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 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.
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.
The protocol presented here requires practice and attention to a few critical steps. First, it is important to prepare the imaging window prior to initiating intubation and surgery. Use a minimal amount of vacuum grease to coat the outer ring of the imaging window, apply the cover glass, and test suction with a drop of distilled water. Preparing this in advance will prevent the exposed lung from drying out during the setup otherwise. While it is possible to flush with warm saline, doing so may risk damaging the fragile p…
The authors have nothing to disclose.
The authors would like to thank Dr. Pina Colarusso, who provided significant expertise in the editing and revising of this manuscript.
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 |