Visualizing Astrocyte Morphology Using Lucifer Yellow Iontophoresis

The animal experiments in this study were performed in accordance with the National Institute of Health Guide for the Care and Use of Laboratory Animals and were approved by the Chancellor's Animal Research Committee at the University of California, Los Angeles. Adult mice (6−8 weeks old) of mixed gender were used in all experiments.

1. Solution Preparation

1. Artificial cerebrospinal fluid (ACSF) solution
   1. Prepare fresh ACSF solution (135 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 14.7 mM NaHCO₃, 11 mM D-glucose, 1.25 mM Na₂HPO₄, and 2 mM CaCl₂) before each experiment. Add MgCl₂ and CaCl₂ in the final step. Dissolve the components in high quality deionized water.
   2. Incubate the ACSF solution at 35 °C in a water bath and bubble with 95% O₂/ 5% CO₂ for at least 30 min before the experiment.
   3. Add 2% lidocaine hydrochloride to reach a final concentration of 0.02% in ACSF (in 100 mL).
2. Fixative solution
   1. Use 10% formalin buffered phosphate.
3. LY dye solution
   1. Prepare 1.5% LY dye solution by dissolving LY CH dilithium salt or LY CH dipotassium salt in 5 mM KCl. Vortex thoroughly. A volume of 1 mL is sufficient for several experiments.
   2. Centrifuge for 10 min at 16,800 x g. Then, filter the supernatant with a 0.2 µm syringe filter into a new tube. Aliquots can be kept at 4 °C for up to 3 months.
      NOTE: It is critical to centrifuge and filter the dye solution to prevent aggregation of the dye particles, which may clog the electrode.

2. Mouse Transcardial Perfusion and Brain Dissection

1. Preparation of equipment and mouse for transcardial perfusion
   NOTE: C57/BL6 mice of either sex can be used. It has been empirically observed that for the method to work reliably, it is important that the mice are not older than 3 months (6−8 weeks old is ideal). The perfusion protocol is described below. An additional reference is provided for more details¹⁴.
   1. Clear perfusion tubing with ACSF solution to remove any air bubbles in the line. Set up surgery tools in order of use (tweezers, curved, blunt scissors, iris scissors).
   2. Deeply anesthetize mouse by putting it into an isoflurane induction chamber, after adding 2−3 mL of isoflurane to the chamber. Allow 1−2 min for anesthetic to take effect. Ensure that breathing does not stop.
   3. Test for toe pinch reflex. Proceed when the mouse is unresponsive to pain stimulus and the reflex is absent. Secure the animal in the supine position with the head placed in the breathing cone with a supply of 5% isoflurane in oxygen and the limbs and tail taped down inside a chemical fume hood.
2. Transcardial perfusion
   1. Using tweezers, lift up skin beneath rib cage. Make a 5−6 cm incision with the curved, blunt scissors through the skin to expose the abdominal cavity. Cut upwards through the abdominal wall until liver and diaphragm are visible.
   2. Gently move the liver away from the diagram. With iris scissors, make a 3−4 cm lateral incision in the diaphragm.
   3. Cut through the rib cage along the both sides of the body to expose the chest cavity. Be careful not to damage heart and avoid the lungs. Lift the sternum up to expose the heart.
   4. Once the heart is visible, inject 0.05 mL of heparin sodium solution (1,000 USP per mL) into the left ventricle as an anticoagulant. Then, insert perfusion needle carefully into the left ventricle. Make sure that the needle stays in the ventricle and does not pierce through to other heart chambers. Make a small incision in the right atrium with iris scissors.
   5. Perfuse with ACSF solution at a rate of approximately 10 mL/min until the fluid existing the body is cleared of blood (1−2 min).
   6. Switch from ACSF solution to fixative solution without introducing any air bubbles. Perfuse with fixative solution for 10 min at 10−20 mL/min.
      CAUTION: Take care that no fixative solution is draining from the nose. This indicates that the needle reached the right ventricle and the fixative is traveling to the lungs rather than through the systemic circuit to the rest of the body.
3. Dissection of the brain
   1. Remove the head of the mouse with scissors and carefully dissect out the mouse brain from the skull. Place in fixative solution for 1.5 h at room temperature for a short post-fixation period.
      NOTE: A good perfusion is necessary for successful LY iontophoresis. Brain should be white-colored and absent of blood in brain vasculature. Body and limbs should appear stiff.

3. Slice Preparation

1. Preparation of hippocampal slices
   1. Wash the brain with 0.1 M phosphate buffered saline (PBS) at room temperature for 5 min. Then, dry off the brain with filter paper and remove the olfactory bulb and cerebellum with a sharp razor blade.
   2. Mount the brain on the vibratome tray using cyanoacrylate glue and fill the tray with PBS at room temperature. Cut hippocampal coronal sections of 110 µm thickness.
      NOTE: Adjust the setting of the vibratome to make sure that slices have the same thickness and even surface. For this experiment, a speed setting of 4.5 and a frequency setting of 8 were used, which are arbitrary settings on the instrument (Table of Materials). Users may need to experiment with the settings on other devices.
   3. Collect the sections from the tray and place in a dish of PBS on ice.

4. Electrode Preparation

1. Prepare a sharp electrode with appropriate resistance.
   1. Use a borosilicate glass single barrel electrode with filament (O.D. 1.0 mm, I.D. 0.58 mm). Pull an electrode on a micropipette puller (Table of Materials). Ideal electrodes filled with 1.5% LY in 5 mM KCl should have a resistance of 200 MΩ when placed into a bath of PBS.
      NOTE: Puller setting varies depending on the apparatus (type of machine used and puller filament). A higher heat setting usually gives longer and finer tips. For the micropipette puller used in this experiment, the settings were: heat: 317, pull: 90, velocity: 70, and delay: 70. A trough type filament was used.
   2. Store electrodes in a closed container to prevent dust from entering the tip. Keep electrodes elevated from the bottom of the box to prevent the tip from breaking.
2. Fill the electrode with LY dye solution.
   1. Place an electrode in vertical position with the tip facing downward. Pipette 1−2 µL of LY solution into the back of the electrode and wait for 5−10 min for the solution to move to the tip via capillary action.
   2. Gently secure the filled electrode in the electrode holder connected to a manipulator.
      NOTE: The silver wire of the electrode holder needs to be in contact with the LY inside the electrode. Adjust volume of the LY solution if needed, depending on the wire length.

5. Filling Astrocytes with Iontophoresis

1. Test the electrode.
   1. Place a brain slice gently into a glass bottom dish filled with 0.1 M PBS at room temperature. Hold the slice in place with a platinum harp with nylon strings.
   2. Ensure that the electrode is connected to a voltage source and place the ground electrode into the bath containing the brain slice.
   3. Move the objective to the brain region of interest.
   4. Lower the electrode into the solution. Under the bright field, move it to the center of the field of view and examine it carefully with the 40x water immersion lens to ensure that it appears clear and without debris or bubbles. If there is anything clogging the electrode tip, replace it with a new one.
      NOTE: Clogging is a concern for the 200 MΩ electrode. With centrifugation and filtration of the dye solution before every experiment, this should not be a frequent problem. However, because the tip of the electrode is small, tissue may sometimes become stuck in the opening. The authors have not found a way to prevent this, but it can be easily dealt with by simply using a new electrode.
   5. Observe the tip of the electrode under the confocal laser-scanning microscope with the 488 nm laser. Then, test dye ejection by turning on the stimulator at 12 V. Note a large fluorescent dye cloud around the tip of the electrode while the stimulator is on. If no or little dye is ejected upon voltage stimulation, replace the electrode.
   6. Under the bright field, slowly lower the electrode toward the slice stopping just above the surface.
2. Fill astrocyte with iontophoresis.
   1. Identify astrocytes 40−50 µm below the slice surface with the infra-red differential interference contrast (IR-DIC). Look for cells with elongated, oval-shaped somata about 10 µm in diameter. Once an astrocyte is chosen, move it to the center of the field of view.
      NOTE: A good cell for iontophoresis has clear, defined borders around it. Do not choose cells too close to the surface of the slice because they cannot be completely filled. It takes some practice over a few days to be able to routinely identify astrocytes for dye filling. Depending on the brain area used for the experiment, the number of astrocytes may vary. This may affect the time needed to identify an astrocyte before filling.
   2. Slowly lower the electrode tip into the slice, navigating through the tissue, until it is on the same plane as the cell body.
      NOTE: Move electrode slowly to avoid damaging the tissue.
   3. Once the cell body of the astrocyte is clearly visible and outlined, slowly and gently advance the electrode forward. Move electrode until the tip impales the soma of the cell. Move the focus of objective slowly up and down to note if the electrode is inside the soma.
      NOTE: The electrode tip must be inside the cell body, and a small indentation on the soma should be observed. Do not move the electrode any further to avoid the tip going through the cell.
   4. Once the electrode tip is inside the cell, turn on the stimulator at ~0.5−1 V and continuously eject current into the cell. Using the confocal microscope, watch the cell fill. Increase the digital zoom to see the details of the cell and make sure that the tip of the electrode is visible inside the cell.
      NOTE: Lower the voltage if it appears that dye is leaking out of the cell or filling other cells in the vicinity. If it appears that dye is still leaking out of the cell, pull electrode out slowly and find another cell. It is important that the dye does not leak to have a high signal/ background ratio in the final image.
   5. Wait for about 15 min until the finer branches and processes appear defined, turn off the voltage and gently withdraw the electrode tip from the cell.
3. Image the filled cell.
   NOTE: Imaging can be performed immediately after filling or following staining with immunohistochemistry. An objective with a higher numerical aperture (NA) results in better resolution.
   1. Wait until the cell returns to original form before imaging (15−20 min) with 40x objective. To image the cell, adjust the setting on the confocal to make sure that the finer branches and processes appear defined.
   2. Set up a z-stack with a step size of 0.3 µm. While imaging, move the objective until there is no signal from the cell and set that as the top. Then, move the objective down (focusing through the cell) until there is no signal, set that as the bottom.
   3. After the imaging is complete, check the electrode for dye ejection. If a large dye cloud appears, it can be used for the next slice. Otherwise, replace it with a new electrode.
      NOTE: Dye filling of a single astrocyte and imaging takes about 45 min to 1 h. For this specific experiment, it was possible to obtain about 3−6 cells per mouse. If needed, multiple cells can be labeled on the same slice. However, to distinguish individual astrocytes, be sure to keep a distance of about 200 µm between cells.

6. Staining with Immunohistochemistry (Optional)

1. Once the imaging is done, immediately place the slice in 10% formalin on ice to preserve for immunohistochemistry. Keep brain slices in the dark and store overnight at 4 °C.
2. Wash brain sections 3x in 0.1 M PBS with 0.5% nonionic surfactant (i.e., Triton X 100) for 5 min each. Then incubate in a blocking solution of 0.1 M PBS with 0.5% nonionic surfactant and 10% normal goat serum (NGS) for 1 h at room temperature with gentle agitation.
3. Incubate the sections with agitation in primary antibodies diluted in 0.1 M PBS with 0.5% nonionic surfactant and 5% NGS for 2 days at 4 °C.
   NOTE: Because of the thickness of the brain slices, the incubation period of the primary antibodies needs to be extended for better penetration and the antibody concentration may be increased. In this experiment, a 1:500 dilution was used for anti GFAP antibody and a 1:500 dilution was used for anti aquaporin-4 antibody.
4. Wash the sections 3x in 0.1 M PBS with 0.5% nonionic surfactant for 10 min each and then incubate with secondary antibodies diluted in 0.1 M PBS with 0.5% nonionic surfactant and 10% NGS for 6 h at room temperature.
   NOTE: Do not choose a secondary antibody excited by 488 nm, which is the wavelength used to visualize the LY dye. In this experiment, a 1:1,000 dilution was used for Alexa Fluor 546 goat anti-chicken IgG (H+L) and Alexa Fluor 647 goat anti-rabbit IgG (H+L).
5. Rinse the sections 3x in 0.1 M PBS for 10 min each. Then mount the sections on glass microscope slides in mounting media suited for fluorescence. Seal slides. Image cells at a step size of 0.3 µm.