Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

All the protocols and methods used in animal subjects have been approved by Institutional Animal Care and Use Committees (IACUC) of Taipei Medical University (Protocol numbers: LAC-2013-0278; LAC-2015-0397)

1. Primary Neuron Culture

1. Preparation before culture
   1. Coat the culture dishes with poly-D-Lysine (Table 1) one day before culture. For a 6-well dish, add 1.5 mL poly-D-Lysine (0.05 mg/mL) into each well. For live-imaging, culture cells on a 12 mm x 12 mm coverslip in a standard coated 6-well plate.
   2. Remove/recycle the poly-D-lysine and wash the dishes twice with 2 mL ddH₂O.
   3. Prepare the surgical tools: a pair of micro-dissecting scissors, curved-tipped forceps, and standard straight-tipped forceps. Sterilize the surgical tools and keep them in 75% ethanol during surgery.
   4. Fill 10 cm petri dishes with 20 mL wash medium (Table 1) and keep on ice.
   5. Fill a 15 mL test tube with 15 mL wash medium and keep the tube on ice.
2. Brain tissue isolation from different brain regions
   1. Sacrifice 16-19 day old pregnant female mice with standard euthanization protocols by placing mice in a non-ventilated cage and then intoxicating with CO₂. E15.5~E16.5 embryos are recommended for hypothalamic neuronal culture and E16.5~E17.5 are more suitable for hippocampal and cortical neuronal culture.
   2. Sterilize the platform, dissecting microscope, and surgery tools with 75% ethanol to avoid contamination.
   3. Cut around the umbilical cord area (dark red area, Figure 1A, 1B dash line) to isolate the pups easily without damaging them (Figure 1C, 1D).
   4. Remove the head portion of each pup with a pair of scissors (Figure 1E, F), then secure the head portion in place using the fine-tipped straight forceps for the eye region (Figure 1G). Use another fine-tipped curved forceps to remove the outer skin and skull from two sides and peel them off from the anterior to the dorsal direction (Figure 1H, black arrow points to the direction). The brain should be isolated without any damage (Figure 1I).
      NOTE: It is essential to maintain the integrity of the brain to ensure accuracy when isolating different regions of the brain.
   5. Keep isolated brains in a clean petri dish with ice-cold wash medium for the following steps.
   6. Flip the brain and keep the ventral side up. The hypothalamus is a round structure in the middle of the brain (Figure 1J, arrow points to the hypothalamic region). Isolate the hypothalamic tissue with fine-tipped curved forceps. Remove the meninges (which have a yellowish-red color) carefully.
      CAUTION: Complete removal of meninges is a critical step to avoid fibroblast contamination.
   7. Hold the olfactory lobe with the sharp forceps and peel off the thin meninges surrounding the whole brain with the curved forceps (Figure 1K, L).
   8. The hippocampus is a banana-like shape which is embedded in the lower part of the cortex. Flip to the underside of the cortex and separate the hippocampus from the cortex by pulling it to the side with the curved forceps (Figure 1M, N, and O). Be sure to remove all the remaining meninges surrounding the brain tissue.
   9. When all regions of the brain have been isolated, chop these tissues in the 15 mL tubes containing ice-cold wash medium with the help of the forceps (step 1.1.5).
      NOTE: Brain tissues can be kept in the ice-cold wash medium for 2-3 h.
3. Tissue digestion, plating and culture
   1. Rinse the tissues by inverting the tissue-containing 15 mL tube 2-3 times, and then keep the tube straight to allow the tissues to settle down (1-3 min).
   2. Remove the upper medium using the glass Pasteur pipette attached to a vacuum system. To wash the tissue, add 15 mL ice cold wash medium and invert the tube 2-3 times. Repeat wash steps 3 times before the next step.
      CAUTION: Be cautious of the tissues on the bottom when removing the upper medium using a vacuum system.
   3. After the final wash, remove the wash medium with the help of a 1 mL pipette.
   4. Incubate tissues with Papain-Trypsin digestion buffer (Table 1) in a 37 °C water bath for 7-14 min. Shake the tubes every two min to ensure all tissues are properly exposed to the digestion buffer.
      NOTE: Incubation time and the volume of digestion buffer can be adjusted based on the tissue size. For hypothalamic tissue collected from 6-8 pups, 300 µL Papain-Trypsin digestion buffer and 7 min incubation time is recommended. More tissue will require more digestion buffer.
   5. Neutralize the enzyme's activity by adding 1 volume of fetal bovine serum. Shake the test tube 3-5 times at room temperature.
   6. Keep the tube on racks and wait for 1-2 min to allow the tissues to settle down; remove the supernatant carefully with a 1 mL pipette.
   7. Add 6 mL plating medium (Table 1) into the test tube and pipet the tissue up-and-down 50 times with a 5 mL pipette gently (for a 6-well plate, 1.5 mL plating medium per well is recommended). Most cells will dissociate into single cells after repeated pipetting.
      NOTE: Try to avoid the formation of bubbles while conducting this step. Most laboratories use flamed glass pasture pipets to dissociate the tissue. A 5 mL pipette with a narrow tip can be a good alternative for this step.
   8. Keep a tube on the rack and wait for 1-2 min to allow the chunky, un-dissociated tissues to settle down. Take the upper phase containing dissociated cells to a new tube and dilute with plating medium (5x volume plating medium per 1x volume of dissociated cells. For example, add 5 mL plating medium for 1 mL dissociated cells).
   9. Calculate the cell density using a hemocytometer and seed the required number of cells into the culture plate coated with poly-D-lysine as in step 1.1.1. We recommend 2-4 x 10⁵ cells per well for a 6 well dish/35 mm dish were recommended for neuron imaging studies. A higher density would be needed for protein collection and analysis.
      NOTE: Do not add too much plating medium, 1-1.5 mL plating medium in one 6-well dish is sufficient for attachment. An excessive amount of medium will prolong the time required for proper cell attachment.
   10. Wait for 2-3 h and check the seeded neurons under the microscope. Neurons will start to grow neuritis when they attach properly.
   11. Remove the plating medium and add 2 mL warmed wash medium (37 °C) into the dish to wash off the plating medium. Repeat this step twice.
   12. Add 2 mL complete culture medium (Table 1) into each well in the 6-well dish.
   13. Change half of the medium every 3-4 days. Prepare the complete medium freshly each time. Neurons cultured from different mouse brain regions will have different morphology and characteristics (Figure 2).

2. Transfection of Plasmid DNA into Primary Neuron with Liposome System

CAUTION: For neuron transfection, an endotoxin-free plasmid DNA purification kit (Materials Table) is recommended for DNA preparation. Additional ethanol precipitation can remove excess solvent and enhances DNA concentration.

1. Liposome-based DNA transfection in primary neurons.
   NOTE: The transfection time depends on the maturation status required in experiments. The hypothalamic neurons were transfected at DIV 7-10 (DIV, days in vitro).
   1. DNA: Liposome mixture preparation: Dilute GFP-GLUT4 plasmid DNA¹⁶ (2 µg) in a microcentrifuge tube containing 300 µL low serum culture medium. Prepare another microcentrifuge tube with 2 µL liposome in 300 µL low serum culture medium, and incubate for 5 min at room temperature.
   2. Combine the contents of both tubes and incubate for 20-30 min at room temperature.
   3. Remove the neuron culture medium from the culture dish. Add 600 µL DNA-liposome mixture into each well and incubate at 37 °C for 4-6 h.
   4. After 4-6 h, remove DNA-liposome mixture and add 2 mL culture medium.
   5. Fluorescent signals, such as GFP alone and GLUT4 protein conjugated with GFP tag (Figure 3 and Figure 4), can be observed after 18-72 h.

3. Live Image Recording

1. Preparing Cells for Live Imaging
   1. Culture WT and CCR5^-/- hypothalamic neuron cells expressing green fluorescence protein labeled glucose transporter 4 (GFP-GLUT4) on the #1.5 (or 0.17mm thickness) glass coverslip which has been pre-coated with poly-D-Lysine in step 1.1.1 in the 6-well plate.
   2. Carefully remove the coverslips with neurons using forceps and place it on the glass slides with caution.
   3. Remove excess medium/liquid with delicate task wipes. Fold the wipes twice, and carefully place them on top of the coverslip. Gently press down the wipes without moving the coverslip.
      CAUTION: Do not press the coverslip against the glass slide forcefully. The purpose of this step is to ensure that the coverslip does not move/drift during image acquisition caused by buoyant force.
   4. Place coverslips and slides on the deconvolution microscope stage, with the coverslip facing down, and secured properly. This step is to ensure that the slide position remains constant so users can track the same set of target cells later.
   5. Observe WT and CCR5^-/- hypothalamic neuron cells with a 60x/1.42 NA oil immersion objective lens.
   6. Treat the selected cell samples with CCL5 (10 ng/mL) or insulin (10 U/mL) for one min. Add 1.5 µL of diluted insulin on the edge of coverslip.
   7. Visualize and record the selected cell samples immediately. Program the video recording software to record for 30 min.
2. Deconvolution microscopy and analysis
   NOTE: This part of the protocol requires the use of a deconvolution microscope and specialized software for analysis.
   1. Turn on the power of the imaging system, allow microscope stage to initialize properly, and then turn on the LED light source.
   2. Add immersion oil (refractive index 1.520 for live samples at 37 °C) on a 60x 1.42 NA objective lens. Place the sample slide on the microscope with the coverslip facing toward the objective lens, and secure the slide properly.
   3. Use bright-field or fluorescence illumination to identify target cells. Adjust the focus until target cells can be clearly observed. Do not move the objective lens outside of the coverslip area to avoid unnecessary scratch marks.
   4. Identify green fluorescence protein conjugated Glucose Transporter 4 (GFP-GLUT4) by the GFP signal. Identify desired target cells for image acquisition. Selected target cell position can be memorized for future reference (Figure 4).
   5. Setup proper experimental parameters (including pixel number, excitation wavelength, transmission percentage, exposure time, stack thickness, time interval, and total imaging time) on each target cell. For this experiment, the image pixel number was set at 512 x 512 (it can be set at 1,024 x 1,024 for higher resolution) for GFP signals. The exposure time was set between 0.025 to 0.05 s for every 5 min. Minor lateral x, y, and z adjustments can be controlled by the recommended software (Materials Table).
   6. Set the exposure parameter to approximately 2,000 to 3,000 counts to achieve maximum pixel intensity. To minimize fluorescence photobleaching, reduce the percentage of excitation light transmission as much as possible while keeping exposure time less than 1 s. Repeat these steps for each additional fluorescence channel(s) and each individual area of interest.
   7. Set the upper and lower limit of the Z-stack on each target cell. This can be achieved by moving the microscope stage until the top and bottom of the target cell are both slightly out of focus. The users can adjust the resolution of the Z-axis by setting the number of images between the upper and lower limit (which can be done by setting the distance between each image).
   8. Stacks of images were deconvolved and later analyzed with the help of the respective software – Velocity from PerkinElmer in this case.