Making, Testing, and Using Potassium Ion Selective Microelectrodes in Tissue Slices of Adult Brain

Summary

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.

Abstract

Potassium ions significantly contribute to the resting membrane potential of cells and, therefore, extracellular K⁺ concentration is a crucial regulator of cell excitability. Altered concentrations of extracellular K⁺ affect the resting membrane potential and cellular excitability by shifting the equilibria between closed, open and inactivated states for voltage-dependent ion channels that underlie action potential initiation and conduction. Hence, it is valuable to directly measure extracellular K⁺ dynamics in health and diseased states. Here, we describe how to make, calibrate and use monopolar K⁺-selective microelectrodes. We deployed them in adult hippocampal brain slices to measure electrically evoked K⁺ concentration dynamics. The judicious use of such electrodes is an important part of the tool-kit needed to evaluate cellular and biophysical mechanisms that control extracellular K⁺ concentrations in the nervous system.

Introduction

Potassium ion concentrations are tightly regulated in the brain, and their fluctuations exert a powerful influence on the resting membrane potential of all cells. In light of these critical contributions, an important goal of biology is to determine the cellular and biophysical mechanisms that are used to tightly regulate the concentration of K⁺ in the extracellular space in different organs of the body^1,2. An important requirement in these studies is the ability to measure K⁺ concentrations accurately. Although many components which contribute to potassium homeostasis in the brain in healthy and diseased states have been identified^3,4,5, further progress has been slowed due to the specialized nature of preparing ion selective microelectrodes for potassium measurement. Microelectrode sensors represent the gold standard for measuring K⁺ concentrations in vitro, in tissue slices and in vivo.

Newer approaches for K⁺ monitoring are under development using optical sensors, however these do not detect a biologically relevant range of K⁺ concentrations or have not been fully vetted in biological systems, although initial results appear promising^6,7,8. Compared to optical sensors, microelectrodes are fundamentally limited to a point source measurement of ions, although electrode arrays could improve the spatial resolution⁹. This article focuses on the single-barreled microelectrode sensors for monitoring K⁺ dynamics.

In this work, we report detailed stepwise procedures to make K⁺ selective microelectrodes, using a valinomycin-based potassium ionophore that permits highly selective (10⁴ fold K⁺ to Na⁺ selectivity) K⁺ movement over membranes¹⁰. A naturally occurring polypeptide, valinomycin acts as a K⁺ permeable pore and facilitates the flow of K⁺ down it's electrochemical gradient. We also describe how to calibrate the electrodes, how to store and use them and finally how to deploy them to measure K⁺ concentration dynamics in acute hippocampal brain slices from adult mice. The use of such electrodes together with genetically modified mice that lack specific ion channels proposed to regulate extracellular K⁺ dynamics should reveal the cellular mechanisms used by the nervous system to control the ambient concentration of K⁺ in the extracellular milieu.

Protocol

All animal experiments were conducted in accordance with the National Institute of Health Guide for the Care and Use of Laboratory Animals and were approved by the Chancellor's Animal Research Committee at the University of California, Los Angeles. All mice were housed with food and water available ad libitum in a 12 h light-dark environment. All animals were healthy with no obvious behavioral changes, were not involved in previous studies, and were sacrificed during the light cycle. Data for experiments were collected from adult mice (6-8 weeks old for all experiments).

1. Preparation of K⁺ selective microelectrodes

1. Silanization of borosilicate glass
   1. Remove sufficient glass capillaries from packaging and place into a 50 mL conical tube. Fill conical tube to top with 1 M HCl. Wash electrodes with gentle agitation in HCl overnight or for a minimum of 6 hours.
   2. Briefly rinse capillaries with 70% ethanol and then dry completely at 100-120 °C for 6-8 hours. Store washed capillaries in containers with anhydrous calcium sulfate desiccant for up to 4 weeks before further use.
   3. Prior to silanization, pull capillaries to a fine tip using a microelectrode puller. The microelectrodes that we use are approximately 2-5 microns in diameter. Always handle washed capillaries with gloves, as oils from skin can interfere with silanization.
   4. Place microelectrodes into a glass container so that the electrodes are elevated from the bottom to prevent tip breakage. Fix the microelectrodes to the container using autoclavable tape or similar adhesive tape.
   5. Remove approximately 0.5 mL of 5% dichlorodimethylsilane (DDS) silanization solution from its container using the nitrogen replacement method (see Figure 2). Fill a balloon with nitrogen gas and attach a syringe or tube and needle to the balloon. Insert the needle into the DDS container while drawing up DDS into a separate syringe via a longer needle.
   6. Apply the silanization solution dropwise to the tips of the pipettes and immediately cover. Place the container holding the microelectrodes with silanization solution into a pre-warmed (170-180 °C) laboratory oven for 10-12 hours or at 200-220 °C for 30 minutes
   7. After the incubation, turn off the oven and then remove the plate from the oven. Be careful when removing the plate from the incubator, as it is extremely hot. Place the plate on a bench at room temperature for 10-15 minutes to allow the glassware to cool.
   8. Remove the microelectrodes from the plate (using a razor blade or scalpel blade to cut the tape) and place them into a desiccant filled airtight container. Silanized microelectrodes kept free of moisture can be used for up to 1 week following silanization.
2. Priming the electrodes
   1. Prepare a stock solution of K⁺ ionophore cocktail: 5% w/v valinomycin, 93% v/v 1,2-dimethyl-3-nitrobenzene, 2% w/v potassium tetrakis(4-chlorophenyl)borate¹¹. This solution is a faint yellow color. Keep in an airtight, opaque container at room temperature. If properly stored this solution can last many months.
   2. Prepare a stock solution of 10 mM HEPES buffered 300 mM NaCl at pH 7.4. Fix the electrode into a clamp and backfill with the buffered NaCl using a 28G microfil tip connected to a syringe. Observe that the saline solution has reached the end of the microelectrode tip. Confirm that the microelectrode is free of large bubbles that could interfere with the flow of current.
   3. Break the tip of the electrode to approximately 10-20 µm wide, using the blunt side of a scalpel or razor blade.
   4. Using a micropipette, apply a small droplet (~0.1 µl) of the K⁺ 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 1-2 mm with the K⁺ ionophore, and remove excess using tissue paper.

2. Calibration of K⁺ Selective Microelectrodes

1. Preparation of calibration solutions
   1. Prepare solutions of various concentrations of KCl in equal osmolarity artificial cerebrospinal fluid (ACSF) by replacing NaCl with KCl. We used 0.1, 1, 4.5, 10 and 100 mM K⁺ ACSF, for calibration of the electrodes. The recipes for these calibration solutions are listed in Table 1.

Chemical	MW	final mM	0.1 mM [K+]	1 mM [K+]	4.5 mM [K+]	10 mM [K+]	100 mM [K+]
	(g / mol)
NaCl	58.44	varies	1.51 g	1.50 g	1.44 g	1.4 g	0.345 g
KCl	1 M stock	varies	20 µl	200 µl	900 µl	2 ml	20 ml
CaCl2	1 M stock	2	400 µl
MgCl2	1 M stock	1	200 µl
NaH2PO4	119.98	1.2	0.29 g
NaHCO3	84.01	26	0.437 g
D-Glucose	180.16	10	0.360 g
Water	q.s. 200 ml

Table 1. Potassium calibration solutions

2. Microelectrode calibration
   1. Bubble all solutions with 95% O₂/5% CO₂ for at least 20 minutes before beginning the experiment. Begin perfusing the bath with 4.5 mM [K⁺] ACSF at a rate of 3 mL per minute. Place the K⁺ selective electrode into electrode holder attached to the electrode headstage on the manipulator. This headstage is connected to an appropriate amplifier. Insert the tip of the electrode into the bath perfusate.
   2. Ensure the Ag/AgCl ground electrode is bathed in the same solution and that the flow is steady. Apply calibration solutions in a stepwise fashion and record the potential changes in mV across the electrode tip. Wait for the potential at the electrode tip to reach a stable value before switching to the next solution
   3. 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 response is at least 52 and no greater than 58 mV per log change in [K⁺].

3. Preparation of Acute Hippocampal Brain Slices

1. Preparation of slice solutions
   1. Prepare 500 mL sucrose cutting solution composed of: 194 mM sucrose, 30 mM NaCl, 4.5 mM KCl, 10 mM D-glucose, 1 mM MgCl₂, 1.2 mM NaH₂PO₄, and 26 mM NaHCO₃, 290-300 mOsm, saturated with 95% O₂ and 5% CO₂.
   2. Prepare 1-2 liters of recording solution (ACSF) composed of: 124 mM NaCl, 4.5 mM KCl, 1 mM MgCl₂, 10 mM D-glucose, 2 mM CaCl₂, 1.2 mM NaH₂PO₄, and 26 mM NaHCO₃; pH 7.3 - 7.4 (after bubbling), 290 - 300 mOsm, saturated with 95% O₂ and 5% CO₂. Fill a beaker containing a brain slice holder chamber with recording solution and keep it at 32-34 °C. Fill the vibratome chamber with ice-water slush.
2. Acute slice preparation
   1. Deeply anesthetize a mouse by placing it in a bell jar precharged with 2-3 mL isoflurane. Check for toe pinch reflex, and if non-responsive, rapidly decapitate it using a pair of sharp shears or guillotine as your animal protocol requires.
   2. Make a 2-3 cm incision using shears from the caudal portion of the skull to cut the scalp along the midline. While manually retracting the scalp, make two 1 cm horizontal incisions from the foramen magnum along the sides of the skull. Then, using fine shears, cut an incision the length of the skull, along the midline from the back of the skull to the nose.
   3. Using fine forceps inserted near the midline, retract the incised skull in two portions. Extract the mouse brain from the skull and use a blade to remove the cerebellum and olfactory bulbs, which are respectively located at the caudal and rostral portions of the brain. These can be identified by the large fissures, which separate them from the cortex.
   4. Mount the brain block onto the vibratome tray using super glue. Fill the vibratome tray with ice cold cutting solution.
   5. Cut tissue sections on the coronal plane at 300 µm thickness. Usually 6 coronal hippocampal slices can be collected.
   6. After each section is cut, immediately transfer the slices to the slice holding beaker warmed to 32-34°C. Keep the sections at this temperature for 20 min before removing the beaker containing the sections and place this at room temperature for at least 20-30 minutes prior to recording.

4. Measurement of Electrically Evoked K⁺ Dynamics

1. Setting up the slice preparation
   1. 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.
   2. Ensure the tips of the bipolar stimulating electrode are parallel to one another and are level with the plane of the slice. Slowly, over the course of 6-7 seconds, insert the electrodes into CA3 stratum radiatum approximately 40-50 µm deep to stimulate Schaffer collaterals. In coronal sections, CA3 can be approximately identified as the portion of the hippocampus proper lateral to the granule cell layer at the hippocampal genu, with the stratum radiatum falling medial and ventral to the pyramidal cell layer.
   3. Carefully insert the K⁺-selective electrode into CA1 stratum radiatum approximately 50 µm deep, by slowly lowering the electrode over approximately 3-4 seconds. Allow the potential to stabilize across the electrode before applying stimulations to the slice: this usually takes 5 to 10 minutes. If the slice exhibits spontaneous changes in extracellular K⁺ then discard and repeat this process with a new slice.
2. Measure evoked K⁺ release
   1. Apply trains of electrical stimulation (8 pulses) by manually depressing the trigger on the stimulator while digitally recording responses. Apply stimulation at 10 Hz and 1 ms pulse width, starting at 10 µA stimulus amplitude.
   2. Apply increasing stimulation amplitudes by a factor of 2 until a maximum K⁺ response amplitude is detected. If you do not see any response, move the position of the K⁺ electrode closer to the stimulation site in 100 µm in increments
   3. Determine the stimulus amplitude that produces the half maximal response. In our experience, this is between 40-160 µA, depending upon the preparation quality, age of the animal, and the distance between stimulation electrodes and K⁺ selective electrode.
   4. In the same slice, using a stimulus amplitude at one step lower than the amplitude that produces the half maximal response (e.g. if 80 µA produces a half-maximal response, use 40 µA) apply stimulus trains of increasing number of pulses. Initially, we have used trains of 1, 2, 4, 8, 16, 32, 64 and 128 pulses.
   5. To confirm that K⁺ signals are mediated by action potential firing of the electrically stimulated Schaffer collaterals bath, apply 0.5 µM TTX in ACSF for 10 minutes and repeat the stimulation protocol. No evoked responses should be observed.
   6. After finishing the slice experiment, confirm the electrode has maintained its responsivity by re-calibrating the electrode in the calibration solutions and ensuring the response has not deviated by more than 10% from the initial calibration

Results

For selective measurement of extracellular K⁺, we prepared ion-selective microelectrodes coated with a hydrophobic layer through silanization of clean borosilicate glass pipettes (Figure 1A). This coating enables the K⁺ ionophore containing valinomycin to rest at the tip of the electrode and permit only K⁺ flux through a narrow opening at the electrode tip (Figure 1B). After priming the electrodes...

Discussion

The method that we describe here has allowed us to assess K⁺ dynamics in response to electrical stimulation of Schaffer collaterals in acute hippocampal slices from adult mice. Our method of preparing K⁺ ion selective microelectrodes is similar to earlier described procedures^12,13,14,15. However, this method has advantages over alternative electrode configurations in that ...

Materials

Name	Company	Catalog Number	Comments
Vibratome	DSK	Microslicer Zero 1
Mouse: C57BL/6NTac inbred mice	Taconic	Stock#B6
Microscope	Olympus	BX51
Electrode puller	Sutter	P-97
Ag/AgCl ground pellet	WPI	EP2
pCLAMP10.3	Molecular Devices	n/a
Custom microfil 28G tip	World precision instruments	CMF28G
Tungsten Rod	A-M Systems	716000
Bipolar stimulating electrodes	FHC	MX21XEW(T01)
Stimulus isolator	World precision instruments	A365
Grass S88 Stimulator	Grass Instruments Company	S88
Borosilicate glass pipettes	World precision instruments	1B150-4
A to D board	Digidata 1322A	Axon Instruments
Signal Amplifier	Multiclamp 700A or 700B	Axon Instruments
Headstage	CV-7B Cat 1	Axon Instruments
Patch computer	Dell	n/a
Sodium Chloride	Sigma	S5886
Potassium Chloride	Sigma	P3911
HEPES	Sigma	H3375
Sodium Bicarbonate	Sigma	S5761
Sodium Phosphate Monobasic	Sigma	S0751
D-glucose	Sigma	G7528
Calcium Chloride	Sigma	21108
Magnesium Chloride	Sigma	M8266
valinomycin	Sigma	V0627-10mg
1,2-dimethyl-3-nitrobenzene	Sigma	40870-25ml
Potassium tetrakis (4-chlorophenyl)borate	Sigma	60591-100mg
5% dimethyldichlorosilane in heptane	Sigma	85126-5ml
TTX	Cayman Chemical Company	14964
Hydrochloric acid	Sigma	H1758-500mL
Sucrose	Sigma	S9378-5kg
Pipette Micromanipulator	Sutter	MP-285 / ROE-200 / MPC-200
Objective lens	Olympus	PlanAPO 10xW