Increasing Durability of Dissociated Neural Cell Cultures Using Biologically Active Coralline Matrix

Summary

Dissociated hippocampal cell culture is a pivotal experimental tool in neuroscience. Neural cell survival and function in culture is enhanced when coralline skeletons are used as matrices, due to their neuroprotective and neuromodulative roles. Hence, neural cells grown on coralline matrix show higher durability, and thereby are more adequate for culturing.

Abstract

Cultures of dissociated hippocampal neuronal and glial cells are a valuable experimental model for studying neural growth and function by providing high cell isolation and a controlled environment. However, the survival of hippocampal cells in vitro is compromised: most cells die during the first week of culture. It is therefore of great importance to identify ways to increase the durability of neural cells in culture.

Calcium carbonate in the form of crystalline aragonite derived from the skeleton of corals can be used as a superior, active matrix for neural cultures. By nurturing, protecting, and activating glial cells, the coral skeleton enhances the survival and growth of these cells in vitro better than other matrices.

This protocol describes a method for cultivating hippocampal cells on a coralline matrix. This matrix is generated by attaching grains of coral skeletons to culture dishes, flasks, and glass coverslips. The grains assist in improving the environment of the cells by introducing them to a fine three-dimensional (3D) environment to grow on and to form tissue-like structures. The 3D environment introduced by the coral skeleton can be optimized for the cells by grinding, which enables control over the size and density of the grains (i.e., the matrix roughness), a property that has been found to influence glial cells activity. Moreover, the use of grains makes the observation and analysis of the cultures easier, especially when using light microscopy. Hence, the protocol includes procedures for generation and optimization of the coralline matrix as a tool to improve the maintenance and functionality of neural cells in vitro.

Introduction

Cultures of dissociated neural cells, in this case hippocampal cells, are a valuable experimental model for studying neural growth and function by providing high cell isolation and accessibility^1,2,3. This type of culture is frequently used in neuroscience, drug development, and tissue engineering due to the large amount of information that can be collected, such as rates of growth and viability, neurotoxicity, neurite outgrowth and networking, synaptic connectivity and plasticity, morphological modifications, neurites organization and wiring, etc.^1,4,5,6,7.

Despite the significance of the cultures, the cultivated cells are usually forced to grow on glass coverslips in a two-dimensional monolayer. These strict environmental modifications significantly decrease the ability of neural cells to survive over time, because glass coverslips are non-nurturing substrates with a low adhesion strength, exhibiting a lower capacity to support cell growth^8,9,10,11.

Because cultivated neural cells are forced to grow in challenging conditions, an essential approach to enhance their survival would be to imitate their natural environment as much as possible^12,13. This could be achieved by using biomaterials that will act as matrices and mimic the extracellular matrix of the cells, enabling them to form a tissue-like structure and assist in their nourishment¹⁴.

The use of biomaterials is a promising approach in improving cell cultures, because they act as biocompatible scaffolds, providing mechanical stability and enhancing a variety of cell properties, including adhesion, survival, proliferation, migration, morphogenesis, and differentiation^15,16,17. Several types of biomaterials are used to improve the conditions of the cells in vitro. Among them are biopolymers, or biological components that are usually part of the extracellular matrix of the cells. These biomaterials are mostly used as a form of polymerized coating agents or hydrogels^18,19,20. On the one hand, the matrices mentioned above give the cells a familiar 3D environment to grow in, encourage their adhesion to the dish, and give them mechanical support^21,22. On the other hand, their polymerized form and the confinement of the cells within hydrogels disturbs the access of the cells to nurturing components present in the growth media and also makes the follow up of the cells by microscopic methods more difficult²³.

Coral exoskeletons are biological marine-originated matrices. They are made of calcium carbonate, have mechanical stability, and are biodegradable. Previous studies using the coral skeleton as a matrix for growing neural cells in culture have shown much greater adhesion, compared to glass coverslips^24,25. In addition, neural cells grown on coral skeleton demonstrated their capability to intake the calcium the skeleton is composed of, which protects the neural cells in conditions of nutrient deprivation²⁶. Moreover, the coral skeleton is a supportive and nurturing matrix that increases the survival of neural cells, encourages the formation of neural networks, elevates the rate of synaptic connectivity, and enables the formation of tissue-like structures^27,28. Recent studies have also shown that the surface topography of the coral skeleton matrix plays a crucial role in the distribution and activation of glial cells^8,29. Also, coral skeleton is effective as a matrix for cultivation of other cell types, such as osteocytes^30,31, hepatocytes, and cardiomyocytes in culture (unpublished data).

Hence, coral skeleton is a promising matrix for cultivation of cells in vitro. Thus, the protocol detailed below describes the technique of cultivating neural cells on coral skeleton for producing more stable and prosperous neural cultures than those achieved by existing methods. This protocol may also be useful for cultivation of cardiomyocytes, hepatocytes, and other cell types.

Protocol

The use of animals in this protocol was approved by the National Animal Care and Use Committee.

NOTE: Calcium carbonated coral skeletons should be used in the crystalline form of aragonite. The coral types tested so far for neural cultures are Porites Lutea, Stylophora Pistillata, and Trachyphyllia Geoffroyi. The skeletons can be purchased whole or ground.

1. Cleaning the coral skeleton pieces

CAUTION: The following steps should be performed in a chemical hood at room temperature, because the solutions described below are hazardous and may cause burns and irritations.

1. Use a hammer to break the coral skeleton and divide it into 0.5–2 cm fragments. In order to dissolve organic and non-organic residues, soak the coral skeleton fragments in 10% sodium hypochlorite solution for 10 min, then wash once with double distilled water (DDW).
2. To remove the remaining organic residues, soak the fragments in a 1M NaOH solution for 5 min, then wash once with DDW. Continue with the removal of the organic deposits by soaking the fragments in 30% H₂O₂ solution for 10 min, then wash the fragments 3x with DDW.
3. Remove as much excess DDW as possible and leave the coral fragments in the hood to dry (1–8 h).

2. Cleaning the glass coverslips

1. Transfer the glass coverslips into a 100 mm glass Petri dish. Add 10 mL of 95% ethanol for 15 min.
2. Remove the ethanol and wash with DDW 3x, waiting 10 min between each wash. Place the dish on an 80 °C pre-warmed heating plate until the DDW evaporates. Gently stir the coverslips within the plate several times while drying to keep them from sticking to each other.
3. Autoclave the coverslips.

3. Preparation of coral skeleton grains

1. Grind the coral skeleton fragments using a mortar and pestle (manual grinding) until complete breakdown. The outcome is a mixture of grains with sizes ranging from 20 µm–200 µm.
2. Alternatively, grind the coral skeleton fragments using an electrical grinding machine at a velocity of 1,000 rpm for 30 s (blade length = 6 cm; width = 0.5 cm–1.0 cm). The resulting grain size is similar to the size range produced by the manual grinding.

4. Purification of grains of a specific size range

NOTE: If control over the size of the grains in a matrix is desired, use the following filtration-based grain purification procedure.

1. Transfer the grains onto a manual or electrical 40 µm filter mesh strainer.
2. Divide the grains into two specific ranges by sieving the grains through the strainer (if using an electrical strainer, the following conditions are recommended: shaking = 600 amplitudes/min, bouncing = 6/s). This procedure produces two groups of grain sizes, one <40 µm, and the second >40 µm.
   NOTE: The two sizes can be determined by using strainers of varying meshes. If a more restricted range is desired, then re-sieve each group from step 4.2 through strainers with different meshes.
3. Autoclave the grains.

5. Preparation of coral grain-coated dishes or coverslips

1. For coating flasks, plates, or Petri dishes
   NOTE: The following steps should be performed under sterile conditions.
   1. Add the coral skeleton grains into a 20 µg/mL poly-D-lysine (PDL) solution dissolved in Hanks' solution. The concentration recommended is 5 mg/10 mg of grains per 1 mL PDL solution.
   2. Pour the solution into flasks and dishes (approximately 2 mL/25 cm²) and incubate overnight at 4 °C. The grains sink and attach to the bottom.
   3. The next day, wash the flasks and dishes once with sterile DDW. Let the flasks and dishes dry in the hood.
      NOTE: It is preferable to use freshly coated flasks and dishes. The coated flasks and dishes can be used up to a week after coating if preserved at 4 °C. However, the effectiveness of the matrix may be compromised.
2. Coating glass coverslips (12 mm diameter, 0.17 mm thickness)
   1. Add coral skeleton grains to DDW at any desired concentration. The common densities used for neural cells are 5 mg/mL and 10 mg/mL. Pour 40 µL of the grain solution onto the center of the coverslip (Figure 4).
   2. Place the coverslips on a heating plate prewarmed to 80 °C and wait for complete evaporation (usually 15 min). Under these conditions, the grains adhere to the coverslip. Autoclave the coated coverslips. Store in sterile conditions.
   3. A day before culture, place the coverslips on the lid of a sterile 24 well plate. Add 100 µL of 20 µg/mL PDL solution to each coverslip. Use the tip to ensure that the liquid covers the entire grain region and the rest of the coverslip surface.
   4. Cover the lid with the bottom of the plate. Wrap the sides of the plates with paraffin film. Incubate overnight at 4 °C.
   5. The next day, wash once with DDW and dry in the hood.
      NOTE: It is preferable to use freshly coated coverslips.

6. Cultivation of hippocampal dissociated cells on coral skeleton grain-coated glass coverslips

NOTE: The method for hippocampal dissociated cell culture was modified from previously published procedures^24,27. The preparation of the culture is described for four rat pups. The expected yield from each hippocampus is 1–1.5 x 10⁶ cells.

1. Setup
   1. Prepare an empty 60 mm Petri dish. Pour 2 mL of minimal essential medium (MEM) into a 60 mm Petri dish and put on ice.
   2. Pour 2 mL of MEM into a 35 mm dish and put it in an incubator at 37 °C, 5–10% CO₂. Pour 2 mL of MEM into a 15 mL tube and keep it at 4 °C.
   3. Pour 1 mL of First Day Medium into a 15 mL tube and put it in an incubator at 37 °C, 5-10% CO₂.
      NOTE: The composition of the First Day Medium is described in the Table of Materials.
   4. Thaw 200 µL of a 2.5% trypsin solution and incubate at 37 °C.
   5. Prepare three glass Pasteur pipettes with 0.5 mm, 0.75 mm, and 1.0 mm diameter heads using a flame.
   6. Prepare adequate surgical tools: one big scissor, two small scissors, one big tweezer, four small tweezers, and one scalpel.
   7. Prepare a stereomicroscope in the hood.
2. Using a big scissor, sacrifice 0–3 day old Sprague Dawley rat pups by separating their heads from their bodies into an empty 60 mm Petri dish (step 6.1.1). Dispose of the bodies.
3. Pick up the head by holding it from the mouth with a big tweezer. Cut the skin covering the skull with a small scissor.
4. With a clean scissor, enter beneath the skull (on the side of the cerebellum) and cut the skull to the right and to the left to enable its removal. Using a small tweezer, peel the skull off from the brain.
5. With a clean tweezer, separate the brain from the head and put it in a 60 mm Petri dish containing 2 mL of cold MEM (see step 6.1.2).
6. Using two small tweezers, dissect the hippocampi under a stereomicroscope. Transfer the hippocampi into the prepared 35 mm dish (from step 6.1.2) containing MEM.
   NOTE: Steps 6.3–6.6 should be repeated for each pup.
7. Cut the hippocampi to approximately 1 mm slices using the scalpel. Add 200 µL of the trypsin solution (diluted to a final concentration of 0.25%). Mix gently and incubate at 37 °C, 5–10% CO₂ for 30 min.
8. After incubation, add 2 mL of cold MEM (step 6.1.2) to the dish to deactivate the trypsin. Transfer the trypsinized tissue to a 15 mL tube containing 1 mL First Day Medium preheated to 37 °C (step 6.1.3) using the glass Pasteur pipette with the largest diameter (step 6.1.5).
   NOTE: The best medium to tissue (v:v) ratio for further processing of the tissue is 1 mL/8 hippocampi. Avoid transferring the medium with the tissue as much as possible.
9. Triturate the tissue by passing it through the largest diameter glass pipette 10–15 times. Then, repeat this process using the glass pipette with the medium diameter. Continue triturating with the smallest diameter pipette. Avoid bubbling to reduce cell death.
10. Let the remaining tissue pieces sink for 2–5 min, then transfer the supernatant into another 15 mL tube. Count the cells using a hemocytometer.
    NOTE: The preferable cell density is 200,000–400,000 cells/mL. Use First Day Medium to dilute or centrifugation at 470 x g to concentrate the cells.
11. Seed 100 µL of cells on each glass coverslip. While seeding, make sure to cover the entire coverslip with cells. Incubate at 37 °C, 5–10% CO₂.
12. The next day, add 500 µL of Neuronal Growth Medium to the wells.
    NOTE: The composition of the Neuronal Growth Medium is described in the Table of Materials.
13. Gently transfer each coverslip to its appropriate well using a tweezer while removing the First Day Medium by tilting the coverslip. Suction the remaining media from the lid.
14. Incubate at 37 °C, 5–10% CO_2. Avoid replacing the medium throughout incubation. Cultures can be maintained under these conditions up to 1 month. The major concern is humidity. Therefore, make sure to maintain the incubator maximally humidified.

Results

In order to prepare the coral skeleton matrix, the entire coral skeleton (Figure 1A) was broken into 0.5–2 cm fragments using a hammer (Figure 1B) and thoroughly cleaned from organic residues through three steps (step 1 in the protocol) using 10% hypochlorite solution, 1M NaOH solution, and 30% H₂O₂ solution (Figure 1C). Coral fragments were well-cleaned when the skeleton color changed from brown (

Discussion

The technique presented here describes a way to improve the maintenance and functionality of neural cells in culture. This is achieved by adhering the cells to a matrix made of coral skeleton grains that nurtures the cells and promotes their growth and activity. Using this technique increases the capacity of the neural culture model to mimic the cells' environment in the brain.

The introduction of the matrix as a culture substrate has several advantages over other substrates used in classi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
24-well plates	Greiner	#60-662160
B-27	Gibco	#17504-044
Bovine Serum Albumin (BSA)	Sigma	#A4503
D – glucose	Sigma	#G8769
Dulbecco's Minimal Essential Eagle (DMEM)	Sigma	#D5796
Electrical sieve	Ari Levy	#3700
Fetal Bovine Serun (FBS)	Biological Industries	#04-007-1A
First Day Medium			85.1% Minimum Essential Eagle’s medium (MEM), 11.5% heat-inactivated fetal bovine serum, 1.2% L-Glutamine and 2.2% D-Glucose.
Flasks	Greiner	#60-690160	25cm^2, Tissue culture treated
Fluoro-deoxy-uridine	Sigma	#F0503
Glass Coverslips	Menzel-Glaser	#BNCB00120RA1
H2O2	Romical	#007130-72-19	Hazardous
Ham's F-12 Nutrient Mixture	Sigma	#N4888
HANK'S solution	Sigma	#H6648
Kynurenic acid	Sigma	#K3375
L - glutamine	Sigma	#G7513
Manual strainer (40µm)	VWR	#10199-654
Minimun Essential Eagle (MEM)	Sigma	#M2279
Mortar and pestle	De-Groot	4-P090
NaClO (Sodium Hypochlorite)	Sigma	#425044	Hazardous
NaOH	Sigma	#S8045	Hazardous
Neuronal Growth Medium			45% MEM, 40% Dulbecco's modified eagle's medium (DMEM), 10% Nutrient mixture F-12 Ham, 0.25% (w/v) bovine serum albumin (BSA), 0.75% D-glucose, 0.25% L-Glutamine, 0.5% B-27 supplement, 0.1% kynurenic acid, 0.01% of 70 % uridine and 30% fluoro-deoxy-uridine.
Petri dish	Greiner	#60-628160, #60-627160	60mm, 35mm, respectively.
Poly D – Lysine	Sigma	#P7280
Smart Dentin Grinder	KometaBio	#GR101
Trypsin	Gibco	#15-090-046
Uridine	Sigma	#U3750