Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

1. Animals

1. All procedures were approved by the Institutional Animal Care and Use Committee at the University of Medicine and Dentistry of New Jersey.
2. Group house male C57BL/6 mice on a 12 hr light/12 hr dark schedule and allow ad libitum access to water and food. Sacrifice at 4-5 months old. Euthanasia of the mice was performed using the surgical plane of anesthesia and a secondary form of euthanasia (i.e. penetrating incision into the chest cavity via the diaphragm). This is consistent with the AVMA Guidelines on Euthanasia.

2. Preparation of Perfusion Solution, Coverslips, Glass Pipettes, and Media

1. Make 1L perfusion solution consisting of 2.5 mM KCl, 7 mM MgCl₂, 1.25 mM NaH₂PO₄, 28 mM NaHCO₃, 0.5 mM CaCl₂, 7 mM glucose, 1 mM ascorbate, and 3 mM pyruvate in distilled dH₂O. Adjust the osmolarity to ~300 mOsm using approximately 80 g/L sucrose. Oxygenate by bubbling with 95% O₂ /5% CO₂ and adjust the pH to 7.4. Each experiment will require approximately 200 ml of perfusion solution. Aliquots can be stored at -20 °C for up to 2 months.
2. Make 250 ml Neurobasal stock containing Neurobasal media without glucose, 2.5 mM glucose, and 100 U/ml penicillin streptomycin. Adjust the osmolarity of the Neurobasal stock to 280 mOsm using sucrose. Make 500 ml Hibernate stock containing Hibernate-A media without glucose, 2.5 mM glucose, and 100 U/ml penicillin streptomycin.
3. Stir both stocks well and filter-sterilize using Stericup vacuum filter units. Be careful not to introduce glucose or other contaminants through the glassware or stir bars. Wrap the bottles in foil to protect the media from light and store at 4 °C for up to 2 months.
4. Flame the tips of 4 autoclaved glass pipettes (9 in) so that the tips are no longer sharp and the opening diameters are large (~0.9 mm), medium-large (~0.7 mm), medium (~0.5 mm), and small (~0.3 mm). The largest should be barely polished and the smallest should be about a third of that diameter.
5. On the day before harvesting neurons, clean four 25 mm glass coverslips using 70% ethanol and passing over a Bunsen burner flame. Place each coverslip in a sterile 35 mm culture dish and add 2 ml sterile 1% poly-D-lysine in dH₂O. Put the dishes in an incubator (37 °C) overnight. The next day, wash with sterile dH₂O 3x and allow coverslips to fully dry.
6. Make 75 µl aliquots of 1 M lactic acid and 200 µl aliquots of GlutaMAX; store at -20 °C. Fully thaw aliquots of lactic acid and GlutaMAX before use to ensure homogeneity.
7. On the day of harvesting neurons, make 30 ml of fresh culture media consisting of Hibernate-A stock containing 1mM lactic acid, 0.5mM GlutaMAX, and 2% B27 without insulin. Vortex and adjust pH to 7.4 using 1 N NaOH.
8. Put 10 ml of culture media in a 60 mm glass Petri dish on ice, put 2 ml in a 15 ml conical tube on ice and keep the remaining media at room temperature.
9. On the day of harvesting neurons, make 25 ml of fresh growth media consisting of Neurobasal stock containing 1 mM lactic acid, 0.5 mM GlutaMAX, and 2% B27 without insulin. Vortex and adjust pH to 7.4 using 1 N NaOH. Allow the fresh growth media to equilibrate in an incubator (37 °C, 5% CO₂) for at least 30 min.

3. Cardiac Perfusion

1. Fully thaw 200 ml perfusion solution and oxygenate with 95% O₂/ 5% CO₂ on ice for at least 15 min.
2. Wash a vibratome blade with acetone, 70% ethanol, and dH₂O. Position the blade in the vibratome.
3. Rinse the vibratome cooling chamber and perfusion tubing with dH₂O. Fill the cooling chamber (4 °C) with perfusion solution and continue to oxygenate.
4. Anesthetize a mouse with pentobarbital sodium 50 mg/ml, diluted 1:1 with sterile water, injected intraperitoneally. Verify the mouse is fully anesthetized by pinching the tail and observing no response.
5. Pour cold perfusion solution into the perfusion reservoir and tubing just prior to dissection. Save 20-30 ml for brain dissection and continue to oxygenate on ice. Gravity flow from an elevated 60 ml syringe through tubing and a 20 G needle provides sufficient flow. Allow solution to run until air bubbles are washed out. Continue to oxygenate.
6. Cut back skin above the ribcage of the mouse. Lift the ribcage and carefully make a horizontal cut through the diaphragm. Make two lateral cuts through the ribcage up toward the arms to expose the heart. Use forceps to hold back the ribcage.
7. Make a small 2 mm cut in the right atrium to provide an exit point for blood to be washed out of the vascular system.
8. Puncture the left ventricle with the perfusion needle, just enough to insert the needle opening. Hold the needle in place with one hand and start the flow of perfusion solution with the other hand. After 10-15 ml of perfusion solution, the blood should be washed out and the effluent will clear.

4. Brain Slicing and Dissection

1. After cardiac perfusion, transfer oxygenated cold perfusion solution to a Petri dish on ice just prior to use.
2. Quickly decapitate, remove the brain from the skull and place the brain into perfusion solution. To remove the brain: pull skin forward, make a midline cut in the skull toward the eye sockets, make 2 lateral cuts toward each eye, use forceps to pull back each side of the skull, make a coronal cut between the eye sockets, and use a spatula to gently coax the brain out into perfusion solution. Use scissors to cut the optic tracts if necessary. Do not to touch the hypothalamic area and do not pull the optic tracts because this puts too much pressure on the median eminence and underlying hypothalamic tissue.
3. Use a razor blade and needle to laterally trim the brain tissue approximately 3 mm posterior and anterior to the hypothalamus while the tissue is submerged (bregma -3.52 mm). It is especially important for the anterior side cut to be level so that the brain will sit straight on the vibratome chuck.
4. Remove the remaining brain tissue from solution and use filter paper to wick away solution from the mounted brain section.
5. Place a dab of super glue on a vibratome chuck and spread the glue to the size of the brain. Put the brain tissue on top of the glue with the anterior side facing down and the hypothalamus facing the blade. Wick away excess glue and place the chuck into the vibratome cooling chamber. Use caution not to leave much glue as it will bubble up around the brain when placed into solution.
6. With the vibratome set to a slow speed (level 2) and high amplitude (level 9), make 300-500 µm slices until reaching the hypothalamic area. Now make thin 100 µm slices cutting from the posterior to the anterior. The visual cues are as follows. Posterior to the hypothalamus the third ventricle will be very small (bregma -2.70 to -2.30 mm). At bregma -2.30 mm, the third ventricle is separated into dorsal and ventral sections on the coronal slice.
7. Once sections of the third ventricle fuse, make two 500 µm slices which will contain the correct region of the VMH (bregma -2.18 to -1.22 mm). Anterior to the VMH, the third ventricle no longer extends to the ventral floor of the brain (bregma -1.06 to -0.82)⁸.
8. Transfer these slices to the Petri dish on ice containing culture media. Dissect the VMH (VMN + ARC) as shown in Figure 1. Use a pipette to gently transfer the VMH pieces in 2 ml of culture media on ice.

5. Dissociation and Culture

1. Remove the VMH from ice and place at room temperature. Add 20 U/ml papain to 4 ml of culture media. Invert to mix and place in a 34 °C water bath. Check and mix by inversion every minute until the digestion media is no longer cloudy. Filter (0.22 μm) into a sterile flask and transfer the tissue to the digestion media.
2. Digest the tissue by shaking at 100 rpm at 34 °C for 30 min.
3. Prepare 1 ml of media containing 8% bovine serum albumin (BSA) during the digestion. Dissolve 80 mg BSA in 1 ml of culture media and filter (0.22 μm) into a conical tube.
4. Wash the tissue by transferring to 5 ml of culture media in a conical tube and slowly inverting once.
5. Add 30 µl DNase enzyme to 6 ml of culture media and mix to make trituration media. Aspirate the wash media and add 3 ml of trituration media.
6. Use the glass pipettes to triturate in the order of largest to smallest. Gently triturate 10x and then wait 4 min for larger pieces to settle. Use the second pipette to transfer the top 2 ml containing a dissociated cell suspension to a new conical tube. Add 2 ml of trituration media, triturate 10x and wait 3 min. Use the third pipette to transfer the top 2 ml to the cell suspension. Add 1 ml of trituration media, triturate 5x and wait 2 min. Use the fourth pipette to transfer the top 2ml to the cell suspension.
7. Layer the dissociated cell suspension on top of the 8% BSA, being careful not to mix. Centrifuge at 1,000 rpm for 5 min.
8. Place autoclaved 6 mm x 8 mm cloning cylinders in the center of each coverslip. Coverslips must be completely dry. Aspirate and resuspend the pellet in 440 µl warm growth media. Fill cloning cylinders with the neuronal suspension and place in the incubator for 20 min.
9. Remove the cloning cylinders and very gently wash away debris. Fill each dish with 2 ml of growth media. Allow cells to recover for at least 1 hr before any assays are performed.

6. Preparation for MPD Imaging

1. Make recording solution consisting of 121 mM NaCl, 4.7 mM KCl, 2 mM CaCl₂, 0.1 mM MgCl₂, 1.2 mM MgSO₄, 0.97 mM K₂HPO₄, 0.23 mM KH₂PO₄, 5 mM NaHCO₃, and 25 mM HEPES. Adjust the pH to 7.4.
2. Add glucose to make the desired low glucose test solution (0.1-2.0 mM glucose). Mix thoroughly and reserve 400 ml low glucose recording solution. Add glucose to the remaining 600 ml to make 2.5 mM glucose recording solution and mix thoroughly.
3. Protect dye from light as much as possible. Add 4 ml of room temperature Buffer to a Blue MPD vial. Mix thoroughly and add 5 µl dye per ml recording solution. Keep solution homogenous by mixing with pipette in between solutions. The dye contains fluorescent molecules, which enter cells with depolarization, and quencher molecules, which cannot cross the cell membrane. Aliquots may be stored at -20 °C and used within 4 days. Do not freeze-thaw more than once.
4. Incubate cells in 2 ml of 2.5 mM glucose recording solution plus dye at 34 °C for 30 min. A dry heating block can be used. Protect from light.
5. Prepare syringe pumps and perfusion system with 2.5 mM and 0.1 mM recording solutions plus dye. Tubing from 60 ml syringes should be connected to a manifold.
6. After stopping any pump, wait for ~10 sec before switching solutions at the manifold to prevent pressure buildup in the syringe. Pressure changes alter fluid delivery to the closed chamber containing the neurons, causing changes in microscope focus during the experiment and distortion of images.
7. The manifold output tubing should be connected to an in-line heater and then polyethylene tubing , which connects to a closed chamber. Bubbles should be cleared and the perfusion rate should be set to 0.5 ml/min. Protect from light.
8. Transfer 25 mm coverslip with adherent neurons to the closed chamber, being careful not to allow coverslip to dry and not to introduce air bubbles. The slip is aligned in the grooves of the bottom piece, a couple drops of recording solution is placed along the sides, and the top piece is gently fitted to hold the coverslip in place.
9. Use a dropper to fill the chamber with recording solution and then place a 18 mm coverslip on top. Gently fit white ring into the chamber to hold the smaller coverslip in place. Press down very slowly and lightly to prevent air bubbles from being introduced into the closed chamber.
10. Start the syringe pump for the 2.5 mM glucose recording solution, press down on the white ring, and connect to the closed chamber. Slowly release pressure from white ring and connect to tubing leading to a waste container.

7. MPD Imaging

1. Center the neurons using low bright field light. Try to minimize the amount of time the cells are being exposed to light.
2. Allow the perfusion system to run for 10 min to allow for stabilization of temperature, focus, and dye equilibrium.
3. While priming, open the MetaMorph program and take a bright field image (10X) of the neurons. Use the Auto-Focus function to take 3 stacks of fluorescent images (150 msec, Narrow Cy3 filter, 10X) and determine the focus within 5 μm. At the end of the 10 min priming period, check the focus once more.
4. Begin to record fluorescent images every 30 sec for 40 min. Establish a baseline at 2.5 mM glucose for 10 min, then decrease glucose for 15 min, and finally return to 2.5 mM glucose for 15 min. Solutions are changed by stopping a syringe pump, waiting 10 sec, turning a port on the manifold, and starting another syringe pump. Changes should occur ahead of desired time points based upon the lag between the manifold and cells.
5. After recording all fluorescent images, take a bright field image of the neurons.

8. MPD Imaging Analysis

1. Use MetaMorph to create oval regions around each cell in the bright field image taken at the end of the experiment. Regions should be slightly larger than each neuron and can be made 1 pixel outside of the cell's dark edge. Do not choose cells that are unhealthy, overlapping, or close to debris. Unhealthy cells often appear dark. Lastly, create 5 large oval regions in areas with no cells or debris for background measurements.
2. Transfer the regions to all fluorescent images and inspect to verify that no movement/shift has occurred. Use the Measure function to record the fluorescence intensity of each region for all images in an Excel file. The use of Journals in Metamorph is advised.
3. In Excel, create an average of the 5 background regions for each fluorescent image. This represents the average background value at each time point. For each image/time point, subtract the background from all region measurements. The background-subtracted fluorescence intensity will be referred to as Intensity hereafter.
4. For each cell, calculate the average Intensity during minutes 2-8 of recording. This represents the Baseline of the cell in 2.5 mM glucose. A 2 minute buffer on both ends of each treatment is used to minimize noise.
5. For each cell, calculate the percent change from baseline: (Intensity-Baseline)/Baseline
6. Create a line graph with the percent change from baseline vs time.
7. Calculate the average percent change from baseline during minutes 18-23. Calculate the average percent change from baseline during minutes 35-40. While image processing is not used, noise is reduced through the averaging of intensity within each cell region, background subtraction, and the averaging of region intensity over 5 min or longer.
8. Control experiments in which 2.5 mM glucose recording solution is exchanged with 2.5 mM glucose recording solution must be performed to determine the noise threshold. Calculate the average percent change from baseline during minutes 18-23 for at least 6 dishes and 3 mice. Determine the mean and standard deviation of these values.
9. Set the threshold for a positive depolarization response to the mean plus 2 standard deviations. The mean plus 2 standard deviations corresponds to a difference with p<0.05. Our control experiments revealed a 10% change from baseline as the appropriate threshold value.
10. For each cell, assess the reversible depolarization response based upon 2 criteria. First, the average percent change from baseline during minutes 18-23 must be greater than 10%, the threshold determined in the previous step. Second, the average percent change from baseline during minutes 35-40 must be less than half of the average percent change from baseline during minutes 18-23. The second criterion was chosen since it was found to be more rigorous than stimulation with glutamate or KCl. Unhealthy neurons often respond to stimulation with glutamate or KCl even though their responses to decreased glucose are not reversible.
11. For each dish, calculate the % of depolarized neurons. This value is used to quantify the % of GI neurons among different treatment groups.