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

Stefan Hall; Sufyan Faridi; Irene Euodia; Sophie Tanner; Andrew Krzysztof Chojnacki; Kamala D. Patel; Juan Zhou; Christian Lehmann

doi:10.3791/63733

JoVE Journal > Immunology and Infection

Inmunología y contagio

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

Published: April 06, 2022

doi:

10.3791/63733

Stefan Hall¹, Sufyan Faridi², Irene Euodia^2,3, Sophie Tanner⁴, Andrew Krzysztof Chojnacki⁵, Kamala D. Patel^6,7, Juan Zhou^2,4, Christian Lehmann^1,2,4,8

¹Department of Physiology and Biophysics,Dalhousie University, ²Department of Microbiology and Immunology,Dalhousie University, ³Department of Neuroscience,Dalhousie University, ⁴Department of Anesthesia, Pain Management, and Perioperative Medicine,Dalhousie University, ⁵Live Cell Imaging Center,University of Calgary, ⁶Department of Physiology and Pharmacology,University of Calgary, ⁷Department of Biochemistry and Molecular Biology,University of Calgary, ⁸Department of Pharmacology,Dalhousie University

Summary

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.

Abstract

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.

Introduction

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.

Protocol

All the procedures described here were performed with prior approval by the Dalhousie University Committee on Laboratory Animals (UCLA).

1. Preparation

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.
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).
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 µm filter.
Ventilator: Set a small rodent ventilator to provide pressure-controlled ventilation at a rate and volume calculated based upon the mouse'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.
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.

2. Anesthesia

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.
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.
Tighten the suture to secure the snout inside the nose cone. Begin gas flow through the nose cone at a concentration of 2.5%.
Confirm adequate depth of anesthesia via toe pinch before moving to the next step.

3. Intubation

Rotate the stand such that the back of the stand and dorsal side of the mouse face toward the surgeon.
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.
Using blunt forceps, lift the lower jaw and displace the tongue to provide clear passage into the respiratory tract.
Insert a modified otoscope (~60° 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.
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.
Push the cannula along the fiber-optic cable, passing between the vocal cords and into the trachea.

4. Ventilation

Retrieve the mouse from the intubation stand and place it on a heating pad in the right lateral decubitus position.
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%.
Place tear gel in the mouse's eyes to prevent drying.
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'clock position. Extend the left hind paw caudally and secure at, approximately, the 6 o'clock position.
Using a cloth tape, lightly stretch the left forepaw to the 12 o'clock position and secure the other end of the tape to the top of the IVM platform as shown in Figure 2A (maintaining slight tension here facilitates subsequent thoracotomy).
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.
Once the temperature is stable at 37.0 °C ± 0.1 °C, proceed to perform the thoracotomy.

5. Thoracotomy

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.
With blunt forceps and straight scissors, make a small longitudinal incision near the bottom of the ribcage to expose the underlying muscle layer.
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.
Repeat this process dorsally until ~5 mm lateral to the vertebral column.
Moving cranially, use blunt dissection to expose the rib cage. Cauterize any exposed blood vessels to preserve hemodynamic stability.
Once the risk of blood loss has been mitigated, extend the incision from the xiphoid process to the axilla.
Repeat this process on the dorsal side of the incision until ~1 cm inferior to the left ear.
Using hemostatic forceps, grasp the dissected epithelial and adipose tissue and place clear of the surgical area (Figure 2B).
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.
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.
Extend the incision laterally along the intercostal muscle in both directions, taking care not to touch the exposed lung surface.
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.
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.
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).
Remove any excess fluid accumulation in the thoracic cavity via capillary action using small strips of gauze.
While proceeding to microscopy, allow ~5 min for intrapleural fluid to dissipate for a more secure interface between the lung and the imaging window.

6. Microscopy

Turn on the vacuum pump and adjust the pressure to ~50–60 mmHg.
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'clock position.
Using the micromanipulator, carefully lower the imaging window until it adheres to and stabilizes the lung surface (Figure 3A).
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.
1. Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.
2. Repeat the previous step until five pulmonary venules have been imaged.
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.
1. Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.
2. Repeat the previous step until five pulmonary arterioles have been imaged.
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.
1. Switch to the 530-550 nm bandpass excitation filter and record 30 s of video in the same field of view.
2. Repeat the previous step until five capillary regions have been imaged.

7. Euthanasia and cleaning protocol

Remove the IVM platform from the microscope stage and adjust isoflurane delivery to 5% for 5 min to euthanize the mouse.
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.
After 5 min has elapsed, stop the ventilator, and ensure complete euthanasia via cervical dislocation.

Representative Results

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 naïve 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 µm. 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 naïve ones (Figures 4C,D). This trend was recapitulated by arteriolar leukocyte adhesion, although both levels of rolling and adhesion were highly variable in the naïve 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 naïve 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.

Figure 1: Lung imaging system. Custom-order system includes (1) an anodized metal base, (2) imaging window, (3) vacuum inlet, (4) micromanipulator. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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) naïve and (B) LPS-treated mice. (C,D) Intranasal LPS administration impacts leukocyte rolling and adhesion in pulmonary venules. Values are given as mean ± SD. Please click here to view a larger version of this figure.

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) naïve and (B) LPS-treated mice. (C,D) Intranasal LPS administration impacts leukocyte rolling and adhesion in pulmonary arterioles. Values are given as mean ± SD. Please click here to view a larger version of this figure.

Figure 6: Adherent leukocytes in pulmonary capillaries. Excitation of rhodamine 6G permits visualization of leukocytes within ROIs in (A) naïve and (B) LPS-treated mice. (C) Intranasal LPS administration impacts leukocyte adhesion in LPS-treated mice. Values are given as mean ± SD. Please click here to view a larger version of this figure.

Figure 7: Pulmonary capillary function. Excitation of FITC-albumin permits visualization of capillary blood flow within ROIs in (A) naïve and (B) LPS-treated mice. (C) Intranasal LPS administration impacts FCD in pulmonary capillaries. Values are given as mean ± SD. Please click here to view a larger version of this figure.

Supplemental File: File for 3D-printable IVM platform. Please click here to download this File.

Supplemental Figure 1: Diagram of the vacuum system. Please click here to download this File.

Supplemental Movie 1: Sample video of blood flow in the pulmonary venule. The green arrow denotes the direction of the blood flow. Please click here to download this Movie.

Supplemental Movie 2: Sample video of leukocyte trafficking in the pulmonary venule. Red arrows denote adherent leukocytes within the vessel. Please click here to download this Movie.

Supplemental Movie 3: Sample video of blood flow in the pulmonary arteriole. The green arrow denotes the direction of the blood flow. Please click here to download this Movie.

Supplemental Movie 4: Sample video of leukocyte trafficking in pulmonary arteriole. Red arrows denote adherent leukocytes within a vessel. Please click here to download this Movie.

Supplemental Movie 5: Sample video of blood flow in pulmonary capillaries. Green arrows denote several well-visualized areas of red blood cell-perfused capillaries. Please click here to download this Movie.

Supplemental Movie 6: Sample video of leukocyte trafficking in pulmonary capillaries. Red arrows denote adherent leukocytes within the field of view. Please click here to download this Movie.

Discussion

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.

Divulgaciones

The authors have nothing to disclose.

Acknowledgements

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

Materials

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

Referencias

Alizadeh-Tabrizi, N., Hall, S., Lehmann, C. Intravital imaging of pulmonary immune response in inflammation and infection. Frontiers in Cell and Developmental Biology. 8, 620471 (2021).
Gaertner, M., et al. Toward a comprehensive interpretation of intravital microscopy images in studies of lung tissue dynamics. Journal of Biomedical Optics. 20 (6), 066009 (2015).
Kreisel, D., et al. In vivo two-photon imaging reveals monocyte-dependent neutrophil extravasation during pulmonary inflammation. Proceedings of the National Academy of Sciences of the United States of America. 107 (42), 18073-18078 (2010).
Looney, M., et al. Stabilized imaging of immune surveillance in the mouse lung. Nature Methods. 8 (2), 91-96 (2011).
Bennewitz, M. F., Watkins, S. C., Sundda, P. Quantitative intravital two-photon excitation microscopy reveals absence of pulmonary vasoocclusion in unchallenged sickle cell disease mice. IntraVital. 3 (2), 29748 (2014).
Blueschke, G., et al. Automated measurement of microcirculatory blood flow velocity in pulmonary metastases of rats. Journal of Visualized Experiments: JoVE. (93), e51630 (2014).
Tabuchi, A., et al. Precapillary oxygenation contributes relevantly to gas exchange in the intact lung. American Journal of Respiratory and Critical Care Medicine. 188 (4), 474-481 (2013).
Rodriguez-Tirado, C., et al. Long-term high-resolution intravital microscopy in the lung with a vacuum stabilized imaging window. Journal of Visualized Experiments: JoVE. (116), e54603 (2016).
Fiole, D., et al. Two-photon intravital imaging of lungs during anthrax infection reveals long-lasting macrophage-dendritic cell contacts. Infection and Immunity. 82 (2), 864-872 (2014).
Thanabalasuriar, A., Neupane, A. S., Wang, J., Krummel, M. F., Kubes, P. iNKT cell emigration out of the lung vasculature requires neutrophils and monocyte-derived dendritic cells in inflammation. Cell Reports. 16 (12), 3260-3272 (2016).
Neupane, A. S., et al. Patrolling alveolar macrophages conceal bacteria from the immune system to maintain homeostasis. Cell. 183 (1), 110-125 (2020).
Tschernig, T., et al. Direct visualisation of microparticles in the living lung. Experimental and Toxicologic Pathology. 65 (6), 883-886 (2013).
Mertens, M., et al. Alveolar dynamics in acute lung injury: Heterogeneous distension rather than cyclic opening and collapse. Critical Care Medicine. 37 (9), 2604-2611 (2009).
Matuszak, J., Tabuchi, A., Kuebler, W. M. Ventilation and perfusion at the alveolar level: Insights from lung intravital microscopy. Frontiers in Physiology. 11, 291 (2020).
Margraf, A., et al. 6% Hydroxyethyl starch (HES 130/0.4) diminishes glycocalyx degradation and decreases vascular permeability during systemic and pulmonary inflammation in mice. Critical Care. 22 (1), 1-12 (2018).
Roller, J., et al. Direct in vivo observations of P-selectin glycoprotein ligand-1-mediated leukocyte-endothelial cell interactions in the pulmonary microvasculature in abdominal sepsis in mice. Inflammation Research. 62 (3), 275-282 (2012).
Marques, P., et al. Cigarette smoke increases endothelial CXCL16-leukocyte CXCR6 adhesion in vitro and in vivo. Potential consequences in chronic obstructive pulmonary disease. Frontiers in Immunology. 8, 1766 (2017).
Condon, M. R., Kim, J. E., Deitch, E. A., Machiedo, G. W., Spolarics, Z. Appearance of an erythrocyte population with decreased deformability and hemoglobin content following sepsis. American Journal of Physiology-Heart and Circulatory Physiology. 284 (6), 2177-2184 (2003).
Trzeciak, S., et al. Early microcirculatory perfusion derangements in patients with severe sepsis and septic shock: Relationship to hemodynamics, oxygen transport, and survival. Annals of Emergency Medicine. 49 (1), 88-98 (2007).
Kim, Y. M., Jeong, S., Choe, Y. H., Hyun, Y. M. Two-photon intravital imaging of leukocyte migration during inflammation in the respiratory system. Acute and Critical Care. 34 (2), 101-107 (2019).
Faust, N., Varas, F., Kelly, L. M., Heck, S., Graf, T. Insertion of enhanced green fluorescent protein into the lysozyme gene creates mice with green fluorescent granulocytes and macrophages. Blood. 96 (2), 719-726 (2000).
Orthgiess, J., et al. Neurons exhibit Lyz2 promoter activity in vivo: Implications for using LysM-Cre mice in myeloid cell research. European Journal of Immunology. 46 (6), 1529-1532 (2016).
Schindelin, J., et al. Fiji: An open-source platform for biological-image analysis. Nature Methods. 9 (7), 676-682 (2012).
Tabuchi, A., Mertens, M., Kuppe, H., Pries, A. R., Kuebler, W. M. Intravital microscopy of the murine pulmonary microcirculation. Journal of Applied Physiology. 104 (2), 338-346 (2008).
Butt, Y., Kurdowska, A., Allen, T. C. Acute lung injury: A clinical and molecular review. Archives of Pathology and Laboratory Medicine. 140 (4), 345-350 (2016).
Park, I., et al. Neutrophils disturb pulmonary microcirculation in sepsis-induced acute lung injury. European Respiratory Journal. 53 (3), 1800786 (2019).
Kim, J. K., et al. In vivo imaging of tracheal epithelial cells in mice during airway regeneration. American Journal of Respiratory Cell and Molecular Biology. 47 (6), 864-868 (2012).
Pieper, M., Schulz-Hildebrandt, H., Mall, M. A., Hüttmann, G., König, P. Intravital microscopic optical coherence tomography imaging to assess mucus-mobilizing interventions for muco-obstructive lung disease in mice. American Journal of Physiology – Lung Cellular and Molecular Physiology. 318 (3), 518-524 (2020).
Lamm, W. J. E., Bernard, S. L., Wiltz, W., Wagner, J., Glenny, R. W. Intravital microscopic observations of 15-µm microspheres lodging in the pulmonary microcirculation. Journal of Applied Physiology. 98 (6), 2242-2248 (2005).
Entenberg, D., et al. A permanent window for the murine lung enables high-resolution imaging of cancer metastasis. Nature Methods. 15 (1), 73-80 (2018).
Amato, M. B. P., et al. Effect of a protective-ventilation strategy on mortality in the acute respiratory distress syndrome. New England Journal of Medicine. 338 (6), 347-354 (2009).
Looney, M. R., Bhattacharya, J. Live imaging of the lung. Annual Review of Physiology. 76, 431-445 (2013).

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Hall, S., Faridi, S., Euodia, I., Tanner, S., Chojnacki, A. K., Patel, K. D., Zhou, J., Lehmann, C. Intravital Widefield Fluorescence Microscopy of Pulmonary Microcirculation in Experimental Acute Lung Injury Using a Vacuum-Stabilized Imaging System. J. Vis. Exp. (182), e63733, doi:10.3791/63733 (2022).