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

The significance of this protocol is that it provides a new way to increase the durability and growth of neurons, glia, and stem cells in vitro. It is also applicable for other cells. The main advantage of the technique is the usage of a matrix made of coral skeleton.

This matrix nurtures and protects neuro cells and it promotes regrowth. Being supportive for neural cells in vitro, the coral skeleton may function similarly as a regenerative implant in brain injury sites. To begin, use a hammer to break the coral skeleton, and divide it into 0.5 to two centimeter fragments.

In order to dissolve organic and non-organic residues, soak the coral skeleton fragments in 10%Sodium hypochlorite solution for 10 minutes. Then wash once with double distilled water. To remove the remaining organic residues, soak the fragments in a one molar sodium hydroxide solution for five minutes, then wash once with double distilled water.

Continue with the removal of the organic deposits by soaking the fragments in 30%hydrogen peroxide solution for 10 minutes, and wash the fragments three times with double distilled water. Remove as much excess double distilled water as possible and leave the coral fragments in the hood to dry. Then clean the glass cover slips by soaking them in 10 milliliters of 95%ethanol in a 100 millimeter glass Petri dish.

After 15 minutes, remove the ethanol and wash with double distilled water three times, waiting 10 minutes between each wash. Remove the double distilled water and place the dish on an 80 degree Celsius prewarmed heating plate. Then gently stir the cover slips until the double distilled water evaporates.

Autoclave the cover slips. Grind the coral skeleton fragments using a mortar and pestle until complete breakdown. A mixture of grains with sizes ranging from 20 to 200 micrometers is obtained.

Alternatively, grind the coral skeleton fragments using an electrical grinding machine with a blade length of six centimeters and width of 0.5 to one centimeters, at a velocity of 1000 RPM for 30 seconds. To purify the grains, transfer the grains onto an electrical 40 micron filter mesh strainer. Adjust the electrical strainer settings to shaking at 600 amplitudes per minute, and bouncing at six times per second.

Then autoclave the grains. Under sterile conditions, add the coral skeleton grains into a 20 microgram per milliliter PDL, dissolved in Hank's solution to achieve the concentration of five milligrams or 10 milligrams of grains per one milliliter PDL solution. Pour the solution into flasks and dishes at the concentration of approximately two milliliters per 25 square centimeters, and incubate overnight at four degrees Celsius.

The grains sink and attach to the bottom. The next day, wash the flasks and dishes once with sterile double distilled water. Let the flasks and dishes dry in the hood.

To coat the glass cover slip, first add coral skeleton grains to double distilled water at a concentration of five or 10 milligrams per milliliter. Place the cover slips on a heating plate prewarmed to 80 degrees Celsius. Then drip 40 microliters of the grain solution onto the center of the cover slip, and wait 15 minutes for complete evaporation.

This will result in the grains adhering to the cover slip. Autoclave the coated cover slips, and store in sterile conditions. Next, place the cover slips on the lid of a sterile 24 well plate, add 100 microliters of 20 microgram per milliliter PDL solution to each cover slip.

Use the tip to ensure that the liquid covers the entire grain region and the rest of the cover slip surface. Cover the lid with the bottom of the plate, and wrap the sides of the place with paraffin film. Incubate overnight at four degrees Celsius.

The next day, wash the cover slips once with double distilled water and dry in the hood. After preparing rat pup heads, hold it from the mouth with big tweezers. Cut the skin covering the skull with a small scissor.

With clean scissors, 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 small tweezers, peel the skull off from the brain. Separate the brain from the head and put it in a 60 millimeter Petri dish containing two milliliters of cold MEM.

Under a stereo microscope, use two small tweezers to dissect the hippocampi. Transfer the hippocampi into the prepared 35 millimeter dish containing MEM. Cut the hippocampi to approximately one millimeter slices using the scalpel.

Add 200 microliters of trypsin solution. Mix gently and incubate at 37 degrees Celsius and five to 10%carbon dioxide for 30 minutes. After incubation, add two milliliters of cold MEM to the dish to deactivate the trypsin.

Then use a glass Pasteur pipette with the largest diameter to transfer the trypsinized tissue to a 15 milliliter tube containing one milliliter of first day medium, preheated to 37 degrees Celsius. Triturate the tissue by passing it through the largest diameter glass pipette 10 to 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. Let the disassociated tissue pieces sink for two to five minutes.

Then transfer the supernatant into another 15 milliliter tube. Count the cells using a hemocytometer. To achieve the preferable cell density between 200, 000 and 400, 000 cells per milliliter.

Use first day medium to dilute. Next, seed 100 microliters of cells on each glass cover slip. Make sure to cover the entire cover slip with cells.

Incubate at 37 degrees Celsius and five to 10%carbon dioxide. The next day, add 500 microliters of neuronal growth medium to the wells. Using tweezers, remove the first day medium by tilting the cover slip, and gently transferring each cover slip to its appropriate well.

Suction the remaining media from the lid. Incubate the plate at 37 degrees Celsius and five to 10%carbon dioxide. Maintain the incubator maximally humidified using a tray of water in the bottom of the incubator.

Do not replace the medium. Cultures can be maintained under these conditions up to one month. In this study, manually grinding and electrical grinding machines both yielded similar outcomes, with a mixture of grains ranging between 20 to 200 micrometers in size.

The manual or electrical strainers produced two types of grain mixtures. One with grains ranging from 40 to 200 micrometers in size, and the second with the size of 40 micrometers or less. Phase contrast micrographs show that the cultivation of hippocampal cells on cover slips coated with coral skeleton grains formed complex neuronal networks in the absence and presence of the matrix.

Cell nuclei staining indicated that 14 days after seeding, cell density was higher on the matrix than on the uncoated cover slips. Staining with microtubule associated protein two showed that a complex neuronal and dendritic network formed on the matrix compared to the control. In addition, glial cells grown on the coral skeleton matrix visualized using the glial fibrillary acidic protein underwent significant morphological changes.

They acquired a spikier appearance, with longer processes, than the glial cells grown on the control substrate, where they appeared rounder and flatter. While isolating the cells, It is important to remember to work fast, maintain the pH by adding CO2 if necessary, and work as gently as possible in order to prevent detachment of coral grains.