Fig. 1
Ventilation system/monitoring setup. A typical setup for intravital microscopy is indicated. Animals are supplied with medical oxygen and isoflurane via a volume-controlled animal ventilator. Physiological parameters are continuously checked via patient monitors and a thermocontroller. Fluid supply is ensured during the entire experiment by an infusion pump
3.4 Surgical Procedure: Spinal Cord Window Preparation
1.
Use a pet trimmer to shave off a patch of the animal’s fur at the level of the thoracic spinal cord. Be sure that no hair fragments remain at the desired spot since these will create strong autofluorescence. For sterile surgery use 70 % ethanol to disinfect the skin before proceeding and use autoclaved instruments.
2.
Perform a midline skin incision of 2–3 cm at the level of the thoracic spinal cord using straight fine iris scissors and straight forceps with serrated tips.
All the following steps should be performed by using a stereomicroscope with fiber-optic illumination for optimal specimen visualization during surgery.
3.
Dissect the paravertebral musculature (m. erector spinae) from the exposed vertebras using straight serrated tip forceps and straight spring scissors. Control bleeding with gauze. Be careful not to touch the intervertebral space.
4.
Fixate three contiguous vertebras laterally (at the level of the pedicle) with steel needles in a rigid custom-built frame. This frame can be directly mounted on the microscopy stage (Fig. 2a, b). For this procedure it may be helpful to use slim tissue forceps to carefully grab the spinous processes of the exposed vertebras. Ensure that the vertebras are properly fixated before proceeding.
Fig. 2
Preparation of a spinal cord window for intravital TPLSM. (a) Indicated is a LEW rat in prone position on a heated custom-built microscopy stage. The animal is intratracheally intubated and ventilated. Physiological conditions are assessed by a rectal thermoprobe connected to a thermocontroller (not shown) and an SpO2 sensor attached to the left leg. Three thoracic vertebras are fixated laterally by steel needles within a custom-built stereotactic frame. Fluid supply is ensured via an intravenous catheter at the tail. (b) Detailed view on a fixated, laminectomized thoracic vertebra with exposed spinal cord channel in the center. Since the dura (mater) spinalis is removed, the subarachnoideal blood vessels are visible. (c) Side view of an animal on the microscopy stage mounted on a motorized (x, y) table. A 20× water immersion objective is located above the spinal cord window
5.
Use a vibration-free dental micromotor with a round tungsten carbide bur to flatten the spinous process of the central vertebra. Be sure to frequently flush the region with 0.9 % isotonic sodium chloride solution to avoid heating during drilling.
6.
Create a ring of low-melting agarose (2.5 % in 0.9 % isotonic sodium chloride solution) around the flattened central vertebra. Be careful to ensure that the temperature of the agarose does not exceed 37–38 °C.
7.
Perform a laminectomy by using a dental micromotor with a fine round tungsten carbide bur to incise two sagittal grooves in a distance of at least 3 mm away from the lateral edges of the vertebra. Immediately afterwards, flush the region with 0.9 % isotonic sodium chloride solution to prevent dehydration of the tissue.
8.
Use medical forceps with curved, serrated tips to grip the far edges of the loosened bone flap. Be careful while peeling the latter away from the underlying dura (mater) spinalis. Carefully flush the exposed spinal cord with 0.9 % isotonic sodium chloride solution. In this way, the integrity of the agarose ring can be easily checked.
9.
Carefully remove the dura (mater) spinalis by using straight forceps with ultrafine tips. Be careful not to perforate the underlying arachnoidea (mater) spinalis, especially during tissue swelling due to (auto)immune inflammation (Fig. 2b). Control bleeding carefully with gauze without generating any pressure on the underlying tissue.
10.
Transfer the microscopy stage carefully onto a motorized (x, y) table and place a water immersion objective above the spinal cord window (Fig. 2c).
11.
Select a spot of interest under the microscope and start TPLSM. Clean the spinal cord window frequently by exchanging the isotonic sodium chloride solution. Ensure that animal parameters are reflecting physiological conditions during the entire experiment.
3.5 Two-Photon Laser Scanning Microscopy (TPLSM) for Monitoring the Behavior of Encephalitogenic GFP+ Effector T Cells Within the Lumen of Leptomeningeal Blood Vessels
Here we use an upright LSM710/Axio Examiner.Z1 microscope combined with a >2.5 W Ti:Sapphire Chameleon Vision II Laser device. The excitation wavelength of the laser is software-controlled and tunable from 690 to 1,040 nm. This allows a flexible handling according to the experimental setup. In the following a typical experimental design is described as an example.
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1.
For visualization of leptomeningeal blood vessels, inject a 400 μg/kg fluorescence-conjugated dextran (e.g., Texas Red dextran, 70,000 MW) intravenously directly before starting TPLSM.
2.
Start the acquisition software and tune the laser wavelength to 880 nm. Here, filter cubes are equipped with 442/46 nm, 525/50 nm, and 624/40 nm for simultaneous detection of collagen (by second harmonic generation), eGFP, and Texas Red dextran, respectively.
3.
Set the stack height by scanning through the tissue in z-direction. For time-lapse recordings, z-stacks should be kept as minimal as possible as the number of z-sections correlates directly with the scanning time and therefore with the temporal resolution of the image. Here, we typically use stacks between 100 and 150 μm, with an approximate single stack size of 4–6 μm. For high-quality imaging, the resolution of the single x, y planes should be at least 512 × 512 pixels (here, 424.27 × 427.27 μm). Thereby, the acquisition rate during bidirectional scanning is approximately 1.3 s per z-plane including two times line averaging. Importantly, for reproducible motility analyses, the interval time has to be kept at an exact value. We typically record 30 min time-lapse videos composed of 58 cycles and 32 s time intervals between two cycles.
4.
Start recording several spots. Ensure that the selected spots are comparable according to cell numbers, vessel size, etc.
5.
Obtain 2D movies by generating maximum intensity projections with a suitable software solution and store them typically as .avi or .tiff series for further processing, e.g., modulate brightness and contrast, stabilization, cell counting, etc.
6.
Analyze 4D (3D + time) raw data with a suitable software solution. Automated tracking modules can be used to trace individual cells within a 30 min recording interval. Note: according to your raw data, it might be essential to manually revise obtained trajectories.
7.
Calculate motility parameters (velocity, track duration, track length, direction of movement, etc.) from the coordinates obtained by the software (Fig. 3). We define vascular crawling cells as individual lymphocytes that move in an amoeboid manner strictly within the lumen of a leptomeningeal blood vessel (therefore, 4D analysis is essential!) and visible for more than two framaes. In contrast, rolling lymphocytes are defined as several round-shaped dots moving strictly with the direction of the blood flow as described (11). Correlation between both migratory phenotypes is performed by quantifying absolute numbers of crawling versus rolling cells within an observation period of 30 min (Fig. 3).
