Whole cell patch clamp electrophysiology in human neuronal cells

Abstract

Whole cell patch clamp recording techniques are commonly used to assay membrane excitability, ion channel function, and synaptic activity in neurons. However, assaying these functional properties of human neurons remains difficult because of the difficulty in obtaining human neuronal cells. Recent advents in stem cell biology, especially the development of the induced pluripotent stem cells, made it possible to generate human neuronal cells in both 2-dimensional (2D) monolayer cultures and 3D brain-organoid cultures. Here, we describe the whole cell patch clamp methods of recording neuronal physiology from human neuronal cells.

1. Introduction

Elucidating the functional properties of human neurons is crucial for understanding aberrant neuron physiology, including membrane excitability, ion channel and synaptic activities, underlying neurodevelopmental and neurodegenerative disorders. Patch clamp techniques (1–3) revolutionized our ability to investigate these functional properties by allowing researchers to dissect the ionic components of synaptic transmission and neuronal excitation (4, 5). This is achieved by placing a micropipette in extremely close contact with a neuronal cell membrane forming a high resistance seal (Gigaseal) to allow the recording of tiny current-voltage changes caused by ionic fluxes across cell membranes via “single” ion channels. Taken a step further, by applying brief negative pressure through the micropipette, one can rupture the membrane and create a continuous flow between the internal solution of the pipette and the cellular cytosol. This so called “whole cell” configuration allows for stable, intracellular recording of ion channel activities across the whole cell membranes. Whole-cell recordings also allow diffusional exchange of material from the micropipette into the cell cytosol for pharmacological manipulation and labeling of the cell morphology.

The whole cell patch clamp recordings have two different configurations: voltage clamp and current clamp, to record current and voltage changes across the cell membranes, respectively. The voltage clamp mode holds the cell’s membrane voltage constant, allowing one to measure changes in currents generated by ion fluxes across the membrane when channels open or close. Under voltage clamp recording mode, whole cell current responses mediated by ion channels including voltage-dependent sodium, potassium, and calcium channels could be recorded by holding the cell membrane at different voltages. Synaptic currents including both excitatory postsynaptic currents (EPSCs) and inhibitory postsynaptic currents (IPSCs) mediated by neurotransmitters such as glutamate and γ-amino butyric acid (GABA) could also be recorded at certain holding potentials depends on different internal solutions compositions.

The current clamp mode holds the current constant, enabling one to measure membrane voltage changes. Resting membrane potentials, action potentials (either spontaneous or elicited by depolarization of cell membrane), and spontaneous or evoked synaptic responses could be recorded. This powerful technique has greatly contributed to the understanding of neuronal electrophysiology and how it relates to neuronal health and disease (6–12).

The high prevalence of neurodevelopmental, neuropsychiatric, and neurodegenerative disorders necessitates investigation of neuronal physiology in human based systems. Human neuronal models also allow representation of heterogeneous genetic backgrounds potentially relevant to a given disease. The advent of induced pluripotent stem cell (iPSC) technology (13) and then further reprogramming to neurons and other brain cells now allows this investigation. One method of making monolayer 2D human neuron cultures is by ectopic expression of specific transcription factors, such as Ngn2 and Ascl1, that drive a stem cell to become a neuron (12, 14). Alternatively, neurons can also be generated using growth factors and small molecules through a process with greater mimicry to normal development (15–19). Together, these two methods for generating human stem cell derived neurons hold great promise for understanding human-specific physiology and pathology. We have used these genetically-defined iPSC-derived neurons and cerebral organoids to model opioid-receptor variants (20), nicotine addiction (21), ethanol’s effect on synaptic activity (22), neurodevelopmental disorders (23–25), and integrated with microelectromechanical systems (26, 27).

In comparison to 2D cell cultures, cerebral organoids offer increased complexity and physiological relevance because of their 3D structure. Brain/cerebral organoids are derived from human iPSCs that aggregate in suspension, undergo neural induction, and finally differentiation and maturation into functional neural tissue. Furthermore, these organoids can be “patterned” to certain brain areas such as cortical, thalamic, or even fused organoids of two separate regions (28–30). These cortical organoids develop cytoarchitecture and anatomical structures reminiscent of the developing human brain’s ventricular zones (31). Thus, the ability of 3D organoids to recapitulate human neural development during its embryonic stage shows itself to be a very promising and powerful model. Recent work has integrated microglia into these brain organoids to facilitate study of neuroimmune interactions and how they relate to neural function (32). One major challenge with long term organoid culture is hypoxia in the organoid interior. Due to the lack of vascularization, cells within the interior of the organoid suffer from lack of oxygen, nutrient delivery, and waste disposal. Cellular hypoxia can be observed as early as two months into culture leading to cell death in the interior of the developing organoid resulting in limited corticogenesis. To solve this issue a culture system called “sliced neocortical organoids” (SNO) was developed (33), bypassing the diffusion limit to prevent cell death over long-term cultures by slicing organoids and culturing in a manner similar to organotypic brain slice culture. This culture method effectively reduces cell hypoxia and fosters enhanced neural development and cortical lamination. Patch clamp recording techniques can be applied to organoid slices to assess neuron physiology (32, 33). However, because of high cell density, cell type heterogeneity, and a functional maturity gradient throughout the cortical layers, patching organoids can be labor intensive and low throughput.

In this chapter we detail the setup and basic patch clamp technique to achieve whole cell configuration in human neuronal cells in both 2D (Figure 1) and 3D (i.e., organoids) cultures (Figure 2, Adapted from Xu et al (32)). We will describe 4 basic protocols to reveal the membrane properties and synaptic activities in human neuronal cells.

Figure 1: Patch-clamp recording in iPSC-derived neurons.

(A) A representative image showing a patched neuron in an NGN2 induced neuron (iN) 2D culture. (B) A representative image of a Dextran-AlexaFluor488 filled iN. (C) A representative trace of spontaneous post-synaptic currents (PSCs) recorded from an iN. (D) A representative trace of spontaneous action potentials recorded from an iN. (E) Example of repetitive action potentials evoked by stepwise currents injections in an iN.

Figure 2: Patch-clamp recording in iPSC-derived brain organoid slice.

(A) A representative image showing a patched neuron in acute organoid slice. (B) Sample images of a neuron filled with biotin ethylenediamine HBR from a day 92 microglia-containing organoid after whole-cell recording. Complex arborization (B1) and spine-like structures (B2, B3) were identified. The images are inverted to a white background to help visualize neuron morphology. (C) Representative traces of whole-cell currents recorded from neurons in acutely sectioned organoid (90 day old in culture) slices. Insert: fast activation/inactivation voltage-dependent sodium currents. (D) Representative traces of spontaneous action potentials recorded from neurons in organoid (90 day old in culture) slices. (E) Example of repetitive action potentials evoked by stepwise currents injections. (F) Representative traces of spontaneous post-synaptic currents (PSCs) recorded from neurons in sliced collected from a day 92 organoid.

2. Materials

2.1. Solutions

Prepare all solutions with double deionized water (ddH₂0) (Resistivity of 18 MΩ at 25 °C) and laboratory grade reagents.

1. External solution – Artificial Cerebral Spinal Fluid (ACSF) contains (in mM):

NaCl 125, KCl 2.5, NaH₂PO₄∙H₂O 1.25, NaHCO₃ 25, MgCl₂ 1.2, CaCl₂ 2.5, glucose (C₆H₁₂O₆) 2.5, sucrose (C₁₂H₂₂O₁₁) 22.5. Bubbled with 95% oxygen 5% CO₂ and perfused at 1–2 ml per min during recordings. Osmolarity ~300. External solution is prepared before each experiment. A 10x stock solution could be made and stored (up to one week) to speed up the preparation of the external solution (see Note 1).

2. Potassium-Gluconate internal solution contains (in mM):

K-gluconate 126, KCl 4, HEPES 10, ATP-Mg 4, GTP-Na₂ 0.3, Phosphocreatine 10, adjust pH to 7.2 with KOH. Measure osmolarity and ensure final osmolarity = 270–290 MΩs. Aliquot solution and store at −20°C for up to 6 months.

2.2. Equipment

The electrophysiology rig is modular and versatile, allowing multi-patch setups, extracellular recordings, and fluorescent visualization with the simple addition of different equipment. The following list of equipment is the minimum necessary components to build a rig for basic patch-clamp electrophysiology recordings. Adjust your setup accordingly depending on your experimental needs.

1. Air table

2. Faraday cage

3. Microscope (upright scope with a 40x water immersion lens, an inverted microscope can be used for 2D neuronal cells)

4. Micromanipulator

5. Patch clamp amplifier with headstage and micropipette holder

6. Digitizer

7. Computer and acquisition software

8. Inline solution heater

9. Recording chamber

10. Perfusion system

    1. Luer lock couplings
    2. Perfusion reservoir gas bubbler
    3. On/Off valves
    4. Syringe filters
    5. I.V. perfusion accessories
    6. Flexible Tygon tubing
    7. Peristaltic pump

11. Pipette puller

2.3. Additional materials:

1. Thin-walled borosilicate glass capillaries

2. MicroFil flexible needle

3. Eppendorf tubes

4. 10 mL and 1 mL syringes

5. P10, P20, P200, P1000 pipettes and pipette tips

6. Forceps

7. Kimwipes

8. Bleach

3. Methods

3.1. Recording protocols (samples provided using Molecular Devices Multiclamp 700B and pClamp 10 software):

1. Amplifier settings:

1. Bessel: 2 kHz

2. Zap: 25 μs, increase if needed

2. Spontaneous post-synaptic currents (sPSCs) recording protocol:

1. Acquisition mode: Gap free

2. Sampling rate per signal (Hz): 10000

3. 3–5 minutes duration

3. V-clamp step-depolarization protocol (whole-cell current responses):

1. Acquisition mode: Episodic stimulation

2. Holding potential −70 mV (should be adjusted accordingly)

3. Runs / trials: 1

4. Sweeps / run: 20

5. Sweep duration (s): 1

6. Start-to-Start Intervals

   1. Sweep (s): 5

7. First level (mV): −100

8. Delta level (mV): 10 (or 5 depending on the input resistances of the neuronal cell types)

9. First duration (ms): 500

4. I-clamp spontaneous action potential and postsynaptic potential recording protocol (I=0):

1. Acquisition mode: Gap free

2. 3–5 minutes duration

3. Sampling rate per signal (Hz): 10000

5. I-clamp step current injection protocol (to elicit action potentials):

1. Acquisition mode: Episodic stimulation

2. Runs / trials: 1

3. Sweeps / run: 16

4. Sweep duration (s): 2

5. Start-to-Start Intervals

6. Sweep (s): Minimum

7. First level (mV): −50

8. Delta level (mV): 10

3.3. Setup:

Carry out all procedures at room temperature unless specified otherwise. See notes 2-6 for instruction on reducing background electrical noise during recordings.

1. Bleach the recording and ground electrode (silver wires) overnight to provide a chloride coating on the silver wires. Thoroughly rinse the electrodes with ddH₂0 and dry with a Kimwipe prior to reassembling.

2. Prepare ACSF solution for recording by diluting the 10x stock solution to 1x with ddH20 in a volumetric flask obtaining a final volume of 500 mL. Add and dissolve 0.225 g glucose and 3.847 g sucrose. Bubble the patch-solution for 10 minutes with carbogen (95% O2 / 5% CO2). Add bivalent cations: 600 μL 1M MgCl₂ and 1250 μL 1M CaCl₂. Patch solution must be kept bubbling for the duration of the recording. Without proper bubbling, the calcium may precipitate out of solution.

3. Prepare K-Glu internal solution. Thaw an aliquot on ice and filter the internal solution through a 0.22 μm filter system. Transfer the filtered internal solution to a 100 μL stainless steel hypodermic needle. Keep the syringe on ice for the duration of the experiments.

4. Perfuse oxygenated patch ACSF solution at 2 mL/min and a temperature of 34°C (see Note 7.)

5. Pull glass patch pipettes. Aim for a resistance of about 5–7 MΩ (see Note 8).

6. Using a sterile transfer pipet, cut the tip off and gently transfer an organoid slice or coverslip to the rig chamber.

   1. Procedures for organoid slicing can be found in this publication (33).
   2. Ensure the perfusion system is turned off upon transferring to avoid inadvertent aspiration of the slice.
   3. Using a pair of forceps, gently move the slice or coverslip to the center of the perfusion bath.
   4. Using the forceps, set a harp on top of the slice, positioning it so at least two harp strings are anchoring the slice (see Note 9).

3.4. Obtaining the whole cell patch configuration:

1. Using the 4x objective, visualize and capture an image of the whole slice for documentation.

2. Switch to the 40x water immersion objective, ensuring that the objective is immersed in the ACSF solution.

3. Identify a cell to patch and center the cell on your screen (see Note 10).

4. Slightly raise the objective to provide room for the glass pipette to be inserted into the chamber and recording solution. Take care to ensure the objective remains submerged in the ACSF solution.

5. Use a clean MicroFil/microloader tip on a 100 μL or 1 ml syringe, fill a glass pipette with approximately 5 μL of internal solution.

   1. The microloader tip must be inserted as far down to the end of the glass pipette as possible.
   2. Flick your wrist to ensure the solution travels to the tip of the pipette, then gently flick the pipette several times to eliminate any bubbles in the solution (see Note 11).
   3. Internal solution must contact the inner electrode.

6. Attach the glass pipette to the pipette holder on the headstage (see Note 12).

7. Submerge the tip of the pipette into the bath solution and adjust the focus of the objective to visualize the pipette. Apply positive pressure (30–70 mbar) to eliminate any remaining air bubbles within the tip. The pipette resistance without positive pressure should measure to about 6–7 MΩ. Apply constant positive pressure to ensure no debris attaches onto the tip of the pipette prior to patching. A dirty tip will not form a high-quality seal.

   1. Applying positive pressure will push any debris in the internal solution out over time. Check for any debris clogging the internal tip of the pipette. Any clogs will be detrimental to the patch, and the pipette should be discarded and replaced.

8. Ensure that the Multiclamp is set to voltage clamp mode at this point.

9. Slowly lower the pipette in tandem with the objective. Take great care to keep the focus of the objective below the tip of the pipette. (This avoids crashing the pipette tip). However, if the focus goes significantly lower than the tip, you risk crashing the objective to the base of the pipette tip and breaking it off.

10. Lower the pipette to just above the slice surface. Use the ‘Auto Pipette Offset’ function to adjust the pipette offset current back to zero in voltage clamp mode.

11. Still applying positive pressure, approach the cell taking care to not make any contact with other cells or debris. The pressure from the internal solution should clear the way (see Note 13).

12. In one smooth motion, bring the pipette tip in contact with the cell membrane. The membrane will form a small “dimple” from the pressure.

    1. Check for a 0.1–0.2 MΩ access resistance increase. Immediately release the positive pressure.
    2. The dimple should relax and form a seal around the tip of the pipette. The access resistance should quickly rise. If not, apply a small amount of negative pressure.
13. Once the access resistance is over 100 MΩ, apply a holding voltage of −45 mV.

    1. If the access resistance does not increase to over 100 MΩ within a few seconds, abort the patch and try again with a different cell and a fresh pipette.
    2. Continue to apply small amounts of negative pressure until the access resistance reaches 1 GΩ or above, forming a gigaseal.
    3. Gently pull the tip of the pipette slightly backwards and up from the slice until the membrane looks relaxed and in the state it was prior to patching.

14. Once a gigaseal has been achieved, apply a holding voltage of −70 mV.

15. Use Auto Cp Fast on Multiclamp to compensate for pipette capacitance.

16. Apply small amounts of negative pressure in tandem with the zap function (25 – 50 us) to break into the cell (see Note 14).

    1. Upon breaking into the cell, the membrane test should show a capacitive transient resulting from the discharge of the membrane capacitance. Let the cell relax for about 5 minutes. If the current amplitude decreases, the patch is starting to seal back up. Apply a very small amount of negative pressure to open it back up.
    2. The access resistance should be less than 20 MΩ and remain unchanged or change less than 15%. Applying more negative pressure may help decrease the access resistance. Consider compensating for access resistance for whole cell current recordings.
    3. If the access resistance remains too high or if the capacitive current looks unstable, abort the recording, and try again with a different cell and fresh pipette.

3.5. Patch-clamp recordings:

1. After successfully establishing whole cell configuration, run the sPSCs protocol for 3–5 minutes to record any spontaneous currents while on voltage clamp mode (at a holding potential of −70 mV, for example).

   1. Record the membrane properties (membrane capacitance, membrane resistance, access resistance, tau, and the holding current) in a lab notebook.
   2. Abort the protocol if the baseline does not remain stable for the duration of the recording.
2. Check the membrane test again to ensure the patch is still open and the access resistance is still less than 20 MΩ.

   1. Apply negative pressure to reopen the patch if the access resistance is too high. If the access resistance remains too high or the capacitive current becomes unstable, abort the patch and try again with a different cell and fresh pipette.

3. Apply whole cell compensation to adjust for cell membrane capacitance. Consider compensate access resistance for voltage-dependent whole-cell current recordings.

4. Run the V-clamp step-depolarization protocol.

5. Disable whole cell compensation and switch to the I-clamp spontaneous protocol.

6. Switch the configuration to current clamp (I=0) mode.

   1. The amplifier’s internal circuitry must briefly reset switching between current and voltage clamp mode. It is crucial to always go through I=0 mode when switching between current and voltage clamp.
7. Run the I-clamp spontaneous protocol to record spontaneous action potentials as well as postsynaptic potentials.

   1. After switching to I=0, let the cell’s resting membrane stabilize before recording.
   2. If the resting membrane continues to increase for the duration of the recording, the cell is unhealthy or dying and the recording must be aborted.

8. Run the I-clamp step current injection protocol. Inject currents to keep basal membrane potential closer to −65 mV if desired.

9. After all acquisition protocols have finished recording, release the negative pressure, and carefully retract the pipette backwards and replace the pipette to record a new cell. Data collected are to be analyzed using appropriate analysis software.

4. Notes

For your convenience, prepare a 10x stock ACSF solution. The stock ACSF solution must be stored at 4°C and used within one week. The molecular weights of the reagents are as listed (g/mol): NaCl 58.44, KCl 74.55, NaH₂PO₄∙H₂O 137.99, NaHCO₃ 84.01, MgCl2 95.21, CaCl2 110.98, glucose 180.16, sucrose 342.30. For 1L 10 x stock, NaCl 73g, KCl 1.86g, NaH₂PO₄∙H₂O 1.73 g, NaHCO₃ 21g, dissolved in ddH₂O to final volumn of 1000 mL. Note that CaCl₂, MgCl₂, sucrose, and glucose are not added to the stock solution. It is imperative that the stock solution is diluted to 1x ACSF prior to the addition of the sucrose and glucose, and the 1x solution must be bubbled with carbogen prior to the addition of CaCl₂ and MgCl₂, otherwise precipitate will form and the solution must be discarded. For 500 mL ACSF, add 0.225 g glucose and 3.847 g sucrose, 0.6mL 1M MgCl₂ and 1.25 mL 1M CaCl₂.

1. Prior to the start of electrophysiology recordings, ensure that the rig has minimal recording noise to ensure high-quality recordings for the duration of the experiment. Most of the electrical noise will be eliminated by grounding the antivibration air table and the protective Faraday cage but take great care that nearby electrical devices such as cell phones, computers, and nearby power sources are not contributing to noise in your system. Ensure all electrical components are grounded and the grounding wires have not inadvertently formed grounding loops.

2. As the first course of action, hook up a ‘model-cell’ to the amplifier headstage. A model-cell replicates the Resister-Capacitor (RC) circuitry of the microelectrode in bath, the electrode in contact with a cell membrane, and the electrode after successfully breaking into the cell. The traces from all three configurations should be steady, if not, the noise issues originate from the software/rig hardware and all settings should be reset.

3. Switch off or disconnect all peripheral devices on the rig including the perfusion system, pipette holder, camera, and the manipulator. Test each device and watch if they are contributing to noise in the recordings.

4. Ensure the rig is entirely dry aside from the flow into the chamber from the perfusion system. Any solution inside the pipette holder or contacting electrical or metal components on the rig may be transmitting small amounts of current and causing noise.

5. A loose seal within the micropipette electrode holder can also contribute to noise. Check the O-rings to ensure they are not damaged, and the seal is tight. With a micropipette inserted into the holder, apply positive pressure, and visually check to see a constant flow of solution from the pipette into the bath. The stream of pressure should remain stable and not weaken over the course of at least one minute.

6. Take care not to let the bath solution rise too high or fall too low in the recording chamber. Allowing the bath solution level to fluctuate may alter flow, leakage, and/or noise or vibrations while recording. Furthermore, make sure that the input/output of the perfusion system are directly opposite each other to ensure adequate flow over the center of the bath where the slice or coverslip is positioned.

7. The optimal resistance of the glass pipette electrodes depends on the size of the cells in the culture. Organoid cells are generally small and delicate, so a higher resistance of 8–9 MΩ allows for good patch formation. Larger pipettes tend to suck the whole cell into the pipette upon breaking into the membrane. In contrast, mouse brain slices and induced neuronal cultures on coverslips have cells that are larger and well anchored. Larger pipettes with lower resistance (4–7 MΩ) allow for lower access resistance and easier breaking into the cell.

8. Smaller organoid slices may not be large enough to pin down with 2 harp strings. Instead, place the harp string directly through the center of the slice. Rotate the harp to ensure that the harp strings are perpendicular to the micropipette when patching. This is the least obtrusive configuration of the harp, allowing you to patch cells close to the harp without the micropipette contacting the strings and disrupting movement. Remove the harp and rotate the slice to patch other regions, rather than moving the pipette to the other side of the slice.

9. Choosing a healthy neuronal cell is critical for successful patching. Under brightfield 40x magnification or DIC imaging, healthy cells will have a well-defined soma with a 3D cell shape. Avoid cells with a rougher or dark edge or shriveled “raisin” like appearances. These cells are unhealthy or dying and are unlikely to form good seals and produce quality recordings. Cells that look swollen or too circular should also be avoided. If your capture software has the feature, mark the cell so you know where to look for it after you refocus the objective on the slice.

10. It can be difficult to see the bubbles in the solution inside the patch pipette. Raise the pipette above your head and towards a light source to see the bubbles better.

11. Ensure the tip of the silver electrode attached to the headstage is submerged in the internal solution but not contacting the tip of the patch pipette. Tightening the glass seal compression caps and applying positive pressure may move the electrode slightly forward breaking the tip of the pipette.

12. Neuronal cells in 3D organoids are quite fragile. Patching cells deep within the slice requires positive pressure to push away other cells and debris, but at the risk of agitating the cell of interest. Approaching the cell at an angle rather than directly on top of the cell will help keep the cell anchored and prevent rough first contact, greatly increasing the chance of a successful seal and patch. If the cell is deep within the slice, do not bring the pipette directly down near the top of where the cell is. Clearing the tissue on top of the cell for too long will stress the cell and displace synapses on the soma. Clear a path on the slice approximately 50 μm away from the cell of interest, moving the pipette diagonally towards the cell applying positive pressure. Quickly move the pipette back and forth in a prodding motion, clearing cells in the way. The first contact is critical, do not touch the cell of interest until no cells are obstructing the way and the angle is perfect.

13. It may be easier to use mouth suction to break into the cell. Using a p1000 pipette tip inserted into Tygon tubing leading to the side port of the pipette holder, apply a small amount of suction with your mouth and quickly pull out the pipette tip. This action will produce a “kissing” sound. One or two “kisses” should be enough to break into the cell. The zap function can be used in tandem with brief suction to help rupture the membrane. Experiment with the force and duration of suction depending on the cell’s health as observed by the state of the capacitive current.