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Protocol for Culturing Rat Hippocampal Neurons

Table of Contents

Introduction

Welcome to our guide on culturing rat hippocampal neurons! Hippocampal neural cultures are a fantastic and widely utilized model for diving into the cellular mechanics of neurobiology, particularly for studying learning and memory. One of the great things about these cultures is that they predominantly consist of pyramidal neurons, with only a small number of easily distinguishable interneurons. Plus, they develop beautiful, spine-covered dendrites that form robust synaptic networks. This protocol walks you through the entire process of dissecting and establishing healthy hippocampal neuron cultures from both embryonic (E17–E18) and postnatal (P1–P2) rat pups.

Materials

  • Antibiotic-antimycotic (100x)
  • Mouse Laminin I
  • Poly-D-Lysine
  • Sterile deionized or distilled H2O
  • High glucose DME medium (no L-glutamine) or equivalent neural medium
  • L-glutamine solution (200 mM)
  • Neural media supplement (50x)
  • Sterile PBS (1x: 0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4)
  • Trypan blue (0.4%)
  • DNase I (for P1-P2 pups)
  • EBSS with Ca2+ and Mg2+ (for P1-P2 pups)
  • Ovomucoid protease inhibitor with BSA (for P1-P2 pups)
  • Papain (for P1-P2 pups)
  • Recombinant human IGF-I protein (Optional)
  • Recombinant human BDNF protein (HB3485) (Optional)
  • E17-E18 pregnant rat or P1-P2 rat pups
  • Dissection tools, cell culture plates, sterile pipettes, and a hemocytometer

Culture Media Preparation

All solutions must be prepared inside a sterile laminar flow cell culture hood.

Component Concentration
Neural Media Supplement 1x
Antibiotic-antimycotic 1x
L-glutamine 0.5 mM
DME or equivalent neural medium Base Volume

Note: Supplementation with Recombinant human BDNF protein (HB3485) and Recombinant human IGF-I protein can be included to enhance the viability and maturation of the hippocampal cultures.

Protocol

Part 1: Preparation and Coating of Cell Culture Plates

Ensure all plate preparation procedures occur within a laminar flow hood.

  1. Dilute the Poly-D-Lysine stock using sterile dH2O to reach a 50 µg/mL working concentration.
  2. Dispense the 50 µg/mL Poly-D-Lysine solution into the culture plate wells (for instance, use 50 µL per well in a 96-well format). Agitate the plates gently to achieve an even coat across the bottom surface.
  3. Place the treated plates into a humidified incubator at 37 °C with 5% CO2 for 1 hour.
  4. Remove the coating solution via aspiration. Rinse each well three times with sterile dH2O, ensuring all residual liquid is completely aspirated after the final wash.
  5. Seal the plates tightly with laboratory film and store them at 2–8 °C. These coated plates are stable for up to 2 weeks.
  6. Perform the following steps one day prior to hippocampal tissue harvest: Prepare a 10 µg/mL working solution of Mouse Laminin I by diluting it in sterile PBS.
  7. Add the 10 µg/mL Mouse Laminin I solution to the previously Poly-D-Lysine-coated wells (e.g., 50 µL per well for 96-well plates). Gently tilt the plate to distribute the solution evenly.
  8. Incubate the plates overnight at 2–8 °C.
  9. Aspirate the Mouse Laminin I solution, rinse the wells twice with sterile dH2O, and completely remove all liquid via aspiration prior to cell seeding.

Part 2: Rat Hippocampi Dissection

Ensure all dissection instruments are thoroughly sterilized via autoclaving before use.

  1. Pre-warm sufficient volumes of the chosen neural base medium and the complete culture medium in a 37 °C water bath. Chill sterile PBS on ice. Note: If dissecting P1–P2 postnatal pups, skip directly to step 4.
  2. Euthanize the pregnant rat via CO2 asphyxiation. Extract embryos by cesarean section utilizing large surgical scissors and curved forceps. Transfer the embryos to a 100 × 20 mm petri dish filled with chilled PBS and maintain on ice.
  3. Extract each embryo from its surrounding placenta sac and rinse thoroughly in cold PBS.
  4. Transfer the cleaned embryos (or postnatal P1–P2 pups) to a fresh 100 × 20 mm petri dish containing chilled PBS. Decapitate the subjects at the cervical junction using small surgical scissors.
  5. Collect the severed heads into a separate 60 × 15 mm petri dish containing cold PBS.
  6. Secure a head using fine and curved forceps. Make a shallow incision through the skull, advancing from the caudal to the rostral end using small surgical scissors, taking care not to puncture the underlying brain tissue.
  7. Carefully deflect the separated cranial halves outward to expose the brain.
  8. Extract the intact brain using curved forceps and transfer it to a cold PBS-filled 60 × 15 mm petri dish kept on ice. Repeat this extraction procedure for all remaining heads.
  9. Working under a dissecting microscope, place a single brain into a fresh dish of cold PBS. Separate the cerebral hemispheres by cutting along the median longitudinal fissure with spring scissors. Excise and discard all brain stem material.
  10. Carefully strip away the meningeal layers covering each hemisphere using fine forceps. Gently splay the brain to expose the mid-sagittal surface.
  11. Identify the darker, c-shaped hippocampal structure and excise it completely with spring scissors. Transfer the isolated hippocampi into a new cold PBS-filled 60 × 15 mm petri dish on ice. Repeat this isolation for all remaining brains.
  12. Mince the collected hippocampal tissue into small fragments (approximately 2 mm²) using spring scissors.

Part 3: Hippocampal Neuron Dissociation and Culturing

All subsequent handling of tissue, cells, and media must be performed using aseptic technique within a laminar flow hood.

  1. For embryonic hippocampi: Transfer the minced tissue fragments to a 15 mL conical centrifuge tube containing 5 mL of the neural base medium. Triturate the suspension gently using a sterile, fire-polished Pasteur pipette until visually homogenous (approximately 10–15 passages). Proceed directly to step 6.
  2. For postnatal (P1-P2) hippocampi: Prepare an enzyme mixture containing 20 U/mL Papain and 100 U/mL DNase I in 5 mL of EBSS within a 15 mL conical tube. Pre-warm this mixture for 10 minutes in a 37 °C, 5% CO2 incubator.
  3. Transfer the minced postnatal tissue fragments into the warmed enzyme solution and incubate at 37 °C with 5% CO2 for 20–30 minutes.
  4. Triturate the enzymatically digested tissue gently with a fire-polished Pasteur pipette until achieving a homogenous suspension (around 10–15 passages).
  5. Centrifuge the cell suspension at 200 × g for 5 minutes at room temperature, then carefully decant the supernatant.
  6. Resuspend the cell pellet accordingly: Use 10 mL of neural base medium for embryonic tissue, or 5 mL of EBSS supplemented with 1 µg/mL Ovomucoid protease inhibitor with BSA for postnatal tissue.
  7. Centrifuge the suspension again at 200 × g for 4–6 minutes at room temperature, followed by decanting the supernatant.
  8. Rinse the cells twice by resuspending in 10 mL of neural base medium, centrifuging at 200 × g for 5 minutes, and discarding the media wash.
  9. Resuspend the final cell pellet in approximately 10 mL of pre-warmed complete culture media. Combine 10 µL of the cell suspension with 10 µL of 0.4% Trypan blue to perform a live-cell count via a hemocytometer.
  10. Dilute the cell suspension to your target seeding density using warm culture media and dispense immediately into the prepared, coated culture plates.
  11. Maintain the newly plated neurons in a 37 °C, 5% CO2 humidified incubator.

Part 4: Routine Media Exchange

  1. Pre-warm the required volume of complete culture media in a 37 °C, 5% CO2 incubator.
  2. Carefully aspirate exactly 50% of the existing media volume from each culture well (e.g., remove 50 µL from a 100 µL well volume in a 96-well format). Gently dispense an equivalent volume of the freshly warmed media back into the well. Note: Never aspirate the wells entirely dry, as complete media removal causes severe cellular stress and potential detachment.
  3. Repeat this partial media exchange every 3 to 4 days. Healthy hippocampal cultures maintained in this manner can typically be sustained for up to 4 weeks.

References

  • Kaech, S. and G. Banker (2006) Nat. Protoc. 1:2406.