube and store at -80 &#176;C.</li>
	<li>Evaluate the serum cytokine levels by ELISA or another similar assay. Dilute the serum 1:5 -&#160;1:20 in an appropriate dilution buffer, depending on the assay, following the manufactures instructions. These dilutions should be empirically determined prior to running the bulk of the samples.</li>
	<li>Due to the low volume of serum collected, reduce the sample volume loaded onto the ELISA plate by half. For example, most commercial ELISAs utilize 100 &#956;l volumes of standards and samples. To conserve specimens, load 50 &#956;l of standards and diluted serum per well.</li>
	<li>Centrifuge the BALF in a refrigerated tabletop centrifuge at 200 x g for 5 min. Transfer the cell free supernatant into two prelabeled 1.5 ml microcentrifuge tubes and store at -80 &#176;C.</li>
	<li>Evaluate BALF cytokines by ELISA or another similar assay without diluting.</li>
	<li>Store unused serum and BALF at -80 &#176;C.</li>
</ol>
5. Differential Staining and BAL Cellularity Evaluation
<ol>
	<li>Following BALF centrifugation and complete HBSS removal, lyse the pelleted red blood cells using hypotonic saline. Resuspend the cells in 900 &#956;l of distilled water. Immediately add 100 &#956;l of 10x PBS.</li>
	<li>Individually lyse each sample.&#160;If samples contain excessive amounts of red blood cells, the cells can be pelleted in the table top centrifuge at 400 x g for 5 min and the red blood cell lysis protocol can be repeated.</li>
	<li>Determine the total BALF cellularity in the 1 ml suspension using a hemacytometer under 10X or 20 X magnification with Trypan Blue staining. Evaluate and present these data as cells/ml.</li>
	<li>Prepare materials for the cytospin and differential staining. Label standard microscope slides using a pencil or solvent resistant pen. Secure the slides into a cytospin bracket and funnel. Secure the slide assembly in the cytospin rotor.</li>
	<li>Cytospin 150 &#956;l of BALF at 100 x g for 5 min. If the cell density is too great to effectively evaluate cell morphology, reduce the volume spun down onto the slides. Allow the slides to air dry overnight.</li>
	<li>Differential stain the slides following the manufactures protocols. Allow the slides to air dry overnight. Coverslip the slides using Permount and evaluate the slides using a microscope equipped with a 20X and 40X objective.</li>
	<li>Harvest the remaining cells for subsequent analysis, such as FACS, electron microscopy, confocal microscopy, RNA extraction for gene expression evaluation, and/or protein extraction for western blot.</li>
</ol>
6. Histopathology Evaluation
<ol>
	<li>Prepare the lungs for histopathology evaluation. After 24-48 hr of formalin fixation, ventrally orient the whole inflated lungs and embedded in paraffin. Cut the resultant blocks to expose the main conducting airway.</li>
	<li>To improve scoring accuracy, position the lungs in the same position and trim each block to yield the maximum longitudinal visualization of the intrapulmonary main axial airway. From this point, cut 5 micron serial sections and stain with hematoxylin and eosin (H&#38;E). Additional sections can be cut and prepared for in situ hybridization using standard protocols.</li>
	<li>Evaluate histopathology using a semi-quantitative scoring system based on the following inflammatory parameters, which are scored between 0 (absent) and 3 (severe): mononuclear and polymorphonuclear cell infiltration; airway epithelial cell hyperplasia and injury; extravasation; perivascular and peribroncheolar cuffing; and the percent of the lung involved with inflammation. Lung histopathology should be evaluated by an experienced pathologist.</li>
	<li>Average all of the parameter scores to generate a total histopathology score or use individual scores to quantify specific aspects of disease progression. Conduct all scoring in a double blind fashion, with reviewers blinded to both genotype and treatment. This scoring system has been previously described6,7,10.</li>
</ol>

<ol>
	<li>Matute-Bello, G., et al. <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+American+Thoracic+Society+workshop+report:+features+and+measurements+of+experimental+acute+lung+injury+in+animals.">An official American Thoracic Society workshop report: features and measurements of experimental acute lung injury in animals.</a> Am. J. respir. Cell Mol. Biol. 44, 725-738 (2011).</li><li>Egger, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Administration+of+bleomycin+via+the+oropharyngeal+aspiration+route+leads+to+sustained+lung+fibrosis+in+mice+and+rats+as+quantified+by+UTE-MRI+and+histology.">Administration of bleomycin via the oropharyngeal aspiration route leads to sustained lung fibrosis in mice and rats as quantified by UTE-MRI and histology.</a> PloS one. 8, (2013).</li><li>Rayamajhi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonsurgical+intratracheal+instillation+of+mice+with+analysis+of+lungs+and+lung+draining+lymph+nodes+by+flow+cytometry.">Nonsurgical intratracheal instillation of mice with analysis of lungs and lung draining lymph nodes by flow cytometry.</a> J. Vis. Exp. , (2011).</li><li>Revelli, D. A., Boylan, J. A., Gherardini, F. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+non-invasive+intratracheal+inoculation+method+for+the+study+of+pulmonary+melioidosis.">A non-invasive intratracheal inoculation method for the study of pulmonary melioidosis.</a> Front. Cell. Infect. Microbiol. 2, 164 (2012).</li><li>Allen, I. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+NLRP3+in+the+development+of+allergic+airway+disease+in+mice.">Analysis of NLRP3 in the development of allergic airway disease in mice.</a> J. Immunol. 188, 2884-2893 (2012).</li><li>Allen, I. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+NLRP12+during+the+in+vivo+host+immune+response+to+Klebsiella+pneumoniae+and+Mycobacterium+tuberculosis.">Characterization of NLRP12 during the in vivo host immune response to Klebsiella pneumoniae and Mycobacterium tuberculosis.</a> PloS one. , (2013).</li><li>Allen, I. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+and+function+of+NPSR1/GPRA+in+the+lung+before+and+after+induction+of+asthma-like+disease.">Expression and function of NPSR1/GPRA in the lung before and after induction of asthma-like disease.</a> Am. J. Physiol. Lung Cell. Mol. Physiol. 291, 1005-1017 (2006).</li><li>Kebaier, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Staphylococcus+aureus+alpha-hemolysin+mediates+virulence+in+a+murine+model+of+severe+pneumonia+through+activation+of+the+NLRP3+inflammasome.">Staphylococcus aureus alpha-hemolysin mediates virulence in a murine model of severe pneumonia through activation of the NLRP3 inflammasome.</a> J. Infect. Dis. 205, 807-817 (2012).</li><li>Roberts, R. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+murine+immune+response+to+pulmonary+delivery+of+precisely+fabricated+nano-+and+microscale+particles.">Analysis of the murine immune response to pulmonary delivery of precisely fabricated nano- and microscale particles.</a> PloS one. 8, (2013).</li><li>Willingham, S. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NLRP3+(NALP3,+Cryopyrin)+facilitates+in+vivo+caspase-1+activation,+necrosis,+and+HMGB1+release+via+inflammasome-dependent+and+-independent+pathways.">NLRP3 (NALP3, Cryopyrin) facilitates in vivo caspase-1 activation, necrosis, and HMGB1 release via inflammasome-dependent and -independent pathways.</a> J. Immunol. 183, 2008-2015 (2009).</li><li>Kline, J. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variable+airway+responsiveness+to+inhaled+lipopolysaccharide.">Variable airway responsiveness to inhaled lipopolysaccharide.</a> Am. J. Respir. Crit. Med. 160, 297-303 (1999).</li><li>Cressman, V. L., Hicks, E. M., Funkhouser, W. K., Backlund, D. C., Koller, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+relationship+of+chronic+mucin+secretion+to+airway+disease+in+normal+and+CFTR-deficient+mice.">The relationship of chronic mucin secretion to airway disease in normal and CFTR-deficient mice.</a> Am. J. Respir. Cell Mol. Biol. 19, 853-866 (1998).</li></ol>

The authors declare&#160;no competing financial interests.

The most critical steps for successfully evaluating the host immune response in mouse lungs is as follows: 1) choose the appropriate mouse strain and sex for the model being evaluated; 2) optimize PAMP delivery to the lungs; 3) correctly collect and process the BALF; and 4) properly fix and prepare of the lungs for histopathological assessments.
The choice of mouse strain is an important factor in evaluating the host immune response. C57Bl/6 mice are typically considered the optimal mouse back...

Pulmonary infections associated with pathogenic bacteria species are a common cause of global morbidity and mortality. Determining the mechanisms that drive the host immune response to these pathogens will promote the development of novel prevention strategies and therapeutic agents that will attenuate the impact of these infections. The overall goal of the protocol described here is to provide the user with a flexible method to evaluate the host innate immune response to pathogen infection using a pathogen associated molecular pattern (PAMP) as a surrogate for live bacteria. The majority of previous studies evaluating the host innate immune response to bacteria have focused on peritoneal models due to the relative ease of execution. While these models are highly useful and have resulted in significant advances in the field of host-pathogen interactions and systemic inflammation, the data generated from these models are not always appropriate for studies involving the respiratory system. Here, a pulmonary model of acute lung inflammation is proposed as a practical and clinically relevant expansion of the classical intraperitoneal (i.p.) injection models. The proposed technique allows for the local assessment of the innate immune response in an organ specific model system.
The methods described here are designed to provide a simple and robust technique to allow users to evaluate the host immune response to LPS, which is a common PAMP. The methods are based on intratracheal (i.t.) instillation of LPS, which induces a robust innate immune response in the lungs of mice and mimics many of the pathophysiological features observed in human patients suffering from respiratory infections and acute lung injury1. A primary advantage of this technique is that it allows the user to evaluate the host immune response without the confounding factors and safety concerns associated with conducting in vivo studies using live bacteria. Likewise, the oropharyngeal i.t. administration route of exposure described in this protocol has significant advantages over other commonly utilized techniques, including intranasal (i.n.) administration and surgical i.t. administration. For example, oropharyngeal i.t. administration allows relatively accurate dosaging and lung deposition compared to i.n. administration, which typically suffers from increased variability of lung deposition due to the loss of agents in the nasal cavity and sinuses2-4. The i.t. administration route circumvents these cavities and allows direct access to the trachea and airway. Likewise, the surgical i.t. approach is a significantly more morbid administration method and requires extensive training to master. The protocols described here also include a description the common techniques and surrogate markers used to evaluate inflammation progression and end with a protocol describing the proper techniques for preparing the lungs for histopathology assessments. These protocols are focused on minimizing the number of mice required for each study by maximizing the data generated from each individual animal.
The protocols described are highly flexible and can be readily modified to evaluate a diverse range PAMPs and damage associated molecular patterns (DAMPs). Furthermore, with a few additional modifications, these protocols can also be applied to studies evaluating allergic airway disease progression or host-pathogen interactions with live bacteria, viruses or fungi5-10.

<table border="0" cellpadding="0" cellspacing="0" width="596">
	<colgroup>
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="27">
			<td height="27" style="height:27px;width:269px;">C57Bl/6J</td>
			<td style="width:181px;">The Jackson Laboratory</td>
			<td style="width:147px;">Stock 000664</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:269px;">Compact Scale</td>
			<td style="width:181px;">Ohaus Scale Corporation</td>
			<td style="width:147px;">71142845</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Small Animal Rectal Thermometer</td>
			<td style="width:181px;">Braintree Scientific</td>
			<td style="width:147px;">TH 5</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Rectal Probe for Rodents</td>
			<td style="width:181px;">Braintree Scientific</td>
			<td style="width:147px;">RET 3</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Ear Punch</td>
			<td style="width:181px;">Braintree Scientific</td>
			<td style="width:147px;">EP-S 901</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Lipopolysaccharide from E. coli 0111:B4</td>
			<td style="width:181px;">InvivoGen</td>
			<td style="width:147px;">LPS-EB</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">1x Phosphate Buffered Saline</td>
			<td style="width:181px;">Life Technologies</td>
			<td style="width:147px;">10010-023</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Isoflurane</td>
			<td style="width:181px;">Baxter</td>
			<td style="width:147px;">40032609</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:269px;">Intratrachael Administration and Lung Inflation Stand</td>
			<td style="width:181px;">ICAP Manufacturing</td>
			<td style="width:147px;">n/a</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Rodent Intubation Stand</td>
			<td style="width:181px;">Braintree Scientific</td>
			<td style="width:147px;">RIS 100</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scissors (blunt/sharp)</td>
			<td style="width:181px;">Fisher Scientific</td>
			<td style="width:147px;">13-806-2</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">forceps (straight)</td>
			<td style="width:181px;">Fisher Scientific</td>
			<td>22-327-379</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">forceps (45&#186;, curved)</td>
			<td style="width:181px;">Fisher Scientific</td>
			<td>10-275</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scissors (blunt/blunt)</td>
			<td style="width:181px;">Fisher Scientific</td>
			<td>08-940</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Pipette (200 &#181;l Capacity)</td>
			<td style="width:181px;">Gilson</td>
			<td style="width:147px;">F123601</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Ethanol</td>
			<td style="width:181px;">Sigma</td>
			<td style="width:147px;">459844</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">&#160;1 ml Syringe</td>
			<td style="width:181px;">BD Medical</td>
			<td style="width:147px;">301025</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">10 ml Syringe</td>
			<td style="width:181px;">BD Medical</td>
			<td style="width:147px;">301604</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">27 G x 0.5 in. needle</td>
			<td style="width:181px;">BD Medical</td>
			<td style="width:147px;">305109</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:269px;">Refrigerated Microcentrifuge</td>
			<td style="width:181px;">Fisher Scientific</td>
			<td style="width:147px;">13-100-676</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:269px;">1.2 mm Tracheal Cannulae with Luer-adapter</td>
			<td style="width:181px;">Harvard Apparatus</td>
			<td style="width:147px;">732836</td>
		</tr>
		<tr height="20">
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			<td height="20" style="height:20px;width:269px;">Permount Mounting Medium</td>
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			<td style="width:147px;">SP15-500</td>
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</table>

the utilization of oropharyngeal intratracheal pamp administration and bronchoalveolar lavage to evaluate the host immune response in mice

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen interactions take advantage of intraperitoneal injections of select bacteria or pathogen associated molecular patterns (PAMPs) in mice. While these techniques have yielded tremendous data associated with infectious disease pathobiology, intraperitoneal injection models are not always appropriate for host-pathogen interaction studies in the lung. Utilizing an acute lung inflammation model in mice, it is possible to conduct a high resolution analysis of the host innate immune response utilizing lipopolysaccharide (LPS). Here, we describe the methods to administer LPS using nonsurgical oropharyngeal intratracheal administration, monitor clinical parameters associated with disease pathogenesis, and utilize bronchoalveolar lavage fluid to evaluate the host immune response. The techniques that are described are widely applicable for studying the host innate immune response to a diverse range of PAMPs and pathogens. Likewise, with minor modifications, these techniques can also be applied in studies evaluating allergic airway inflammation and in pharmacological applications.

The cell walls of gram-negative bacteria are composed of LPS, which is highly abundant in the environment. Inhalation of LPS in sensitive human populations exacerbates airway reactivity and is capable of triggering a robust immune response11. LPS is also a common PAMP used in mouse models to elicit a robust innate immune response. In the protocol described here, the mice received an i.t. dose of LPS isolated from E. coli (serotype 0111:B4) using oropharyngeal i.t. administration. In models of LPS exposure, both ...

Watch this Scientific Journal Video about The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice at JoVE.com

The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

The host immune response to pathogen infection is a tightly regulated process. Utilizing a lipopolysaccharide lung exposure model in mice, it is possible to conduct high resolution evaluations of the complex mechanisms associated with disease pathogenesis.

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen ...

the-utilization-oropharyngeal-intratracheal-pamp-administration

Research

JoVE Journal

Immunology and Infection

16.5K Views.  Virginia Polytechnic Institute and State University. The host immune response to pathogen infection is a tightly regulated process. Utilizing a lipopolysaccharide lung exposure model in mice, it is possible to conduct high resolution evaluations of the complex mechanisms associated with disease pathogenesis.

Video: The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

Measuring Bacterial Load and Immune Responses in Mice Infected with Listeria monocytogenes

Listeria monocytogenes (Listeria) is a Gram-positive facultative intracellular pathogen1. Mouse studies typically employ intravenous injection of Listeria, which results in systemic infection2. After injection, Listeria quickly disseminates to the spleen and liver due to uptake by CD8&alpha;+ dendritic cells and Kupffer cells3,4. Once phagocytosed, various bacterial proteins enable Listeria to escape the phagosome, survive within the cytosol, and infect neighboring cells5. During the first three days of infection, different innate immune cells (e.g. monocytes, neutrophils, NK cells, dendritic cells) mediate bactericidal mechanisms that minimize Listeria proliferation. CD8+ T cells are subsequently recruited and responsible for the eventual clearance of Listeria from the host, typically within 10 days of infection6.
Successful clearance of Listeria from infected mice depends on the appropriate onset of host immune responses6	. There is a broad range of sensitivities amongst inbred mouse strains7,8. Generally, mice with increased susceptibility to Listeria infection are less able to control bacterial proliferation, demonstrating increased bacterial load and/or delayed clearance compared to resistant mice. Genetic studies, including linkage analyses and knockout mouse strains, have identified various genes for which sequence variation affects host responses to Listeria infection6,8-14. Determination and comparison of infection kinetics between different mouse strains is therefore an important method for identifying host genetic factors that contribute to immune responses against Listeria. Comparison of host responses to different Listeria strains is also an effective way to identify bacterial virulence factors that may serve as potential targets for antibiotic therapy or vaccine design.
We describe here a straightforward method for measuring bacterial load (colony forming units [CFU] per tissue) and preparing single-cell suspensions of the liver and spleen for FACS analysis of immune responses in Listeria-infected mice. This method is particularly useful for initial characterization of Listeria infection in novel mouse strains, as well as comparison of immune responses between different mouse strains infected with Listeria. We use the Listeria monocytogenes EGD strain15 that, when cultured on blood agar, exhibits a characteristic halo zone around each colony due to &beta;-hemolysis1 (Figure 1). Bacterial load and immune responses can be determined at any time-point after infection by culturing tissue homogenate on blood agar plates and preparing tissue cell suspensions for FACS analysis using the protocols described below. We would note that individuals who are immunocompromised or pregnant should not handle Listeria, and the relevant institutional biosafety committee and animal facility management should be consulted before work commences.

Measuring Bacterial Load and Immune Responses in Mice Infected with Listeria monocytogenes

Listeria monocytogenes is a model organism for studying immune responses and genetic susceptibility to intracellular bacteria in mice. This method enables one to measure bacterial load and generate single-cell suspensions of the liver and spleen from mice for FACS analysis to determine changes in immune cells due to Listeria infection.

Listeria monocytogenes (Listeria) is a Gram-positive facultative intracellular pathogen1. Mouse studies typically employ intravenous injection of ...

An Introduction to Parasitic Wasps of Drosophila and the Antiparasite Immune Response

Most known parasitoid wasp species attack the larval or pupal stages of Drosophila. While Trichopria drosophilae infect the pupal stages of the host (Fig. 1A-C), females of the genus Leptopilina (Fig. 1D, 1F, 1G) and Ganaspis (Fig. 1E) attack the larval stages. We use these parasites to study the molecular basis of a biological arms race. Parasitic wasps have tremendous value as biocontrol agents. Most of them carry virulence and other factors that modify host physiology and immunity. Analysis of Drosophila wasps is providing insights into how species-specific interactions shape the genetic structures of natural communities. These studies also serve as a model for understanding the hosts' immune physiology and how coordinated immune reactions are thwarted by this class of parasites.
The larval/pupal cuticle serves as the first line of defense. The wasp ovipositor is a sharp needle-like structure that efficiently delivers eggs into the host hemocoel. Oviposition is followed by a wound healing reaction at the cuticle (Fig. 1C, arrowheads). Some wasps can insert two or more eggs into the same host, although the development of only one egg succeeds. Supernumerary eggs or developing larvae are eliminated by a process that is not yet understood. These wasps are therefore referred to as solitary parasitoids.
Depending on the fly strain and the wasp species, the wasp egg has one of two fates. It is either encapsulated, so that its development is blocked (host emerges; Fig. 2 left); or the wasp egg hatches, develops, molts, and grows into an adult (wasp emerges; Fig. 2 right). L. heterotoma is one of the best-studied species of Drosophila parasitic wasps. It is a "generalist," which means that it can utilize most Drosophila species as hosts1. L. heterotoma and L. victoriae are sister species and they produce virus-like particles that actively interfere with the encapsulation response2. Unlike L. heterotoma, L. boulardi is a specialist parasite and the range of Drosophila species it utilizes is relatively limited1. Strains of L. boulardi also produce virus-like particles3 although they differ significantly in their ability to succeed on D. melanogaster1. Some of these L. boulardi strains are difficult to grow on D. melanogaster1 as the fly host frequently succeeds in encapsulating their eggs. Thus, it is important to have the knowledge of both partners in specific experimental protocols.
In addition to barrier tissues (cuticle, gut and trachea), Drosophila larvae have systemic cellular and humoral immune responses that arise from functions of blood cells and the fat body, respectively. Oviposition by L. boulardi activates both immune arms1,4. Blood cells are found in circulation, in sessile populations under the segmented cuticle, and in the lymph gland. The lymph gland is a small hematopoietic organ on the dorsal side of the larva. Clusters of hematopoietic cells, called lobes, are arranged segmentally in pairs along the dorsal vessel that runs along the anterior-posterior axis of the animal (Fig. 3A). The fat body is a large multifunctional organ (Fig. 3B). It secretes antimicrobial peptides in response to microbial and metazoan infections.
Wasp infection activates immune signaling (Fig. 4)4. At the cellular level, it triggers division and differentiation of blood cells. In self defense, aggregates and capsules develop in the hemocoel of infected animals (Fig. 5)5,6. Activated blood cells migrate toward the wasp egg (or wasp larva) and begin to form a capsule around it (Fig. 5A-F). Some blood cells aggregate to form nodules (Fig. 5G-H). Careful analysis reveals that wasp infection induces the anterior-most lymph gland lobes to disperse at their peripheries (Fig. 6C, D).
We present representative data with Toll signal transduction pathway components Dorsal and Sp&auml;tzle (Figs. 4,5,7), and its target Drosomycin (Fig. 6), to illustrate how specific changes in the lymph gland and hemocoel can be studied after wasp infection. The dissection protocols described here also yield the wasp eggs (or developing stages of wasps) from the host hemolymph (Fig. 8).

An Introduction to Parasitic Wasps of Drosophila and the Antiparasite Immune Response

Parasitoid (parasitic) wasps constitute a major class of natural enemies of many insects including Drosophila melanogaster. We will introduce the techniques to propagate these parasites in Drosophila spp. and demonstrate how to analyze their effects on immune tissues of Drosophila larvae.

Most known parasitoid wasp species attack the larval or pupal stages of Drosophila. While Trichopria drosophilae infect the pupal stages of the host (Fig. ...

Quantitative Measurement of the Immune Response and Sleep in Drosophila

A complex interaction between the immune response and host behavior has been described in a wide range of species. Excess sleep, in particular, is known to occur as a response to infection in mammals 1 and has also recently been described in Drosophila melanogaster2. It is generally accepted that sleep is beneficial to the host during an infection and that it is important for the maintenance of a robust immune system3,4. However, experimental evidence that supports this hypothesis is limited4, and the function of excess sleep during an immune response remains unclear. We have used a multidisciplinary approach to address this complex problem, and have conducted studies in the simple genetic model system, the fruitfly Drosophila melanogaster. We use a standard assay for measuring locomotor behavior and sleep in flies, and demonstrate how this assay is used to measure behavior in flies infected with a pathogenic strain of bacteria. This assay is also useful for monitoring the duration of survival in individual flies during an infection. Additional measures of immune function include the ability of flies to clear an infection and the activation of NF&kappa;B, a key transcription factor that is central to the innate immune response in Drosophila. Both survival outcome and bacterial clearance during infection together are indicators of resistance and tolerance to infection. Resistance refers to the ability of flies to clear an infection, while tolerance is defined as the ability of the host to limit damage from an infection and thereby survive despite high levels of pathogen within the system5. Real-time monitoring of NF&kappa;B activity during infection provides insight into a molecular mechanism of survival during infection. The use of Drosophila in these straightforward assays facilitates the genetic and molecular analyses of sleep and the immune response and how these two complex systems are reciprocally influenced.

Quantitative Measurement of the Immune Response and Sleep in Drosophila

To understand a link between the immune response and behavior, we describe a method to measure locomotor behavior in Drosophila during bacterial infection as well as the ability of flies to mount an immune response by monitoring survival, bacterial load, and real-time activity of a key regulator of innate immunity, NF&kappa;B.

A complex interaction between the immune response and  host behavior has been described in a wide range of species. Excess sleep, in  particular, is known ...

Quantification of the Respiratory Burst Response as an Indicator of Innate Immune Health in Zebrafish

The phagocyte respiratory burst is part of the innate immune response to pathogen infection and involves the production of reactive oxygen species (ROS). ROS are toxic and function to kill phagocytized microorganisms. In vivo quantification of phagocyte-derived ROS provides information regarding an organism's ability to mount a robust innate immune response. Here we describe a protocol to quantify and compare ROS in whole zebrafish embryos upon chemical induction of the phagocyte respiratory burst. This method makes use of a non-fluorescent compound that becomes fluorescent upon oxidation by ROS. Individual zebrafish embryos are pipetted into the wells of a microplate and incubated in this fluorogenic substrate with or without a chemical inducer of the respiratory burst. Fluorescence in each well is quantified at desired time points using a microplate reader. Fluorescence readings are adjusted to eliminate background fluorescence and then compared using an unpaired t-test. This method allows for comparison of the respiratory burst potential of zebrafish embryos at different developmental stages and in response to experimental manipulations such as protein knockdown, overexpression, or treatment with pharmacological agents. This method can also be used to monitor the respiratory burst response in whole dissected kidneys or cell preparations from kidneys of adult zebrafish and some other fish species. We believe that the relative simplicity and adaptability of this protocol will complement existing protocols and will be of interest to researchers who seek to better understand the innate immune response.

The innate immune response protects organisms against pathogen infection. A critical component of the innate immune response, the phagocyte respiratory burst, generates reactive oxygen species that kill invading microorganisms. We describe a respiratory burst assay that quantifies reactive oxygen species produced when the innate immune response is chemically induced.

The phagocyte respiratory burst is part of the innate immune response to  pathogen infection and involves the production of reactive oxygen species (ROS). ...

Using RNA-interference to Investigate the Innate Immune Response in Mouse Macrophages

Macrophages are key phagocytic innate immune cells. When macrophages encounter a pathogen, they produce antimicrobial proteins and compounds to kill the pathogen, produce various cytokines and chemokines to recruit and stimulate other immune cells, and present antigens to stimulate the adaptive immune response. Thus, being able to efficiently manipulate macrophages with techniques such as RNA-interference (RNAi) is critical to our ability to investigate this important innate immune cell. However, macrophages can be technically challenging to transfect and can exhibit inefficient RNAi-induced gene knockdown. In this protocol, we describe methods to efficiently transfect two mouse macrophage cell lines (RAW264.7 and J774A.1) with siRNA using the Amaxa Nucleofector 96-well Shuttle System and describe procedures to maximize the effect of siRNA on gene knockdown. Moreover, the described methods are adapted to work in 96-well format, allowing for medium and high-throughput studies. To demonstrate the utility of this approach, we describe experiments that utilize RNAi to inhibit genes that regulate lipopolysaccharide (LPS)-induced cytokine production.

In this protocol, we describe methods to efficiently transfect murine macrophage cell lines with siRNAs using the Amaxa Nucleofector 96-well Shuttle System, stimulate the macrophages with lipopolysaccharide, and monitor the effects on inflammatory cytokine production.

Macrophages are key phagocytic innate immune cells.  When macrophages encounter a pathogen, they produce antimicrobial proteins and compounds to kill the ...

In Vitro Analysis of Myd88-mediated Cellular Immune Response to West Nile Virus Mutant Strain Infection

An attenuated West Nile virus (WNV), a nonstructural (NS) 4B-P38G mutant, induced higher innate cytokine and T cell responses than the wild-type WNV in mice. Recently, myeloid differentiation factor 88 (MyD88) signaling was shown to be important for initial T cell priming and memory T cell development during WNV NS4B-P38G mutant infection. In this study, two flow cytometry-based methods &#8211; an in vitro T cell priming assay and an intracellular cytokine staining (ICS) &#8211; were utilized to assess dendritic cells (DCs) and T cell functions. In the T cell priming assay, cell proliferation was analyzed by flow cytometry following co-culture of DCs from both groups of mice with carboxyfluorescein succinimidyl ester (CFSE) - labeled CD4+ T cells of OTII transgenic mice. This approach provided an accurate determination of the percentage of proliferating CD4+ T cells with significantly improved overall sensitivity than the traditional assays with radioactive reagents. A microcentrifuge tube system was used in both cell culture and cytokine staining procedures of the ICS protocol. Compared to the traditional tissue culture plate-based system, this modified procedure was easier to perform at biosafety level (BL) 3 facilities. Moreover, WNV- infected cells were treated with paraformaldehyde in both assays, which enabled further analysis outside BL3 facilities. Overall, these in vitro immunological assays can be used to efficiently assess cell-mediated immune responses during WNV infection.

In Vitro Analysis of Myd88-mediated Cellular Immune Response to West Nile Virus Mutant Strain Infection

Two flow cytometry-based methods &#8211; an in vitro T cell priming assay and intracellular cytokine staining were utilized to measure antigen presenting capacity of dendritic cells and antigen-specific T cell responses to a West Nile virus mutant infection in mice.

An attenuated West Nile virus (WNV), a nonstructural (NS) 4B-P38G mutant, induced higher innate cytokine and T cell responses than the wild-type WNV in ...

Protective Efficacy and Pulmonary Immune Response Following Subcutaneous and Intranasal BCG Administration in Mice

Despite global coverage of intradermal BCG vaccination, tuberculosis remains one of the most prevalent infectious diseases in the world. Preclinical data have encouraged pulmonary tuberculosis vaccines as a promising strategy to prevent pulmonary disease, which is responsible for transmission. In this work, we describe the methodology used to demonstrate in the mouse model the benefits of intranasal BCG vaccination when compared to subcutaneous. Our data revealed greater protective efficacy following intranasal BCG administration. In addition, our results indicate that pulmonary vaccination triggers a higher immune response in lungs, including Th1 and Th17 responses, as well as an increase of immunoglobulin A (IgA) concentration in respiratory airways. Our data show correlation between protective efficacy and the presence of IL17-producing cells in lungs post-Mycobacterium tuberculosis challenge, suggesting a role for this cytokine in the protective response conferred by pulmonary vaccination. Finally, we detail the global workflow we have developed to study respiratory vaccination in the mouse model, which could be extrapolated to other tuberculosis vaccines, apart from BCG, targeting the mucosal response or other pulmonary routes of administration such as the intratracheal or aerosol.

We herein detail the methodology followed to compare protective efficacy and lung immune response induced by intranasal and subcutaneous immunization with BCG in mouse model. Our results show the benefits of pulmonary vaccination and suggest a role for IL17-mediated response in vaccine-induced protection.

Despite global coverage of intradermal BCG vaccination, tuberculosis remains one of the most prevalent infectious diseases in the world. Preclinical data ...

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is based on the injection of non-pathogenic particles into insects, as the particles will be detected by the immune system and will induce the production of immune effectors. We focus here on the stimulation of the melanization response in the mosquito Anopheles gambiae. The melanization response results in the encapsulation of foreign particles and parasites with a dark layer of melanin. To stimulate this response, mosquitoes are inoculated with beads in the thoracic cavity using microcapillary glass tubes. Then, after 24 hr, the mosquitoes are dissected to retrieve the beads. The degree of melanization of the bead is measured using image analysis software. Beads do not have the pathogenic effects of parasites, or their capacity to evade or suppress the immune response. These injections are a way to measure immune efficacy and the impact of immune stimulations on other life history traits, such as fecundity or longevity. It is not exactly the same as directly studying host-parasite interactions, but it is an interesting tool to study immunity and its evolutionary ecology.

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

Through inoculation with beads, the described technique enables the stimulation of the mosquito melanization response in the hemolymph circulating system. The amount of melanin covering the beads can be measured after dissection as a measure of the immune response.

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is ...

Bronchoalveolar Lavage of Murine Lungs to Analyze Inflammatory Cell Infiltration

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain insight into an ongoing disease state.
Here, a simple and efficient method is described to perform BAL on murine lungs without the need of special tools or equipment. BAL fluid is isolated by inserting a catheter in the trachea of terminally anesthetized mice, through which a saline solution is instilled into the bronchioles. The instilled fluid is gently retracted to maximize BAL fluid retrieval and to minimize shearing forces. This technique allows the viability, function, and structure of cells within the airways and BAL fluid to be preserved.
Numerous techniques may be applied to gain further understanding of the disease state of the lung. Here, a commonly used technique for the identification and enumeration of different types of immune cells is described, where&#160;flow cytometry is combined with a select panel of fluorescently labeled cell surface-specific markers. The BAL procedure presented here can also be used to analyze infectious agents, fluid constituents, or inhaled particles within murine lungs.

The health status of the lung is reflected by the type and number of immune cells that are present in the bronchioles of the lung. We describe a bronchoalveolar lavage technique that allows the isolation and study of nonadherent cells and soluble factors from the lower respiratory tract of mice.

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain ...

Isolation of Extracellular Vesicles from Murine Bronchoalveolar Lavage Fluid Using an Ultrafiltration Centrifugation Technique

Extracellular vesicles (EVs) are newly discovered subcellular components that play important roles in many biological signaling functions during physiological and pathological states. The isolation of EVs continues to be a major challenge in this field, due to limitations intrinsic to each technique. The differential ultracentrifugation with density gradient centrifugation method is a commonly used approach and is considered to be the gold standard procedure for EV isolation. However, this procedure is time-consuming, labor-intensive, and generally results in low scalability, which may not be suitable for small-volume samples such as bronchoalveolar lavage fluid. We demonstrate that an ultrafiltration centrifugation isolation method is simple and time- and labor-efficient yet provides a high recovery yield and purity. We propose that this isolation method could be an alternative approach that is suitable for EV isolation, particularly for small-volume biological specimens.

Here, we describe two extracellular vesicle isolation protocols, ultrafiltration centrifugation and ultracentrifugation with density gradient centrifugation, to isolate extracellular vesicles from murine bronchoalveolar lavage fluid samples. The extracellular vesicles derived from murine bronchoalveolar lavage fluid by both methods are quantified and characterized.