This protocol describes in detail, the surgical preparation and use of a vacuum-stabilized system to image pulmonary leukocyte adhesion and capillary perfusion in vivo. The main advantage of this technique is that it allows stable high resolution intravital imaging of the intact mirroring lung with minimal physical trauma. To begin, prepare the imaging window by administering a thin layer of vacuum grease to the top of the outer ring while avoiding contamination of the vacuum channel.
Place a clean 8-millimeter glass cover slip on the window and gently press down to create a seal. Connect the imaging window to a vacuum pump fitted with a digital pressure gauge and capable of providing 50 to 60 millimeters of mercury-constant suction. Pass a length of fiber optic cable through a 20-gauge endotracheal cannula and submerge the tip in lidocaine HCL.
After anesthesia, insert a modified otoscope, such that the upper incisors fit within the gap in the speculum. Adjust the scope and tongue position until the epiglottis and the vocal cords are clearly visible. Insert the fiber optic cable through the gap in the speculum and use small circular movements to pass the cable between the vocal cords, then using the fiber optic cable as a guide, gently slide the cannula into the trachea to intubate the mouse.
After starting ventilation with 1.5%isoflurane, confirm the depth of anesthesia by testing for pedal reflex. Secure the cannula to the snout with medical tape. Use labeling tape to secure the right forepaw to the heating pad at the nine o'clock position and extend the left hind paw caudally and secure it at the six 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 intravital microscopy platform, then apply a rectal temperature probe and paw pulse oximeter to monitor the vital signs throughout the experiment. To prepare the mouse for surgery, sterilize the thorax and abdomen with a 70%alcohol wipe and apply light coat of mineral oil to dampen the hair on the left side of the mouse. Make a small incision near the bottom of the rib cage to expose the underlying muscle layer, then extend the initial incision and use blunt dissection to expose the rib cage.
Using hemostatic forceps, grasp the dissected epithelial and adipose tissue and place clear of the surgical area. To fluorescently label leukocytes and blood flow, administer a 2.5-milliliter per kilogram bolus of Rhodamine 6G and FITC-albumin via tail vein. Using toothed forceps, grasp the rib immediately inferior to the position of the base of the lung at end inspiration.
Slightly retract the forceps to pull the rib away from the lung and then cut the rib to induce a pneumothorax. Extend the incision laterally in both directions, taking care not to touch the lung. Grasp the next highest rib with blunt forceps 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 eventually 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 rib cage to expose the 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 the spinal column, then cut the adjoining rib. Moving cranially, and eventually, use a repeated pattern of cauterization and cutting to excise an approximately one-by-two centimeter portion of the ribcage. To prepare for imaging, transfer the intravital microscopy platform to the microscope stage and position the imaging window directly above the exposed lung.
Use the micro manipulator to carefully lower the imaging window until it adheres to and stabilizes the lung surface. To visualize the pulmonary microcirculation, use a wide field fluorescence microscope fitted with a 20x objective and standard FITC and Rhodamine filter sets. Use a black and white CCD camera to record the videos with optimal contrast.
Using the FITC filter set, identify a pulmonary venue based on the convergent pattern of blood flow and record a 30-second video with the venue in focus, then repeat the recording in the same field of view using the Rhodamine filter set to visualize leukocytes. To locate a pulmonary arterial, use the FITC filter set to identify a vessel with a divergent pattern of blood flow and record a 30-second video. Repeat the recording in the same field of view using the Rhodamine filter set to visualize leukocytes.
To locate the capillary regions of interest, use the FITC filter set to identify an area of alveoli and capillaries not intersected by larger vessels and record a 30-second video. Repeat the recording in the same field of view using the Rhodamine filter set to visualize leukocytes. This video depicts FITC imaging of blood flow within a pulmonary venue as indicated by the green arrow.
Rhodamine imaging permits visualization of leukocytes in the same venue with two adherent leukocytes indicated by the red arrows. These methods were also applied to visualize the same phenomena and pulmonary arterials and capillary regions of interest. To demonstrate the quantification of microcirculatory parameters, mice were treated with intranasal LPS to induce pulmonary inflammation.
Leukocyte trafficking in pulmonary venues is shown here with outlined areas representing the analyzed endothelial regions in naive and LPS-treated mice. Leukocyte adhesion and rolling were increased in LPS-treated mice. Leukocyte trafficking in pulmonary arterials is shown here with outlined areas representing the analyzed endothelial regions in naive and LPS-treated mice.
Leukocyte adhesion was higher in LPS-treated mice. This figure shows leukocyte adhesion within pulmonary capillaries in naive and LPS-treated mice. Leukocyte adhesion was shown to be higher in LPS-treated mice versus naive.
This figure shows perfusion of pulmonary capillaries in naive and LPS-treated mice. Functional capillary density was shown to be lower in LPS-treated mice versus naive. There are a lot of challenging aspects of this protocol.
Optimizing the mouse's positioning on the platform is a simple step that can make the surgical and imaging steps much more reproducible. This procedure is adaptable to a wide range of study parameters and microscopy approaches. Therefore, it should facilitate further research of pulmonary microcirculation in both healthy and diseased states.
