Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Summary

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Abstract

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Introduction

Glutamate receptors mediate the majority of excitatory synaptic transmission at central nervous system synapses. The two major subtypes of ionotropic glutamate receptors localized at the spine head of the postsynaptic membrane are N-methyl-D-aspartate (NMDA) and α-amino-3-hydroxy-5-methylisoxazole-4-proprionic acid (AMPA) receptors. At resting membrane potentials, AMPA receptors carry most of the postsynaptic current during synaptic transmission. In the hippocampus, the NMDA receptor plays a key role in triggering changes in the number of AMPA receptors in the postsynaptic membrane: by acting as a ”coincidence detector”¹ to initiate changes in synaptic strength¹, the NMDA receptor participates in the synaptic mechanisms that are thought to underpin learning and memory at a subcellular level. In response to depolarization of the postsynaptic neuron in parallel with presynaptic transmitter release, calcium enters via the NMDA receptor to initiate AMPA receptor insertion or removal². These receptor dynamics underlie synapse plasticity: an increase in synaptic strength is long-term potentiation^2,3 (LTP), while a decrease in synaptic strength is long-term depression⁴ (LTD). Therefore AMPA receptor movement is thought to be responsible for synaptic plasticity expression, while NMDA receptors are thought to control its induction.

Determining the precise mechanisms underlying synaptic transmission and plasticity requires studying small populations of synapses, ideally single synapses. While some synapses are highly suited for study at this level, e.g., the Calyx of Held⁵, for most synaptic populations this is extremely difficult due to the small and diffuse nature of the synaptic connections. Two major electrophysiology techniques have been developed to examine single synaptic connections: The first is minimal stimulation, where one presynaptic fiber is presumed stimulated extracellularly. The second technique is paired recordings, where two simultaneous whole cell recordings from synaptically connected neurons is performed. A major advantage of minimal stimulation is that it is rapid and relatively simple to perform, involving placement of an extracellular stimulating electrode into the axonal tract while simultaneously recording from a postsynaptic neuron. The primary concern when using this technique is that reliable stimulation of a single cell can rarely be guaranteed trial after trial.

Over the past fifteen years we have routinely used paired whole-cell recordings from two synaptically connected pyramidal neurons^6-17. The major advantage of this technique is that only one presynaptic neuron is consistently and reliably stimulated. It also allows not only electrophysiological characterization but also pharmacological manipulation of the presynaptic neuron^6,18. However, the probability of synaptic connectivity between neurons is low, making connected pairs difficult to obtain¹⁹. The use of organotypic brain slice cultures circumvents this obstacle as synaptic connectivity can re-establish in vitro and moreover the nature of the resulting connectivity is similar to that in native brain tissue²⁰. In addition, organotypic cultures express LTP, LTD^{7-10,12-15,21} and additional forms of short-term synaptic plasticity including paired-pulse facilitation (PPF) and depression (PPD)^6,22,23, enabling plasticity mechanisms to be studied in pairs of neurons. Here we describe the detailed methodology involved in successfully attaining paired recordings in this in vitro system. This information can readily be adapted to other experimental systems, including acute slices and other brain regions.

Protocol

Animal Ethics Statement:

The protocols described in this manuscript follow the animal care guidelines established by The University of Auckland and Stanford University. P7 rat pups were euthanized by rapid decapitation. Hippocampal dissection is then immediately performed as described below.

1. Organotypic Hippocampal Slice Culture

1. Preparation
   1. Prepare dissection Medium (used only for dissecting the brain). Combine 200 ml Minimum Essential Medium, 2 ml penicillin-streptomycin solution (10,000 units of each, in 0.85% NaCl), 5 ml HEPES buffer solution, 2 ml 1 M Tris stock solution (pH 7.2), and filter sterilize with 0.22 μm filter. Carry out hippocampal dissection in ice-cold dissection medium.
   2. Cool the dissection medium. Place the dissection media in the freezer approximately 1 hr prior to beginning the dissection until the liquid is very cold. Do not allow large ice crystals to form. Store on ice until required.
   3. Prepare culture medium (used for everything except dissection of hippocampi). Combine 100 ml Minimum Essential Medium, (1x concentration, liquid) w/Hank’s salts, w/ L-glutamine, 2 ml penicillin-streptomycin solution (liquid, 10,000 units of each, in 0.85% NaCl ), 2.5 ml HEPES 1 M buffer solution, 50 ml Hank’s Balanced Salt Solution, 50 ml Horse Serum (defined, heat inactivated), and filter sterilize with 0.22 μm filter.
   4. Prepare the culture dishes. Place 1 ml of culture media per 35 mm culture dish, and add a membrane insert to each dish. Put up to seven of these dishes into one 150 mm Petri dish (referred to hereafter as a ‘plate’). Place the plate in a CO₂ incubator for at least an hour before the dissection begins so that the culture medium in the dishes attains the proper temperature and pH.
2. Dissection of the Hippocampi from Rat Pups at Postnatal Day 7 (P7)
   1. Pre-sterilize all dissection tools under UV light before the procedure.
   2. After rapid decapitation, remove the brain and place into chilled medium in one dish, and then remove it to a piece of moistened filter paper for the dissection. Tease the cortex away from the midbrain using blunt smooth plastic-coated miniature spatulas, exposing the hippocampus. Cut the fornix, and then gently work the spatula underneath the hippocampus to flip it out (Figure 1A).
      NOTE: Successful slice cultures can be prepared using animals up to P10.
   3. Trim the isolated hippocampus away from the rest of the brain. Transfer the hippocampi into a new dish containing chilled dissection media using a moistened soft paintbrush (e.g., white sable #4).
   4. Clean the underside (i.e. the flatter side) of the hippocampus of choroid plexus during dissection, as these spongy, meninge-like tissues make it difficult to separate hippocampal slices from one another later on.
      NOTE: Leave these tissues in place if they cannot be teased away gently.
   5. Perform the entire dissection as quickly as possible without damaging the hippocampi.
      NOTE: A careful dissection is more important than a fast one, as long as the experimental conditions are chilled. Slice the hippocampi from three rats simultaneously. The slice health is not affected as long as the total time from start to when the plated slices go into the incubator is under 30 min.
3. Slicing
   1. Slice the hippocampi transversely into 400 μm cross-sections using a manual tissue slicer. The method by which the slicing is conducted is not important, so long as it is not overly damaging to the tissue and produces slices of consistent thickness with readily visible laminar architecture (i.e. Ammon’s Horn is easily seen to be preserved in the tissue).
   2. Lie the hippocampi on the stage of the tissue chopper on top of a triple thickness of #2 filter paper. The hippocampi are sliced entirely without removing any slices individually (i.e. the entire hippocampus is left on the stage of the chopper until all cuts have been made; rather like loaves of sliced bread at this point; Figure 1B).
   3. Transfer the hippocampi into dissection medium using a soft paintbrush laid next to each hippocampus (white sable #2). Gently push each hippocampus to the side to break the adhesion between the hippocampus and the underlying filter paper. Roll the brush underneath each hippocampus to pick it up off the stage. Place all hippocampi into the same 35 mm Petri dish of chilled dissection medium. Separate the hippocampi into individual slices by manually agitating the dish in alternating clockwise and counterclockwise motions.
   4. Inspect the slices under a dissecting microscope and discard any which are damaged, small, or which do not possess clearly visible cell body layers.
      NOTE: It will often be the case that not all the slices are successfully separated from their siblings in this manner. In this case, they can be separated by turning them on edge and prodding them with the tips of fine forceps. “Good” slices are then transferred to another clean Petri dish containing chilled dissection medium using a cut off and fire-polished Pasteur pipette (Figures 1C-E).
4. Storage
   1. Place the individual slices onto the membrane filter inserts. Using a Pasteur pipette cut off and fire polished to give a larger bore, individually transfer slices to the culture inserts.
      NOTE: The size of the bore created when cutting and fire-polishing the pipette is important. Too small and the slices will not easily leave the pipette for the membrane. Too large and excess fluid is placed on the surface of the membrane along with the slice. The best bore diameter is typically about half the diameter of the barrel of the pipette. When cutting, this diameter can be chosen by cutting at the proper place on the taper of the pipette.
   2. Slice Transfer
      1. Fill the pipette with medium first, and then suck up a single slice with small suction forces. This keeps the slice near the opening of the pipette and prevents too much medium expulsion into the insert to place the slice.
      2. Apply a slight pressure on the bulb to form a small hanging droplet, allow the slice to settle into that droplet, and then touch that droplet to the membrane and let the slice “fall” onto the membrane. Generally three slices are placed onto each membrane insert.
         NOTE: If the fluid droplet is too large, the droplets from the three slices tend to merge on the membrane surface, and the slices will thus all gather to the center of the membrane, making it more difficult to separate them for electrophysiological recording later.
   3. Fluid Removal
      1. Aspirate any excess medium from the top of the culture plate insert for all slices in that plate (3 slices per insert = 21 slices per plate), so that slices are not immersed in or surrounded by a pool of dissection medium.
      2. Perform this is with an uncut, but fire-polished Pasteur pipette. Allow the slice to settle and adhere to the membrane for a minute before pipetting. Take care not to suck the slice up into the pipette. Carry out fluid removal for the inserts that are plated first, to allow sufficient time for the slices to settle.
   4. Slice Storage
      1. Store slice cultures in a 5.0% CO₂ incubator at 37 °C. Observe the final arrangement of the cultures in Figure 1E. The individual slices rest upon a culture plate insert which has a porous membrane for a bottom, and this insert fits inside a Petri dish filled with a small amount of medium; by this means, the slices are not immersed in the medium, but have access to it only through the membrane at the bottom of the culture plate insert.
      2. Optionally, remove the slices may for study either by removing the culture plate insert or by cutting out a section of insert membrane containing a slice.
5. Maintenance
   1. Exchange the medium in the Petri dishes the day after making cultures. Transfer the culture plate insert to a new petri dish containing 1 ml of fresh culture medium that is equilibrated in the incubator for at least an hour prior to transfer.
   2. Change the medium again in the same manner on the third day after making cultures. On the third day, transfer the cultures to an incubator set at 34 °C. Change the medium twice each week (every 3-4 days).
   3. Identify healthy cultures. Select healthy cultures for paired whole cell recordings.
      NOTE: Healthy cultures have a well-defined edge and a clearly defined pyramidal cell layer. Cultures with dark (likely necrotic) areas present or a vacuolated appearance with flattened borders are rejected. Employing these criteria, usually two-thirds of slice cultures are sufficiently healthy for recording.
   4. Utilize these cultures within a fairly small window of time. Do not used tissues that are cultured for more than two weeks since the neurons begin to display slightly epileptiform behavior which slowly worsens with time. The exact upper limit of neuronal survival is not known but it is possible to record from functioning pyramidal cells as late as 16 weeks in culture.

2. Paired Whole Cell Recordings

1. ACSF and Intracellular Solutions
   1. Prepare presynaptic electrode solution by mixing (in mM) 120 potassium gluconate, 40 HEPES, 5 MgCl₂, 2 NaATP, and 0.3 NaGTP (pH 7.2 with KOH; osmolarity: 290 mOsm). Whilst the same electrode solution is used for postsynaptic neurons, cesium gluconate (120 mM) is typically used as the major salt in the postsynaptic electrode, plus 5 mM QX314.
      NOTE: This enables stable voltage clamping at positive potentials to record postsynaptic NMDAR-mediated currents^8-10,13. It also prevents the potassium-induced hyperexcitability of the presynaptic neuron by the internal solution flowing from the postsynaptic recording electrode before a giga ohm seal is made.
   2. Prepare postsynaptic electrode solution composing (in mM) 120 cesium gluconate, 40 HEPES, 5 MgCl₂, 5 QX314, 2 NaATP, and 0.3 NaGTP (pH 7.2 with CsOH; osmolarity: 290 mOsm). Pull glass microelectrodes at 5-10 MΩ resistance and fill with filtered internal solution.
   3. Prepare artificial cerebrospinal fluid (ACSF) composing (in mM) 119 NaCl, 2.4 KCl, 1.3 MgSO₄, 2.4 CaCl₂, 1 Na₂HPO₄, 26.2 NaHCO₃, 11 glucose, pH 7.4, saturated with 95% O₂, 5% CO₂. NOTE: If AMPAR-mediated responses need to be blocked for examination of NMDAR EPSCs, include 10 μM CNQX or NBQX in the ACSF.
2. Obtain Paired Whole Cell Recordings
   1. To examine synaptic transmission between two individual neurons, designate one neuron as the presynaptic neuron and hold in current clamp to induce action potentials and initiate synaptic transmission.
   2. Designate the second neuron, as the postsynaptic neuron, and hold it in current or voltage clamp depending on the information required by the researcher. Obtain detailed examination of the glutamatergic currents, with AMPAR-mediated EPSCs examined at -65 mV and NMDAR-mediated EPSCs examined at +30 mV^6-16 by holding the postsynaptic neuron in voltage clamp
   3. Establish a paired recording. To establish a paired recording, the presynaptic whole cell recording is always obtained first, enabling multiple sequential postsynaptic neurons to be obtained until one that is synaptically connected is obtained. If performing synaptic plasticity experiments, it is especially critical to obtain the presynaptic neuron first as the induction of LTP in the postsynaptic neuron must be initiated within 10 min of obtaining the whole cell recordings to prevent washout of cytoplasmic factors required for LTP^8,19.
   4. Avoid movement-induced cell loss. A major challenge when performing paired whole cell recordings is avoiding vibration and movement-induced disruption of the first whole cell recording whilst obtaining the second recording.
      1. Obtain the first whole cell recording and then move the microscope lens 10-200 μm in the x-axis so that the established recording is located at the edge of the area visible on the monitor (Figure 2A).
      2. Raise the lens of the microscope ~5-10 mm while ensuring that the ACSF still maintains contact with the lens. Mount the second electrode and guide into the fluid meniscus (Figure 2B).
      3. Move the electrode in the x-y plane until it is directly under the light path through the lens but still significantly above the established recording electrode. Once the electrode is visible under the microscope, ensure the tip is at the far edge of the monitor away from the first electrode.
      4. Move the second electrode down in sequence with the microscope focus until both electrodes are in the same focal plane. It is important that the level of positive pressure is strong enough to avoid tip blockage but not too strong so as to disrupt the first whole cell recording. This translates to the positive pressure inducing movement in 2-3 neurons from the electrode when it first enters the slice.
      5. Locate a postsynaptic partner in a similar focal plane to the first presynaptic recording (Figure 2C). Recording is generally Obtain paired recordings between CA3 pyramidal neurons the second whole cell from a 0-200 mm neuron of the first recording (Figures 2C,D).
         NOTE: Synaptic transmission between neuronal pairs are stable over time periods of up to 3-4 hr when using these standard whole cell recording techniques.
3. Synaptic plasticity: Once Upon obtaining a successful synaptically-connected pyramidal cell pair (Figures 3A,B), examine the characteristics of synaptic transmission and plasticity.
   1. LTP induction.
      1. Induce LTP by pairing presynaptic action potentials (1 Hz) with postsynaptic depolarization to -10 to 0 mV for 1 min (Figure 3C)^7-9,12-14. Initiate pairing within 10 min of break-in to the postsynaptic neuron.
         NOTE: LTP can also be induced with both neurons held in current clamp by pairing presynaptic and postsynaptic action potentials at 1 Hz for 1 min, with postsynaptic action potentials elicited 10 msec following injection of current into the presynaptic neuron⁸.
   2. Further induce LTD by low frequency stimulation (LFS) at 1 Hz combined with a slight depolarization of the postsynaptic neuron to -55 mV for 5-10 min (Figure 3C)⁹.

Results

Synaptic connectivity is evident by stimulating the presynaptic neuron to fire an action potential by passing a depolarizing current pulse (typically 20-50 pA for 20 msec) via the recording electrode. The postsynaptic current trace is then examined for the presence of a monosynaptic EPSC evoked at short (<5 msec) and consistent latencies after the peak of the presynaptic action potential (Figure 3A). In most experiments multiple postsynaptic neurons are tested before a synaptically-connected pair can ...

Discussion

Here we have described the requirements for establishing successful paired whole cell recordings in organotypic hippocampal slice cultures. Paired recordings can also be performed in multiple preparations, including acute slices and dissociated culture systems^26,27. While the focus here has been on the induction of longer forms of synaptic plasticity (namely LTP and LTD), it is important to highlight that paired whole cell recordings in organotypic, acute slice and dissociated cell preparations have provided i...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Minimum Essential Medium			Stable motorized micromanipulators
Penicillin-Streptomycin solution			Shallow tissue bath
HEPES buffer solution			DIC camera
1 M Tris stock solution			Amplifier
Hank’s Balanced Salt Solution			Computer
Horse Serum			Vibration isolation table
Plastic-coated miniature spatulas			Upright microscope
Soft paintbrush			Data acquistion and analysis software
Manual tissue chopper			Electrode puller
#2 Filter paper			Faraday cage
#5 Forceps
Membrane inserts
CO2 incubator
Dissection hood
Class II hood