Slice Preparation and General Use
For an illustrated method, download the PDF
Daniel H. Chun
Tensor Biosciences
Step 1. MED Probe Coating.
The surface of the MED Probe is relatively hydrophobic. Therefore, treatment with polyethylenimine is necessary for tissue adhesion. This coating process should be done only once to new probes, as repetitive exposure to ACSF and tissue will enhance future slice adhesion.
Fill a new probe with 1ml of 0.1% polyethylenimine in 25 mM borate buffer (PEI). Cover with parafilm and store for at least 12hrs. Probes with PEI can be stored for several days. Rinse 3 times with sterile distilled water before use.
Step 2. Warming up the MED 64 Setup.
In general, the first thing that should be done when beginning your day is to turn on various heat sources such as water baths, incubators, and humidified O2 / CO2. Temperatures should be set so that the ambient temperature inside the probe is approximately ~31 degrees-32 degrees C.
Step 3. ACSF.
Prepare 1 liter of ACSF and oxygenate immediately and continuously throughout the day. pH= ~7.4. Composition (in mM): NaCl 124, KCl 3, KH2PO4 1.25, D-Glucose 10, MgSO4 1, NaHCO3 36, CaCl2 2.
After oxygenating the ACSF for at least 10 min, remove 150 ml (two beakers of 75ml + 75ml to speed up freezing) and place in an -80 degrees C freezer or dry ice. Complete the checklist in Step 4 and then remove the solution when 1/3 of the volume has turned to ice. Crush the partially frozen ACSF into a fine pulp/slush and oxygenate until ready to use. 75ml should be used for removing the brain and dissecting, and the other 75ml should be placed into the vibratome chamber.
Step 4. Brain Slices
The following is a checklist of tools and procedures that should be completed prior to removing the brain:
Decapitation tools:
- Razor blade
- Small scissor
- Forceps / Tweezers
- Spatula
- Guillotine
- Disposal bag
- Paper towels
Brain Dissecting Tools:
- Razor blade
- Plastic spoon
- Curved forceps
- Petri dish with filter paper lining
- Clean vibratome blade
- Extra filter paper (to soak up excess glue and ACSF)
- Suctioning pipette (transferring slices from vibratome to recovery chamber)
- Cyanoacrylate glue (Crazy Glue‰)
- Remove frozen ACSF from freezer, crush and oxygenate.
- Place half of the frozen ACSF slush into the vibratome chamber (mixed with some room temp ACSF), this must be constantly oxygenated.
- Fill outer area of the vibratome chamber with crushed ice
- Block of Agar glued onto vibratome base plate
- Constantly oxygenated slice recover /storage chamber with mesh nylon bottom.
Decapitation
Remove the brain and place into the remaining 75ml of ice-cold ACSF. This step should be done quickly and with the least amount of damage / pressure onto the brain. Times range from 25 to 45 seconds. Times greater than 90 seconds could necessitate cutting again.
Dissection
1. Using the plastic spoon, place the brain into the petri dish so that the ventral side of the brain is touching the filter paper (also filled with frozen slush ACSF).
2. Remove the Cerebellum (coronal).
3. Remove the frontal lobe of the brain (coronal section). Approximately 1/3 of the brain.
4. Tilt the brain onto the rostral / frontal end (now flat from the previous cut).
5. Remove both ventral-lateral areas. 20 degrees to 30 degrees off the horizontal axis. 6. Remove the brain from the ice-cold ACSF onto a dry filter paper. The ventral side of the brain should again be on the bottom.
6. While the brain is drying, place a few drops of glue onto the vibratome base plate near the block of agar. 8. Spread the glue out so that it forms a thin layer covering a few square centimeters.
7. Lift the brain off of the filter paper and place onto the base plate so that the back / caudal portion of the brain is against the agar and the ventral side is on the glue.
8. Perform a medial sagittal cut.
9. Make sure the ventral-lateral areas are firmly glued onto the plate, and then remove any excess glue and ACSF.
10. Wait an additional 10 seconds for the glue to dry and place into the 0 degree C vibratome chamber.
11. The first cut should be 4 to 5 mm. thick, such that the hippocampus becomes visible. The thickness of hippocampal slices is typically 350 microns. The slice includes hippocampus and surrounding cortical areas.14. Place the freshly cut slices into the recovery / storage container for at least 50min.
Step 5. Perfusion, Air, and Temperature Settings.
The cover for the probe has three openings: one for inflow of fluid, one for outflow of fluid, and one for supplying oxygen. The needles for fluid inflow and outflow may be raised or lowered accordingly, to control fluid level.
A perfusion system is used to circulate fresh, oxygenated ACSF into the probe. The ACSF solution as well as the oxygen entering the probe chamber should be maintained at 32 degrees C.
An IV dripper should be placed somewhere in the delivery line. Flow rates can vary from static to up to 2 ml/min. Typically, 0.5 to 1.0 ml/min is used.
Humidified oxygen should enter the probe at no less than 0.10 liters/min and no greater than 0.30 liters/min.
Step 6. Placing Slices onto the MED64 Probe.
There are two methods for placing slices onto the probe. The first method involves an adhesion process, and the second utilizes small weights to anchor down the slice.
Adhesion Method
Following recovery, choose a slice exhibiting obvious laminar structure. Transfer the slice to the MED probe chamber, containing culture medium with sera (10% fetal bovine serum + 10% horse serum). Using a small paintbrush, position the slice on the electrode array in the center of the MED probe. Inspect the probe under the microscope, and align the slice further such that electrodes are positioned in the desired stimulation and recording sites.
For example, in a typical LTP experiment, Schaffer-commissural fibers are stimulated. Align the slice such that electrodes make contact with Schaffer-commissural fibers. Recording sites are most likely the areas indicated here.
After positioning the slice on the electrode array, completely remove the culture medium with sera used for incubation and replace with 250 microliters of fresh culture medium with sera.
During the adhesion process, it is critical that the slice receives sufficient oxygen. Prepare an airtight container with a secure lid. Flow oxygen into the container, optionally through a Teflon plate with very fine orifices. Pour a small amount of water into the bottom of the container to ensure sufficient humidity. Place the MED probe in the container, and close the lid tightly.
Place the container securely in a water bath at 38 degrees C with 95% oxygen, 5% carbon dioxide flow for one hour. Make sure the airflow pressure is positive, typically 20 mmHg.
The incubation period should not exceed one hour since the culture medium contains 20% sera, which may cause damage to the slices.
Weight Method
One way to guarantee contact between the probe and slice is to weigh it down. One method is to twist several wires (100-200mg). Be sure to form an opening in the middle so that no pressure is directly on the electrodes.
- Once the weight is made, remove one slice from the recovery chamber and place into the probe.
- Suction out most of the ACSF in the probe (~200 ul remaining).
- Under the microscope, position the slice into the desired orientation.
- Apply weight. The weight should not be touching any areas critical for recording.
- Take picture.
Step 7. Recording
Before the experiment begins, take a microscopic photograph of the position of the slice on the probe. Import the photomicrograph to the MED conductor software, and confirm the relative placement of the slice on the electrode array.
Place the MED probe on the MED connector, replace the connector cover, and, finally, place the probe cover on the probe. Make sure the needles of the probe cover are properly grounded (two yellow cables in the picture).
Referring to the photomicrograph, select channels for stimulation. Adjust the stimulation magnitude with the MED amplifier. MED Conductor sends the stimulation signal, while simultaneously recording and storing evoked responses from all 64 channels.
Step 8. Cleaning Up
- Turn off all electrical sources.
- Shut off the oxygen tank.
- Obtain the probe from the connector.
- Thoroughly rinse the probe under DI water (remember to keep the electrodes moist at all times, especially over night).
- Thoroughly rinse all glassware under hot water (Do not use soap).
- Use the perfusion pump and rinse out the tubing with DI water.
- Tubing and IV drips should be replaced on a monthly basis.
For an illustrated method, download the PDF.
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