The overall goal of this procedure is to isolate and associate E17 mouse neuronal tissue, in order to culture, record and stimulate neuronal networks on microelectrode arrays. This method can help answer key questions in the study of network dynamics, as they relate to the basic mechanisms of learning and memory. The main advantage of this technique is that micro-electrode arrays are able to record from multiple sites simultaneously and therefore can give better spatial, temporal resolution, compared to the more traditional glass pipette on single site recording.

Though this method can provide insight into the network dynamics of learning and memory, it can also be used to investigate drug toxicity or design paradigms for next-generation, personalized medicine. Visual demonstration of this process is critical, as manipulation of the tissue, during the dissection and dissociation process is difficult to learn from simply reading protocols alone. To prepare the array, add 40 to 50 microliters of thawed laminin to the center of each MEA, and 0.2 milliliters of laminin to the control plates, using sterile pipette tips.

Cover all the MEAs and the control dishes in the biohood, and transfer them to an incubator at 37 degrees Celsius for one hour. After one hour, carefully remove the excess laminin from the center of the MEAs using suction with sterile Pasteur pipettes, and let the surface air-dry before plating. Next, pour L-15 medium into four 100 millimeter Petri dishes, cover them, and place them in a minus 20 degree Celsius freezer for about 40 to 60 minutes, until the medium has a slushy consistency, but is not frozen solid.

To prepare for the dissection, place a glass Petri dish face down in a tray full of ice. Now spray the animals lower abdomen with 70%ethanol. Using small surgical scissors, make a V-shaped cut through the skin and subcutaneous fat of the lower abdomen, extending the cut to the distal ends of the thoracic cavity, and exposing the uterus.

Using forceps, carefully lift the uterus between the embryos. Cut away the connective tissue with dissection scissors until the entire uterus is free. Then briefly rinse the uterus with 70%ethanol to remove any blood, and place it in one of the four Petri dishes filled with cold L-15.

Subsequently, release each embryo from the uterus and the interior embryonic sac, using two pairs of fine-tipped forceps. Place the freed embryos in the second dish full of cold L-15, then transfer the heads to the third Petri dish, and the bodies to the fourth one. In the biohood, place a wedge of autoclaved filter paper on the chilled glass Petri dish stage, and place one embryo head on the filter paper.

Next place the cutting edge of the lower shear of iris scissors into the base of the skull, keeping the lower shear against the inner surface of the skull and away from the brain, cut through the occipital plate and then along the midline between the parietal plates. Continue cutting rostrally between the cartilaginous frontal skull plates. Then, starting at the center of the occipital plate, make a perpendicular cut to the left and to the right of the center cut.

After that, carefully slide a small spatula between the ventral surface of the brain and the bottom skull plates, until it is completely under the brain. Then lift the spatula up to remove the brain. Using a clean spatula, first remove the olfactory bulb, then dissect the frontal lobe in a trapezoidal pattern.

Next, transfer the tissue to a 15 milliliter centrifuge tube containing storage medium. In this procedure, use two sterile scalpel blades to mince the tissue. Then add 2.5 milliliters of the Dnase papain mixture to the minced tissue in the Petri dish.

Gently swirl the Petri dish to ensure that all tissue is free in the solution, and not adhered to the bottom of the dish. Afterward place the dish in an incubator at 37 degrees Celsius for 15 minutes. In the biohood, use a sterile wide-bore transfer pipette to transfer all the media and tissue to a five milliliter sterile cryogenic tube.

Place the tip of the same transfer pipette close to the bottom of the tube, and gently chiterate the mixture 10 to 15 times. Then add two milliliters of warmed DMEM 5/5 to the dissociated cell mixture, and cap the centrifuge tube. Gently mix it by inversion, and centrifuge it at 573 times g for five minutes at room temperature.

In the biohood, discard all the supernatant without breaking the pallet. Subsequently, add one milliliter of warmed DMEM 5/5 to the pallet in the tube to resuspend the cells. Use a sterile small-bore transfer pipette to break up the pallet by gently pipeting up and down until the mixture is homogenous.

Then transfer 10 microliters of the cell suspension to a microcentrifuge tube. Outside of the biohood, add 10 microliters of trypan blue to the 10 microliters of cell suspension in the microcentrifuge tube. Then load 10 microliters of the trypan blue cell suspension onto a disposal hemocytometer chip in order to count the cells.

In the biohood, use a sterile micropipette to transfer 50 microliters of cell suspension to the center of each array, and each control Petri dish. Then place the covered Petri dishes in an incubator set to 37 degrees Celsius for three to four hours. Next, gently add one milliliter of warmed DMEM 5/5 to each MEA.

Carefully add the medium one drop at a time around the inside edges of the MEA, and avoid washing the cells away from the array center. Afterward, place a cap containing a gas-permeable FEP membrane on each MEA before returning them to the incubator. After two days, perform a complete medium replacement with warmed DMEM+by drawing out the medium from the dish.

Carefully place the tip of the pipette on the inside wall, and dispense one milliliter of warmed DMEM+For the subsequent feedings, draw out 500 microliters of medium from the dish, and dispense 500 microliters of warmed DMEM+again. In this procedure, inspect the dish samples every other day under a microscope, to look for cell coverage over the array, and contamination by either bacteria or fungi. Before taking the cultures out of the incubator, plug in the temperature controller, and turn on the system, allowing the heated baseplate of the preamplifier to reach 35 degrees Celsius.

Then place the capped culture in the preamplifier, so that the black line in the MEA well aligns with the reference ground. Ensure that the preamplifier pins line up so that the top of the preamplifier is secured, and the culture cap is still on. Next, press start in the software environment to start visualizing the signals.

In the main window select Spikes, Detection, Automatic, change the standard deviation value to minus five, and click Refresh to reset the threshold. After visualizing the signals, click Stop, Record, and Play to begin recording activity. Embryonic mouse neurons are plated on 60 channel MEAs, which allow for the simultaneous recording of the neuronal activity across the network from each electrode.

Here is a representative raster plot of activity from eight electrodes in response to stimulation before, and after training. The vertical red line indicates the time of the stimulus, and the black tick marks indicate action potentials. In pre-training there is an immediate response to the stimulus pulse across channels.

In post-training, the network exhibits a more prolonged activity response, as well as the immediate response to the stimulation. Shown here is the average of 12 trained, and 10 control networks, the trained networks indicate significantly altered spike frequencies. After watching this video you should have a good understanding of how to isolate and associate neuronal tissue from embryonic mice, and be able to culture, record and stimulate neuronal networks from microelectrode arrays.

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