Name: intravital widefield fluorescence microscopy of pulmonary microcirculation in experimental acute lung injury using a vacuum-stabilized imaging system
Uploaded: 2022-04-06T14:40:00.000+00:00
Duration: 9 min 28 s
Description: 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.

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 pulmonary tissue.
After intubation, while transferring the mouse to the IVM platform, the cannula may occasionally become displaced. To prevent this, consider tying the cannula to the mouse's front teeth or suturing it to the skin around its mouth. If using pressure-controlled ventilation, the tidal volume should be carefully monitored throughout the procedure. It should remain steady at approximately 0.20 mL, as significantly lower values (e.g., ~0.10 mL) may indicate single lung ventilation. If this occurs, slightly retracting the endotracheal cannula may resolve the issue. The use of inhalant anesthesia (isoflurane) facilitates control of anesthetic depth while the mouse is ventilated. Other means of anesthesia (e.g., intravenous ketamine/xylazine10,11) have been employed by other laboratories, and each carry their respective advantages and drawbacks.
During the surgery, blunt dissection is employed to minimize the risk of severing blood vessels. This is of particular importance when dissecting the thick and heavily vascularized adipose tissue near the shoulder. Resection of the ribcage requires both speed and accuracy. The heat from the cauterizer is intense enough to burn the lung if used excessively. When resecting the ribcage, there should be empty space between the lung and the chest wall. If the lung adheres to the chest wall, lightly pressing down on the outside of the rib cage will encourage adhesion to the underlying parietal pleura. Alternatively, use a blunt needle to inject a small amount of warm saline between the lung and the ribcage to facilitate release. Positioning the window arm at the 3 o'clock position is advised to avoid unwanted pressure on the ribs during imaging (Figure 3A). It will also leave more clearance between the micromanipulator and the microscope objective, allowing easier access for manipulations. When lowering the imaging window, it is critical to target the central region of the lung as contact with the edge will result in an inadequate seal and excessive motion artifact during imaging. Also, multiple attempts to lower the window can damage the lung. Likewise, cycles of detachment and re-stabilization will contribute to lung injury and may compromise experimental accuracy. When performed correctly, however, the preparation presented here is stable enough to enable high-resolution intravital imaging with a 20x objective.
A rigorous cleaning protocol is necessary to prevent contamination and blockage of the vacuum channel within the imaging window. Using a 30 G needle immediately following experiments to repeatedly flush with distilled water is effective for removing most contaminants. This is followed by a flush with concentrated ethanol and a final flush with distilled water before reattaching to the vacuum line to remove moisture. Using acetone or strong detergents may be adequate to resolve serious blockages, should they occur.
Notably, this protocol employs a higher readout of vacuum pressure versus some other reported methods4,11, and this may raise concerns of damage to lung tissue. An important distinction, however, is that the digital gauge used here measures pressure only at the vacuum pump. This pressure is delivered through narrow tubing and the extremely narrow channel of the imaging window itself prior to distribution over a larger surface area of the lung. As such, no evidence of damage to the imaged region was observed in these experiments following intravital sessions. Furthermore, imaged alveoli exhibited tidal distension and contraction, indicating that this physiological phenomenon was not significantly disrupted.
Despite the increasing popularity of lung IVM as a tool for studying disease in vivo, there are limitations to this technique. First, the invasive and terminal nature of the surgery induces a non-negligible effect on the mouse's physiological state and limits the procedure to a single imaging session. However, it should be noted that several longitudinal lung IVM approaches have been developed30. Second, the use of mechanical ventilation may induce a degree of ventilator-associated lung injury (VALI)31, although this is limited by the short duration of the procedure. Third, contact between the visceral pleura and glass coverslip and the application of vacuum pressure may result in altered microvascular blood flow. Finally, perhaps the most significant limitation of this approach is that imaging is restricted to subpleural alveoli in non-dependent pulmonary regions, which are not representative of the entire lung32.
In summary, this protocol can be used to study leukocyte-endothelial interactions in the pulmonary microvasculature using intravital fluorescence microscopy. While these experiments employ endotoxin-induced lung injury, an acute model which was selected based on previous research, this protocol may also be adapted to study other pathological and physiological processes in the lung. Furthermore, the lung imaging system employed here is applicable to a range of microscopy approaches, and the imaging window is large enough to accommodate high numerical aperture oil-immersion objectives. Thus, the described procedures should facilitate further research into the impact of various disease states on pulmonary microcirculation.

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.

<table><tbody><tr><td>1 mL BD Luer Slip Tip Syringe sterile, single use</td><td>Becton, Dickinson and Company</td><td>309659</td><td>1 mL syringe</td></tr><tr><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></tr><tr><td>Albumin-Fluorescein Isothiocyanate</td><td>Sigma-Aldrich</td><td>A9771-1G</td><td>FITC-albumin</td></tr><tr><td>Alcohol Swab Isopropyl Alcohol 70% v/v</td><td>Canadian Custom Packaging Company</td><td>80002455</td><td>Alcohol wipe</td></tr><tr><td>AVDC110 Advanced Digital Video Converter</td><td>Canopus</td><td>00631069602029</td><td>Digital video converter</td></tr><tr><td>B/W - CCD - Camera</td><td>Horn Imaging</td><td>BC-71</td><td>Camera</td></tr><tr><td>Bovie Deluxe High Temperature Cautery Kit</td><td>Fine Science Tools</td><td>18010-00</td><td>Cauterizer</td></tr><tr><td>C57BL/6 Mice</td><td>Charles River Laboratories International</td><td>C57BL/6NCrl</td><td>C57BL/6 Mice</td></tr><tr><td>Cotton Tipped Applicators</td><td>Puritan</td><td>806-WC</td><td>Cotton applicator</td></tr><tr><td>CS-8R 8mm Round Glass Coverslip</td><td>Warner Instruments</td><td>64-0701</td><td>Glass coverslip</td></tr><tr><td>Digital Pressure Gauge</td><td>ITM Instruments Inc.</td><td>DG2551L0NAM02L0IM&amp;amp;V</td><td>Digital Pressure Gauge</td></tr><tr><td>Dr Mom Slimline Stainless LED Otoscope</td><td>Dr. Mom Otoscopes</td><td>1001</td><td>Otoscope</td></tr><tr><td>Ethyl Alchohol 95% Vol</td><td>Commercial Alcohols</td><td>P016EA95</td><td>95% ethanol</td></tr><tr><td>Fine Scissors - Martensitic Stainless Steel</td><td>Fine Science Tools</td><td>14094-11</td><td>Scissors</td></tr><tr><td>Fisherbrand Colored Labeling Tape</td><td>Fisher Scientific</td><td>1590110</td><td>Labeling tape</td></tr><tr><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></tr><tr><td>Hartman Hemostats</td><td>Fine Science Tools</td><td>13003-10</td><td>Hemostatic forceps</td></tr><tr><td>High Vacuum Grease</td><td>Dow Corning</td><td>DC976VF</td><td>Vacuum grease</td></tr><tr><td>Isoflurane USP</td><td>Fresenius Kabi</td><td>CP0406V2</td><td>Isoflurane</td></tr><tr><td>LIDOcaine HCl Injection 1% 50 mg/5 mL</td><td>Teligent Canada</td><td>0121AD01</td><td>Lidocaine HCl 1%</td></tr><tr><td>Lung SurgiBoard</td><td>Luxidea, Inc.</td><td>IMCH-0001</td><td>Designed for intravital microscopy of the lung</td></tr><tr><td>Mineral Oil</td><td>Teva Canada</td><td>00485802</td><td>Mineral oil</td></tr><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></tr><tr><td>MST49 Fluorescence Microscope</td><td>Leica Microsystems</td><td>10 450 022</td><td>Fluorescence Microscope</td></tr><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></tr><tr><td>Non-Woven Sponges 2&quot; x 2&quot;</td><td>AMD-Ritmed</td><td>A2101-CH</td><td>Gauze</td></tr><tr><td>Optixcare Eye Lube Plus</td><td>Aventix</td><td>5914322</td><td>Tear gel</td></tr><tr><td>Original Prusa i3 MK3S+ 3D Printer</td><td>Prusa Research</td><td>PRI-MK3S-KIT-ORG-PEI</td><td>3D printer</td></tr><tr><td>Oxygen, Compressed</td><td>Linde Canada Inc.</td><td></td><td>Oxygen</td></tr><tr><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></tr><tr><td>Pyrex 5340-2L 5340 Filtering Flasks, 2000 mL</td><td>Cole-Parmer Canada Company</td><td>5340-2L</td><td>Vacuum flask</td></tr><tr><td>Rhodamine 6 G</td><td>Sigma-Aldrich</td><td>252433</td><td>Rhodamine 6G</td></tr><tr><td>Secure Soft Cloth Medical Tape - 3&quot;</td><td>Primed</td><td>PM5-630709</td><td>Cloth tape</td></tr><tr><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></tr><tr><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></tr><tr><td>Tissue Forceps, 12.5cm long, Curved, 1 x 2 Teeth</td><td>World Precision Instruments</td><td>501216</td><td>Toothed forceps</td></tr><tr><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><td>Medical tape</td></tr><tr><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></tr><tr><td>Whatman 6720-5002 50 mm In-Line Filters, PTFE, 0.2 &amp;micro;m</td><td>Cole-Parmer Canada Company</td><td>6720-5002</td><td>Inline 0.2&amp;micro;m filter</td></tr></tbody></table>

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 successful surgical preparation is identifiable by several factors. The lung should be relatively stable with respiration causing cyclic frameshifts no greater than 25 &#181;m. Alveoli should be clearly visible and may exhibit tidal distension/contraction. Excitation by blue light (450-490 nm wavelength) will permit visualization of blood flow directionality, and it may be possible to distinguish individual red blood cells (Supplemental Movie 1, Supplemental Movie 3, and Supplemental Movie 5). Leukocytes will be clearly identifiable upon excitation by green light (530-560 nm wavelength, Supplemental Movie 2, Supplemental Movie 4, and Supplemental Movie 6). After completion of imaging and removal of the suction window, there may be slight bruising of the lung surface, although not within the imaged area, as shown in Figure 3B.
Several technical challenges may interfere with experimental viability. Accumulation of blood on the lung surface will compromise vacuum stabilization and may even clog the channel. To avoid this, extreme caution should be exercised during each surgical step. Excessively high vacuum pressure may damage the lung and affect microcirculation. This may be identifiable by alveolar stasis or excessive bruising on the pulmonary surface (Figure 3C) and can be remedied by reducing pressure from the vacuum pump. As well, errors in intravenous fluorophore injection may lead to poor visualization of leukocyte trafficking and blood flow.
Following completion of the protocol, blinded manual analysis was performed using Fiji21 in a manner adapted from the previous literature28. Five venules, arterioles, and capillary regions-of-interest (ROIs) were analyzed from each animal. Leukocyte adhesion in venules and arterioles was defined as the number of cells that remain adhered to the vascular endothelium during 30 s of observation per area of endothelial surface. This is relayed in cells/mm2. Leukocyte adhesion in capillaries was defined as the number of cells within ROI that remain adhered to the vascular endothelium during 30 s of observation per total analyzed area. This is also relayed in cells/mm2. Leukocyte rolling was defined as twice the number of cells passing a reference point in the vessel during the 30 s observation period. Free-flowing leukocytes were excluded by comparing the speed of passage with that of red blood cell flow, and this is relayed in cells/min. To measure microcirculatory perfusion, FCD was defined as the sum of the lengths of red blood cell-perfused capillaries per observation area. This is relayed in cm/cm2. Each parameter is reported as a mean value for each animal.
A common trend of leukocyte recruitment was observed in pulmonary venules, with increased adhesion and rolling in LPS-treated mice versus na&#239;ve ones (Figures 4C,D). This trend was recapitulated by arteriolar leukocyte adhesion, although both levels of rolling and adhesion were highly variable in the na&#239;ve group (Figures 5C,D). Notably, LPS administration resulted in a substantial increase in leukocyte adhesion in pulmonary capillary ROIs (Figure 6C). LPS mice also demonstrated a reduced FCD versus na&#239;ve mice (Figure 7C). These effects within the pulmonary capillaries are consistent with previous literature identifying increases in immune cells per field of view and derangement of normal capillary perfusion following various inflammatory stimuli4,5,16.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63733/63733fig01.jpg" /> 
Figure 1: Lung imaging system. Custom-order system includes (1) an anodized metal base, (2) imaging window, (3) vacuum inlet, (4) micromanipulator. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63733/63733fig02.jpg" /> 
Figure 2: Surgical preparation. (A) Mouse is secured to the IVM platform in the right lateral decubitus position. (B) Ribcage is exposed using blunt dissection to preserve hemodynamics. (C) Thoracotomy is performed to expose the left lung. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63733/63733fig03.jpg" /> 
Figure 3: Vacuum stabilization. (A) Application of vacuum imaging window stabilizes pulmonary surface. (B) Utilization of vacuum pressures below 75 mmHg minimizes damage to the lung, particularly within the imaged area. (C) Higher vacuum pressures may cause significant bruising. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63733/63733fig04.jpg" /> 
Figure 4: Leukocyte trafficking in pulmonary venules. Excitation of rhodamine 6G permits visualization of adherent and rolling leukocytes. Outlined areas represent analyzed portions of vascular endothelium as confirmed by excitation of FITC-albumin in (A) na&#239;ve and (B) LPS-treated mice. (C,D) Intranasal LPS administration impacts leukocyte rolling and adhesion in pulmonary venules. Values are given as mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63733/63733fig05.jpg" /> 
Figure 5: Leukocyte trafficking in pulmonary arterioles. Excitation of rhodamine 6G permits visualization of adherent and rolling leukocytes. Outlined areas represent analyzed portions of vascular endothelium as confirmed by excitation of FITC-albumin in (A) na&#239;ve and (B) LPS-treated mice. (C,D) Intranasal LPS administration impacts leukocyte rolling and adhesion in pulmonary arterioles. Values are given as mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63733/63733fig06.jpg" /> 
Figure 6: Adherent leukocytes in pulmonary capillaries. Excitation of rhodamine 6G permits visualization of leukocytes within ROIs in (A) na&#239;ve and (B) LPS-treated mice. (C) Intranasal LPS administration impacts leukocyte adhesion in LPS-treated mice. Values are given as mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/63733/63733fig07.jpg" /> 
Figure 7: Pulmonary capillary function. Excitation of FITC-albumin permits visualization of capillary blood flow within ROIs in (A) na&#239;ve and (B) LPS-treated mice. (C) Intranasal LPS administration impacts FCD in pulmonary capillaries. Values are given as mean &#177; SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/63733fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File: File for 3D-printable IVM platform. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/Supplemental File 1.stl" target="_blank">Please click here to download this File.</a>
Supplemental Figure 1: Diagram of the vacuum system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/Supplemental Figure 1.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Movie 1: Sample video of blood flow in the pulmonary venule. The green arrow denotes the direction of the blood flow. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S1 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>
Supplemental Movie 2: Sample video of leukocyte trafficking in the pulmonary venule. Red arrows denote adherent leukocytes within the vessel. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S2 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>
Supplemental Movie 3: Sample video of blood flow in the pulmonary arteriole. The green arrow denotes the direction of the blood flow. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S3 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>
Supplemental Movie 4: Sample video of leukocyte trafficking in pulmonary arteriole. Red arrows denote adherent leukocytes within a vessel. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S4 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>
Supplemental Movie 5: Sample video of blood flow in pulmonary capillaries. Green arrows denote several well-visualized areas of red blood cell-perfused capillaries. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S5 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>
Supplemental Movie 6: Sample video of leukocyte trafficking in pulmonary capillaries. Red arrows denote adherent leukocytes within the field of view. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63733/S6 Annotated.m4v" target="_blank">Please click here to download this Movie.</a>

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

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

2.8K Views.  Dalhousie University. Dit protocol beschrijft in detail de chirurgische voorbereiding en het gebruik van een vacuümgestabiliseerd systeem om pulmonale leukocytenadhesie en capillaire perfusie in vivo in beeld te brengen. Het belangrijkste voordeel van deze techniek is dat het stabiele intravitale beeldvorming met hoge resolutie van de intacte spiegelende long mogelijk maakt met minimaal fysiek trauma. Bereid om te beginnen het beeldvormingsvenster voor door een dunne laag vacuümvet aan de bovenkant van de buiten...

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

Long-term High-Resolution Intravital Microscopy in the Lung with a Vacuum Stabilized Imaging Window

Metastasis to secondary sites such as the lung, liver and bone is a traumatic event with a mortality rate of approximately 90% 1. Of these sites, the lung is the most difficult to assess using intravital optical imaging due to its enclosed position within the body, delicate nature and vital role in sustaining proper physiology. While clinical modalities (positron emission tomography (PET), magnetic resonance imaging (MRI) and computed tomography (CT)) are capable of providing noninvasive images of this tissue, they lack the resolution necessary to visualize the earliest seeding events, with a single pixel consisting of nearly a thousand cells. Current models of metastatic lung seeding postulate that events just after a tumor cell&#39;s arrival are deterministic for survival and subsequent growth. This means that real-time intravital imaging tools with single cell resolution 2 are required in order to define the phenotypes of the seeding cells and test these models. While high resolution optical imaging of the lung has been performed using various ex vivo preparations, these experiments are typically single time-point assays and are susceptible to artifacts and possible erroneous conclusions due to the dramatically altered environment (temperature, profusion, cytokines, etc.) resulting from removal from the chest cavity and circulatory system 3. Recent work has shown that time-lapse intravital optical imaging of the intact lung is possible using a vacuum stabilized imaging window 2,4,5 however, typical imaging times have been limited to approximately 6 hr. Here we describe a protocol for performing long-term intravital time-lapse imaging of the lung utilizing such a window over a period of 12 hr. The time-lapse image sequences obtained using this method enable visualization and quantitation of cell-cell interactions, membrane dynamics and vascular perfusion in the lung. We further describe an image processing technique that gives an unprecedentedly clear view of the lung microvasculature.

This protocol describes the use of multiphoton microscopy to perform long-term high-resolution, single cell imaging of the intact lung in real time using a vacuum stabilized imaging window.

Metastasis to secondary sites such as the lung, liver and bone is a traumatic event with a mortality rate of approximately 90% 1. Of these sites, the lung ...

Cancer Research

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

Neuroscience

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly hydrated, forming a substantial endothelial surface layer (ESL) that contributes to the maintenance of endothelial function. As the endothelial glycocalyx is often aberrant in vitro and is lost during standard tissue fixation techniques, study of the ESL requires use of intravital microscopy. To best approximate the complex physiology of the alveolar microvasculature, pulmonary intravital imaging is ideally performed on a freely-moving lung. These preparations, however, typically suffer from extensive motion artifact. We demonstrate how closed-chest intravital microscopy of a freely-moving mouse lung can be used to measure glycocalyx integrity via ESL exclusion of fluorescently-labeled high molecular weight dextrans from the endothelial surface. This non-recovery surgical technique, which requires simultaneous brightfield and fluorescent imaging of the mouse lung, allows for longitudinal observation of the subpleural microvasculature without evidence of inducing confounding lung injury.

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx/endothelial surface layer is ideally studied using intravital microscopy. Intravital microscopy is technically challenging in a moving organ such as the lung. We demonstrate how simultaneous brightfield and fluorescent microscopy may be used to estimate endothelial surface layer thickness in a freely-moving in vivo mouse lung.

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly ...

Medicine

Automated Measurement of Microcirculatory Blood Flow Velocity in Pulmonary Metastases of Rats

Because the lung is a major target organ of metastatic disease, animal models to study the physiology of pulmonary metastases are of great importance. However, very few methods exist to date to investigate lung metastases in a dynamic fashion at the microcirculatory level, due to the difficulty to access the lung with a microscope. Here, an intravital microscopy method is presented to functionally image and quantify the microcirculation of superficial pulmonary metastases in rats, using a closed-chest pulmonary window and automated analysis of blood flow velocity and direction. The utility of this method is demonstrated to measure increases in blood flow velocity in response to pharmacological intervention, and to image the well-known tortuous vasculature of solid tumors. This is the first demonstration of intravital microscopy on pulmonary metastases in a closed-chest model. Because of its minimized invasiveness, as well as due to its relative ease and practicality, this technology has the potential to experience widespread use in laboratories that specialize on pulmonary tumor research.

A method is presented to measure microcirculatory blood flow velocity in pulmonary cancer metastases of the pleural surface in rats in an automated fashion, using closed-chest pulmonary intravital microscopy. This model has potential to be used as a widespread tool to perform physiologic research on pulmonary metastases in rodents.

Because the lung is a major target organ of metastatic disease, animal models to study the physiology of pulmonary metastases are of great importance. ...

A Multimodal Imaging Approach Based on Micro-CT and Fluorescence Molecular Tomography for Longitudinal Assessment of Bleomycin-Induced Lung Fibrosis in Mice

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which causes significant deterioration in lung function and subsequent death from respiratory failure.
The pathogenesis of IPF in experimental animal models has been induced by bleomycin administration. In this study, we investigate an IPF-like mouse model induced by a double intratracheal bleomycin instillation. Standard histological assessments used for studying lung fibrosis are invasive terminal procedures. The goal of this work is to monitor lung fibrosis through noninvasive imaging techniques such as Fluorescent Molecular Tomography (FMT) and Micro-CT. These two technologies validated with histology findings could represent a revolutionary functional approach for real time non-invasive monitoring of IPF disease severity and progression. The fusion of different approaches represents a step further for understanding the IPF disease, where the molecular events occurring in a pathological condition can be observed with FMT and the subsequent anatomical changes can be monitored by Micro-CT.

We describe a non-invasive multimodal imaging approach based on Micro-CT and fluorescence molecular tomography for longitudinal assessment of the mouse lung fibrosis model induced by double intratracheal instillation of bleomycin.

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which ...

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or fungal) inflammatory conditions. An overwhelming host response may result in compromised respiration and acute lung injury, which is characterized by lung neutrophil recruitment as a result of the patho-logical host immune, coagulative and tissue remodeling response. Sensitive microscopic methods to visualize and quantify murine lung cellular adaptations, in response to low-dose (0.05 ppm) ozone, a potent environmental pollutant in combination with bacterial lipopolysaccharide, a TLR4 agonist, are crucial in order to understand the host inflammatory and repair mechanisms. We describe a comprehensive fluorescent microscopic analysis of various lung and systemic body compartments, namely the broncho-alveolar lavage fluid, lung vascular perfusate, left lung cryosections, and sternal bone marrow perfusate. We show damage of alveolar macrophages, neutrophils, lung parenchymal tissue, as well as bone marrow cells in correlation with a delayed (up to 36-72 h) immune response that is marked by discrete chemokine gradients in the analyzed compartments. In addition, we present lung extracellular matrix and cellular cytoskeletal interactions (actin, tubulin), mitochondrial and reactive oxygen species, anti-coagulative plasminogen, its anti-angiogenic peptide fragment angiostatin, the mitochondrial ATP synthase complex V subunits, &#945; and &#946;. These surrogate markers, when supplemented with adequate in vitro cell-based assays and in vivo animal imaging techniques such as intravital microscopy, can provide vital information towards understanding the lung response to novel immunomodulatory agents.

Combined ozone and bacterial endotoxin exposed mice show wide-spread cell death, including that of neutrophils. We observed cellular adaptations such as disruption of cytoskeletal lamellipodia, increased cellular expression of complex V ATP synthase subunit &#946; and angiostatin in broncho-alveolar lavage, suppression of the lung immune response and delayed neutrophil recruitment.

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or ...

Advanced Imaging of Lung Homing Human Lymphocytes in an Experimental In Vivo Model of Allergic Inflammation Based on Light-sheet Microscopy

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an attractive therapeutic target in the clinical management of affected patients. In order to further optimize strategies aiming at therapeutic regulation of pathologically imbalanced tissue infiltration of pro-inflammatory immune cells, it will be of particular importance to achieve improved insights into disease- and organ-specific homing properties of peripheral lymphocytes. The here described experimental protocol allows to monitor lung accumulation of fluorescently labeled and adoptively transferred human lymphocytes in the context of papain-induced pulmonary inflammation. In contrast to standard in vitro assays frequently used for the analysis of immune cell migration and chemotaxis, the now introduced in vivo setting takes into account lung-specific aspects of tissue organization and the influence of the complex inflammatory scenario taking place in the living murine organism. Moreover, three-dimensional cross-sectional light-sheet fluorescence microscopic imaging does not only provide quantitative data on infiltrating immune cells, but also depicts the pattern of immune cell localization within the inflamed lung. Overall, we are able to introduce an innovative technique of high value for immunological research in the field of chronic inflammatory lung diseases, which can be easily applied by following the provided step-by-step protocol.

The here introduced protocol allows characterization of the lung homing capacity of primary human lymphocytes under in vivo inflammatory conditions. Pulmonary infiltration of adoptively transferred human immune cells in a mouse model of allergic inflammation can be imaged and quantified by light-sheet fluorescence microscopy of chemically cleared lung tissue.

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an ...

Quantifying Single Microvessel Permeability in Isolated Blood-perfused Rat Lung Preparation

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with real time imaging, it becomes feasible to determine permeability changes in individual pulmonary microvessels. Herein we describe steps to isolate rat lungs and perfuse them with autologous blood. Then, we outline steps to infuse fluorophores or agents via a microcatheter into a small lung region. Using these procedures described, we determined permeability increases in rat lung microvessels in response to infusions of bacterial lipopolysaccharide. The data revealed that lipopolysaccharide increased fluid leak across both venular and capillary microvessel segments. Thus, this method makes it possible to compare permeability responses among vascular segments and thus, define any heterogeneity in the response. While commonly used methods to define lung permeability require postprocessing of lung tissue samples, the use of real time imaging obviates this requirement as evident from the present method. Thus, the isolated lung preparation combined with real time imaging offers several advantages over traditional methods to determine lung microvascular permeability, yet is a straightforward method to develop and implement.

The isolated blood-perfused lung preparation makes it feasible to visualize microvessel networks on the lung surface. Here we describe an approach to quantify permeability of single microvessels in isolated lungs using real time fluorescence imaging.

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with ...

Biology

Air-Inflation of Murine Lungs with Vascular Perfusion-Fixation

Lung histology is often used to investigate the contributions provided by airspace cells during lung homeostasis and disease pathogenesis. However, commonly used instillation-based fixation methods can displace airspace cells and mucus into terminal airways and can alter tissue morphology. In comparison, vascular perfusion-fixation techniques are superior at preserving the location and morphology of cells within airspaces and the mucosal lining. However, if positive airway pressure is not simultaneously applied, regions of the lungs may collapse and capillaries may bulge into the alveolar spaces, leading to distortion of the lung anatomy. Herein, we describe an inexpensive method for air-inflation during vascular perfusion-fixation to preserve the morphology and location of airway and alveolar cells and interstitium in murine lungs for downstream histologic studies. Constant air pressure is delivered to the lungs via the trachea from a sealed, air-filled chamber that maintains pressure via an adjustable liquid column while fixative is perfused through the right ventricle.

Presented is a method for air-inflation with vascular perfusion-fixation of the lungs that preserves the location of cells within airways, alveoli and interstitium for structure-function analyses. Constant airway pressure is maintained with an air-inflation chamber while fixative is perfused via the right ventricle. Lungs are processed for histologic studies.

Lung histology is often used to investigate the contributions provided by airspace cells during lung homeostasis and disease pathogenesis. However, ...

Real-Time Monitoring of Cellular Dynamics in Mouse Lung Transplantation

Intravital Two-Photon Microscopy of the Transplanted Mouse Lung

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by crosstalk between donor and recipient cells. A better understanding of these processes relies on the use of preclinical animal models and is aided by an ability to study intra-graft immune cell trafficking in real-time. Intravital two-photon microscopy can be used to image tissues and organs for depths up to several hundred microns with minimal photodamage, which affords a great advantage over single-photon confocal microscopy. Selective use of transgenic mice with promoter-specific fluorescent protein expression and/or adoptive transfer of fluorescent dye-labeled cells during intravital two-photon microscopy allows for the dynamic study of single cells within their physiologic environment. Our group has developed a technique to stabilize mouse lungs, which has enabled us to image cellular dynamics in na&#239;ve lungs and orthotopically transplanted pulmonary grafts. This technique allows for detailed assessment of cellular behavior within the vasculature and in the interstitium, as well as for examination of interactions between various cell populations. This procedure can be readily learned and adapted to study immune mechanisms that regulate inflammatory and tolerogenic responses after lung transplantation. It can also be expanded to the study of other pathogenic pulmonary conditions.

Author Spotlight: Advancing Lung Transplant Immunology Through Intravital Imaging

Here, we present a protocol to intravitally image the transplanted mouse left lung using two-photon microscopy. This represents a valuable tool for studying cellular dynamics and interactions in real-time following murine lung transplantation.

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by ...

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

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