Name: murine oropharyngeal aspiration model of ventilator-associated and hospital-acquired bacterial pneumonia
Uploaded: 2018-06-28T14:00:00.000+00:00
Duration: 4 min 32 s
Description: Watch this Scientific Journal Video about Murine Oropharyngeal Aspiration Model of Ventilator-associated and Hospital-acquired Bacterial Pneumonia at JoVE.com

This work was supported by the National Institute of Allergy and Infectious Diseases at the National Institutes of Health [Grant Numbers R01 AI117211, R01 AI130060, R21 AI127954, and R42 AI106375 to BS] and US Food and Drug Administration [Contract HHSF223201710199C to BML].

All procedures involving animals must be approved by the researcher&#8217;s Institutional Animal Care and Use Committee (IACUC).
1. Preparation of Bacterial Inoculum
<ol>
	<li>Isolate bacterial colonies.
	<ol>
		<li>Streak a bacterial strain (e.g., A. baumannii &#8220;HUMC1&#8221;) on appropriate sterile agar medium (e.g., Tryptic soy agar), being careful to generate isolated colonies.</li>
		<li>Incubate at appropriate conditions (e.g., overnight at 37 &#176;C).</li>
	</ol>
	</li>
	<li>Grow overnight culture.
	<ol>
		<li>Select a representative, isolated colony from the agar plate with a sterile inoculating loop and use it to inoculate 10 mL of appropriate sterile broth medium (e.g., Tryptic soy broth).</li>
		<li>Allow the sample to reach the stationary phase at the proper conditions (e.g., 37 &#176;C with shaking at 200 rpm overnight in a vented-cap, 50-mL conical vial).</li>
	</ol>
	</li>
	<li>Grow the subculture.
	<ol>
		<li>Transfer 100 &#956;L of overnight culture to 10 mL of fresh sterile broth using a pipette.</li>
		<li>Allow to reach exponential/log phase at the proper conditions (e.g., 37 &#176;C with shaking at 200 rpm for 3 h in a vented-cap, 50-mL conical vial).</li>
	</ol>
	</li>
	<li>Wash the subculture.
	<ol>
		<li>Remove the subculture from its incubating environment.</li>
		<li>Centrifuge at 4,000 &#215; g for 5 min to pellet the bacteria.</li>
		<li>Aspirate and discard the supernatant.</li>
		<li>Add 10 mL of sterile phosphate-buffered saline (PBS) to the pellet and vortex vigorously until fully resuspended.</li>
		<li>Repeat steps 1.4.2 - 1.4.4 twice, for a total of 3 washes.</li>
	</ol>
	</li>
	<li>Adjust the concentration of bacterial suspension.
	<ol>
		<li>Using a spectrophotometer that measures optical density at 600 nm (OD600), measure the optical density of the bacterial suspension.</li>
		<li>Add sterile PBS to the bacterial suspension until the optical density of the inoculum drops to an OD600 of 0.5, using the equation Ci&#215;Vi = Cf&#215;Vf, where &#8220;C&#8221; is concentration, &#8220;V&#8221; is volume, &#8220;i&#8221; is initial, and &#8220;f&#8221; is final. *Cf should always be 0.5; Vf is the variable to solve.</li>
		<li>If the bacterial suspension drops below an OD600 of 0.5, centrifuge it at 4,000 &#215; g for 5 min to pellet the bacteria and remove an appropriate amount of the supernatant to reach an OD600 of 0.5.</li>
		<li>Perform serial dilutions in sterile PBS (e.g., three 1:100 dilutions to achieve 1 &#215; 10-6) and plate on sterile agar to determine the correlation coefficient that reveals the bacterial concentration in colony-forming units per mL (CFUs/mL) at an OD600 of 0.5. 
		Note: The value of this correlation coefficient will vary for each strain of each species and must be determined prior to inoculum preparation for an infection.</li>
		<li>After determining the correlation coefficient, use it to create an inoculum of the desired concentration (e.g., 2 &#215; 108-&#160; 1 &#215; 109 CFUs/mL); if the desired inoculum is greater than the OD600 of 0.5, centrifuge the bacterial suspension at 4,000 &#215; g for 5 min to pellet the bacteria and remove an appropriate amount of the supernatant to reach the desired concentration.</li>
		<li>Perform in vivo pilot studies to determine the virulence of each bacterial isolate in each strain of mouse as these values will vary between bacterial isolates of the same species and mice of different genetic backgrounds. For various Gram-negative species, the LD100 in mice is generally 1 - 5 &#215; 108 CFUs/mouse, although certain strains are lethal at lower inocula.</li>
	</ol>
	</li>
	<li>Freeze identical aliquots for future use as on-demand, precise inocula, as previously published8 (optional).
	<ol>
		<li>Prepare 1 L of bacterial inoculum as described above, transfer to two 500-mL conical vials, and centrifuge at 4,000 &#215; g for 5 min.</li>
		<li>Discard 450 mL of the supernatant from each conical vial and resuspend the pellets of bacteria in the remaining supernatant.</li>
		<li>Transfer the concentrated bacterial inoculum to a 250 mL beaker containing a magnetic stir bar and continuously mix on top of a stir plate at 300 rpm.</li>
		<li>Carefully transfer exactly 600 &#956;L of concentrated bacterial inoculum (e.g., 1 &#215; 1010 CFUs/mL) using a pipette to a 1.6-mL cryogenic vial containing exactly 300 &#956;L of sterile H2O and 300 &#956;L of sterile glycerol; mix and store at -80 &#176;C. 
		Note: The glycerol necessary for storage can confound the experimental results and cause excess mortality so it is necessary to wash it out.</li>
		<li>When ready for use, thaw the sample for 10 min at room temperature.</li>
		<li>Transfer 1 mL to a 50-mL conical vial, add 9 mL of sterile PBS, and centrifuge at 4,000 &#215; g for 5 min to pellet the bacteria. Aspirate the supernatant and resuspend the pre-determined quantity of CFUs (1 mL &#215; frozen stock concentration in CFUs/mL) in an appropriate volume of PBS to reach the desired concentration for the infectious inoculum.</li>
	</ol>
	</li>
</ol>
2. Anesthetizing Mice
<ol>
	<li>Ketamine/Xylazine 
	Note: Anesthesia with ketamine/xylazine has a long duration, which offers enough time for the researcher who is new at this technique to complete the inoculation procedure before the mouse awakens.
	<ol>
		<li>Prepare 10 mL of injectable solution by combining 8.5 mL of pharmaceutifcal grade PBS, 1 mL of 100 mg/mL Ketamine stock solution (final concentration 10 mg/mL), and 0.5 mL of 20 mg/mL Xylazine stock solution (final concentration 10 mg/mL).</li>
		<li>Administer 10 &#956;L/g by intraperitoneal (IP) injection (e.g., 250 &#956;L for a 25-g mouse).</li>
		<li>Apply ophthalmic ointment to the eyes of the mice anesthetized with Ketamine/Xylazine because their eyes are prone to damage during extended periods of anesthesia.</li>
		<li>Place the mouse in a cage and monitor until it wakes up from anesthesia. 
		Note: The animal should not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
	</ol>
	</li>
	<li>Isoflurane 
	Note: Mice quickly wake up from isoflurane-induced anesthesia after being removed from the induction chamber, so this method of anesthesia should only be used after becoming proficient at the inoculation procedure.
	<ol>
		<li>Place the na&#239;ve mice in an appropriately sized induction chamber with oxygen flowing.</li>
		<li>After the chamber is sealed shut, anesthetize the mice using a precision vaporizer to add isoflurane at 4% (v/v) for at least 5 min; confirm the anesthesia by removing a mouse from the induction chamber and measuring how long it takes to wake up from anesthesia (~60 s). 
		Note: Be aware that anesthesia times may vary based on institutional guidelines and some animals may require more than 5 min to become fully anesthetized.</li>
		<li>Maintain the anesthesia for up to 10 min by adjusting the isoflurane concentration to 2 - 3%; if total duration of anesthesia is &#62;15 min, apply ophthalmic ointment to the eyes of anesthetized mice.</li>
		<li>Place the mouse in a cage and monitor until it wakes up from anesthesia, as in step 2.1.</li>
	</ol>
	</li>
</ol>
3. Inoculating/Infecting Mice
<ol>
	<li>Suspend an anesthetized mouse, hanging it by its top incisors on a strong, thin string (e.g., dental floss) secured to a fixed object at a height approximately twice the mouse&#8217;s body length (e.g., 20 cm) above the operating surface (e.g., laboratory bench).</li>
	<li>Gently pull the mouse&#39;s tongue out of its mouth using sterile, blunt-ended forceps. Transfer the tongue to a sterile, gloved finger to prevent accidental crushing of the mouse&#8217;s tongue. Hold the tongue outside of the mouth, allowing access to the oropharynx and preventing swallowing of the inoculum to the side.</li>
	<li>While holding the mouse&#8217;s tongue, place the 50-&#956;L inoculum (bacterial suspension) into the oropharynx using a micropipette; the oropharynx is located at the junction of the oral cavity and the pharynx toward the back of the mouth. 
	Note: It is very important to only deliver liquid by pipetting to the first stop on the micropipette. The mouse will stop breathing for a few seconds when the inoculum is placed in the oropharynx; eventually, reflexive aspiration will cause the mouse to inhale the bacterial suspension, indicated by the distinctive crackling noise of liquid entering the lungs, which results in pneumonia infection.</li>
	<li>If the inoculum is not placed far back enough to block normal respiration, pinch the nares with forceps to force reflexive aspiration through the mouth and subsequent inhalation of the inoculum.</li>
	<li>After inoculation, release the tongue, remove the mouse from the suspension string, place it in a cage, and monitor until it wakes up from anesthesia, as in step 2.1.</li>
</ol>
4. Monitoring Disease Progression
Note: Due to animal suffering, various indicators should be used to indicate when mice become moribund; euthanasia should be performed following this determination, in accordance with a previously approved IACUC protocol; various markers of moribundity include body temperature, weight loss, appearance, gait, and other biomarkers that can be obtained from blood (e.g., via iSTAT)9,10,11,12,13,14,15,16.
<ol>
	<li>Euthanize the mice according to previously approved IACUC protocol (e.g., CO2 followed by cervical dislocation).</li>
	<li>Remove the lungs from euthanized mouse by cutting the trachea, pulmonary artery, and pulmonary vein close to the lungs.
	<ol>
		<li>Microscopy
		<ol>
			<li>Place the lungs (or portion) in a specimen mold.</li>
			<li>Fill the specimen mold with Optimal Cutting Temperature (O.C.T.) compound, completely submerging the tissue.</li>
			<li>Freeze the sample at -80 &#176;C.</li>
			<li>Send the sample to a pathology lab to section and mount on slides for microscopy.</li>
		</ol>
		</li>
		<li>Bacterial burden
		<ol>
			<li>Weigh the lung tissue on an analytical balance.</li>
			<li>Transfer the lung tissue to 50-mL conical vial containing 2 - 5 mL of sterile PBS (depending on the size of the tissue).</li>
			<li>Homogenize the lung tissue with a tissue homogenizer.</li>
			<li>Perform serial dilutions of the homogenate in sterile PBS to achieve approximately 1,000 CFUs/mL, based on expected bacterial burden in lungs.
			<ol>
				<li>For example, if 1 &#215; 109 CFUs/mg is expected, perform three 1:100 dilutions: Transfer 100 &#956;L of homogenate to 9.9 mL of PBS and vortex; this is the first 1:100 dilution. Transfer 100 &#956;L of the first 1:100 dilution to 9.9 mL of PBS and vortex; this is the second 1:100 dilution. Transfer 100 &#956;L of the second 1:100 dilution to 9.9 mL of PBS and vortex; this is the third and final 1:100 dilution.</li>
				<li>If bacterial burden in the lungs is unknown, perform a range of dilutions to capture the expected range.</li>
			</ol>
			</li>
			<li>Plate dilution(s) on sterile agar.
			<ol>
				<li>Prepare sterile Tryptic Soy Broth (TSB) with agar (TSA) plates: Combine 30 g of TSB, 15 g of agar, and 1 L of water, mix with a magnetic stirrer, and heat until boiling at ~100 &#176;C for 5 min. Allow to cool and then autoclave on liquid cycle for 15 min. Allow to cool to ~55 &#176;C, transfer 10 mL freshly autoclaved TSA to sterile Petri dishes, and allow to cool to room temperature. Let dry on benchtop for 2 - 5 d or uncovered under a biosafety cabinet for 10 min.</li>
				<li>Transfer 100 &#956;L of the dilution to a Petri dish containing sterile TSA and spread across TSA using a spreader or glass beads.</li>
				<li>Store the Petri dish overnight at 37 &#176;C in a humidified incubator (i.e., 37 &#176;C incubator containing an uncovered 1-L beaker filled with water)</li>
			</ol>
			</li>
			<li>Determine CFUs/mL lung homogenate based on agar plating (e.g., 100 CFUs on TSA plate with 100 &#956;L of third and final dilution described above would be 100 CFUs/100 &#956;L &#215; 100dil#1 &#215; 100dil#2 &#215; 100dil#3 = 1&#215;109 CFUs/mL).</li>
			<li>Calculate CFUs/mg lung tissue based by dividing CFUs/mL lung homogenate by mg lung tissue/mL homogenate (e.g., if sample described above used 100 mg of lung tissue homogenized in 5 mL of PBS, then 1 &#215; 109 CFUs/mL &#247; 100 mg/5 mL = 5 &#215;&#160;107 CFUs/mg).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Spellberg, B., Talbot, G. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recommended+Design+Features+of+Future+Clinical+Trials+of+Antibacterial+Agents+for+Hospital-Acquired+Bacterial+Pneumonia+and+Ventilator-Associated+Bacterial+Pneumonia.">Recommended Design Features of Future Clinical Trials of Antibacterial Agents for Hospital-Acquired Bacterial Pneumonia and Ventilator-Associated Bacterial Pneumonia.</a> Clinical Infectious Diseases. 51 (S1), S150-S170 (2010).</li><li>Kalil, A. 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=Management+of+Adults+With+Hospital-acquired+and+Ventilator-associated+Pneumonia:+2016+Clinical+Practice+Guidelines+by+the+Infectious+Diseases+Society+of+America+and+the+American+Thoracic+Society.">Management of Adults With Hospital-acquired and Ventilator-associated Pneumonia: 2016 Clinical Practice Guidelines by the Infectious Diseases Society of America and the American Thoracic Society.</a> Clinical Infectious Diseases. 63 (5), e61-e111 (2016).</li><li>Medina, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+model+of+pneumococcal+pneumonia.">Murine model of pneumococcal pneumonia.</a> Methods in Molecular Biology. , 405-410 (2010).</li><li>Azoulay-Dupuis, E., 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=In+vivo+efficacy+of+a+new+fluoroquinolone,+sparfloxacin,+against+penicillin-susceptible+and+-resistant+and+multiresistant+strains+of+Streptococcus+pneumoniae+in+a+mouse+model+of+pneumonia.">In vivo efficacy of a new fluoroquinolone, sparfloxacin, against penicillin-susceptible and -resistant and multiresistant strains of Streptococcus pneumoniae in a mouse model of pneumonia.</a> Antimicrobial Agents and Chemotherapy. 36 (12), 2698-2703 (1992).</li><li>Mizgerd, J. P., Skerrett, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+human+pneumonia.">Animal models of human pneumonia.</a> American Journal of Physiology-Lung Cellular and Molecular Physiology. 294 (3), L387-L398 (2008).</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=Non-surgical+intratracheal+instillation+of+mice+with+analysis+of+lungs+and+lung+draining+lymph+nodes+by+flow+cytometry.">Non-surgical intratracheal instillation of mice with analysis of lungs and lung draining lymph nodes by flow cytometry.</a> Journal of Visualized Experiments. (51), (2011).</li><li>Nielsen, T. B., Bruhn, K. W., Pantapalangkoor, P., Junus, J. L., Spellberg, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+virulent+Acinetobacter+baumannii+to+reduce+variability+of+in+vivo+studies.">Cryopreservation of virulent Acinetobacter baumannii to reduce variability of in vivo studies.</a> BMC Microbiology. 15, 252 (2015).</li><li>Trammell, R. A., Toth, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Markers+for+predicting+death+as+an+outcome+for+mice+used+in+infectious+disease+research.">Markers for predicting death as an outcome for mice used in infectious disease research.</a> Comparative Medicine. 61 (6), 492-498 (2011).</li><li>Bast, D. J., 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=Novel+murine+model+of+pneumococcal+pneumonia:+use+of+temperature+as+a+measure+of+disease+severity+to+compare+the+efficacies+of+moxifloxacin+and+levofloxacin.">Novel murine model of pneumococcal pneumonia: use of temperature as a measure of disease severity to compare the efficacies of moxifloxacin and levofloxacin.</a> Antimicrobial Agents and Chemotherapy. 48 (9), 3343-3348 (2004).</li><li>Hankenson, F. 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=Weight+loss+and+reduced+body+temperature+determine+humane+endpoints+in+a+mouse+model+of+ocular+herpesvirus+infection.">Weight loss and reduced body temperature determine humane endpoints in a mouse model of ocular herpesvirus infection.</a> Journal of the American Association for Laboratory Animal Science. 52 (3), 277-285 (2013).</li><li>Adamson, T. W., Diaz-Arevalo, D., Gonzalez, T. M., Liu, X., Kalkum, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypothermic+endpoint+for+an+intranasal+invasive+pulmonary+aspergillosis+mouse+model.">Hypothermic endpoint for an intranasal invasive pulmonary aspergillosis mouse model.</a> Comparative Medicine. 63 (6), 477-481 (2013).</li><li>Nielsen, T. 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=Diabetes+Exacerbates+Infection+via+Hyperinflammation+by+Signaling+through+TLR4+and+RAGE.">Diabetes Exacerbates Infection via Hyperinflammation by Signaling through TLR4 and RAGE.</a> mBio. 8 (4), (2017).</li><li>Nielsen, T. 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=Monoclonal+Antibody+Protects+Against+Acinetobacter+baumannii+Infection+by+Enhancing+Bacterial+Clearance+and+Evading+Sepsis.">Monoclonal Antibody Protects Against Acinetobacter baumannii Infection by Enhancing Bacterial Clearance and Evading Sepsis.</a> Journal of Infectious Diseases. 216 (4), 489-501 (2017).</li><li>Cheng, B. L., 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=Evaluation+of+serotypes+5+and+8+capsular+polysaccharides+in+protection+against+Staphylococcus+aureus+in+murine+models+of+infection.">Evaluation of serotypes 5 and 8 capsular polysaccharides in protection against Staphylococcus aureus in murine models of infection.</a> Human Vaccine Immunotherapy. 13 (7), 1609-1614 (2017).</li><li>Wong, D., 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=Clinical+and+Pathophysiological+Overview+of+Acinetobacter+Infections:+a+Century+of+Challenges.">Clinical and Pathophysiological Overview of Acinetobacter Infections: a Century of Challenges.</a> Clinical Microbiology Reviews. 30 (1), 409-447 (2017).</li></ol>

The authors declare that they have no competing financial interests.

To be sure, mice are not miniature humans. Results obtained from mouse models must be considered in context and subsequently interpreted for applicability to humans, based on differences and similarities between the two species6. It is also important to choose the appropriate mouse strain as certain are more susceptible to some infections than others; the same applies to the pathogen strain of choice16.
It is essential to perform infections in an...

Lower respiratory infection is the world&#39;s deadliest communicable disease and the most common cause of death in developing countries1. Globally, these infections account for more than 3.2 million deaths1. In addition, nosocomial pneumonia is among the most common and deadly forms of healthcare acquired infections, and is caused by the most antibiotic-resistant pathogens2,3. The typical route of acquisition of bacterial pneumonia for both community-acquired and nosocomial pneumonia is the aspiration of oropharyngeal contents into the alveoli. Murine models used to study these diseases often use intranasal inoculation4, depositing much of the bacteria outside the lung, causing off-target complications and symptoms like sinusitis and physical trauma, which are incongruent with the disease progression in human that the models were designed to emulate. Other models may use inhalation chambers and micromisting devices, which more accurately mimic viral, tuberculous, and fungal pneumonias, but do not accurately recapitulate the normal route of acquisition for typical bacterial pneumonias.
The murine oropharyngeal aspiration pneumonia model may be utilized to simulate the natural route and pathogenesis of bacterial pneumonia. By inoculating 50 &#181;L of the bacterial suspension into the oropharynx of anesthetized mice using a pipette, reflexive aspiration ensues, which results in infectious pneumonia. Using this model, one can examine the pathogenesis of pneumonia-causing pathogens and new treatments to combat these diseases with a higher fidelity model, more analogous to aspiration pneumonia infections observed in human. Additionally, unlike similar models that infect through the oral cavity5,6, this model ensures that the full inoculum reaches the lungs instead of the gut, where it can cause off-site inflammation and infections, such as gastritis and enteritis. Finally, unlike another published model that requires a laryngoscope and inoculates through the trachea7, this model does not obstruct the airway with a gavage needle and does not require injection for inoculum delivery. Instead, inoculation relies on the natural aspiration reflex of the mouse.

<table><tbody><tr><td>Agar</td><td>BD</td><td>214530</td><td>Combine with TSB to make TSA</td></tr><tr><td>Beads, Borosilicate Glass</td><td>Kimble</td><td>135003</td><td>Sterilize by baking or autoclaving before each use</td></tr><tr><td>Beaker, 250 mL</td><td>Pyrex</td><td>1003</td><td>Used during precise aliquoting of concentrated bacterial inocula</td></tr><tr><td>Centrifuge</td><td>Sorvall</td><td>ST 40R</td><td>Capable of 4,000&amp;times;g at 4&amp;deg;C</td></tr><tr><td>Chamber for Anesthesia</td><td>Kent Scientific Corporation</td><td>VetFlo-0720</td><td>Accommodates up to 5 mice</td></tr><tr><td>Cryomold, Intermediate Size</td><td>Sakura Tissue-Tek</td><td>4566</td><td>Disposable vinyl specimen molds, 15&amp;times;15&amp;times;5 mm</td></tr><tr><td>Dental Floss</td><td>Oral-B</td><td>37000469537</td><td>Tie to stable post approx. 6&quot; above table height</td></tr><tr><td>Forceps</td><td>VWR</td><td>82027-440</td><td>Used to gently pull tongue out of mouse&#039;s mouth</td></tr><tr><td>Homogenizer for Lung Tissue</td><td>Omni International</td><td>TM125-115</td><td>Autoclave before first use; rinse between samples</td></tr><tr><td>Isoflurane for Anesthesia</td><td>Abbott</td><td>10015516</td><td>Alternative drug can be used; modify procedure accordingly</td></tr><tr><td>iSTAT Cartridge</td><td>Abbott</td><td>03P79-25</td><td>Various cartridges are available to suit your needs</td></tr><tr><td>Ketamine, 100 mg/mL</td><td>Western Medical Supply</td><td>4165</td><td>Dilute 1:10 in PBS to 1 mg/mL and combine with Xylazine at 1 mg/mL</td></tr><tr><td>Ointment for Eyes</td><td>Akorn</td><td>Tears Renewed</td><td>Avoid touching eye with tip of dispenser</td></tr><tr><td>Optimal Cutting Temperature (O.C.T.) Compound</td><td>Fisher Scientific</td><td>23-730-571</td><td>Used to freeze lung samples at -80 &amp;deg;C to prepare for pathology sectioning</td></tr><tr><td>Petri Dish</td><td>VWR</td><td>25384-302</td><td>Polystyrene, disposable, sterilized, 100&amp;times;15 mm</td></tr><tr><td>Phosphate-Buffered Saline (PBS)</td><td>Corning</td><td>21-031-CM</td><td>Dulbecco&#039;s PBS without calcium and magnesium</td></tr><tr><td>Pipette Tips, 200-&amp;mu;L</td><td>VWR</td><td>10017-044</td><td>Autoclave before use</td></tr><tr><td>Pipetter, 200-&amp;mu;L</td><td>Gilson</td><td>Pipetman P200</td><td>Autoclave and calibrate before use</td></tr><tr><td>Spreader, Bacterial Cell</td><td>Bel-Art</td><td>F377360006</td><td>Sterilize by baking or autoclaving before each use</td></tr><tr><td>Stir Bar, Magnetic, 7.9 mm Diameter &amp;times; 38.1 mm Length</td><td>VWR</td><td>58948-150</td><td>Used for stiring concentrated bacterial inocula during aliquoting</td></tr><tr><td>Stir Plate, Magnetic</td><td>Corning</td><td>PC-620D</td><td>Used for stiring concentrated bacterial inocula during aliquoting</td></tr><tr><td>Tryptic Soy Broth (TSB)</td><td>BD</td><td>211822</td><td>Combine with Agar to make TSA</td></tr><tr><td>Vial, Conical, Sterile, 50 mL</td><td>Corning</td><td>431720</td><td>Used for preparing bacterial inocula</td></tr><tr><td>Vial, Conical, Sterile, 500 mL</td><td>Corning</td><td>431123</td><td>Used to concentrate inocula for preparing frozen inocula</td></tr><tr><td>Vial, Cryogenic, 2.0 mL</td><td>Corning</td><td>430659</td><td>Used for cryogenic storage of concentrated bacterial inocula</td></tr><tr><td>Xylazine, 20 mg/mL</td><td>Akorn</td><td>AnaSed Injection</td><td>Dilute 1:20 in PBS to 1 mg/mL and combine with Ketamine at 1 mg/mL</td></tr></tbody></table>

murine oropharyngeal aspiration model of ventilator-associated and hospital-acquired bacterial pneumonia

Murine infection models are critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics designed to combat causative pathogens. Infectious pneumonia is among the most common infections presented by patients in the clinic and thus warrants an appropriate in vivo model. Typical pneumonia models use intranasal inoculation, which deposits excessive organisms outside the lung, causing off-target complications and symptoms, such as sinusitis, gastritis, enteritis, physical trauma, or microparticle misting to mimic aerosol spread more typical of viral, tuberculous, or fungal pneumonia. These models do not accurately reflect the pathogenesis of typical community- or healthcare-acquired bacterial pneumonia. In contrast, this murine model of oropharyngeal aspiration pneumonia mimics the droplet route in healthcare-acquired pneumonia. Inoculating 50 &#181;L of the bacteria suspension into the oropharynx of anesthetized mice causes reflexive aspiration, which results in pneumonia. With this model, one can examine the pathogenesis of pneumonia-causing pathogens and new treatments to combat these diseases.

By carefully following the protocol, reproducible and robust data can be easily obtained. It is critical to strictly adhere to one&#39;s customized inoculum preparation protocol for experiments to be compared to one each other. It is also important to properly handle mice during the infection procedure. Be sure to place mice into an anesthesia chamber devoid of isoflurane. Mice will panic if they are placed into a chamber that has been pre-filled with isoflurane and may experience excess ...

Watch this Scientific Journal Video about Murine Oropharyngeal Aspiration Model of Ventilator-associated and Hospital-acquired Bacterial Pneumonia at JoVE.com

Murine Oropharyngeal Aspiration Model of Ventilator-associated and Hospital-acquired Bacterial Pneumonia

Infectious pneumonia is among the most common infections in human. An appropriate in vivo model is critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics. With this murine oropharyngeal aspiration pneumonia model, one can examine the pathogenesis and new treatments against these deadly infections.

Murine infection models are critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics designed to combat causative ...

murine-oropharyngeal-aspiration-model-ventilator-associated-hospital

Research

JoVE Journal

Medicine

11.6K Views.  University of Southern California. Infectious pneumonia is among the most common infections in human. An appropriate in vivo model is critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics. With this murine oropharyngeal aspiration pneumonia model, one can examine the pathogenesis and new treatments against these deadly infections.

Video: Murine Oropharyngeal Aspiration Model of Ventilator-associated and Hospital-acquired Bacterial Pneumonia

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 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 ...

Immunology and Infection

Pseudomonas aeruginosa Induced Lung Injury Model

In order to study human acute lung injury and pneumonia, it is important to develop animal models to mimic various pathological features of this disease. Here we have developed a mouse lung injury model by intra-tracheal injection of bacteria Pseudomonas aeruginosa (P. aeruginosa or PA). Using this model, we were able to show lung inflammation at the early phase of injury. In addition, alveolar epithelial barrier leakiness was observed by analyzing bronchoalveolar lavage (BAL); and alveolar cell death was observed by Tunel assay using tissue prepared from injured lungs. At a later phase following injury, we observed cell proliferation required for the repair process. The injury was resolved 7 days from the initiation of P. aeruginosa injection. This model mimics the sequential course of lung inflammation, injury and repair during pneumonia. This clinically relevant animal model is suitable for studying pathology, mechanism of repair, following acute lung injury, and also can be used to test potential therapeutic agents for this disease.

Pseudomonas aeruginosa Induced Lung Injury Model

We have developed a mouse lung injury model by intra-tracheal injection of bacteria Pseudomonas aeruginosa. This model mimics lung injury during pneumonia and is clinically relevant.

In order to study human acute lung injury and pneumonia, it is important to develop animal models to mimic various pathological features of this disease. ...

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

Studying host-pathogen interaction enables us to understand the underlying mechanisms of the pathogenicity during microbial infection. The prognosis of the host depends on the involvement of an adapted immune response against the pathogen1. Immune response is complex and results from interaction of the pathogens and several immune or non-immune cellular types2. In vitro studies cannot characterise these interactions and focus on cell-pathogen interactions. Moreover, in the airway3, particularly in patients with suppurative chronic lung disease or in mechanically ventilated patients, polymicrobial communities are present and complicate host-pathogen interaction. Pseudomonas aeruginosa and Candida albicans are both problem pathogens4, frequently isolated from tracheobronchial samples, and associated to severe infections, especially in intensive care unit5. Microbial interactions have been reported between these pathogens in vitro but the clinical impact of these interactions remains unclear6. To study the interactions between C. albicans and P. aeruginosa, a murine model of C. albicans airways colonization, followed by a P. aeruginosa-mediated acute lung infection was performed.

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

While in vitro study of host-pathogen interactions allow the characterization of specific immune responses, in vivo models are required to observe the effects of complex responses. Using Candida albicans exposure followed by Pseudomonas aeruginosa-mediated lung infection, we established a murine model of microbial interactions involved in ventilator-associated pneumonia pathogenicity.

Studying host-pathogen interaction enables us to understand the underlying mechanisms of the pathogenicity during microbial infection. The prognosis of ...

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant preclinical advances in our understanding of the underlying cellular and molecular pathogenesis. In vivo mouse models capture the integrated physiology and resilience of the host defense response in a manner not revealed by alternative, simplified ex vivo approaches. Several methods have been described in the literature for intrapulmonary inoculation of bacteria in mice, including aerosolization, intranasal delivery, peroral endotracheal cannulation under &#39;blind&#39; and visualized conditions, and transcutaneous endotracheal cannulation. All methods have relative merits and limitations. Herein, we describe in detail a non-invasive, technically non-intensive, inexpensive, and rapid method for intratracheal delivery of bacteria that involves aspiration (i.e., inhalation) by the mouse of an infectious inoculum pipetted into the oropharynx while under general anesthesia. This method can be used for pulmonary delivery of a wide variety of non-caustic biological and chemical agents, and is relatively easy to learn, even for laboratories with minimal prior experience with pulmonary procedures. In addition to describing the aspiration pneumonia method, we also provide step-by-step procedures for assaying the subsequent in vivo pulmonary innate immune response of the mouse, in particular, methods for quantifying bacterial clearance and the cellular immune response of the infected airway. This integrated and simple approach to pneumonia assessment allows for rapid and robust evaluation of the effect of genetic and environmental manipulations upon pulmonary innate immunity.

Several methods have been described in the literature for modeling bacterial pneumonia in mice. Herein, we describe a non-invasive, inexpensive, rapid method for inducing pneumonia via aspiration (i.e., inhalation) of a bacterial inoculum pipetted into the oropharynx. Downstream methods for assessment of the pulmonary innate immune response are also detailed.

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant ...

A Robust Pneumonia Model in Immunocompetent Rodents to Evaluate Antibacterial Efficacy against S. pneumoniae, H. influenzae, K. pneumoniae, P. aeruginosa or A. baumannii

Efficacy of candidate antibacterial treatments must be demonstrated in animal models of infection as part of the discovery and development process, preferably in models which mimic the intended clinical indication. A method for inducing robust lung infections in immunocompetent rats and mice is described which allows for the assessment of treatments in a model of serious pneumonia caused by S. pneumoniae, H. influenzae, P. aeruginosa, K. pneumoniae or A. baumannii. Animals are anesthetized, and an agar-based inoculum is deposited deep into the lung via nonsurgical intratracheal intubation. The resulting infection is consistent, reproducible, and stable for at least 48 h and up to 96 h for most isolates. Studies with marketed antibacterials have demonstrated good correlation between in vivo efficacy and in vitro susceptibility, and concordance between pharmacokinetic/pharmacodynamic targets determined in this model and clinically accepted targets has been observed. Although there is an initial time investment when learning the technique, it can be performed quickly and efficiently once proficiency is achieved. Benefits of the model include elimination of the neutropenic requirement, increased robustness and reproducibility, ability to study more pathogens and isolates, improved flexibility in study design and establishment of a challenging infection in an immunocompetent host.

A Robust Pneumonia Model in Immunocompetent Rodents to Evaluate Antibacterial Efficacy against S. pneumoniae, H. influenzae, K. pneumoniae, P. aeruginosa or  A. baumannii

A method for establishing lung infections in immunocompetent rodents is described. With proficiency, this method can be performed quickly and easily to induce stable infection with many isolates of S. pneumoniae, H. influenzae, P. aeruginosa, K. pneumoniae and A. baumannii.

Efficacy of candidate antibacterial treatments must be demonstrated in animal models of infection as part of the discovery and development process, ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

A Precise Pathogen Delivery and Recovery System for Murine Models of Secondary Bacterial Pneumonia

Secondary bacterial pneumonias following influenza infections consistently rank within the top ten leading causes of death in the United States. To date, murine models of co-infection have been the primary tool developed to explore the pathologies of both the primary and secondary infections. Despite the prevalence of this model, considerable discrepancies regarding instillation procedures, dose volumes, and efficacies are prevalent among studies. Furthermore, these efforts have been largely incomplete in addressing how the pathogen may be directly influencing disease progression post-infection. Herein we provide a precise method of pathogen delivery, recovery, and analysis to be used in murine models of secondary bacterial pneumonia. We demonstrate that intratracheal instillation enables an efficient and accurate delivery of controlled volumes directly and evenly into the lower respiratory tract. Lungs can be excised to recover and quantify the pathogen burden. Following excision of the infected lungs, we describe a method to extract high quality pathogen RNA for subsequent transcriptional analysis. This procedure benefits from being a non-surgical method of delivery without the use of specialized laboratory equipment and provides a reproducible strategy to investigate pathogen contributions to secondary bacterial pneumonia.

Here, we present methods to improve secondary bacterial pneumonia studies by providing a non-invasive route of instillation into the lower respiratory tract followed by pathogen recovery and transcript analysis. These procedures are reproducible and can be performed without specialized equipment such as cannulas, guide wires, or fiber optic cables.

Secondary bacterial pneumonias following influenza infections consistently rank within the top ten leading causes of death in the United States. To date, ...

A Mouse Model for the Transition of Streptococcus pneumoniae from Colonizer to Pathogen upon Viral Co-Infection Recapitulates Age-Exacerbated Illness

Streptococcus pneumoniae (pneumococcus) is an asymptomatic colonizer of the nasopharynx in most individuals but can progress to a pulmonary and systemic pathogen upon influenza A virus (IAV) infection. Advanced age enhances host susceptibility to secondary pneumococcal pneumonia and is associated with worsened disease outcomes. The host factors driving those processes are not well defined, in part due to a lack of animal models that reproduce the transition from asymptomatic colonization to severe clinical disease. 
This paper describes a novel mouse model that recreates the transition of pneumococci from asymptomatic carriage to disease upon viral infection. In this model, mice are first intranasally inoculated with biofilm-grown pneumococci to establish asymptomatic carriage, followed by IAV infection of both the nasopharynx and lungs. This results in bacterial dissemination to the lungs, pulmonary inflammation, and obvious signs of illness that can progress to lethality. The degree of disease is dependent on the bacterial strain and host factors. 
Importantly, this model reproduces the susceptibility of aging, because compared to young mice, old mice display more severe clinical illness and succumb to disease more frequently. By separating carriage and disease into distinct steps and providing the opportunity to analyze the genetic variants of both the pathogen and the host, this S. pneumoniae/IAV co-infection model permits the detailed examination of the interactions of an important pathobiont with the host at different phases of disease progression. This model can also serve as an important tool for identifying potential therapeutic targets against secondary pneumococcal pneumonia in susceptible hosts.

A Mouse Model for the Transition of Streptococcus pneumoniae from Colonizer to Pathogen upon Viral Co-Infection Recapitulates Age-Exacerbated Illness

This paper describes a novel mouse model for the transition of pneumococcus from an asymptomatic colonizer to a disease-causing pathogen during viral infection. This model can be readily adapted to study polymicrobial and host-pathogen interactions during the different phases of disease progression and across various hosts.

Streptococcus pneumoniae (pneumococcus) is an asymptomatic colonizer of the nasopharynx in most individuals but can progress to a pulmonary and systemic ...

Experimental Model to Evaluate Resolution of Pneumonia

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to high morbidity and mortality. Currently, there are no pre-clinical models that recapitulate the complexity of human ARDS. However, infectious models of pneumonia (PNA) can replicate the main pathophysiological features of ARDS. Here, we describe a model of PNA induced by the intratracheal instillation of live Streptococcus pneumoniae and Klebsiella pneumoniae in C57BL6 mice. In order to evaluate and characterize the model, after inducing injury, we carried out serial measurements of body weight and bronchoalveolar lavage (BAL) for measuring markers of lung injury. Additionally, we harvested lungs for cell count and differentials, BAL protein quantification, cytospin, bacterial colony-forming unit counts, and histology. Lastly, high dimensional flow cytometry was performed. We propose this model as a tool to understand the immune landscape during the early and late resolution phases of lung injury.

This manuscript describes the establishment of an infectious model of pneumonia in mice and the respective characterization of injury resolution along with methods for growing bacteria and intratracheal instillation. A novel approach using high-dimensional flow cytometry to evaluate the immune landscape is also described.

Acute respiratory distress syndrome (ARDS) causes acute lung injury, characterized by rapid alveolar damage and severe hypoxemia. This, in turn, leads to ...

Establishing a Lung-on-Chip Model for Respiratory Infection Studies

Immunocompetent Alveolus-on-Chip Model for Studying Alveolar Mucosal Immune Responses

We introduce an advanced immunocompetent lung-on-chip model designed to replicate the human alveolar structure and function. This innovative model employs a microfluidic-perfused biochip that supports an air-liquid interface mimicking the environment in the human alveoli. Tissue engineering is used to integrate key cellular components, including endothelial cells, macrophages, and epithelial cells, to create a representative tissue model of the alveolus. The model facilitates in-depth examinations of the mucosal immune responses to various pathogens, including viruses, bacteria, and fungi, thereby advancing our understanding of lung immunity. The primary goal of this protocol is to provide details for establishing this alveolus-on-chip model as a robust in vitro platform for infection studies, enabling researchers to closely observe and analyze the complex interactions between pathogens and the host's immune system within the pulmonary environment. This is achieved through the application of microfluidic-based techniques to simulate key physiological conditions of the human alveoli, including blood flow and biomechanical stimulation of endothelial cells, alongside maintaining an air-liquid interface crucial for the realistic exposure of epithelial cells to air. The model system is compatible with a range of standardized assays, such as immunofluorescence staining, cytokine profiling, and colony-forming unit (CFU)/plaque analysis, allowing for comprehensive insights into immune dynamics during infection. The Alveolus-on-chip is composed of essential cell types, including human distal lung epithelial cells (H441) and human umbilical vein endothelial cells (HUVECs) separated by porous polyethylene terephthalate (PET) membranes, with primary monocyte-derived macrophages strategically positioned between the epithelial and endothelial layers. The tissue model enhances the ability to dissect and analyze the nuanced factors involved in pulmonary immune responses in vitro. As a valuable tool, it should contribute to the advancement of lung research, providing a more accurate and dynamic in vitro model for studying the pathogenesis of respiratory infections and testing potential therapeutic interventions.

Author Spotlight: Developing a Microfluidic Lung-on-Chip Model for In-Depth Study of Human Immune Response and Infection Mechanisms

Lung-on-chip models surpass traditional 2D cultures by mimicking the air-liquid interface and endothelial cell perfusion, simulating blood flow and nutrient exchange crucial for lung physiology studies. This enhances lung research relevance, offering a dynamic, physiologically accurate environment to advance the understanding and treatment of respiratory infections.

We introduce an advanced immunocompetent lung-on-chip model designed to replicate the human alveolar structure and function. This innovative model employs ...

Murine Oropharyngeal Aspiration Model of Ventilator-associated and Hospital-acquired Bacterial Pneumonia

Summary

Explore More Videos

Chapters in this video

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

Pseudomonas aeruginosa Induced Lung Injury Model

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

A Robust Pneumonia Model in Immunocompetent Rodents to Evaluate Antibacterial Efficacy against S. pneumoniae, H. influenzae, K. pneumoniae, P. aeruginosa or A. baumannii

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

A Precise Pathogen Delivery and Recovery System for Murine Models of Secondary Bacterial Pneumonia

A Mouse Model for the Transition of Streptococcus pneumoniae from Colonizer to Pathogen upon Viral Co-Infection Recapitulates Age-Exacerbated Illness

Experimental Model to Evaluate Resolution of Pneumonia

Immunocompetent Alveolus-on-Chip Model for Studying Alveolar Mucosal Immune Responses