Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

All animal protocols were reviewed and approved by the Animal Care and Use Committee at the University of South Carolina (assurance number: A3049-01).

1. Low-density Embryonic Neuronal Culture

1. Preparation for primary cortical neuron culture
   1. Solutions:
      1. Prepare Poly-L-Lysine stock solution by dissolving 5 mg of Poly-L-Lysine in 10 ml Borate buffer.
      2. Prepare working solution by diluting 1 ml Poly-L-Lysine Stock in 49 ml Borate Buffer.
      3. Prepare borate Buffer, pH 8.4 by starting with 395 ml dH₂O and then adding all ingredients (1.24 g boric acid + 1.9 g of Borax). Adjust pH to 8.4 by 1 M NaOH. Adjust to 400 ml and filter with 0.2 µm nylon membrane filter under hood.
      4. Prepare HBSS solution, pH 7.2 by starting with 445 ml dH₂O and adding all other ingredients (50 ml 10X HBSS Stock + 1.2 g HEPES). Adjust pH to 7.2 by 1 N HCl, bring to 500 ml and filter with 0.2 µm nylon membrane filter under hood.
      5. Prepare plating medium: DMEM/F12 with 10% fetal bovine serum. Filter with 0.2 µm nylon membrane filter under hood.
      6. Prepare complete growth medium: To 50 ml Neurobasal medium, add the supplements (500 µl GlutaMax (100X) + 500 µl Glucose + 1 ml B-27 (50X) + 500 µl of Antibiotic-Antimycotic solution (100X) + 100 µl 7.5% Sodium Bicarbonate. Filter with 0.2 µm nylon membrane filter under hood.
   2. Two days before culture:
      1. Coat twelve 35 mm glass-bottom dishes with 2 ml of Poly-L-Lysine working solution, and keep under the hood O/N.
   3. One day before culture:
      1. Empty coating agent from the dishes and rinse with ddH₂O. Allow dishes to dry for an hour under hood.
      2. Prepare plating medium (DMEM/F12 with 10% fetal bovine serum (10% of total solution). Pipette 2 ml medium into each dish, and incubate O/N at 37 ^oC, 5% CO₂.
         Note: It is important to avoid plastic culture dishes when culturing. Use of glass-bottom dishes (for inverted microscopes) or glass coverslips (for upright microscopes) yields sharp and clear images. Compared to a plastic-bottom dish, the glass-bottom dish avoids unwanted hues or glare during fluorescent microscopy.
2. Cell culture protocol:
   1. Put refrigerated HBSS Buffer (100-150 ml), 100 mm petri dishes, forceps, scissors, 50 ml tubes, and 15 ml tubes under UV light in hood.
   2. Euthanize pregnant rat with lethal inhalation of 5% sevoflurane under the hood outside of culture room.
   3. Sterilize with 70% EtOH, tent the skin of the lower abdomen using forceps while cutting from the tail up in the middle of abdominal cavity.Open the uterus using scissors to expose the placenta.
   4. Remove 8-10 E18 fetuses. Place 8-10 fetuses in petri dishes containing HBSS solution.
   5. Hold the back of the head of the fetus with forceps, then use scissors to sever the head from body. Place it in another petri dish filled with 5-7 ml of HBSS. Transfer dish to hood.
   6. Fill two additional 100 mm petri dish with cold HBSS.
   7. Peel aside skull, and scoop brain into a new petri dish filled with HBSS.
   8. Use sharp curved forceps, separate cerebellum and brain stem from brain, then transfer brain to a new petri dish with cold HBSS in it.
   9. Use tweezers to secure, separate the hemispheres with curved forceps. Remove meninges; isolate frontal cortex and transfer pieces into marked 15 ml centrifuge tubes.
   10. Fill the 15 ml tube with fresh 2 ml HBSS and add 20 µl Trypsin EDTA (10x concentrated mixture with 0.5% Trypsin and 5.3 mM EDTA.) to each 15 ml tube.
   11. Incubate 10-15 min at RT. Gently swirl tube every few minutes, do not allow settling.
   12. After 10-15 min, use glass pipette to remove old HBSS and rinse twice with new HBSS.
   13. Immerse in trypsin inhibitor (1 mg/ml, 2 mg Trypsin inhibitor in 2 ml HBSS), mix well and let it sit 5 min. Wash twice with fresh HBSS.
   14. Using glass pipette with rubber bulb, triturate brain pieces 10-15 times very slowly so as to avoid bubbles.Triturate again using a calibrated pipette and a sterile tip with a reduced tip diameter very slowly until homogeneous.
   15. Transfer dissociated cells at desired density to pre-prepared coated culture dishes (50 cells/mm2). Incubate O/N at 37 ^oC, 5% CO2.
   16. After 24 hr, replace DMEM/F12 medium with freshly made complete growth serum-free Neurobasal medium.
   17. Maintain the cells at all times in a 5% CO₂/95% room air humidified incubator for at least 12-14 days before experimentation. Supplement the cells every 5-6 days with fresh Neurobasal medium replacing approximately 50% of the old medium.

2. Fluorescent Labeling and Immunocytochemistry

Note: The immunofluorescent labeling of primary cortical cell cultures was carried out in glass bottom 35 mm cell culture dishes with a working volume of 1 ml.

1. Wash the glass-bottom dish twice with phosphate-buffered saline pH 7.4 (PBS).
2. Fix cells with 4% paraformaldehyde for 15 min at RT.
3. Wash cells twice with PBS and permeabilize with 0.1% Triton X-100 in PBS for 5 min.
4. Treat cells for 20 min at RT with an F-actin specific stain, AlexaFluor 488 Phalloidin (1:40, 25 µl of Phalloidin in 1 ml PBS).
5. Rinse cells twice with PBS and block with 10% normal goat serum at RT for 1-2 hr.
6. Dilute the chicken polyclonal anti-MAP2 antibody (1:2,500) in PBS with 2% normal goat serum and incubate O/N at 4 ^oC.
7. Following incubation, rinse in PBS twice.
8. Dilute the secondary antibody (Alexa Red 594-conjugated goat anti-chicken IgG (1:500) with 2% normal goat serum and incubate for 2 hr at RT.
9. Rinse with PBS and add 10 µl/dish of Hoescht dye.
10. Incubate 3 min at RT and wash with PBS twice.
    Note: The labeled cells now ready for step 3 (F-actin puncta counting).
11. Preserve cell sample with 100 µl of antifade reagent as a coverslipping agent and keep at 4 ^oC in the dark for long term storage.
    Note: Phalloidin is relatively stable in PBS at +4 °C in the dark for up to 1 week and fluorescence can be preserved with antifade coverslipping reagents. However, optimal images are obtained from 1-3 days after staining since Phalloidin may start to diffuse out with extended storage.

3. F-actin Puncta Counting

1. Turn on fluorescent microscope and switch to 20× objective. Open microscope software, and set up program at 1280×960 pixel image size, and 0.17 µm/pixel image resolution at 1× zoom.
2. Acquire images of co-labeled F-actin/MAP2 neurons under green (495 nm)/red (613 nm) fluorescent channels.
3. Choose 5 Green (F-actin)/Red (MAP2) immunolabeled/Blue (Hoescht) fluorescent images of individual neuron with clearly defined dendritic arbors.
4. Identify the F-actin rich structures in second order dendritic segment (length range 25-75 µm) with continuous MAP2 immunofluorescence.
5. Rotate the selected region of images to a horizontal level. Copy and paste as a new image.
6. Subtract the background of images, using a consistent adjustments for each dish.
7. Count the bright green F-actin puncta and trace the length of selected dendritic segment manually by trained independent observers. Export the data to a spreadsheet file.
8. Calculate the density by dividing total F-actin labeled puncta (N) by the length (L) of the MAP2 labeled dendrites. Express data as number of F-actin puncta per 10 µm of dendrite
   Note: Puncta (size ≤1.5 µm) of F-actin fluorescence with a peak intensity of at least 50% above the average intensity of staining in the dendritic shaft were included in each selected dendritic segment.