This work was funded by departmental support from the Department of Anesthesia and Perioperative Care, University of California San Francisco and San Francisco General Hospital, as well as by an NIH R01 award (to AP): 1R01HL146753.

All procedures and steps described below were approved by the institutional animal care and use committee (IACUC) at the University of California San Francisco. Any mouse strain can be used, though some strains have a more robust lung IR inflammatory response compared to others12. Mice that are approximately 12-15 weeks of age (30-40 g) or older tolerate and survive the lung IR surgery better than younger mice. Both male and female mice can be used for these surgeries.
1. Mouse Intubation Protocol
<ol>
	<li>Anesthesia and preparation for intubation
	<ol>
		<li>Wipe the mouse abdomen with an ethanol swab. Anesthetize the mouse with an intraperitoneal injection of tribromoethanol (250-400 mg/kg). Assess the appropriate depth of anesthesia by the lack of pedal withdrawal reflex. Place eye lubricating ointment now or later (step 2.1.4). 
		NOTE: For this procedure, tribromoethanol (and etomidate as an alternative option) provides a stable anesthetic plane without affecting the hemodynamic conditions required for this surgery. This anesthetic is only used once to avoid the risk of peritoneal adhesions. Isoflurane could also be used, but we do not use it here. The practitioner is free to use whatever anesthetic recipe they see fit.</li>
		<li>Place the anesthetized mouse on an intubation stand or plastic support in a supine position, suspended by its upper incisors on looped 4-0 sutures (silk or other) across two support anchors.</li>
		<li>To keep the mouse immobilized during the intubation procedure, loosely tape the lower part of the chest (or both upper limbs) to the platform.</li>
		<li>Place the fiberoptic flexible light gently on the trachea of the mouse, slightly below the vocal cords. Adjust the level of illumination so that only a dark field is visible when looking into the mouse oropharynx except for red light emanating from below the vocal cords, demonstrating the target for the eventual placement of the endotracheal tube. Note that vocal cord movements should be visible with the naked eye or, if needed, under magnification.</li>
	</ol>
	</li>
	<li>Intubation procedure
	<ol>
		<li>Hold the tweezers with the dominant hand and use them to gently grip and draw the tongue out of the oral cavity.</li>
		<li>Open the lower jaw using forceps held by the non-dominant hand, and then push the forceps into the larynx to gently lift the epiglottis. At this time, release the tongue from the tweezers.</li>
		<li>Look for the vocal cords. They should open and close according to each breath. Holding the cannula with the guide wire pre-loaded, insert the tip of the wire through the vocal cords.</li>
		<li>Being very careful not to move the wire by holding a portion of it that is outside the cannula but just above the vocal cords, withdraw the cannula, leaving just the wire in place with its distal end within the trachea.</li>
		<li>At this point, perform a second visualization of the vocal cords to confirm that the wire distal tip remains passed through the illuminated vocal cords and into the trachea, and is not in the unlit esophagus.</li>
		<li>Hold the wire outside the mouth with the curved forceps in the left hand, stabilized against a hard surface, and carefully advance the 20G catheter with tape wings over the wire.</li>
		<li>Once the distal end of the wire emerges from the back end of the 20G catheter or endotracheal tube, hold that end with the curved forceps and smoothly advance the 20G catheter into the trachea.</li>
		<li>Carefully remove the wire from the distal end of the 20G catheter with the curved forceps without dislodging the placement of the catheter.</li>
		<li>Briefly connect the catheter to the ventilator before securing it to confirm proper placement into the trachea and not the esophagus. Confirm tracheal placement by observation of mechanical ventilation-dependent bilateral chest wall movements and the absence of inflation of the stomach.</li>
	</ol>
	</li>
	<li>Post-intubation
	<ol>
		<li>Disconnect the catheter from the ventilator. Fix the tape wings (attached to the catheter) through the lower lip of the mouse using a 4-0 vicryl suture to firmly secure the endotracheal tube (ETT) to the mouse during all subsequent procedures/manipulations. 
		NOTE: Alternatively, silk tape or other tape can be used to secure the ETT, however care should be taken to avoid dislodgement of the ETT during movement of the animal from the intubation sled to the surgical surface.</li>
		<li>Carefully remove the mouse from the intubation sled. Briefly connect the catheter to the ventilator set at a tidal volume 0.2-0.225 mL and a respiratory rate of 120-150 breaths per min to confirm correct tracheal placement of the orotracheal tube and then disconnect with the mouse breathing spontaneously through the orotracheal tube.</li>
		<li>Do not leave the animal unattended from this point onward until it has regained sufficient consciousness to maintain sternal recumbency at the end of the procedure.</li>
	</ol>
	</li>
</ol>
2. Lung ischemia and reperfusion (IR) surgery protocol
<ol>
	<li>Analgesia and preparation of the surgical site
	<ol>
		<li>Wipe the mouse abdomen with an ethanol swab and inject buprenorphine (0.05-0.1 mg/kg) intraperitoneally.</li>
		<li>Shave the hair over the left thorax area up to the left scapula. Remove excess shaved hair using alcohol swabs. 
		NOTE: Steps 2.1.1 and 2.1.2 can also be performed before intubation if there concern for dislodgement of the ETT when secured with silk tape.</li>
		<li>Place the mouse on a warming pad in a left lateral or 3/4 turned position and connect the tracheal tube on the ventilator with a tidal volume of 0.2-0.225 mL (~8 mg/kg) and a respiratory rate of 120-150 breaths per min. Do not use supplemental oxygen for this procedure.</li>
		<li>Apply eye lubricant with a sterile cotton-tip swab. Turn the mouse to 3/4 left side up and immobilize all four limbs and the tail with laboratory tape.</li>
		<li>Disinfect the shaved skin area and surrounding fur with povidone-iodine and wait for the solution to dry. Then cover the surgical field with a sterile drape or clear plastic film and create a rectangular opening in the drape or plastic film for the surgical field.</li>
	</ol>
	</li>
	<li>Surgical procedure
	<ol>
		<li>Confirm the appropriate level of anesthesia (provided by the administration of tribromoethanol and buprenorphine as described earlier) by testing response to toe pinch.</li>
		<li>Using a pair of sharp scissors and a pair of larger forceps (narrow pattern forceps or similar), make a 2 cm transverse skin incision below the inferior angle of the scapula in the left lateral thorax. Use the scissors and a finer pair of forceps (extra fine graefe forceps or similar) to cut into the muscular layer and dissect down to the ribs.</li>
		<li>Identify the second intercostal space and hold the second rib with the extra fine forceps. Pulling the rib upward, use a sterile #11 or #12 (curved) scalpel blade (no handle necessary) to enter the pleural space by separating and cutting across the 2nd-3rd space&#39;s intercostal muscles. Consider pausing ventilation to reduce injury to the left lung apex.</li>
		<li>Insert three sterilized retractors. Use the smallest/narrowest retractor cephalad along the orientation of the ribs, the medium size retractor to the left along the 2nd rib, and the largest retractor to the right along the surface of the 3rd rib.</li>
		<li>Open the chest with slow and progressive retraction using the elastic retractor cords. Expose and identify the left pulmonary artery (PA) by moving the left lung apex away with a sterile cotton-tip swab.</li>
		<li>Use the micro forceps, ultrafine forceps in the right hand and PA or vessel dilating forceps in the left hand, to gently expose and create the field in which the left PA and bronchus are both visible.</li>
		<li>Using the PA forceps, pick up the left PA and pull gently but firmly upward and cephalad to visualize the transparent bronchus below. Increase magnification on the dissection microscope (see equipment list for more details) at this point to maximum (2x). 
		NOTE:&#160;Sterilize all equipment before use. Additionally, to maintain sterility, only the tips of surgical instruments should enter the sterile surgical field.</li>
		<li>While retracting the PA away from the bronchus, carefully pass the closed ultrafine forceps through the space between the left PA and bronchus. Then, use these forceps to hold and pull a 7-0 or 8-0 prolene suture through the space between the left pulmonary artery (above) and bronchus (below).</li>
		<li>Encircle the left PA by tying a slipknot to create an occlusion in the PA. Blood flow interruption is easily visualized under the microscope. This marks the initiation of the ischemic period.</li>
		<li>Externalize the free end of the knot through a different entry point in the anterior left thorax using a 24G-28G needle and secure the end of the suture with a small piece of tape for easier identification later on.</li>
		<li>Reinflate the lung to expel as much air out of the chest cavity as possible using a PEEP valve/tubing on the rodent ventilator. Then, close the ribcage with two interrupted 4-0 nylon sutures.</li>
		<li>Close the muscle and subcutaneous layer with a running 4-0 nylon suture. Then apply two or three drops of topical bupivacaine (0.5%) to the incision. Use a 4-0 nylon suture to close the skin layer with a running suture.</li>
	</ol>
	</li>
	<li>Post-operative care
	<ol>
		<li>When spontaneous ventilation has resumed, disconnect the endotracheal tube from the ventilator and extubate the mouse.</li>
		<li>Place the mouse on the warming pad to maintain body temperature during early post-anesthesia recovery.</li>
		<li>Carefully monitor the mouse while recovering from general anesthesia. Pull the externalized slipknot gently at the end of the ischemic period (30 min or 1 h).</li>
		<li>Move the mouse from the warming pad to a cage once it has exhibited signs of recovery: self-righting and/or movement.</li>
		<li>After the period of reperfusion (1 h or 3 h), euthanize the animal and collect blood by cardiac puncture and lung tissue for further analysis. For 1 h reperfusion, collect plasma for ELISA, tissue for RNA, and protein analysis; for 3 h reperfusion, additionally collect tissue for histology.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

This manuscript details the steps involved in performing the ventilated lung IR model developed by Dodd-o et al.9. This model has helped identify molecular pathways involved in the generation and resolution of inflammation from lung IR in isolation14,15,16,17, lung IR in combination with co-existing infection18, and lung IR in relation to the gut-...

Ischemia reperfusion (IR) injury can occur when blood flow is restored to an organ or tissue bed after some period of interruption. In the lung, IR can occur in isolation or in association with other injurious processes such as infection, hypoxia, atelectasis, volutrauma (from high tidal volumes during mechanical ventilation), barotrauma (high peak or sustained pressures during mechanical ventilation), or blunt (non-penetrating) lung contusion injury1,2,3. There remain several gaps in our knowledge about the mechanisms of LIRI and the impact of concurrent processes (e.g., infection) on LIRI outcomes, and also the treatment options for LIRI are limited. An in vivo model of pure LIRI is required to identify the pathophysiology of lung IR injury in isolation and to study its contribution to any multi-hit process of which lung injury is a component.
Murine lung IR models can be used to study the lung-specific pathophysiology of multiple processes, including lung transplantation3, pulmonary embolism4, and lung injury following hemorrhagic trauma with resuscitation5. Currently used models include surgical lung transplantation6, hilar clamping7, ex vivo lung perfusion8, and ventilated lung IR9. Here, we provide a detailed protocol for a murine ventilated lung IR model of sterile lung injury. There are multiple benefits of this approach (Figure 2), including that it induces minimal hypoxia and minimal atelectasis, and it is a survival surgery model that allows for long-term studies.
Reasons to choose this model of LIRI over other models such as the hilar clamping and ex vivo perfusion models are the following: this model minimizes the inflammatory contributions of atelectasis, mechanical ventilation, and hypoxia; it preserves cyclical ventilation; it maintains an intact in vivo circulatory immune system that can respond to the IR injury; and finally, as a survival procedure, it permits the longer-term analysis of the mechanisms of secondary injury generation (2-hit models) and injury resolution. Overall, we believe this ventilated lung IR model provides the &#34;purest&#34; form of IR injury that can be studied experimentally.
Other publications have described the use of orotracheal intubation of mice to perform IT injections or installations10,11, but not as the starting point for a survival surgery as it is in this model. The placement of an orotracheal tube permits the performance of lung surgery by allowing the collapse of the operative lung. It also allows for the reinflation of the lung at the end of the procedure, which is critical for the pneumothorax and for the ability of the mouse to return to spontaneous ventilation at the conclusion of the procedures. Finally, the removal of the secured orotracheal tube is a simple procedure that, unlike an invasive tracheotomy, is compatible with a survival surgery. This allows for longer term research studies focused on understanding the progression and resolution of LIRI and associated disorders, as well as the creation of chronic injury models.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fiber Optic Light Pipe</td>
			<td>Cole-Parmer</td>
			<td>UX-41720-65</td>
			<td>Fiberoptic light pipe</td>
		</tr>
		<tr>
			<td>Fiber Optic Light Source</td>
			<td>AmScope</td>
			<td>SKU: CL-HL250-B</td>
			<td>Light source for fiberoptic lights</td>
		</tr>
		<tr>
			<td>Germinator 500</td>
			<td>Cell Point Scientific, Inc.</td>
			<td>No.5-1450</td>
			<td>Bead Sterilizer</td>
		</tr>
		<tr>
			<td>Heating Pad</td>
			<td>AIMS</td>
			<td>14-370-223</td>
			<td>Alternative option</td>
		</tr>
		<tr>
			<td>Lithium.Ion Grooming Kits(hair clipper)</td>
			<td>WAHL home products</td>
			<td>SKU 09854-600B</td>
			<td>To remove mouse hair on surgical site</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon</td>
			<td>SMZ-10</td>
			<td>Other newer options available at the company website</td>
		</tr>
		<tr>
			<td>MiniVent Ventilator</td>
			<td>Havard Apparatus</td>
			<td>Model 845</td>
			<td>Mouse ventilator</td>
		</tr>
		<tr>
			<td>Ultrasonic Cleaner</td>
			<td>Cole-Parmer</td>
			<td>UX-08895-05</td>
			<td>Clean tools that been used in operation</td>
		</tr>
		<tr>
			<td>Warming Pad</td>
			<td>Kent Scientific</td>
			<td>RT-0501</td>
			<td>To keep mouse warm while recovering from surgery</td>
		</tr>
		<tr>
			<td>Weighing Scale</td>
			<td>Cole-Parmer</td>
			<td>UX-11003-41</td>
			<td>Weighing scale</td>
		</tr>
		<tr>
			<td>Surgery Tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4-0 Silk Suture</td>
			<td>Ethicon</td>
			<td>683G</td>
			<td>For closing muscle layer</td>
		</tr>
		<tr>
			<td>7-0 Prolene Suture</td>
			<td>Ethicon Industry</td>
			<td>EP8734H</td>
			<td>Using for making a slip knot of left pulmonary artery</td>
		</tr>
		<tr>
			<td>Bard-Parker (11) Scalpel (Rib-Back Carbon Steel Surgical Blade, sterile, single use)</td>
			<td>Aspen Surgical</td>
			<td>372611</td>
			<td>For entering thoracic cavity (option 1)</td>
		</tr>
		<tr>
			<td>Bard-Parker (12) Scalpel</td>
			<td>Aspen Surgical</td>
			<td>372612</td>
			<td>For entering thoracic cavity (option 2)</td>
		</tr>
		<tr>
			<td>Extra Fine Graefe Forceps</td>
			<td>FST</td>
			<td>11150-10</td>
			<td>Muscle/rib holding forceps</td>
		</tr>
		<tr>
			<td>Magnetic Fixator Retraction System</td>
			<td>FST</td>
			<td>1. Base Plate (Nos. 18200-03) 
			2. Fixators (Nos. 18200-01) 
			3. Retractors (Nos. 18200-05 through 18200-12) 
			4. Elastomer (Nos.18200-07) 5. Retractor(No.18200-08)</td>
			<td>Small Animal Retraction System</td>
		</tr>
		<tr>
			<td>Monoject Standard Hypodermic Needle</td>
			<td>COVIDIEN</td>
			<td>05-561-20</td>
			<td>For medication delivery IP</td>
		</tr>
		<tr>
			<td>Narrow Pattern Forceps</td>
			<td>FST</td>
			<td>11002-12</td>
			<td>Skin level forceps</td>
		</tr>
		<tr>
			<td>Needle holder/Needle driver</td>
			<td>FST</td>
			<td>12565-14</td>
			<td>for holding needles</td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>BD</td>
			<td>305110</td>
			<td>26 gauge needle for externalizing slipknot (24 or 26 gauge needle okay too)</td>
		</tr>
		<tr>
			<td>PA/Vessel Dilating forceps</td>
			<td>FST</td>
			<td>00125-11</td>
			<td>To hold PA; non-damaging gripper</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>14060-09</td>
			<td>Used for incision and cutting into the muscular layer durging surgery</td>
		</tr>
		<tr>
			<td>Ultra Fine Dumont micro forceps</td>
			<td>FST</td>
			<td>11295-10 (Dumont #5 forceps, Biology tip, tip dimension:0.05*0.02mm,11cm)</td>
			<td>For passing through the space between the left pulmonary artery and bronchus</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.25% Bupivacaine</td>
			<td>Hospira, Inc.</td>
			<td>0409-1159-02</td>
			<td>Topical analgesic used during surgical wound closure</td>
		</tr>
		<tr>
			<td>Avertin (2,2,2-Tribromoethanol)</td>
			<td>Sigma-Aldrich</td>
			<td>T48402-25G</td>
			<td>Anesthetic, using for anesthetize the mouse for IR surgery, the concentration used in IR surgery is 250-400 mg/kg.</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Covetrus North America</td>
			<td>59122</td>
			<td>Analgesic: concentration used for surgery is 0.05-0.1 mg/kg</td>
		</tr>
		<tr>
			<td>Eye Lubricant</td>
			<td>BAUSCH+LOMB</td>
			<td>Soothe Lubricant Eye Ointment</td>
			<td>Relieves dryness of the eye</td>
		</tr>
		<tr>
			<td>Povidone-Iodine 10% Solution</td>
			<td>MEDLINE INDUSTRIES INC</td>
			<td>SKU MDS093944H (2 FL OZ, topical antiseptic)</td>
			<td>Topical liquid applied for an effective first aid antiseptic at beginning of surgery</td>
		</tr>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol Swab</td>
			<td>BD brand</td>
			<td>&#160;BD 326895</td>
			<td>for sterilzing area of injection and surgery</td>
		</tr>
		<tr>
			<td>Plastic film</td>
			<td>KIRKLAND</td>
			<td>Stretch-Tite premium</td>
			<td>Alternative for covering the sterilized surgical field (more cost effective)</td>
		</tr>
		<tr>
			<td>Rodent Surgical Drapes</td>
			<td>Stoelting</td>
			<td>50981</td>
			<td>Sterile field or drape for surgical field</td>
		</tr>
		<tr>
			<td>Sterile Cotton Tipped Application</td>
			<td>Pwi-Wnaps</td>
			<td>703033</td>
			<td>used for applying eye lubricant</td>
		</tr>
		<tr>
			<td>Top Sponges</td>
			<td>Dukal Corporaton</td>
			<td>Reorder # 5360</td>
			<td>Stopping bleeding from skin/muscle</td>
		</tr>
	</tbody>
</table>

a mouse model of orotracheal intubation and ventilated lung ischemia reperfusion surgery

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

Inflammation generated by unilateral ventilated sterile lung ischemia reperfusion (IR) injury: Following 1 h of ischemia, we observed increased levels of cytokines in the serum and within the lung tissue by both ELISA and qRT-PCR that peaked at 1 h following reperfusion and rapidly returned to baseline within 12-24 h after reperfusion13. For samples collected at 3 h following reperfusion, we observed intense neutrophil infiltration within the left lung tissue and noted that the intensity of the in...

Watch this Scientific Journal Video about A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery at JoVE.com

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR ...

a-mouse-model-orotracheal-intubation-ventilated-lung-ischemia

Research

JoVE Journal

Immunology and Infection

3.5K Views.  University of California San Francisco and San Francisco General Hospital. A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Video: A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Isolation of Mouse Lung Dendritic Cells

Lung dendritic cells (DC) play a fundamental role in sensing invading pathogens 1,2 as well as in the control of tolerogenic responses 3 in the respiratory tract. At least three main subsets of lung dendritic cells have been described in mice: conventional DC (cDC) 4, plasmacytoid DC (pDC) 5 and the IFN-producing killer DC (IKDC) 6,7. The cDC subset is the most prominent DC subset in the lung 8. 
The common marker known to identify DC subsets is CD11c, a type I transmembrane integrin (&beta;2) that is also expressed on monocytes, macrophages, neutrophils and some B cells 9. In some tissues, using CD11c as a marker to identify mouse DC is valid, as in spleen, where most CD11c+ cells represent the cDC subset which expresses high levels of the major histocompatibility complex class II (MHC-II). However, the lung is a more heterogeneous tissue where beside DC subsets, there is a high percentage of a distinct cell population that expresses high levels of CD11c bout low levels of MHC-II. Based on its characterization and mostly on its expression of F4&#47;80, an splenic macrophage marker, the CD11chiMHC-IIlo lung cell population has been identified as pulmonary macrophages 10 and more recently, as a potential DC precursor 11. 
In contrast to mouse pDC, the study of the specific role of cDC in the pulmonary immune response has been limited due to the lack of a specific marker that could help in the isolation of these cells. Therefore, in this work, we describe a procedure to isolate highly purified mouse lung cDC. The isolation of pulmonary DC subsets represents a very useful tool to gain insights into the function of these cells in response to respiratory pathogens as well as environmental factors that can trigger the host immune response in the lung.

A highly purified preparation of mouse lung dendritic cells is described. Specific emphasis is given to the isolation of conventional dendritic cell subset.

Lung dendritic cells (DC) play a fundamental role in sensing invading pathogens 1,2 as well as in the control of tolerogenic responses 3 in the ...

Induction of Graft-versus-host Disease and In Vivo T Cell Monitoring Using an MHC-matched Murine Model

Graft-versus-host disease (GVHD) is the limiting barrier to the broad use of bone marrow transplant as a curative therapy for a variety of hematological deficiencies. GVHD is caused by mature alloreactive T cells present in the bone marrow graft that are infused into the recipient and cause damage to host organs. However, in mice, T cells must be added to the bone marrow inoculum to cause GVHD. Although extensive work has been done to characterize T cell responses post transplant, bioluminescent imaging technology is a non-invasive method to monitor T cell trafficking patterns in vivo. 
Following lethal irradiation, recipient mice are transplanted with bone marrow cells and splenocytes from donor mice. T cell subsets from L2G85.B6 (transgenic mice that constitutively express luciferase) are included in the transplant. By only transplanting certain T cell subsets, one is able to track specific T cell subsets in vivo, and based on their location, develop hypotheses regarding the role of specific T cell subsets in promoting GVHD at various time points. At predetermined intervals post transplant, recipient mice are imaged using a Xenogen IVIS CCD camera. Light intensity can be quantified using Living Image software to generate a pseudo-color image based on photon intensity (red = high intensity, violet = low intensity). 
Between 4-7 days post transplant, recipient mice begin to show clinical signs of GVHD. Cooke et al.1 developed a scoring system to quantitate disease progression based on the recipient mice fur texture, skin integrity, activity, weight loss, and posture. Mice are scored daily, and euthanized when they become moribund. Recipient mice generally become moribund 20-30 days post transplant. 
Murine models are valuable tools for studying the immunology of GVHD. Selectively transplanting particular T cell subsets allows for careful identification of the roles each subset plays. Non-invasively tracking T cell responses in vivo adds another layer of value to murine GVHD models.

Induction of Graft-versus-host Disease and In Vivo T Cell Monitoring Using an MHC-matched Murine Model

Murine bone marrow transplantation is a widely used technique to study immunological mechanisms governing graft-versus-host disease in humans. The ability to monitor T cell trafficking patterns in vivo allows for detailed analysis of the development and perpetuation of T cell responses during graft-versus-host disease.

Graft-versus-host disease (GVHD) is the limiting barrier to the broad use of bone marrow transplant as a curative therapy for a variety of hematological ...

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. ...

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and pneumothorax from pleural disruption can result in a significant mortality rate. This protocol is a simple method of rat thymectomy that utilizes a mini-sternotomy and endotracheal intubation. Intubation is accomplished with a non-invasive and easily reproducible method and allows for positive pressure ventilation to prevent pneumothorax and a controlled airway that allows sufficient time for careful thymus dissection to minimize pleural disruption. A 1.5 cm sternal incision decreases contact with mediastinal vessels and pleura, while still providing full visualization of the thymus. Following exposure of the mediastinum, the thymus is removed by blunt dissection under magnification. The pleural space is then sealed by suture closure of the pre-tracheal muscles followed by the application of surgical glue. The thorax is then closed by suture closure of the sternum, followed by suture closure of the skin. All thymectomies were complete as evidenced by immunohistochemical (IHC) staining of mediastinal tissue, and absence of na&#239;ve T-cells by flow cytometry, and the procedure had a 96% survival rate. This method is suitable when complete thymectomy with minimal complications is desired for further immunological studies in athymic adult rats.

Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described.

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and ...

A 3D Human Lung Tissue Model for Functional Studies on Mycobacterium tuberculosis Infection

Tuberculosis (TB) still holds a major threat to the health of people worldwide, and there is a need for cost-efficient but reliable models to help us understand the disease mechanisms and advance the discoveries of new treatment options. In vitro cell cultures of monolayers or co-cultures lack the three-dimensional (3D) environment and tissue responses. Herein, we describe an innovative in vitro model of a human lung tissue, which holds promise to be an effective tool for studying the complex events that occur during infection with Mycobacterium tuberculosis (M. tuberculosis). The 3D tissue model consists of tissue-specific epithelial cells and fibroblasts, which are cultured in a matrix of collagen on top of a porous membrane. Upon air exposure, the epithelial cells stratify and secrete mucus at the apical side. By introducing human primary macrophages infected with M. tuberculosis to the tissue model, we have shown that immune cells migrate into the infected-tissue and form early stages of TB granuloma. These structures recapitulate the distinct feature of human TB, the granuloma, which is fundamentally different or not commonly observed in widely used experimental animal models. This organotypic culture method enables the 3D visualization and robust quantitative analysis that provides pivotal information on spatial and temporal features of host cell-pathogen interactions. Taken together, the lung tissue model provides a physiologically relevant tissue micro-environment for studies on TB. Thus, the lung tissue model has potential implications for both basic mechanistic and applied studies. Importantly, the model allows addition or manipulation of individual cell types, which thereby widens its use for modelling a variety of infectious diseases that affect the lungs.

A 3D Human Lung Tissue Model for Functional Studies on Mycobacterium tuberculosis Infection

Human tuberculosis infection is a complex process, which is difficult to model in vitro. Here we describe a novel 3D human lung tissue model that recapitulates the dynamics that occur during infection, including the migration of immune cells and early granuloma formation in a physiological environment.

Tuberculosis (TB) still holds a major threat to the health of people worldwide, and there is a need for cost-efficient but reliable models to help us ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Influenza A Virus Studies in a Mouse Model of Infection

Influenza viruses cause over 500,000 deaths worldwide1 and are associated with an annual cost of 12 - 14 billion USD in the United States alone considering direct medical and hospitalization expenses and work absenteeism2. Animal models are crucial in Influenza A virus (IAV) studies to evaluate viral pathogenesis, host-pathogen interactions, immune responses, and the efficacy of current and/or novel vaccine approaches as well as antivirals. Mice are an advantageous small animal model because their immune system is evolutionarily similar to that found in humans, they are available from commercial vendors as genetically identical subjects, there are multiple strains that can be exploited to evaluate the genetic basis of infections, and they are relatively inexpensive and easy to manipulate. To recapitulate IAV infection in humans via the airways, mice are first anesthetized prior to intranasal inoculation with infectious IAVs under proper biosafety containment. After infection, the pathogenesis of IAVs is determined by monitoring daily the morbidity (body weight loss) and mortality (survival) rate. In addition, viral pathogenesis can also be evaluated by assessing virus replication in the upper (nasal mucosa) or lower (lungs) respiratory tract of infected mice. Humoral responses upon IAV infection can be rapidly evaluated by non-invasive bleeding and secondary antibody detection assays aimed at detecting the presence of total or neutralizing antibodies. Here, we describe the common methods used to infect mice intranasally (i.n) with IAV and evaluate pathogenesis, humoral immune responses and protection efficacy.

Influenza A viruses (IAVs) are important human respiratory pathogens. To understand the pathogenicity of IAVs and to perform preclinical testing of novel vaccine approaches, animal models mimicking human physiology are required. Here, we describe techniques to evaluate IAV pathogenesis, humoral responses and vaccine efficacy using a mouse model of infection.

Influenza viruses cause over 500,000 deaths worldwide1 and are associated with an annual cost of 12 - 14 billion USD in the United States alone ...

Assessment of the Cytotoxic and Immunomodulatory Effects of Substances in Human Precision-cut Lung Slices

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new therapeutics. Additionally, registration of new substances requires appropriate risk assessment with adequate testing systems to avoid the risk of individuals being harmed, for example, in the working environment. Such risk assessments are usually conducted in animal studies. In view of the 3Rs principle and public skepticism against animal experiments, human alternative methods, such as precision-cut lung slices (PCLS), have been evolving. The present paper describes the ex vivo technique of human PCLS to study the immunomodulatory potential of low-molecular-weight substances, such as ammonium hexachloroplatinate (HClPt). Measured endpoints include viability and local respiratory inflammation, marked by altered secretion of cytokines and chemokines. Pro-inflammatory cytokines, tumor necrosis factor alpha (TNF-&#945;), and interleukin 1 alpha (IL-1&#945;) were significantly increased in human PCLS after exposure to a sub-toxic concentration of HClPt. Even though the technique of PCLS has been substantially optimized over the past decades, its applicability for the testing of immunomodulation is still in development. Therefore, the results presented here are preliminary, even though they show the potential of human PCLS as a valuable tool in respiratory research.

In view of the 3Rs principle, respiratory models as alternatives to animal studies are evolving. Especially for risk assessment of respiratory substances, there is a lack of appropriate assays. Here, we describe the use of human precision-cut lung slices for the assessment of airborne substances.

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new ...

Oral Intubation of Adult Zebrafish: A Model for Evaluating Intestinal Uptake of Bioactive Compounds

Most pathogens invade organisms through their mucosa. This is particularly true in fish as they are continuously exposed to a microbial-rich water environment. Developing effective methods for oral delivery of immunostimulants or vaccines, which activate the immune system against infectious diseases, is highly desirable. In devising prophylactic tools, good experimental models are needed to test their performance. Here, we show a method for oral intubation of adult zebrafish and a set of procedures to dissect and prepare the intestine for cytometry, confocal microscopy and quantitative polymerase chain reaction (qPCR) analysis. With this protocol, we can precisely administer volumes up to 50 &#181;L to fish weighing approximately 1 g simply and quickly, without harming the animals. This method allows us to explore the direct in vivo uptake of fluorescently labelled compounds by the intestinal mucosa and the immunomodulatory capacity of such biologics at the local site after intubation. By combining downstream methods such as flow cytometry, histology, qPCR and confocal microscopy of the intestinal tissue, we can understand how immunostimulants or vaccines are able to cross the intestinal mucosal barriers, pass through the lamina propria, and reach the muscle, exerting an effect on the intestinal mucosal immune system. The model could be used to test candidate oral prophylactics and delivery systems or the local effect of any orally administered bioactive compound.

The protocol describes intubating adult zebrafish with a biologic; then dissecting and preparing the intestine for cytometry, confocal microscopy and qPCR. This method allows administration of bioactive compounds to monitor intestinal uptake and the local immune stimulus evoked. It&#160;is relevant for testing the intestinal dynamics of oral prophylactics.

Most pathogens invade organisms through their mucosa. This is particularly true in fish as they are continuously exposed to a microbial-rich water ...

Induction of Mouse Lung Injury by Endotracheal Injection of Bleomycin

Pulmonary fibrosis is a hallmark of several human lung diseases with a different etiology. Since current therapies are rather limited, mouse models continue to be an essential tool for developing new antifibrotic strategies. Here we provide an effective method to investigate in vivo antifibrotic activity of human mesenchymal stromal cells obtained from whole umbilical cord (hUC-MSC) in attenuating bleomycin-induced lung injury. C57BL/6 mice receive a single endotracheal injection of bleomycin (1.5 U/kg body weight) followed by a double infusion of hUC-MSC (2.5 x 105) into the tail vein, 24 h and 7 days after the bleomycin administration. Upon sacrifice at days 8, 14, or 21, inflammatory and fibrotic changes, collagen content, and hUC-MSC presence in explanted lung tissue are analyzed. The injection of bleomycin into the mouse&#160;trachea allows the direct targeting of the lungs, leading to extensive pulmonary inflammation and fibrosis. The systemic administration of a double dose of hUC-MSC results in the early blunting of the bleomycin-induced lung injury. Intravenously infused hUC-MSC are transiently engrafted into the mouse lungs, where they exert their anti-inflammatory and antifibrotic activity. In conclusion, this protocol has been successfully applied for the preclinical testing of hUC-MSC in an experimental mouse model of human pulmonary fibrosis. However, this technique can be easily extended both to study the effect of different endotracheally administered substances on the pathophysiology of the lungs and to validate new anti-inflammatory and antifibrotic systemic therapies.

Here we present an effective method to investigate the antifibrotic activity of intravenously infused human mesenchymal stromal cells obtained from the whole umbilical cord following the induction of lung injury by an endotracheal injection of bleomycin in C57BL/6 mice. This protocol can be easily extended to the preclinical testing of other therapeutics.

Pulmonary fibrosis is a hallmark of several human lung diseases with a different etiology. Since current therapies are rather limited, mouse models ...