The authors would like to thank Dr. Pina Colarusso, who provided significant expertise in the editing and revising of this manuscript.

All the procedures described here were performed with prior approval by the Dalhousie University Committee on Laboratory Animals (UCLA).
1. Preparation
<ol>
	<li>Lung imaging system: To prepare the window, administer a thin layer of vacuum grease to the top of the outer ring while avoiding contamination of the vacuum channel. Place a clean 8 mm glass coverslip on the window and gently press down to create a seal.</li>
	<li>Widefield Fluorescence Microscope: Perform imaging with a conventional widefield fluorescence microscope fitted with a 20x/0.40 long working distance objective and a black and white charge-coupled device (CCD) camera with a frame-rate of 25 FPS. Apply a 530-550 nm bandpass excitation filter to excite Rhodamine-6G and a 460-490 nm bandpass filter to excite Fluorescein Isothiocyanate (FITC).</li>
	<li>Vacuum system: Connect the imaging window to a vacuum pump fitted with a digital pressure gauge capable of providing 50-60 mmHg constant suction, as shown in Supplemental Figure 1. In short, connect the imaging window to the pump through 1.0 mm I.D. polyethylene tubing, 1.0 cm I.D. polyethylene tubing, a vacuum flask, and an inline 0.2 &#181;m filter.</li>
	<li>Ventilator: Set a small rodent ventilator to provide pressure-controlled ventilation at a rate and volume calculated based upon the mouse&#39;s weight. Provide a positive end-expiratory pressure (PEEP) at 5 cmH2O for the duration of the experiment, and set the target pressure to 20 cmH2O.</li>
	<li>Anesthetic: Using a low-flow anesthesia delivery system, prime a 5.0 mL syringe with 99.9% isoflurane. Use a waste gas scavenging system to minimize the risk of inhalation by the surgeon.</li>
</ol>
2. Anesthesia
<ol>
	<li>Place a 20-25 g 12-week-old male C57Bl/6 mouse in the anesthesia induction chamber. With the chamber securely closed, begin induction with isoflurane gas at a concentration of 3% and a flow rate of 500 mL/min.</li>
	<li>Once the mouse is anesthetized (visualized by slowed respiration rate), transfer it to the intubation stand and secure the upper incisors to the hanging suture.</li>
	<li>Tighten the suture to secure the snout inside the nose cone. Begin gas flow through the nose cone at a concentration of 2.5%.</li>
	<li>Confirm adequate depth of anesthesia via toe pinch before moving to the next step.</li>
</ol>
3. Intubation
<ol>
	<li>Rotate the stand such that the back of the stand and dorsal side of the mouse face toward the surgeon.</li>
	<li>Pass the tip of a 20 cm length of fiber-optic cable through a 20 G endotracheal cannula and submerge the tip in Lidocaine HCl (1%) to facilitate the passage of the cable through the larynx.</li>
	<li>Using blunt forceps, lift the lower jaw and displace the tongue to provide clear passage into the respiratory tract.</li>
	<li>Insert a modified otoscope (~60&#176; of speculum circumference removed) such that the upper incisors fit within the gap in the speculum. Adjust the scope and tongue position until the epiglottis, and vocal cords are clearly visible.</li>
	<li>Insert the fiber-optic cable loaded with the endotracheal cannula through the gap in the speculum and into the larynx. Using small circular movements, pass the cable through the vocal cords and into the trachea.</li>
	<li>Push the cannula along the fiber-optic cable, passing between the vocal cords and into the trachea.</li>
</ol>
4. Ventilation
<ol>
	<li>Retrieve the mouse from the intubation stand and place it on a heating pad in the right lateral decubitus position.</li>
	<li>Connect the cannula to ventilator tubing and start the ventilator. Reduce the anesthetic concentration to 1.5% and monitor depth by testing for pedal reflex. If the reflex persists, increase the concentration incrementally to as a high as 2%.</li>
	<li>Place tear gel in the mouse&#39;s eyes to prevent drying.</li>
	<li>Using medical tape, secure the cannula to the snout. Using labeling tape, secure the right forepaw to the heating pad at, approximately, the 9 o&#39;clock position. Extend the left hind paw caudally and secure at, approximately, the 6 o&#39;clock position.</li>
	<li>Using a cloth tape, lightly stretch the left forepaw to the 12 o&#39;clock position and secure the other end of the tape to the top of the IVM platform as shown in Figure 2A&#160;(maintaining slight tension here facilitates subsequent thoracotomy).</li>
	<li>Insert a rectal temperature probe and secure the probe by taping it to the heating pad. Place a pulse oximeter on the right hind paw and secure to the heating pad, taking care not to disrupt circulation.</li>
	<li>Once the temperature is stable at 37.0 &#176;C &#177; 0.1 &#176;C, proceed to perform the thoracotomy.</li>
</ol>
5. Thoracotomy
<ol>
	<li>Sterilize the thorax and abdomen with a 70% alcohol wipe. Apply a light coat of mineral oil to dampen the hair on the left side of the mouse-from the sternum to the vertebral column and from the shoulder to the bottom of the ribcage.</li>
	<li>With blunt forceps and straight scissors, make a small longitudinal incision near the bottom of the ribcage to expose the underlying muscle layer.</li>
	<li>Moving ventrally, use blunt dissection to separate the epithelial and adipose tissue from the muscle layer. Cauterize any exposed blood vessels. Once the risk of blood loss has been mitigated, extend the original incision ventrally until the xiphoid process.</li>
	<li>Repeat this process dorsally until ~5 mm lateral to the vertebral column.</li>
	<li>Moving cranially, use blunt dissection to expose the rib cage. Cauterize any exposed blood vessels to preserve hemodynamic stability.</li>
	<li>Once the risk of blood loss has been mitigated, extend the incision from the xiphoid process to the axilla.</li>
	<li>Repeat this process on the dorsal side of the incision until ~1 cm inferior to the left ear.</li>
	<li>Using hemostatic forceps, grasp the dissected epithelial and adipose tissue and place clear of the surgical area (Figure 2B).</li>
	<li>Inject a solution of Rhodamine-6G (0.5 mg/mL; 1.5 mL/kg) for visualization of leukocytes and Bovine FITC-albumin (50 mg/mL; 1 mL/kg) for visualization of capillary perfusion via the tail vein.</li>
	<li>Using toothed forceps, grasp the rib immediately inferior to the position of the base of the lung at end-inspiration and slightly retract to pull the rib away from the lung. Cut the rib to induce a pneumothorax.</li>
	<li>Extend the incision laterally along the intercostal muscle in both directions, taking care not to touch the exposed lung surface.</li>
	<li>Using blunt forceps, grasp the next highest rib and slightly retract to allow the lung to fall away from the chest wall. If the lung does not detach, press the chest wall lightly against the lung to cause the lung to adhere to the underlying pleura and thus fall away more easily.</li>
	<li>Continue the original incision ventrally until the sternum and cranially until the apex of the lung is exposed. Use cotton applicators and gauze to attenuate any bleeding that arises.</li>
	<li>Raise the ribcage to expose intercostal blood vessels on the dorsal aspect of the thoracic cavity. Taking care not to damage the lung, cauterize the most inferior intercostal vessel near to the spinal column, and then cut the rib. Moving cranially and ventrally, repeat until an approximately 1 cm x 1.5 cm portion of the ribcage is excised (Figure 2C).</li>
	<li>Remove any excess fluid accumulation in the thoracic cavity via capillary action using small strips of gauze.</li>
	<li>While proceeding to microscopy, allow ~5 min for intrapleural fluid to dissipate for a more secure interface between the lung and the imaging window.</li>
</ol>
6. Microscopy
<ol>
	<li>Turn on the vacuum pump and adjust the pressure to ~50&#8211;60 mmHg.</li>
	<li>Transfer the IVM platform to the microscope stage. Position the metal post and micromanipulator such that the imaging window is directly above the exposed lung and the window arm approaches the lung from, approximately, the 3 o&#39;clock position.</li>
	<li>Using the micromanipulator, carefully lower the imaging window until it adheres to and stabilizes the lung surface (Figure 3A).</li>
	<li>Using the 20x objective and the 460-490 nm bandpass excitation filter, identify a pulmonary venule based on the convergent pattern of blood flow. Center the vessel in the field of view and record 30 s of video.
	<ol>
		<li>Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.</li>
		<li>Repeat the previous step until five pulmonary venules have been imaged.</li>
	</ol>
	</li>
	<li>Using the 460-490 nm bandpass excitation filter, identify a pulmonary arteriole based on the divergent pattern of blood flow. Center the vessel in the field of view and record 30 s of video.
	<ol>
		<li>Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.</li>
		<li>Repeat the previous step until five pulmonary arterioles have been imaged.</li>
	</ol>
	</li>
	<li>Using the 460-490 nm bandpass excitation filter, locate a region of alveoli and capillaries not intersected by larger vessels and record 30 s of video.
	<ol>
		<li>Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.</li>
		<li>Repeat the previous step until five capillary regions have been imaged.</li>
	</ol>
	</li>
</ol>
7. Euthanasia and cleaning protocol
<ol>
	<li>Remove the IVM platform from the microscope stage and adjust isoflurane delivery to 5% for 5 min to euthanize the mouse.</li>
	<li>While waiting, discard the cover glass and disconnect the imaging window from the platform. Clean the imaging window with a small brush and use a 30 G syringe inserted into the channel to flush it several times with distilled water. Then, flush with 95% ethanol using the vacuum pump.</li>
	<li>After 5 min has elapsed, stop the ventilator, and ensure complete euthanasia via cervical dislocation.</li>
</ol>

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Dr. Kamala D. Patel is the president and co-founder of Luxidea, the enterprise from which the imaging window utilized in this experiment was purchased.

The protocol presented here requires practice and attention to a few critical steps. First, it is important to prepare the imaging window prior to initiating intubation and surgery. Use a minimal amount of vacuum grease to coat the outer ring of the imaging window, apply the cover glass, and test suction with a drop of distilled water. Preparing this in advance will prevent the exposed lung from drying out during the setup otherwise. While it is possible to flush with warm saline, doing so may risk damaging the fragile p...

Intravital microscopy (IVM) is a useful imaging tool for visualizing and studying various biophysical processes in vivo. The lung is highly challenging to image in vivo due to its enclosed location, the fragile nature of its tissue, and motion artifacts induced by respiration and heartbeat1,2. Various intravital microscopy (IVM) setups have been developed for real-time imaging of leukocyte-endothelial interactions in pulmonary microcirculation to overcome these challenges. Such approaches are based on surgically exposing and stabilizing the lung for imaging.
Animals are typically prepared for lung IVM by surgical procedures. First, animals are intubated and ventilated, which permits surgical excision of a thoracic window and subsequent interventions to stabilize the lung for imaging. One technique involves gluing the parenchyma onto a glass coverslip3, a procedure that risks significant physical trauma to the imaged tissue. More advanced is the utilization of a vacuum system to stabilize the lung under a glass window4. This setup facilitates loose adherence of the lung surface to the coverslip via a reversible vacuum spread over a large local area and expands the lung while still limiting the movement in x, y, and z dimensions4. The vacuum is applied evenly through a channel surrounding the imaging area of the setup and pulls the tissue into a shallow conical region facing the imaging-grade coverslip4. Through this viewing window, the lung microcirculation can be studied using various optical imaging modalities.
Lung IVM enables quantitative imaging of a multitude of microcirculatory parameters. These include measurements such as leukocyte track speed and length5, red blood cell flow velocity6 and oxygenation7, tumor metastases8, the distinction of immune cell subpopulations9,10,11, visualization of microparticles12, alveolar dynamics13,14, vascular permeability15, and capillary function16. The focus here is on leukocyte recruitment and capillary function. Initiation of leukocyte recruitment in the pulmonary microcirculation involves transient rolling interactions and firm adhesive interactions between leukocytes and endothelial cells, both of which are increased under inflammatory conditions16,17. Typically, rolling is quantified by the number of leukocytes that pass an operator-defined reference line, while adhesion is quantified by the number of leukocytes that are immobile on the endothelium16. Capillary function may also be affected in inflammatory states, often resulting in decreased perfusion. This can be attributed to several factors, including a reduction of red blood cell deformability18 and variegated expression of inducible NO synthase by endothelial cells resulting in pathological shunting19. Typically, the aggregate length of perfused capillaries per area is measured and reported as functional capillary density (FCD).
Studying leukocyte recruitment in the lungs in real-time requires labeling biological targets with fluorescent dyes or fluorescent-labeled antibodies20. Alternatively, various transgenic mouse strains such as lysozyme M-green fluorescent protein (LysM-GFP) mice can be utilized to image specific immune cell subsets such as neutrophils21,22. The fluorescent-labeled leukocytes can then be visualized using widefield fluorescence microscopy, confocal microscopy, or multiphoton microscopy. These techniques achieve contrast by utilizing specific excitation wavelengths and detecting emitted fluorescence while simultaneously blocking the detection of the excitation wavelength, thus highlighting the labeled object.
Existing research concerning the quantification of leukocyte rolling, adhesion, and functional capillary density in the murine lung has relied primarily on manual video analysis. This is made possible through open-source software such as Fiji6,23, proprietary software such as CapImage12, or custom-made image processing systems24. Conversely, various proprietary software platforms (e.g., NIS Element, Imaris, Volocity, MetaMorph) enable automated measurement of a wide array of other physiological parameters, including many of those previously mentioned here5,6,7,8,9,10,11,12,13,15.
Important observations have been made regarding the pathology of acute lung injury (ALI) and acute respiratory distress syndrome (ARDS) using lung IVM. ARDS is characterized by a host of pathophysiological processes in the lung, including pulmonary edema and alveolar damage caused by dysfunction of the endothelium and epithelial barrier25. Using a murine model, it has been found that sepsis-induced ALI is associated with significant detrimental changes in immune cell trafficking in the lung environment26. Neutrophils recruited to the capillaries of mice with sepsis-induced ALI were found to impede microcirculation, thereby increasing hypoxia in ALI26. Additionally, IVM has been used to gain insights into the underlying mechanism of repair following the onset of ARDS27. Lung IVM has also been a valuable tool in understanding pathophysiological changes in various obstructive lung diseases. For example, visualization of mucus transport in diseases such as cystic fibrosis (CF) and chronic obstructive pulmonary disease (COPD) has facilitated the study of novel and existing treatments for mucous clearance28. Leukocyte trafficking under these conditions has been analyzed as well17.
This protocol expands on the approach initially described by Lamm et al.29 to study leukocyte-endothelial interactions using conventional fluorescence microscopy. The described procedures employ an in vivo lung imaging system, which includes a 16.5 cm x 12.7 cm metal base, micromanipulator, and vacuum imaging window (Figure 1). The system is mounted in a 20 cm x 23.5 cm 3-D printed platform (Supplemental File 1) to provide secure attachment for the ventilator tubing and heating pad. This method offers reproducible and quantifiable imaging of murine pulmonary microcirculation in vivo. Important aspects of the surgical preparation as well as proper utilization of a vacuum-stabilized lung imaging system are explained in detail. Finally, an experimental model of ALI is used to provide representative imaging and analysis of altered leukocyte rolling, leukocyte adhesion, and capillary perfusion associated with inflammation. The use of this protocol should facilitate further important investigations into pathophysiological changes in pulmonary microcirculation during acute disease states.

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			<td>1 mL BD Luer Slip Tip Syringe sterile, single use</td>
			<td>Becton, Dickinson and Company</td>
			<td>309659</td>
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			<td>ADSON Dressing Forceps, Tip width 0.6 mm, teeth length 11.5 mm, 12 cm</td>
			<td>RWD Life Science Co.</td>
			<td>F12002-12</td>
			<td>Blunt forceps</td>
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			<td>Albumin-Fluorescein Isothiocyanate</td>
			<td>Sigma-Aldrich</td>
			<td>A9771-1G</td>
			<td>FITC-albumin</td>
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			<td>Alcohol Swab Isopropyl Alcohol 70% v/v</td>
			<td>Canadian Custom Packaging Company</td>
			<td>80002455</td>
			<td>Alcohol wipe</td>
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			<td>AVDC110 Advanced Digital Video Converter</td>
			<td>Canopus</td>
			<td>00631069602029</td>
			<td>Digital video converter</td>
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			<td>B/W - CCD - Camera</td>
			<td>Horn Imaging</td>
			<td>BC-71</td>
			<td>Camera</td>
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			<td>Bovie Deluxe High Temperature Cautery Kit</td>
			<td>Fine Science Tools</td>
			<td>18010-00</td>
			<td>Cauterizer</td>
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			<td>C57BL/6 Mice</td>
			<td>Charles River Laboratories International</td>
			<td>C57BL/6NCrl</td>
			<td>C57BL/6 Mice</td>
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			<td>Cotton Tipped Applicators</td>
			<td>Puritan</td>
			<td>806-WC</td>
			<td>Cotton applicator</td>
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			<td>CS-8R 8mm Round Glass Coverslip</td>
			<td>Warner Instruments</td>
			<td>64-0701</td>
			<td>Glass coverslip</td>
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			<td>Digital Pressure Gauge</td>
			<td>ITM Instruments Inc.</td>
			<td>DG2551L0NAM02L0IM&#38;V</td>
			<td>Digital Pressure Gauge</td>
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			<td>Dr Mom Slimline Stainless LED Otoscope</td>
			<td>Dr. Mom Otoscopes</td>
			<td>1001</td>
			<td>Otoscope</td>
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			<td>Ethyl Alchohol 95% Vol</td>
			<td>Commercial Alcohols</td>
			<td>P016EA95</td>
			<td>95% ethanol</td>
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			<td>Fine Scissors - Martensitic Stainless Steel</td>
			<td>Fine Science Tools</td>
			<td>14094-11</td>
			<td>Scissors</td>
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			<td>Fisherbrand Colored Labeling Tape</td>
			<td>Fisher Scientific</td>
			<td>1590110</td>
			<td>Labeling tape</td>
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			<td>Gast DOA-P704-AA High-Capacity Vacuum Pump</td>
			<td>Cole-Parmer Canada Company</td>
			<td>ZA-07061-40</td>
			<td>Vacuum pump</td>
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			<td>Hartman Hemostats</td>
			<td>Fine Science Tools</td>
			<td>13003-10</td>
			<td>Hemostatic forceps</td>
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			<td>High Vacuum Grease</td>
			<td>Dow Corning</td>
			<td>DC976VF</td>
			<td>Vacuum grease</td>
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		<tr>
			<td>Isoflurane USP</td>
			<td>Fresenius Kabi</td>
			<td>CP0406V2</td>
			<td>Isoflurane</td>
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		<tr>
			<td>LIDOcaine HCl Injection 1% 50 mg/5 mL</td>
			<td>Teligent Canada</td>
			<td>0121AD01</td>
			<td>Lidocaine HCl 1%</td>
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			<td>Lung SurgiBoard</td>
			<td>Luxidea, Inc.</td>
			<td>IMCH-0001</td>
			<td>Designed for intravital microscopy of the lung</td>
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			<td>Mineral Oil</td>
			<td>Teva Canada</td>
			<td>00485802</td>
			<td>Mineral oil</td>
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		<tr>
			<td>Mouse Endotracheal Intubation Kit</td>
			<td>Kent Scientific Corporation</td>
			<td>ETI-MSE</td>
			<td>Intubation stand, anesthesia mask, 20 G endotracheal cannula, fibre optic cable</td>
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			<td>MST49 Fluorescence Microscope</td>
			<td>Leica Microsystems</td>
			<td>10 450 022</td>
			<td>Fluorescence Microscope</td>
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		<tr>
			<td>N Plan L 20x/0.40 Long Working Distance Microscope Objective</td>
			<td>Leica Microsystems</td>
			<td>566035</td>
			<td>20x objective</td>
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		<tr>
			<td>Non-Woven Sponges 2&#34; x 2&#34;</td>
			<td>AMD-Ritmed</td>
			<td>A2101-CH</td>
			<td>Gauze</td>
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		<tr>
			<td>Optixcare Eye Lube Plus</td>
			<td>Aventix</td>
			<td>5914322</td>
			<td>Tear gel</td>
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			<td>Original Prusa i3 MK3S+ 3D Printer</td>
			<td>Prusa Research</td>
			<td>PRI-MK3S-KIT-ORG-PEI</td>
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			<td>Linde Canada Inc.</td>
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			<td>PrecisionGlide Needle 30 G x 1/2 (0.3 mm x 13 mm)</td>
			<td>Becton, Dickinson and Company</td>
			<td>305106</td>
			<td>30 G needle</td>
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			<td>Pyrex 5340-2L 5340 Filtering Flasks, 2000 mL</td>
			<td>Cole-Parmer Canada Company</td>
			<td>5340-2L</td>
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			<td>Rhodamine 6 G</td>
			<td>Sigma-Aldrich</td>
			<td>252433</td>
			<td>Rhodamine 6G</td>
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			<td>Secure Soft Cloth Medical Tape - 3&#34;</td>
			<td>Primed</td>
			<td>PM5-630709</td>
			<td>Cloth tape</td>
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			<td>Silastic Medical Grade Tubing .040 in. ID x .085 in. OD</td>
			<td>Dow Corning</td>
			<td>602-205</td>
			<td>1.0 mm I.D. polyethylene tubing</td>
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			<td>Somnosuite Low-Flow Anesthesia System</td>
			<td>Kent Scientific Corporation</td>
			<td>SS-01, SS-04-module</td>
			<td>Small rodent ventilator, Low-flow anesthesia system, Heating pad, Rectal temperature probe, Pulse oximeter</td>
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			<td>Tissue Forceps, 12.5cm long, Curved, 1 x 2 Teeth</td>
			<td>World Precision Instruments</td>
			<td>501216</td>
			<td>Toothed forceps</td>
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			<td>Transpore Medical Tape, 1527-1, 1 in x 10 yd (2.5 cm x 9.1 m)</td>
			<td>3M</td>
			<td>7000002795</td>
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			<td>Tubing,Clear,3/8 in Inside Dia.</td>
			<td>Grainger Canada</td>
			<td>USSZUSA-HT3314</td>
			<td>1.0 cm I.D. polyethylene tubing</td>
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			<td>Whatman 6720-5002 50 mm In-Line Filters, PTFE, 0.2 &#181;m</td>
			<td>Cole-Parmer Canada Company</td>
			<td>6720-5002</td>
			<td>Inline 0.2&#181;m filter</td>
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intravital widefield fluorescence microscopy of pulmonary microcirculation in experimental acute lung injury using a vacuum-stabilized imaging system

Intravital imaging of leukocyte-endothelial interactions offers valuable insights into immune-mediated disease in live animals. The study of acute lung injury (ALI)/acute respiratory distress syndrome (ARDS) and other respiratory pathologies in vivo is difficult due to the limited accessibility and inherent motion artifacts of the lungs. Nonetheless, various approaches have been developed to overcome these challenges. This protocol describes a method for intravital fluorescence microscopy to study real-time leukocyte-endothelial interactions in the pulmonary microcirculation in an experimental model of ALI. An in vivo lung imaging system and 3-D printed intravital microscopy platform are used to secure the anesthetized mouse and stabilize the lung while minimizing confounding lung injury. Following preparation, widefield fluorescence microscopy is used to study leukocyte adhesion, leukocyte rolling, and capillary function. While the protocol presented here focuses on imaging in an acute model of inflammatory lung disease, it may also be adapted to study other pathological and physiological processes in the lung.

To illustrate results achievable through this protocol, acute lung injury (ALI) was induced 6 h prior to imaging using a model of intranasal bacterial lipopolysaccharide (LPS) instillation. Briefly, mice (n = 3) were anesthetized with isoflurane, and small droplets of LPS from Pseudomonas aeruginosa in sterile saline (10 mg/mL) were pipetted into the left naris at a dosage of 5 mg/kg. This was compared to na&#239;ve mice (n = 3; no intranasal administration).
Upon imaging, a successfu...

Watch this Scientific Journal Video about Intravital Widefield Fluorescence Microscopy of Pulmonary Microcirculation in Experimental Acute Lung Injury Using a Vacuum-Stabilized Imaging System at JoVE.

Intravital Widefield Fluorescence Microscopy of Pulmonary Microcirculation in Experimental Acute Lung Injury Using a Vacuum-Stabilized Imaging System

Intravital fluorescence microscopy can be utilized to study leukocyte-endothelial interactions and capillary perfusion in real-time. This protocol describes methods to image and quantify these parameters in the pulmonary microcirculation using a vacuum-stabilized lung imaging system.

Intravital imaging of leukocyte-endothelial interactions offers valuable insights into immune-mediated disease in live animals. The study of acute lung ...

intravital-widefield-fluorescence-microscopy-pulmonary

Research

JoVE Journal

Immunology and Infection

2.7K Views.  Dalhousie University. Intravital fluorescence microscopy can be utilized to study leukocyte-endothelial interactions and capillary perfusion in real-time. This protocol describes methods to image and quantify these parameters in the pulmonary microcirculation using a vacuum-stabilized lung imaging system.

Video: Intravital Widefield Fluorescence Microscopy of Pulmonary Microcirculation in Experimental Acute Lung Injury Using a Vacuum-Stabilized Imaging System

Intravital Imaging of the Mouse Thymus using 2-Photon Microscopy

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic tools and surgical procedures, TPM becomes a powerful technique to identify "niches" inside organs and to document cellular "behaviors" in live animals. While intravital imaging provides information that best resembles the real cellular behavior inside the organ, it is both more laborious and technically demanding in terms of required equipment/procedures than alternative ex vivo imaging acquisition. Thus, we describe a surgical procedure and novel "stereotactic" organ holder that allows us to follow the movements of Foxp3+ cells within the thymus.
Foxp3 is the master regulator for the generation of regulatory T cells (Tregs). Moreover, these cells can be classified according to their origin: ie. thymus-differentiated Tregs are called "naturally-occurring Tregs" (nTregs), as opposed to peripherally-converted Tregs (pTregs). Although significant amount of research has been reported in the literature concerning the phenotype and physiology of these T cells, very little is known about their in vivo interactions with other cells. This deficiency may be due to the absence of techniques that would permit such observations. The protocol described in this paper provides a remedy for this situation.
Our protocol consists of using nude mice that lack an endogenous thymus since they have a punctual mutation in the DNA sequence that compromises the differentiation of some epithelial cells, including thymic epithelial cells. Nude mice were gamma-irradiated and reconstituted with bone marrows (BM) from Foxp3-KIgfp/gfp mice. After BM recovery (6 weeks), each animal received embryonic thymus transplantation inside the kidney capsule. After thymus acceptance (6 weeks), the animals were anesthetized; the kidney containing the transplanted thymus was exposed, fixed in our organ holder, and kept under physiological conditions for in vivo imaging by TPM. We have been using this approach to study the influence of drugs in the generation of regulatory T cells.

We have developed novel laboratory tools and protocols for intravital imaging acquisition of the thymus. Our technique should help in the identification of &#x201C;niches&#x201D; within the thymus where T cell development occurs.

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic ...

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick tissue samples and organs in health and disease. By controlling the scanning pattern in multi-photon microscopy and applying appropriate numerical algorithms, we developed a striped-illumination approach, which enabled us to achieve 3-fold better axial resolution and improved signal-to-noise ratio, i.e. contrast, in more than 100 &#181;m tissue depth within highly scattering tissue of lymphoid organs as compared to standard multi-photon microscopy. The acquisition speed as well as photobleaching and photodamage effects were similar to standard photo-multiplier-based technique, whereas the imaging depth was slightly lower due to the use of field detectors. By using the striped-illumination approach, we are able to observe the dynamics of immune complex deposits on secondary follicular dendritic cells &#8211; on the level of a few protein molecules in germinal centers.

High-resolution intravital imaging with enhanced contrast up to 120 &#181;m depth in lymph nodes of adult mice is achieved by spatially modulating the excitation pattern of a multi-focal two-photon microscope. In 100 &#181;m depth we measured resolutions of 487 nm (lateral) and 551 nm (axial), thus circumventing scattering and diffraction limits.

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick ...

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. ...

Influenza Virus Propagation in Embryonated Chicken Eggs

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous emergence of new influenza virus strains due to mutation and re-assortment complicates the control of the virus and necessitates the permanent development of novel drugs and vaccines. The laboratory-based study of influenza requires a reliable and cost-effective method for the propagation of the virus. Here, a comprehensive protocol is provided for influenza A virus propagation in fertile chicken eggs, which consistently yields high titer viral stocks. In brief, serum pathogen-free (SPF) fertilized chicken eggs are incubated at 37 &#176;C and 55-60% humidity for 10 &#8211; 11 days. Over this period, embryo development can be easily monitored using an egg candler. Virus inoculation is carried out by injection of virus stock into the allantoic cavity using a needle. After 2 days of incubation at 37 &#176;C, the eggs are chilled for at least 4 hr at 4 &#176;C. The eggshell above the air sac and the chorioallantoic membrane are then carefully opened, and the allantoic fluid containing the virus is harvested. The fluid is cleared from debris by centrifugation, aliquoted and transferred to -80 &#176;C for long-term storage. The large amount (5-10 ml of virus-containing fluid per egg) and high virus titer which is usually achieved with this protocol has made the usage of eggs for virus preparation our favorable method, in particular for in vitro studies which require large quantities of virus in which high dosages of the same virus stock are needed.

Fertile chicken eggs are widely used to produce large amounts of human influenza A virus as they provide a convenient and cost-effective system to prove high yields of virus.

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous ...

Long Term Intravital Multiphoton Microscopy Imaging of Immune Cells in Healthy and Diseased Liver Using CXCR6.Gfp Reporter Mice

Liver inflammation as a response to injury is a highly dynamic process involving the infiltration of distinct subtypes of leukocytes including monocytes, neutrophils, T cell subsets, B cells, natural killer (NK) and NKT cells. Intravital microscopy of the liver for monitoring immune cell migration is particularly challenging due to the high requirements regarding sample preparation and fixation, optical resolution and long-term animal survival. Yet, the dynamics of inflammatory processes as well as cellular interaction studies could provide critical information to better understand the initiation, progression and regression of inflammatory liver disease. Therefore, a highly sensitive and reliable method was established to study migration and cell-cell-interactions of different immune cells in mouse liver over long periods (about 6 hr) by intravital two-photon laser scanning microscopy (TPLSM) in combination with intensive care monitoring.
The method provided includes a gentle preparation and stable fixation of the liver with minimal perturbation of the organ; long term intravital imaging using multicolor multiphoton microscopy with virtually no photobleaching or phototoxic effects over a time period of up to 6 hr, allowing tracking of specific leukocyte subsets; and stable imaging conditions due to extensive monitoring of mouse vital parameters and stabilization of circulation, temperature and gas exchange.
To investigate lymphocyte migration upon liver inflammation CXCR6.gfp knock-in mice were subjected to intravital liver imaging under baseline conditions and after acute and chronic liver damage induced by intraperitoneal injection(s) of carbon tetrachloride (CCl4).
CXCR6 is a chemokine receptor expressed on lymphocytes, mainly on Natural Killer T (NKT)-, Natural Killer (NK)- and subsets of T lymphocytes such as CD4 T cells but also mucosal associated invariant (MAIT) T cells1. Following the migratory pattern and positioning of CXCR6.gfp+ immune cells allowed a detailed insight into their altered behavior upon liver injury and therefore their potential involvement in disease progression.

Stable intravital high-resolution imaging of immune cells in the liver is challenging. Here we provide a highly sensitive and reliable method to study migration and cell-cell-interactions of immune cells in mouse liver over long periods (about 6 hours) by intravital multiphoton laser scanning microscopy in combination with intensive care monitoring.

Liver inflammation as a response to injury is a highly dynamic process involving the infiltration of distinct subtypes of leukocytes including monocytes, ...

Imaging Neutrophils and Monocytes in Mesenteric Veins by Intravital Microscopy on Anaesthetized Mice in Real Time

Efficient immune response is dependent on rapid mobilization of blood leukocytes to the site of infection or injury. Investigating leukocyte migration in vivo is crucial for understanding the molecular basis of leukocyte transendothelial migration and interaction with vascular endothelium. One powerful approach involves intravital microscopy on transgenic mice expressing fluorescent proteins in cells of interest.
Here we present a protocol for imaging monocytes and neutrophils in the CX3CR1gfp/wt mouse i.v. injected with orange dye-labeled neutrophils with an inverted confocal microscope. Time-lapse movies gathered from 30 min&#160;to several hours of imaging allow the analysis of leukocyte behavior in mesenteric veins under both steady state and inflammatory conditions. We also describe the steps to locally induce blood vessel inflammation with TLR2/TLR1 agonist Pam3SK4 and monitor the subsequent recruitment of neutrophils and monocytes.
The presented technique can also be used to monitor other populations of leukocytes and investigate molecules implicated in leukocyte recruitment or trafficking using other stimuli or transgenic mice.

We detail a protocol to monitor the behavior of neutrophils and monocytes in mesenteric veins under steady state and inflammatory conditions using intravital confocal microscopy on anaesthetized mice.

Efficient immune response is dependent on rapid mobilization of blood leukocytes to the site of infection or injury. Investigating leukocyte migration in ...

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese ...

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. ...

A Model of Self-limited Acute Lung Injury by Unilateral Intra-bronchial Acid Instillation

Selective intra-bronchial instillation of hydrochloric acid (HCl) to the murine left mainstem bronchus causes acute tissue injury with histopathologic findings similar to human acute respiratory distress syndrome (ARDS). The resulting alveolar edema, alveolar-capillary barrier damage, and leukocyte infiltration predominantly affect the left lung, preserving the right lung as an uninjured control and allowing animals to survive. This model of self-limited acute lung injury enables investigation of tissue resolution mechanisms, such as macrophage efferocytosis of apoptotic neutrophils and restitution of alveolar-capillary barrier integrity. This model has helped identify important roles for resolution agonists, including specialized pro-resolving mediators (SPMs), providing a foundation for the development of new therapeutic approaches for patients with ARDS.

Selective intra-bronchial acid instillation to the left lung in mice results in unilateral and self-limited acute lung injury that models human acute respiratory distress syndrome (ARDS) induced by gastric acid aspiration.

Selective intra-bronchial instillation of hydrochloric acid (HCl) to the murine left mainstem bronchus causes acute tissue injury with histopathologic ...

Studying Effects of Cigarette Smoke on Pseudomonas Infection in Lung Epithelial Cells

Cigarette smoking is the major etiological cause for lung emphysema and chronic obstructive pulmonary disease (COPD). Cigarette smoking also promotes susceptibility to bacterial infections in the respiratory system. However, the effects of cigarette smoking on bacterial infections in human lung epithelial cells have yet to be thoroughly studied. Described here is a detailed protocol for the preparation of cigarette smoking extracts (CSE), treatment of human lung epithelial cells with CSE, and bacterial infection and infection determination. CSE was prepared with a conventional method. Lung epithelial cells were treated with 4% CSE for 3 h. CSE-treated cells were, then, infected with Pseudomonas at a multiplicity of infection (MOI) of 10. Bacterial loads of the cells were determined by three different methods. The results showed that CSE increased Pseudomonas load in lung epithelial cells. This protocol, therefore, provides a simple and reproducible approach to study the effect of cigarette smoke on bacterial infections in lung epithelial cells.

Studying Effects of Cigarette Smoke on Pseudomonas Infection in Lung Epithelial Cells

Described here is a protocol to study how cigarette smoke extract affects bacterial colonization in lung epithelial cells.

Cigarette smoking is the major etiological cause for lung emphysema and chronic obstructive pulmonary disease (COPD). Cigarette smoking also promotes ...