Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

All experimental procedures have been carried out in accordance with the EU Directive for the Protection of Animals, the German Animal Welfare Act (Tierschutzgesetz) and the Guidelines of the Federation of European Laboratory Animal Science Association.

1. Set-up for Electrophysiology

Before commencing with paired recording, an electrophysiology set-up has to be built. A brief outline how such a set-up is assembled is given below:

1. Install an anti-vibration table on which the microscope, the manipulators and all other equipment will be placed.
   NOTE: Vibration or any other kind of movement needs to be as small as possible when recording from synaptic connections because this requires a change of pipettes (searching electrode replaced by recording electrode).
2. Place a Faraday cage around the anti-vibration table to reduce electrical noise. Connect all equipment inside the Faraday cage with the electrical ground.
3. Install a microscope with a motorized focus axis that is based on a motorized x-y table on the anti-vibration table according to the manufacturer’s guidelines. This allows to reliably move the microscope to neurons that are not in the same field of view as it is the case for translaminar connections in the neocortex.
4. Install a video camera on the camera port of the microscope and connect it to analog and/or digital screens to observe the slice preparation and the movement of electrodes. Use a camera, which can be switched between a low- and a high-power magnification to obtain an overview and a close-up image, respectively, of the synaptically coupled neuronal cell pair. This also helps to control the movement of the patch electrodes.
5. Install a specimen table containing an inset for the bath chamber (drilled in-house from a Perspex block) for the brain slice preparation. This specimen table must not be connected to the microscope, i.e., must be fixed to the air table and the microscope should be maneuverable under it.
6. Mount two (or more if required) high-precision micromanipulators that can be moved in all three dimensions. Install patch-clamp pre-amplifier on the manipulators and connect them to the main amplifier.
   NOTE: If required additional manipulators can be installed for e.g., more than two pre-amplifiers, extracellular stimulation electrodes, drug-delivery systems, etc.
7. Place the bath chamber in the specimen table. Install a solution inlet and outlet and connect them to the perfusion system.
8. Use a peristaltic pump to maintain the perfusion of the recording chamber with artificial cerebrospinal fluid (ACSF; Table 1) at a stable flow rate of 4 – 6 ml/min for optimal slice viability.
   NOTE: Do not exceed this flow rate significantly as it will result in turbulence in the ASCF and hence movement of the slice.
9. Install holders for the temperature probe and Ag/AgCl ground electrode onto the specimen table. This is necessary to allow the probe and electrode to sample the bath in the chamber.
10. For a good visibility of neurons in the slice preparation use infrared illumination in the microscope. This reduces light diffraction and hence image blur. Ensure that the microscope is equipped with differential interference contrast optics to achieve a ‘3D’-like visualization. This will help patching neurons and allow targeting neurons deep in the slice (60 µm or deeper).
11. During the experiment, keep brain slices in an experimental chamber with a glass coverslip at the bottom. Place a ‘platinum harp’ on the top of slice to prevent slices from floating. A platinum harp is made from U-shaped, flattened platinum wire with strings of dental floss glued onto it.
12. Keep the ACSF perfusing the slice at a temperature of 32 – 35 °C to ensure optimal control of oxygenation and pH. This can be done by heating the solution with a Peltier device installed close to the ACSF inflow into the bath chamber.
    NOTE: Use ultrathin cover slips so that even microscope condensers with a high numerical aperture and short working distance can be focused on the specimen. This is necessary for an optimal illumination of the slice.
    For an image and a description of an electrophysiology set-up optimized for paired recordings in the brain slice preparation see Figure 4 in Ref. ³⁴.

2. Brain Slice Preparation

1. Mildly anesthetize the animal from which the brain tissue should be taken with isoflurane (final concentration <0.1%).
   NOTE: Other anesthetics can also be used. Use of the anesthetic should comply with the respective animal committee’s recommendations and rules. Always ensure that the animal is not irritated or under stress after adding the anesthetic.
2. Decapitate the animal, open the skull and remove the brain as quickly as possible using the procedure described in Refs. ^34,35.
3. Transfer the brain into cooled (4 °C) artificial cerebrospinal fluid aerated with a mixture of carbogen gas with 95% O₂ and 5% CO₂.
4. Isolate the brain region that should be studied.
   NOTE: The parameters to obtain optimal preservation and minimal truncation of dendrites and axons of pre- and postsynaptic neurons should be determined in advance for each neuronal connection and developmental stage under investigation. In an optimally dissected brain slice, the axon of the presynaptic neuron is not required to be parallel to the slice surface. Rather, they should point into the slices with a shallow angle. The applies for the apical dendrite of postsynaptic pyramidal neurons; this should be checked under the light microscope. This is particularly important for non-local or translaminar synaptic connections, i.e., where pre- and postsynaptic somata are more than 200 µm apart and the probability of dendritic and axonal truncations is likely to be substantially higher than for local connections.
5. Transfer the brain to the microtome chamber. Based on empirical findings, different extracellular slicing solutions are used depending on the age of the animal (see Tables 2 and 3; helpful information can also be found under ‘http://www.brainslicemethods.com/’).
6. Trim the brain until the target region is visible.
7. To obtain slices with a good cell visibility, cut 300 – 400 µm thick brain slices if the animal is mature (>3 weeks). If the animal is immature (1^st to 2^nd postnatal week for mice and rats) slices may be cut up to a thickness of ~600 – 800 µm without substantially impairing cell visibility.
   NOTE: This will improve the stability of ‘immature’ slices and reduce the number of possible truncations of long-range dendritic and axonal collaterals.
8. Transfer slices from the microtome chamber to an incubation chamber filled with slicing solution aerated with a mixture of 95% O₂ and 5% CO₂. Keep the slices in the incubation chamber for at least 30 min to 1 hr either at RT or at ~30 °C, depending on the type of experiment.

3. Paired Patch-Clamp Recording and Biocytin Filling

Depending on the type of synaptic connection three different approaches are used to find synaptically coupled neurons. If the connection probability is low (as can be expected for most excitatory connections), proceed as follows:

1. Neuronal connections with a low connectivity
   1. Fill the patch pipettes with an internal solution of the composition listed in Table 4. For all recordings in the whole-cell configuration add biocytin at concentrations between 3 – 5 mg/ml to the internal (pipette) solution to obtain good staining results for the pre- and postsynaptic neurons. Let biocytin diffuse into the cell for at least 15 – 30 min (depending on the size of the neuron).
      NOTE: Do not add biocytin to the solution used in the ‘searching’ pipettes mentioned below.
   2. Patch a putative postsynaptic neuron in the whole-cell mode. Use patch pipettes with a long and slender shank. This facilitates the maneuverability of the pipette under the objective and reduces movement artifacts that may occur when the electrode is introduced in the slice.
   3. Then, patch a potential presynaptic neuron in a cell-attached configuration using a ‘searching’ patch pipette of 8 – 10 MΩ resistance and a small tip diameter. With these pipettes establish a ‘loose’ cell-attached patch. Fill the ‘searching’ pipette with an internal solution in which K⁺ is replaced by Na⁺ (Table 5) in order not to depolarize neurons during searching for a presynaptic cell.
      NOTE: The ‘loose’ seal resistance is not in the GΩ range but typically around 30 – 300 MΩ.
   4. Hold the membrane potential under the ‘loose’ seal to about -30 to -60 mV in current clamp mode. Then, apply large current pulses (0.2 – 2 nA) to elicit an action potential in the potential presynaptic cell. Observe this action potential as a small spikelet on the voltage response.
   5. Set the stimulation frequency to 0.1 Hz to prevent run-down of a postsynaptic response. Use lower stimulation frequencies in case the brain tissue is very immature or the synaptic connection not very reliable.
   6. In case the ‘postsynaptic’ neuron does not respond to stimulation of the tested ‘presynaptic’ neuron, patch a new neuron in ‘loose’ seal mode. Do not replace the ‘searching’ patch electrode as long as the seal with a resistance of >30 MΩ is established. Test up to 30 potentially presynaptic neurons in this way.
      NOTE: To prevent damaging the neuron during the searching procedure with loose patch clamp, only gently suction should be applied to the searching pipette compared with that applied to the whole-cell patch pipette.
   7. If stimulation in the ‘loose’ seal mode results in an EPSP/IPSP with a latency <5 msec in the postsynaptic neuron remove the ‘searching’ pipette from the presynaptic neuron.
   8. Replace the ‘searching’ patch pipette with a patch electrode filled with biocytin-containing, regular internal solution. Ensure this recording patch pipette has a resistance of 4 – 8 MΩ; the larger the soma size the lower the electrode resistance can be.
   9. Patch the presynaptic neuron and record in whole-cell, current-clamp mode. Elicit action potentials by current injection in the presynaptic neurons and record the postsynaptic response. Store the data on a computer for later off-line analysis.
2. Neuronal connections with a high connectivity
   NOTE: For neuronal microcircuits with a high connectivity ratio (> 30%) use a different procedure to find synaptic connections. This procedure is outlined below and is often used for inhibitory synaptic connection that have on average a higher connectivity ratio than excitatory connections³⁶. It should not be used when the connectivity ratio falls below this value.
   1. Patch a presynaptic neuron directly in the whole-cell mode. Then, patch a potential postsynaptic neuron, also in whole cell mode. Both cells should be under current clamp control. Elicit an action potential in the presynaptic cell. Monitor the prospective postsynaptic neuron for a postsynaptic potential.
   2. In case the neuron does not show a response to presynaptic stimulation, patch a new neuron in whole-cell mode using a patch electrode filled with regular internal solution. If a connection is found, do not change the patch pipette but proceed with recording. Then, proceed as in step 3.1.9.
3. Neuronal connections via gap-junctions
   NOTE: To search for electrical (gap-junction) connections use the following procedure which is a modified version of that used for low-probability connections.
   1. Patch a postsynaptic neuron in the whole-cell patch clamp configuration.
   2. Subsequently, patch a putative presynaptic neuron in the loose-seal configuration (low seal resistance of 30 – 300 MΩ) using a ‘searching’ patch pipette of 7 – 10 MΩ resistance. This pipette should be filled with the modified high Na⁺ internal solution for loose-seal stimulation.
   3. Inject 100 – 300 msec long hyperpolarizing current pulses via the ‘loose’ seal; the pipette command potential should be set to about -60 to -70 mV. Observe a gap junction connection if this stimulation results in a small hyperpolarizing response in the ‘postsynaptic’ neuron.
   4. Repatch the ‘presynaptic’ neuron in the whole-cell mode and proceed as described above.
      NOTE: To ensure a good staining of the axonal and dendritic processes, use the protocol given below. This protocol is described in brief here, however, a detailed protocol for histochemical processing used in our laboratory is given in Ref. ³⁷.

4. Histochemical Processing

1. Transfer the brain slice with the pair of synaptically coupled neurons into a vial containing 4% paraformaldehyde dissolved in 0.1 M phosphate buffer and fixate the slice for 24 hr. For electron microscopy, fixate the slice using 2.5% glutaraldehyde and 1% paraformaldehyde.
2. On the next day, transfer the slices into normal phosphate buffer containing no fixative. Wash slices with phosphate buffer for 6 – 8 times for 10 – 15 min each to remove excess fixative.
3. Incubate slices for 20 min in a 3% H₂O₂ solution in 0.1 M phosphate buffer to minimize the endogenous peroxidase reaction. Observe strong bubble formation. Continue the reaction until no further bubbles are produced. Then, rinse the slices in 0.1 M phosphate buffer for 6 – 8 times (for 10 min each) to remove any remaining H₂O₂.
4. Immunocytochemical and chromogenic reactions
   Visualization of biocytin-filled neurons is based on a streptavidin-biotinylated horseradish peroxidase reaction catalyzed with diaminobenzidine; this produces a dark precipitate that produces a crisp stain with a high spatial resolution. The following procedures are used for the chromogenic reaction:
   1. Prepare the ABC solution as per the manufacture’s protocol. Add to 14.55 ml phosphate buffer supplemented with 150 µl 10% Triton X100 (Table 6; only for light microscopy) and keep the solution for about 30 min in the dark. Incubate slices in this solution O/N before use.
   2. Incubate slices in ABC solution on a shaker for 1 hr at RT and in the dark. Subsequently, keep slices O/N at 4 °C in a fridge. On the next morning, incubate slices at RT for an additional hour.
   3. After this, rinse slices at least four times for 10 min each in phosphate buffer. Then, incubate on a shaker for about 30 min in the dark in 2 ml of a nickel-intensified 3-3’-diaminobenzidine solution (Table 7). Be very careful when handling diaminobenzidine and use only under the fume hood; it is extremely carcinogenic even at very low concentrations.
   4. Add 6.5 µl 3% H₂O₂ to the diaminobenzidine-containing solution. Monitor the chromogenic reaction under a light microscope until brown-black stained neurons are clearly visible. Then, stop the reaction immediately by washing slices several times in phosphate buffer.
   5. Mount slices with stained neurons on adhesive, silane-coated Histobond or gelatinized standard object slides.
   6. Keep the slides in a moist chamber with at least 80% humidity O/N. On the next day, let the slices air-dry for a further 10 min. Prior to embedding, transfer slides in solutions with increasing concentrations of ethanol (20 – 100%) in order to dehydrate them.
      NOTE: The dehydration process should be slow otherwise distortions in the dendritic and axonal processes are likely to occur³⁷. If an alternative embedding procedure is chosen, e.g., one with the glycerol-based Moviol, a dehydration is not required; however this is likely to result in an inhomogeneous compression of the slice and hence a distorted neuronal morphology.
   7. Finally, incubate for 10 min in xylol, embed the slices in a hydrophobic embedding medium for light microscopy and place an ultra-thin coverslips on top. Let slides dry for 20 min at RT.

5. Neuronal Reconstruction and Synaptic Contact Localization

1. Examine slides with biocytin-labeled neurons under a light microscope equipped with a 60X/100X oil immersion objective and a condenser with a high numerical aperture for optimal spatial resolution. Ensure that axonal boutons and dendritic spines are clearly visible.
2. Trace neurons or synaptically coupled neuron pairs using a commercial neuron tracing system (see Table of Specific Materials/Equipment) to obtain 3D neuronal reconstructions. Carry out tracing manually under constant visual inspection to detect even small axonal or dendritic collaterals. For paired recordings from synaptically coupled neurons manual tracing is still the method of choice because the attribution of e.g., an axonal profile to either the pre- or postsynaptic neuron requires substantial reconstruction experience.
3. If required, make counts of axonal boutons and/or dendritic spines. Because spines or even spine subtypes and axonal boutons may exhibit a specific distribution and density with respect to e.g., cortical layer or area, this could help to characterize neuronal cell types.
4. For synaptically coupled neuron pairs, check the presynaptic axon and postsynaptic dendrites for close appositions at the highest magnification available. Observe an axonal bouton and a postsynaptic dendritic spine or shaft is in the same focal plane can be considered as a putative synaptic contact. Mark this contact in the reconstruction.
5. Correct the neuronal reconstructions for shrinkage in all three spatial dimensions in the appropriate section of the tracing program.
   NOTE: During fixation, histochemical processing and dehydration substantial mechanical deformation and shrinkage occur; this will affect axonal and dendritic length measurements and severely distort their spatial arrangement. Shrinkage correction factors for the embedding medium are ~1.1 for x and y directions and ~2.1 for the z-direction³⁷.
6. For a quantitative morphological analysis use a data analysis program for neurophysiology (see Table of Specific Materials/Equipment).