This study was supported by the Defense Threat Reduction Agency (DTRA) under contract HDTRA1-14-C-0116, and the Center for Excellence in Infection Genomics (DOD #W911NF-11-1-0136).

This work was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Animal protocols involving rodents were approved by the University of Texas at San Antonio Institutional Animal Care and Use Committee (IACUC) under protocol MU009(RA).
1. Prepare Catheter, Trachea Indicator, and F. tularensis Inoculum
<ol>
	<li>Prepare Catheter (20 G x 2 in)
	<ol>
		<li>Cut the 20 G x 2 in needle and grind the tip of the needle to a blunt and smooth finish with a rotary tool. 
		NOTE: The length of the blunted needle should allow the metal tip to rescind approximately 3 mm into the catheter sleeve when fully seated onto the blunted needle. The blunted needle gives the catheter structural rigidity to allow placement of the catheter in the trachea, and the absence of the blunted needle tip protruding from the catheter sleeve prevents injury to the trachea.</li>
		<li>Clean the blunted needles and catheters using 70% ethanol and sterilize under UV light for 15 min.</li>
	</ol>
	</li>
	<li>Prepare Trachea Indicator
	<ol>
		<li>Trim a 1,000 &#181;L pipette tip to allow the tip to sit into a catheter opening and form an air-tight seal.</li>
		<li>Remove a filter from a 200 &#181;L aerosol barrier pipette tip and place inside the previously trimmed 1,000 &#181;L pipette tip.</li>
		<li>Ensure the filter can move freely inside the 1,000 &#181;L pipette tip when the tip is pointed tip-down.</li>
	</ol>
	</li>
	<li>F. tularensis Inoculum
	<ol>
		<li>Grow F. tularensis strain overnight on appropriate agar plate at 37 &#176;C. Scrape approximately 100 &#181;L of the bacterial lawn from the agar plate with a sterile inoculating loop and use to inoculate a 500 mL Erlenmeyer flask containing 250 mL of appropriate liquid growth medium (for Fn: tryptic soy broth (TSB) supplemented with 0.1% L-Cysteine hydrochloride monohydrate) and incubate shaking overnight at 37 &#176;C. 
		NOTE: F. novicida used in the video was grown on a tryptic soy agar (TSA) plate supplemented with 0.1% L-Cysteine hydrochloride monohydrate.</li>
		<li>Centrifuge the overnight grown liquid culture at 4,221 x g at room temperature for 10 min and remove the supernatant without disrupting the bacteria pellet. 
		CAUTION: Please follow institution&#39;s biosafety recommendations to discard supernatant.</li>
		<li>Gently re-suspend the bacterial pellet with 25 mL of appropriate liquid medium (Fn: TSB supplemented with 0.1% L-Cysteine hydrochloride monohydrate) containing 10% glycerol, using a 25 mL pipette.</li>
		<li>Using a 1 mL pipette, aliquot the resuspended bacterial culture into 500 &#181;L aliquots in vials, freeze in a dry ice/ethanol bath, and store at -80 &#176;C.</li>
		<li>To determine the titer of the frozen culture vials, remove two frozen vials stored at -80 &#176;C, thaw on ice, and perform serial dilutions in Phosphate-buffered Saline (PBS) followed by plating on appropriate growth medium (Fn: TSA containing 0.1% L-Cysteine hydrochloride). Plates are incubated 24 - 48 h at 37 &#176;C, and colony forming units (CFU) are enumerated to calculate the number of bacterial cells within the frozen culture vials (average of two vials).</li>
		<li>Prepare the bacterial inoculum by thawing a frozen vial on ice, and then dilute the culture with PBS to the final concentration of 107 CFU/100 &#181;L. The inoculum should be at a concentration that will yield the desired CFU in 100 &#181;L (the final concentration of glycerol within the inoculum cannot be more than 3%, to prevent asphyxiation of the rat). Frozen culture vials stored at -80 &#176;C can be used for up to six months after preparation.</li>
	</ol>
	</li>
</ol>
2. Rat Anesthesia
<ol>
	<li>Connect the anesthesia chamber to the properly operating isoflurane vaporizer of an anesthesia machine.</li>
	<li>Connect the gas scavenging tube on the anesthesia chamber to the gas scavenging system.</li>
	<li>Open the oxygen flow on the anesthesia machine to 4 L/min.</li>
	<li>Set the isoflurane vaporizer to 5%.</li>
	<li>Place the rat in the anesthesia chamber. 
	NOTE: We typically utilize rats between 8 - 10 weeks old (130 - 180 g); younger rats are difficult to inoculate via this technique due to the small size of the mouth cavity.</li>
	<li>When the induction of anesthesia has taken place, maintain the rat at 5% isoflurane, 2 L/min oxygen to effect. This takes about 3 - 10 min.</li>
	<li>Determine the depth of anesthesia by the quality and rate of respiration and heartbeat, and reaction to reflex stimulation as in the toe pinch test. Ideal depth of anesthesia is indicated by 1 - 1.5 s count between each breath.</li>
</ol>
3. Intratracheal Inoculation
<ol>
	<li>Remove the rat from the anesthesia chamber and place the rat dorsally on the rodent intubation stand. Attach the front teeth of rat to the holder to keep rat in place. 
	NOTE: If the rat starts to awaken during the procedure, the catheter can be removed and the rat returned to the anesthesia chamber to reach a deeper plane of anesthesia.</li>
	<li>Use dominant hand to move the tongue to same side as support hand with broad point dressing thumb forceps.</li>
	<li>Use support hand and secure the tongue with the laryngoscope against the rat&#39;s lower jaw and visualize the trachea and esophagus of the animal. The trachea will open and close as the rat breathes.</li>
	<li>Insert the catheter containing a 20-gauge blunted-needle, prepared in step 1.1, into the trachea. There may be slight resistance, and insertion into trachea may be &#34;bumpy&#34; due to the catheter rubbing the tracheal cartilages. Due to moisture or shallow respiration, the trachea can be covered by the epiglottis which prevents visualization of the trachea. Gently touching the edge of the epiglottis cartilage will cause the cartilage flap to open and uncover the trachea.</li>
	<li>Remove the blunted-needle from the catheter while ensuring the catheter remains in the trachea.</li>
	<li>Allow a few seconds to pass for rat to be able to breathe around catheter inserted into trachea.</li>
	<li>Firmly seat the trachea indicator onto the catheter opening. Movement of the aerosol barrier will indicate the catheter is inserted correctly into the trachea.</li>
	<li>Ensure that the rat is laying on the intubation stand such that the chest of the animal is facing perpendicular to the plane of the intubation stand.</li>
	<li>Remove the trachea indicator and deliver 100 &#181;L (107 CFU) of the inoculum containing F. tularensis using a 200 &#181;L pipette tip, firmly seating the tip against the catheter opening.</li>
	<li>Attach a 1 mL slip tip tuberculin syringe filled with air and deliver 300 &#181;L of air to ensure inoculum reaches the lungs of the rat.</li>
	<li>Remove the catheter from the trachea and remove the rat from the surgical platform.</li>
	<li>Allow the rat to awaken and return to the cage. Make sure that the breathing has returned to normal.</li>
	<li>Determine the CFU in the F. tularensis inoculum as described in step 1.3.5 to confirm the CFU delivered in the trachea.</li>
	<li>Repeat steps 3.1 - 3.9 on unvaccinated (na&#239;ve) animals with 100 &#181;L PBS in place of the inoculum.</li>
</ol>

<ol>
	<li>Ellis, J., Oyston, P. C. F., Green, M., Titball, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tularemia.">Tularemia.</a> Clin Microbiol Rev. 15 (4), 631-646 (2002).</li><li>Lyons, C. R., Wu, T. H. <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+Francisella+tularensis+infection.">Animal models of Francisella tularensis infection.</a> Ann N Y Acad Sci. 1105, (2007).</li><li>Hutt, J. A., Lovchik, J. A., Dekonenko, A., Hahn, A. C., Wu, T. 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+Natural+History+of+Pneumonic+Tularemia+in+Female+Fischer+344+Rats+after+Inhalational+Exposure+to+Aerosolized+Francisella+tularensis+subspecies+tularensis+Strain+Schu+S4.">The Natural History of Pneumonic Tularemia in Female Fischer 344 Rats after Inhalational Exposure to Aerosolized Francisella tularensis subspecies tularensis Strain Schu S4.</a> Am J Pathol. 187 (2), 252-267 (2017).</li><li>Ray, H. 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=The+Fischer+344+rat+reflects+human+susceptibility+to+Francisella+pulmonary+challenge+and+provides+a+new+platform+for+virulence+and+protection+studies.">The Fischer 344 rat reflects human susceptibility to Francisella pulmonary challenge and provides a new platform for virulence and protection studies.</a> PloS one. 5, e9952 (2010).</li><li>Jemski, J. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Respiratory+tularemia:+comparison+of+selected+routes+of+vaccination+in+Fischer+344+rats.">Respiratory tularemia: comparison of selected routes of vaccination in Fischer 344 rats.</a> Infect Immun. 34 (3), 766-772 (1981).</li><li>Wu, T. H., 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=Vaccination+of+Fischer+344+rats+against+pulmonary+infections+by+Francisella+tularensis+type+A+strains.">Vaccination of Fischer 344 rats against pulmonary infections by Francisella tularensis type A strains.</a> Vaccine. 27 (34), 4684-4693 (2009).</li><li>Mara-Koosham, G., Hutt, J. A., Lyons, C. R., Wu, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antibodies+contribute+to+effective+vaccination+against+respiratory+infection+by+type+A+Francisella+tularensis+strains.">Antibodies contribute to effective vaccination against respiratory infection by type A Francisella tularensis strains.</a> Infect Immun. 79 (4), 1770-1778 (2011).</li><li>Oyston, P. C., Sjostedt, A., Titball, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tularaemia:+bioterrorism+defence+renews+interest+in+Francisella+tularensis.">Tularaemia: bioterrorism defence renews interest in Francisella tularensis.</a> Nat Rev Microbiol. 2 (12), 967-978 (2004).</li><li>Kingry, L. C., Petersen, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+review+of+Francisella+tularensis+and+Francisella+novicida.">Comparative review of Francisella tularensis and Francisella novicida.</a> Front Cell Infect Microbiol. 4, 35 (2014).</li><li>Rohmer, 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=Comparison+of+Francisella+tularensis+genomes+reveals+evolutionary+events+associated+with+the+emergence+of+human+pathogenic+strains.">Comparison of Francisella tularensis genomes reveals evolutionary events associated with the emergence of human pathogenic strains.</a> Genome Biol. 8 (6), R102 (2007).</li><li>Chu, P., 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=Live+attenuated+Francisella+novicida+vaccine+protects+against+Francisella+tularensis+pulmonary+challenge+in+rats+and+non-human+primates.">Live attenuated Francisella novicida vaccine protects against Francisella tularensis pulmonary challenge in rats and non-human primates.</a> PLoS Pathog. 10 (10), e1004439 (2014).</li><li>Signarovitz, A. 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=Mucosal+Immunization+with+Live+Attenuated+Francisella+novicida+U112&#916;iglB+Protects+against+Pulmonary+F.+tularensis+SCHU+S4+in+the+Fischer+344+Rat+Model.">Mucosal Immunization with Live Attenuated Francisella novicida U112&#916;iglB Protects against Pulmonary F. tularensis SCHU S4 in the Fischer 344 Rat Model.</a> PloS one. 7 (10), e47639 (2012).</li><li>Cunningham, A. 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=Enhancement+of+vaccine+efficacy+by+expression+of+a+TLR5+ligand+in+the+defined+live+attenuated+Francisella+tularensis+subsp.+novicida+strain+U112DiglB::fljB.">Enhancement of vaccine efficacy by expression of a TLR5 ligand in the defined live attenuated Francisella tularensis subsp. novicida strain U112DiglB::fljB.</a> Vaccine. 32 (40), 5234-5240 (2014).</li></ol>

The authors have nothing to disclose

The Fischer 344 rat is becoming an important model for tularemia vaccine development3. Exposure to F. tularensis through the pulmonary route is critical for demonstrating efficacy against weaponized forms of F. tularensis, because these are delivered as aerosols. Intratracheal inoculation of the rat facilitates exposure of the rat lungs to F. tularensis without the need for large, expensive, and complicated aerosol generating equipment. All experiments utilizing select agent forms of F. tularensis additionally require BSL3 containment, which typically occurs in severely restricted space. Thus, this technique minimizes the amount of additional equipment that needs to be housed within that work environment.
Because Fn was the F. tularensis strain utilized for videography, all techniques were performed under BSL2 containment. Adaptation of this technique to the BSL3 environment includes all procedures performed within a biosafety cabinet by personnel wearing biosafety gear (full hood, powered air purifying respirator (PAPR), protective cover all with hood, double gloves, booties), and these adaptations reduce mobility, dexterity, and visibility. The trachea indicator is an important component that allows the confirmation that the catheter was correctly placed within the trachea, considering it is often difficult to otherwise make this determination when working under BSL3 conditions.
There are anatomically correct rat &#34;simulators&#34; that have a trachea and esophagus, and these are useful to perfect the technique prior to working with live animals. However, working with the simulator is not identical to working with a live rat. One means to determine if this technique is performed correctly in the live animal is to utilize Trypan blue dye as the inoculum on an anesthetized rat, and after the procedure immediately euthanize the animal. Dissection of the lung tissue and stomach will reveal if the dye was delivered into the lungs and not the esophagus. A rat that has been vaccinated with Fn/Ft by this technique can also be euthanized shortly after inoculation and the lung tissue plated to determine actual deposition within the lung.
Correct intratracheal inoculation will be important for evaluation of tularemia vaccine efficacy in the Fischer 344 rat, but it may also be useful for other vaccine and/or therapeutic applications in rats as well, including biodefense. Thus, this technique may be adaptable to a variety of pulmonary applications utilizing the rat model. While delivery by an aerosol generating device may be more similar to a biothreat scenario, the intratracheal inoculation of F. tularensis represents a relatively simple, cost-effective alternative for tularemia vaccine development.

F. tularensis (Ft) causes the human disease, tularemia. When the bacteria are acquired through the pulmonary route, this leads to pneumonic tularemia, which has high morbidity and mortality1. F. tularensis is considered a biothreat agent due to the danger associated with aerosolized forms, and there is currently no vaccine approved for human use in the U.S. An intensive effort is currently underway to develop vaccines and therapeutic measures against pneumonic tularemia, to protect the human population against the illicit use of this bacterial biothreat.
Much of the tularemia research has focused on the mouse model, due to the extreme sensitivity of mice to F. tularensis infection, and the prevalence of reagents. However, mice have proven to be a difficult model for vaccine development, due to the difficulty of demonstrating vaccine efficacy in this model2. Recently, the Fischer 344 rat has been developed as a model for tularemia vaccine development3. The sensitivity of the Fischer 344 rat to various F. tularensis subspecies mimics human sensitivity4, and rats can be protected against F. tularensis pulmonary challenge by vaccination with a live vaccine strain known to protect humans5,6,7. Because the Fischer 344 rat models some features of F. tularensis infection of humans, it may be an extremely useful model for the development of a vaccine that protects against pulmonary F. tularensis exposure.
An effective vaccine needs to protect humans against pulmonary exposure to F. tularensis. The most likely pulmonary exposure from weaponized F. tularensis would be aerosolized bacteria inhaled into the lungs8. However, aerosol generation of F. tularensis is both dangerous and cumbersome, and requires specialized equipment and containment. An alternate route of pulmonary exposure in the rat that is perhaps more adaptable for multiple laboratories lacking specialized equipment is via intratracheal inoculation6. This technique utilizes a laryngoscope for the correct placement of a catheter within the trachea of an anesthetized rat. Placement within the trachea, rather than the esophagus, is verified by a simple device that visualizes airflow from the lungs. F. tularensis is subsequently delivered into the lungs through the catheter by administration with a syringe, followed by the introduction of air into the catheter to ensure pulmonary delivery of the bacteria. In contrast, Jemski5 previously reported that F. tularensis inoculated into Fischer 344 rats via the intranasal route could not be cultured from the lungs until 3 days post-inoculation, indicating that intranasal inoculation in rats does not result in direct delivery of bacteria into the lungs.
Select agent forms of F. tularensis (F. tularensis subsp. tularensis, F. tularensis subsp. holarctica) require Biosafety Level 3 (BSL3) containment procedures, which would prevent videography. However, F. novicida (Fn) is exempt from select agent status due to its avirulence in healthy humans, and can be utilized safely under Biosafety Level 2 (BSL2) conditions9,10. Moreover, Fn serves as the basis for live attenuated vaccines that can protect against F. tularensis pulmonary exposure when delivered via intratracheal inoculation11,12,13. The technique presented here allows for the study of infections that occur through the pulmonary route utilizing rats as a model for humans. This technique can be performed without the need for specialized aerosol-generating equipment. Fn was used for the techniques filmed here.

<table><tbody><tr><td>GreenLine fiber optic blade size 0</td><td>Carefusion</td><td>5-5231-00</td><td>Macintosh American profile</td></tr><tr><td>GreenLight system laryngoscope handle</td><td>Carefusion</td><td>4559GSP</td><td></td></tr><tr><td>Exel International Safelet I.V. Catheter</td><td>EXEL INTERNATIONAL</td><td>26743</td><td></td></tr><tr><td>Slip Tip Sterile Syringes 1mL</td><td>BD</td><td>309659</td><td></td></tr><tr><td>Broad Point Dressing Thumb Forceps</td><td>Thermo Scientific</td><td>76-302</td><td></td></tr><tr><td>200 &amp;mu;L barrier tip</td><td>GeneseeScientific</td><td>24-142</td><td></td></tr><tr><td>1,000 &amp;mu;L pipette tip</td><td>Olympus Plastics</td><td>24-173</td><td></td></tr><tr><td>Dremel 3000-2/28 Rotary tool kit</td><td>Dremel</td><td>3000228</td><td></td></tr><tr><td>Rodent Intubation Stand</td><td>Braintree Scientific</td><td>RIS 200</td><td></td></tr><tr><td>Isoflurane</td><td>Butler Schein</td><td>NDC 11695-6776-2</td><td></td></tr><tr><td>Rodent anesthesia machine</td><td>Surgivet</td><td>VTC302 Classic T3</td><td></td></tr><tr><td>Rodent Anesthesia chamber</td><td>Braintree Scientific</td><td>AB 1</td><td></td></tr></tbody></table>

intratracheal inoculation of fischer 344 rats with francisella tularensis

Pulmonary infection with the bacterium Francisella tularensis can lead to the serious and potentially fatal disease, tularemia, in humans. Due to the current lack of an approved tularemia vaccine for humans, research is focused on vaccine development utilizing appropriate animal models. The Fischer 344 rat has emerged as a model that reflects human susceptibility to F. tularensis infection, and thus is an attractive model for tularemia vaccine development. Intratracheal inoculation of the Fischer 344 rat with F. tularensis mimics pulmonary exposure in humans. The successful delivery into the rat trachea is critical for pulmonary delivery. A laryngoscope with illumination is used to properly intubate the tracheae of anesthetized rats; the correct placement within the trachea is determined by a simple device to detect breathing. Following intubation, the F. tularensis culture is delivered in a measured dose via syringe. This technique standardizes pulmonary delivery of F. tularensis within the rat trachea to evaluate vaccine efficacy.

The humoral response to intratracheal inoculation of F. tularensis in the rat can be determined by enzyme-linked immunosorbent assay (ELISA) against UV-inactivated bacteria, as described previously11. Total Immunoglobulin G (IgG) response of Fischer 344 rats to inactivated whole cell bacteria was assessed post-intratracheal inoculation with an attenuated strain of Fn (107 CFU inoculum) at day 14 and day 28 (Figure 1). Mock-vaccinated rats received PBS intratracheally. An increase in serum antibody titers against Fn post-inoculation relative to na&#239;ve mock-vaccinated animals indicates the intratracheal vaccination efficacy. Low serum reactivity may indicate incorrect intratracheal placement.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56123/56123fig1.jpg" /> 
Figure 1: Total IgG Responses to Live Attenuated Fn Intratracheal Inoculation in F344 Rats. Sera from F344 rats (n = 5) inoculated intratracheally with a live attenuated Fn strain (107 CFU) were analyzed for total IgG levels reactive with whole cell Fn at day 14 and day 28 post-inoculation. Mock-vaccinated (na&#239;ve) rats (n = 5) were inoculated intratracheally with PBS. Red area denotes reactivity of na&#239;ve sera (F344 rats mock-vaccinated with PBS). The error bars represent the SEM. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56123/56123fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Intratracheal Inoculation of Fischer 344 Rats with Francisella tularensis at JoVE.com

Intratracheal Inoculation of Fischer 344 Rats with Francisella tularensis

This protocol describes intratracheal inoculations of Fischer 344 rats with Francisella tularensis. This procedure mimics pulmonary exposure of humans to this potential biothreat agent and can be used to test vaccine and therapeutic efficacy against pulmonary tularemia.

Intratracheal Inoculation of Fischer 344 Rats with Francisella tularensis

Pulmonary infection with the bacterium Francisella tularensis can lead to the serious and potentially fatal disease, tularemia, in humans. Due to the ...

intratracheal-inoculation-fischer-344-rats-with-francisella

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

7.1K Views.  University of Texas San Antonio. Dit protocol beschrijft intratracheale inentingen van Fischer 344 ratten met Francisella tularensis. Deze procedure bootst pulmonaire blootstelling van mensen aan deze potentiële biothreat agent en kan worden gebruikt voor het testen van vaccins en de therapeutische werkzaamheid tegen pulmonale tularemie.

Video: Intratracheal Inoculation of Fischer 344 Rats with Francisella tularensis

Non-surgical Intratracheal Instillation of Mice with Analysis of Lungs and Lung Draining Lymph Nodes by Flow Cytometry

Phagocytic cells such as alveolar macrophages and lung dendritic cells (LDCs) continuously sample antigens from the alveolar spaces in the 
lungs. LDCs, in particular, are known to migrate to the lung draining lymph nodes (LDLNs) where they present inhaled antigens to T cells initiating an 
appropriate immune response to a variety of immunogens1,2. To model interactions between the lungs and airborne antigens in mice, antigens can be 
administered intranasally1,3,4, intratracheally5 or as aerosols6. Delivery by each route involves distinct technical skills and limitations that need to be 
considered before designing an experiment. For example, intranasal and aerosolized exposure delivers antigens to both the lungs and the upper respiratory 
tract. Hence antigens can access the nasal associated lymphoid tissue (NALT)7, potentially complicating interpretation of the results. In addition, swallowing, 
sneezing and the breathing rate of the mouse may also lead to inconsistencies in the doses delivered. Although the involvement of the upper respiratory tract may 
be preferred for some studies, it can complicate experiments focusing on events specifically initiated in the lungs. In this setting, the intratracheal (i.t) 
route is preferable as it delivers test materials directly into the lungs and bypasses the NALT. Many i.t injection protocols involve either blind intubation of the 
trachea through the oral cavity or surgical exposure of the trachea to access the lungs. Herein, we describe a simple, consistent, non-surgical method for i.t 
instillation. The opening of the trachea is visualized using a laryngoscope and a bent gavage needle is then inserted directly into the trachea to deliver the 
innoculum. We also describe procedures for harvesting and processing of LDLNs and lungs for analysis of antigen trafficking by flow cytometry.

We illustrate non-surgical delivery of test materials into the lungs of anesthetized mice via the trachea. This method permits lung exposure to bacterial and viral pathogens, cytokines, antibodies, beads, chemicals, or dyes. We further describe harvesting and processing of lungs and lung draining lymph nodes (LDLNs) for flow cytometry.

Phagocytic cells such as alveolar macrophages and lung dendritic cells (LDCs) continuously sample antigens from the alveolar spaces in the 
lungs. LDCs, ...

Noninvasive Intratracheal Intubation to Study the Pathology and Physiology of Mouse Lung

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. With the increasing availability of knockout and transgenic derivatives, together with a vast amount of genetic information, mice provide one of the best models to study the molecular mechanisms underlying the pathology and physiology of lung diseases. Inhalation, intranasal instillation, intratracheal instillation, and intratracheal intubation are the most widely used techniques by a number of investigators to administer materials of interest to mouse lungs. There are pros and cons for each technique depending on the goals of a study. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented. This technique was applied to administer bleomycin to mouse lungs as a model to study pulmonary fibrosis.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular ...

Medicine

Sublingual Immunotherapy as an Alternative to Induce Protection Against Acute Respiratory Infections

Sublingual route has been widely used to deliver small molecules into the bloodstream and to modulate the immune response at different sites. It has been shown to effectively induce humoral and cellular responses at systemic and mucosal sites, namely the lungs and urogenital tract. Sublingual vaccination can promote protection against infections at the lower and upper respiratory tract; it can also promote tolerance to allergens and ameliorate asthma symptoms. Modulation of lung&#8217;s immune response by sublingual immunotherapy (SLIT) is safer than direct administration of formulations by intranasal route because it does not require delivery of potentially harmful molecules directly into the airways. In contrast to intranasal delivery, side effects involving brain toxicity or facial paralysis are not promoted by SLIT. The immune mechanisms underlying SLIT remain elusive and its use for the treatment of acute lung infections has not yet been explored. Thus, development of appropriate animal models of SLIT is needed to further explore its potential advantages. 
This work shows how to perform sublingual administration of therapeutic agents in mice to evaluate their ability to protect against acute pneumococcal pneumonia. Technical aspects of mouse handling during sublingual inoculation, precise identification of sublingual mucosa, draining lymph nodes and isolation of tissues, bronchoalveolar lavage and lungs are illustrated. Protocols for single cell suspension preparation for FACS analysis are described in detail. Other downstream applications for the analysis of the immune response are discussed. Technical aspects of the preparation of Streptococcus pneumoniae inoculum and intranasal challenge of mice are also explained.
SLIT is a simple technique that allows screening of candidate molecules to modulate lungs&#8217; immune response. Parameters affecting the success of SLIT are related to molecular size, susceptibility to degradation and stability of highly concentrated formulations.

The present work illustrates the convenience of using sublingual immunotherapy to boost the innate immune response in the lungs and confer protection against acute pneumococcal pneumonia in mouse.

Sublingual route has been widely used to deliver small molecules into the bloodstream and to modulate the immune response at different sites. It has been ...

Intubation-mediated Intratracheal (IMIT) Instillation: A Noninvasive, Lung-specific Delivery System

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences between the murine and human respiratory systems, especially in their upper respiratory tracts (URT). In some models, these differences in the murine nasal cavity can have a significant impact on disease progression and presentation in the lower respiratory tract (LRT) when using intranasal instillation techniques, potentially limiting the usefulness of the mouse model to study these diseases. For these reasons, it would be advantageous to develop a technique to instill bacteria directly into the mouse lungs in order to study LRT disease in the absence of involvement of the URT. We have termed this lung specific delivery technique intubation-mediated intratracheal (IMIT) instillation. This noninvasive technique minimizes the potential for instillation into the bloodstream, which can occur during more invasive traditional surgical intratracheal infection approaches, and limits the possibility of incidental digestive tract delivery. IMIT is a two-step process in which mice are first intubated, with an intermediate step to ensure correct catheter placement into the trachea, followed by insertion of a blunt needle into the catheter to mediate direct delivery of bacteria into the lung. This approach facilitates a &#62;98% efficacy of delivery into the lungs with excellent distribution of reagent throughout the lung. Thus, IMIT represents a novel approach to study LRT disease and therapeutic delivery directly into the lung, improving upon the ability to use mice as surrogates to study human respiratory disease. Furthermore, the accuracy and reproducibility of this delivery system also makes it amenable to Good Laboratory Practice Standards (GLPS), as well as delivery of a wide range of reagents which require high efficiency delivery to the lung.

Intubation-mediated Intratracheal IMIT Instillation: A Noninvasive, Lung-specific Delivery System

Intubation-mediated intratracheal (IMIT) instillation of reagents is an excellent, noninvasive method for studying respiratory disease, as well as a method for instilling therapeutic reagents directly into the lung. It is a rapid and highly reproducible method which is suitable for preclinical testing.

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences ...

Intra-tracheal Administration of Haemophilus influenzae in Mouse Models to Study Airway Inflammation

Here, we describe a detailed procedure to efficiently and directly deliver Haemophilus influenzae into the lower respiratory tracts of mice. We demonstrate the procedure for preparing H. influenzae inoculum, intra-tracheal instillation of H. influenzae into the lung, collection of broncho-alveolar lavage fluid (BALF), analysis of immune cells in the BALF, and RNA isolation for differential gene expression analysis. This procedure can be used to study the lung inflammatory response to any bacteria, virus or fungi. Direct tracheal instillation is mostly preferred over intranasal or aerosol inhalation procedures because it more efficiently delivers the bacterial inoculum into the lower respiratory tract with less ambiguity.

Intra-tracheal Administration of Haemophilus influenzae in Mouse Models to Study Airway Inflammation

We demonstrate the procedure for intra-tracheal inoculation of Haemophilus influenzae into the lower respiratory tracts of mice. This is a very useful tool to study signaling pathways that regulate airway inflammation in mouse models.

Here, we describe a detailed procedure to efficiently and directly deliver Haemophilus influenzae into the lower respiratory tracts of mice. We ...

Method for the Isolation of Francisella tularensis Outer Membranes

Francisella tularensis is a Gram-negative intracellular coccobacillus and the causative agent of the zoonotic disease tularemia. When compared with other bacterial pathogens, the extremely low infectious dose (&lt;10 CFU), rapid disease progression, and high morbidity and mortality rates suggest that the virulent strains of Francisella encode for novel virulence factors. Surface-exposed molecules, namely outer membrane proteins (OMPs), have been shown to promote bacterial host cell binding, entry, intracellular survival, virulence and immune evasion. The relevance for studying OMPs is further underscored by the fact that they can serve as protective vaccines against a number of bacterial diseases. Whereas OMPs can be extracted from gram-negative bacteria through bulk membrane extraction techniques, including sonication of cells followed by centrifugation and/or detergent extraction, these preparations are often contaminated with periplasmic and/or cytoplasmic (inner) membrane (IM) contaminants. For years, the "gold standard" method for the biochemical and biophysical separation of gram-negative IM and outer membranes (OM) has been to subject bacteria to spheroplasting and osmotic lysis, followed by sucrose density gradient centrifugation. Once layered on a sucrose gradient, OMs can be separated from IMs based on the differences in buoyant densities, believed to be predicated largely on the presence of lipopolysaccharide (LPS) in the OM. Here, we describe a rigorous and optimized method to extract, enrich, and isolate F. tularensis outer membranes and their associated OMPs.

Method for the Isolation of Francisella tularensis Outer Membranes

A protocol for separating inner and outer membranes from Francisella tularensis by spheroplasting, osmotic lysis, and sucrose density gradient ultracentrifugation.

Francisella tularensis is a Gram-negative intracellular coccobacillus and the causative agent of the zoonotic disease tularemia. When compared with other ...

Biology

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, ...

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

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

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

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

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

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 ...

Intratracheal Inoculation of Fischer 344 Rats with Francisella tularensis

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