Neuroscience
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A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture
Chapters
Summary March 5th, 2016
Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms.
Transcript
The overall goal of this procedure is to establish healthy longterm cultures of primary mouse hippocampal neurons at ultra-low density, in order to facilitate immunocytochemistry labeling, and the study of neuronal cell autonomous mechanisms. This study can help understand some key questions in the field of neuroscience research such as the studies on the specificity of protein in neuromorphology of function of development. The main advantage of this technique is that it allows neurons to grow at ultra-low density on a cover slip without the support of a glial feeder layer.
Demonstrating the procedure will be Mariel Piechowicz, a student from my laboratory. Prepare two high-density and two low-density 24-well plates. Using a 28 gauge needle, etch the bottom of 24 well plates, so that the displaced plastics will act as a support for the cover slips with low-density neurons growing on them.
Transfer the sterile glass cover slips into the two 24-well plates designated for low-density cultures. Coat all four plates with 300 microliters of 0.1 milligram per milliliter Poly-D-lysine diluted in borate buffer, and return them to the incubator for overnight incubation coating. Before tissue collection, prepare two 10 centimeter petri dishes filled with ice-cold sterile HBSS.
Under the microscope, hold the embryo by the neck with the forceps, and make the first cut right on the midline, below the cerebellum level. Subsequently, make a cut on the left side of the base of the skull. Next, make another cut on the right side of the base of the skull.
Leave a small part of the skull bone and skin tissue at the rostral end uncut so that the whole brain can be flipped aside for extraction easily. Now transfer the brain into the HBSS solution. Inspect the brain to make sure that it is intact without gross cut off regions.
Then, repeat the procedures to collect the rest of the embryo brains in a new petri dish containing 20 milliliters of cold HBSS buffer. Position each brain with the ventral side up. While holding down the brain with forceps at the caudal end, make a cut diagonally between the middle rostral point and the caudal temporal region with a sharp tweezer.
Repeat this for the other hemisphere. Afterward, separate the two hemispheres with intact hippocampus from the rest of the brain's stem tissues. Repeat the procedures for all the brains and pool all the dissected hemispheres together.
Then, remove the meninges using two pairs of tweezers. Identify the developing hippocampus as a curved structure that is folded inside the temporal lobe and separate it from the adjoining cortical tissues. Collect and pool all the hippocampi tissues together.
In this procedure, transfer all the dissected hippocampi into the enzymatic digestion solution in a 15 milliliter conical tube. Next, place the tube inside a water bath and incubate at 37 degrees Celsius for 25 minutes. After 25 minutes, carefully remove the enzyme solution.
Then, add five milliliters of warm wash medium to bring the total volume to 10 milliliters. Gently wash the tissue by slowly inverting the tube five times. Remove the wash medium and add 10 milliliters of wash medium again, for an additional wash.
After that, remove the wash medium and add three milliliters of fresh warm wash medium again. Shake the tube by hand rigorously 15 times to dislodge the digested neurons from the tissue. The medium should become turbid once the cells are separated from the tissue into the wash medium.
Then, collect the cell suspension in a new 15-milliliter conical tube, while leaving the remaining tissue in the tube. After that, add another two milliliters of wash medium to the tissue, and gently separate the remaining tissue by triturating with a plastic pipette 15 times. Let the cell suspension sit for five minutes before pooling it into a new 15-milliliter conical tube that contains the collected shake off cells.
Then, count the density of neurons with a hemocytometer. Calculate the cell suspension volume needed to yield a density of 250, 000 neurons per milliliter, and add it to one of the two conical tubes containing 30 milliliters of complete culture medium, to yield a high-density neuron plating medium. Transfer 1.2 milliliters of this high-density neuron solution to another 30 milliliters of complete culture medium, to yield a concentration of 10, 000 neurons per milliliter, and use this solution to seed the low-density cover slips.
In this procedure, place 24-well plates with etched bottoms for high-density neurons in the culture hood. Remove 300 microliters of the pre-conditioned medium by vacuum, then plate 0.6 milliliters of high-density cell suspension at 250, 000 neurons per milliliter. Place the other two 24-well plates with cover slips in the culture hood.
Remove 300 microliters of the preconditioned medium by vacuum, before plating 0.6 milliliters of low-density cell suspension at 10, 000 neurons per milliliter on the cover slips. Afterward, return all four 24-well plates to the incubator, and incubate for two hours. After that, flip the low-density cover slips with adhering neurons to the high-density culture and make sure the neurons are facing each other.
Subsequently, return the two plates with co-culture to the incubator. Feed the co-cultures by adding 300 microliters of fresh feed medium at DIV five. Following this initial feeding, add 300 microliters of fresh feed medium without cytosine arabinoside to each well every week.
Should the culture be kept for more than one month, replace half of the medium with fresh feed medium every week beyond one month time point. Morphologically, both high-density and low-density neurons should look healthy up to three months. Here, immunocytochemitry labeling is used to reveal the functional glutamatergic synapses in which NMDA receptor subunit NR1 and the ApoE receptor subunit GluR1 are labeled with double staining.
And the functional synapses can be readily identified and quantified using image J.Low-density neurons are also labeled with antibody against dendritic protein marker MAP2 in combination with axonal protein marker p-Tau. This double labeling allows clear distinction of dendrites and axons. Low-density cultures can also be successfully transfected with DNA plasmids using calcium phosphate methods, although the transfection rate is typically very low at later stages.
This figure illustrates that EGFP transfection can reveal neuronal morphology, and render fine dendritic spine structures visible. Lastly, as a proof of concept, ultra-low density neurons can be grown under conditions that promote autapse formation. These ultra-low density autaptic neurons grown on Poly-D-lysine coated microislands, can be sustained by the high-density feeder neurons to beyond two months with this co-culture protocol.
Once mastered, this technique can be done in two to three hours on the day of cell culture if it is performed properly. While attempting this procedure, it is critical to coat the glass cover slips and 24-well plate bottoms with Poly-D-lysine. A simple etch of the plate bottoms is adequate to provide mechanical and nutritional support to the low-density hippocampal neurons to grow on the cover slips.
After watching this video, you should have a good understanding on how to grow ultra-low density primary mouse embryonic hippocampal neurons, in the absence of a glial feeder layer. Hopefully you can utilize this method for your own experimental purpose.
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