Neuroscience
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Making, Testing, and Using Potassium Ion Selective Microelectrodes in Tissue Slices of Adult Brain
Chapters
Summary May 7th, 2018
Potassium ions contribute to the resting membrane potential of cells and extracellular K+ concentration is a crucial regulator of cellular excitability. We describe how to make, calibrate and use monopolar K+-selective microelectrodes. Using such electrodes enables the measurement of electrically evoked K+ concentration dynamics in adult hippocampal slices.
Transcript
The overall goal of this technique is to make, calibrate, and use monopolar potassium ion-selective microelectrodes. Using such electrodes enables the quantitative assessment of evoked potassium ion concentration dynamics in adult brain slices. This method can help answer key questions in physiology, such as identifying the cellular and molecular mechanisms for potassium ion homeostasis in the central nervous system.
The main advantage of this technique is that, in addition to being an easy and straightforward method of fabrication, microelectrode sensors represent the gold standard for the measurement of potassium ion concentration. To prepare the borosilicate glass capillaries for silanization, place them in a 50-milliliter conical tube. Fill the conical tube with one-molar hydrochloric acid, and incubate the glass capillaries in hydrochloric acid with gentle agitation overnight for a minimum of six hours.
Afterward, briefly rinse the capillaries with 70%ethanol and then dry them completely at 100 to 120 degrees Celsius for six to to eight hours. Store the washed capillaries in containers with anhydrous calcium sulfate desiccant for up to four weeks before further use. Prior to silanization, pull the capillaries to a fine tip using a microelectrode puller.
Then, place the microelectrodes in a glass container using autoclavable tape, so that the electrodes are elevated from the bottom to prevent tip breakage. Next, remove approximately 0.5 milliliters of 5%DDS silanization solution from its container using the nitrogen-replacement method. Fill a balloon with nitrogen gas and attach a syringe or tube and needle to the balloon.
Then, insert the needle into the DDS container while drawing up DDS into a separate syringe via a longer needle. Subsequently, apply the silanization solution drop-wise to the tips of the pipettes and immediately cover the container. Place the container holding the microelectrodes with silanization solution in a pre-warmed laboratory oven at 170 to 180 degrees Celsius for 10 to 12 hours or at 200 to 220 degrees Celsius for 30 minutes.
After incubation, remove the plate from the oven. Place the plate on a bench at room temperature for 10 to 15 minutes to allow the glassware to cool. Remove the microelectrodes from the plate and place them into a desiccant-filled airtight container.
Silanized microelectrodes kept free of moisture can be used for up to one week following silanization. To prime the electrodes, prepare a stock solution of potassium ionophore cocktail and keep it in an airtight, opaque container at room temperature. Next, prepare a stock solution of 10-millimolar HEPES-buffered 300-millimolar sodium chloride at pH 7.4.
Fix the electrode in a clamp. Subsequently, using a blunt instrument, rake the tip of the electrode to approximately 10 to 20 micrometers wide, then, backfill the electrode with the HEPES-buffered sodium chloride using a 28 gauge MicroFil tip connected to a syringe. Observe that the saline solution has reached the end of the microelectrode tip and make sure that the microelectrode is free of large bubbles that could interfere with the flow of current.
Using a micropipette, apply a small drop of the potassium ionophore near the tip of the microelectrode. If the electrode has been properly silanized, the droplet will be absorbed into the broken tip. Fill the electrode to approximately 1 millimeter with the potassium ionophore and remove the excess using tissue paper.
To calibrate the microelectrodes, bubble all calibration and slice buffer solutions with 95%oxygen/5%carbon dioxide for at least 20 minutes before beginning the experiment. Begin perfusing the bath with 4.5-millimolar potassium ion-containing ACSF at a rate of three milliliters per minute. Place the potassium ion-selective electrode into the electrode holder attached to the electrode headstage on the manipulator.
Then, insert the tip of the electrode into the bath perfusate. Ensure the silver/silver chloride ground electrode is bathed in the same solution and the flow is steady. Apply calibration solutions in a step-wise fashion and record the potential changes in millivolts across the electrode tip.
Wait for the potential at the electrode tip to reach a stable value before switching to the next solution. Then, measure the steady-state voltage change in response to the application of the calibration solutions to the electrode tip. Confirm that the slope of the electrode voltage response is at least 52 and no greater than 59 millivolts per ten-fold change in potassium concentration.
To prepare hippocampal slices, make a two to three-centimeter incision from the caudal portion of the skull to cut the scalp along the midline. While manually retracting the scalp, make two one-centimeter horizontal incisions from the foramen magnum along the sides of the skull. Then, using fine shears, make an incision along the midline from the back of the skull to the nose.
Next, insert fine forceps near the midline and retract the incised skull in two portions. Extract the mouse brain from the skull and use a blade to remove the cerebellum, prefrontal cortex, and olfactory bulbs, which are respectively located at the caudal and rostral portions of the brain. Then, mount the brain block onto the vibratome tray using Super Glue.
Fill the vibratome tray with ice-cold cutting solution. Afterward, cut tissue sections on the coronal plain at 300-micrometer thickness. Usually four to six coronal hippocampal slices can be collected.
After each section is cut, immediately transfer the slice to the slice-holding beaker warmed to 32 to 34 degrees Celsius. Keep the sections at this temperature for 20 minutes before transferring the beaker with the sections to room temperature for at least 20 to 30 minutes prior to recording. To measure the potassium ion dynamics, gently place the brain slice in the bath using a Pasteur pipette and gently hold it in place with a platinum harp with nylon strings.
Ensure the tips of the bipolar stimulating electrode are approximately parallel to one another and are level with the plain of the slice. Over the course of five to seven seconds, slowly insert the electrodes into CA3 stratum radiatum approximately 40 to 50 micrometers deep to stimulate Schaffer collaterals. Then, carefully insert the potassium ion-selective electrode into CA1 stratum radiatum to approximately 50 micrometers deep by slowly lowering the electrode over approximately three to four seconds.
Allow the potential to stabilize across the electrode before applying stimulations to the slice, which usually takes five to 10 minutes. If the slice exhibits excessive spontaneous changes in extra-cellular potassium ion concentrations, discard and repeat the process with a new slice. To measure evoked potassium ion release, apply trains of electrical stimulation via the stimulus isolator while digitally recording responses.
Apply stimulation at 10 hertz and one millisecond per pulse starting at 10 microamps stimulus amplitude. Apply increasing stimulation amplitudes by a factor of two until a maximum potassium response amplitude is detected. If no response is observed, move the position of the potassium ion-selective electrode closer to the stimulation site in 100-micrometer increments.
To confirm that potassium ion concentration changes are mediated by action potential firing of the electrically-activated Shaffer collaterals, bath apply 0.5 micromolar TTX in ACSF for 10 minutes and repeat the stimulation protocol. No evoked responses should be observed. After priming the electrodes with the backfilled HEPES-buffered saline solution and the potassium ionophore, the electrodes can be tested for their rapid response to stepwise changes in bath potassium ion concentrations and for the linear response to bath potassium ion changes over the 0.1 to 100-millimolar calibration range in a manner predicted by the Nernst equation.
The steady-state potential can be plotted against the bath potassium ion concentration in order to determine the slope of the line, which should be approximately V 58.2 millivolts and no less than 52 millivolts per ten-fold change in potassium ion concentration. The responsiveness of the potassium-selective electrodes was further tested and the response to a 5.5 millimolar increase in potassium was observed with rise and decay time constants of approximately 85 milliseconds. Once the stimulating and potassium ion-selective electrodes have been placed into the tissue and the recording has reached a stable baseline, then pulses of increasing current amplitude can be applied.
The waveform of this activity appears as a rapid increase in potassium with an exponential decay rate, which is abolished with TTX application. Once mastered, this technique can be done in approximately three to four hours if it is performed with care and accuracy. While attempting this procedure, it's important to remember to keep the electrode tip as small as possible to enable accurate recordings but large enough to permit low noise.
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