Whole Cell Recording Analysis Essay

Citation: Schramm AE, Marinazzo D, Gener T, Graham LJ (2014) The Touch and Zap Method for In Vivo Whole-Cell Patch Recording of Intrinsic and Visual Responses of Cortical Neurons and Glial Cells. PLoS ONE 9(5): e97310. https://doi.org/10.1371/journal.pone.0097310

Editor: Liset Menendez de la Prida, Consejo Superior de Investigaciones Cientificas - Instituto Cajal, Spain

Received: November 8, 2013; Accepted: April 18, 2014; Published: May 29, 2014

Copyright: © 2014 Schramm et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by an Agence Nationale de Recherche grant (FUNVISYNIN) and an HFSP grant (RGP0049/2002) to LJG. AES was also supported in part by a Marie Curie grant to Sophie Denève and Boris Gutkin (Ecole Normale Supérieure Paris). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors also confirm that Dr. Marinazzo is a PLOS ONE Editorial Board member. This does not alter the authors' adherence to PLOS ONE editorial policies and criteria.

Introduction

The patch-clamp electrophysiological technique was developed to record currents from single membrane channels [1] and later applied to record macroscopic currents and voltages in the so-called whole-cell configuration [2]. Advances have allowed the study of functional synaptic dynamics, gene transcription at the cellular level, and measurements coupled with histology [2]–[7], for a review see [8]. Patch clamp recordings in intact tissue was made possible by Blanton and colleagues [4], who introduced the blind patch technique in vitro; this work inspired the first in vivo patch clamp recordings of functional responses in the intact animal [9], [10]. The introduction of in vivo two-photon microscopy has allowed visual monitoring of whole-cell recordings in vivo[11]–[15], but since this approach is limited to upper layers of the cortex, the blind patch method remains an important technique in vivo. The refinement of this technique has recently reached a new level, with the automated in vivo protocol described by Kodandaramaiah et. al. [16]. For reviews on the methodological aspect of the technique see [17]–[19].

Whole-cell patch recording is difficult to master, requiring a sequence of delicate maneuvers by the experimentalist. The in vivo setting introduces additional constraints; most importantly, maintaining the healthy physiological state of the animal can severely limit the “windows of opportunity” for recordings, and reducing brain movements is always a problem. In light of increasingly sophisticated in vivo protocols, such as simultaneous imaging with two-photon microscopy and the awake behaving preparation, these factors motivate simplifying the technical aspects of whole-cell patch protocols (e.g. obtaining the rapid access to the cell's interior). The limitation of positive pressure is further motivated when the pipette solution contains a dye, e.g., fluorescent calcium indicator [20], [21]. In this case, dye ejected from the pipette during the approach to the neuron increases the extracellular background fluorescence, reducing the contrast and limiting the number of attempts at a given cortical location [15], [22].

A constant challenge is to improve the fundamental step of obtaining electrical access to the interior of the cell, in particular to improve recording stability and to achieve low access, or “series”, resistance (Ra, the resistance between the amplifier input and the cell interior), a crucial parameter for protocols that perturb membrane voltage with current supplied by the amplifier. Another concern is how the recording method modifies tissue or cell physiology. Previous methods to improve whole-cell patch recordings, for example the “tightness” of the seal, include cleaning the cell with either enzymes [2], or by applying positive pressure from the recording or an adjacent pipette [2], [4], [6], [17], [23], [24]. A similar “washing” is also performed by outflow of the pipette solution due to positive pressure while positioning the pipette on the cell membrane during in vitro or in vivo recordings under visual control (for example the “shadow” patching technique [14], [15]). In general, the standard protocol is to apply some type of “wash” step, obtain a gigaohm-seal by suction, and then achieve whole-cell access by applying a ramp or short pulses of suction to the pipette to stress the membrane patch underneath the pipette tip until it breaks. These hydraulic and mechanical operations may be detrimental: Outflow of intracellular solution with a high potassium concentration may initiate or intensify processes that change the dynamical state of the neuronal circuit, such as spreading depression [25], [26], or modify blood vessel contractility [27]. Histological examination of cortical tissue after in vivo patch recordings often shows significant physical damage due to the patch pipette, which will be exacerbated by solution outflow. Subjecting the membrane to directed flow from the pipette may also alter membrane protein function, if only by physical disruption. Finally, the essentially mechanical step of rupturing the membrane to obtain whole-cell mode by suction is difficult, if not impossible, to control at the microscopic level, compromising reproducibility and risking harm to the recorded cell.

To address these issues for whole-cell patch recordings, thus to simplify the technique, improve recording quality, and be less invasive to the recorded cell and its local network, we have developed a revised protocol, “Touch and Zap”. As presented here this method is a direct modification of the standard blind whole-cell patch method for in vivo cortical recordings, and is applicable to either blind or visually-guided patch clamp protocols in brain tissue, in vitro or in vivo.

Materials and Methods

Ethics statement

Protocols were approved by the “Direction Départementale des Services Vétérinaires de Paris”. All painful manipulations (incisions, pressure points) were preceded by injections of local anesthetic (lidocaine).

Animal preparation

Adult Sprague Dawley Rats (male, 300–500 g) were anesthetized with urethane (1.5 g/kg i.p.). Fully anesthetized (no reaction to paw pinching; corneal reflex absent) rats were placed in a stereotaxic rig (Narishige SN-3N). Dexamethasone, to prevent cerebral edema (1 mg/kg i.m.), and either glycopyrrolate (∼0.03 mg/kg i.m.) or atropine methyl nitrate (0.3 mg/kg, i.m.), to reduce secretions and to prevent bradycardia, were injected at the beginning of the experiment. Rectal temperature was monitored and maintained at 37.5±0.5°C by a controlled heating blanket (CWE TC 1000). The electrocardiogram (ECG) was monitored to indicate the depth of anesthesia and the overall health state of the animal, and supplemental urethane was injected i.p as necessary.

For the cat protocols, anesthesia was induced in young adult males (3.2–4.2 kg) with a ketamine-xylasine mixture (10 and 1 mg/Kg respectively, i.m.). After tracheal intubation and installation of a urinary catheter and a rectal temperature probe the animal was placed in the stereotaxic rig (Narishige SN-3N). Rectal temperature was monitored and maintained at 37.5±0.5°C by a controlled heating blanket (CWE TC 1000). Throughout the experiment the animal was perfused by a propofol-sufentanil solution in Ringer 5% glucose (respectively, 5 mg/kg/h for anesthesia and 4 µg/kg/h for analgesia). Perfusion was adjusted as necessary to maintain anesthesia depth as monitored by the ECG and the pCO2 of expired air. To eliminate eye movements, paralysis was induced by pancuronium (0.3 mg/kg i.v.) followed by a continuous venous perfusion of a Ringer 5% glucose-pancuronium solution (0.3 mg/kg/h), and the animal was ventilated by a respiratory pump. A bilateral pneumothorax was performed to reduce brain movements.

After installation in the stereotaxic rig, craniotomies were performed, either above primary somatosensory cortex (for the rat) or above primary visual cortex (for both animals). The typical duration of the experiments was 4–6 hours for rats and from one to three days for cats. At the end of an experiment euthanasia was accomplished with an overdose injection of sodium pentobarbital (i.p. or i.v.).

Electrophysiology

Patch electrodes were pulled from thin wall (OD 1.5 mm, ID 1.10 mm) borosilicate glass capillaries with filament (Sutter Instruments), using a horizontal puller (Sutter Instruments P-97) in 3 steps in order to have a small tip (∼2 µm) and a long and thin taper to minimize damage to the cortex. Pipette shape was constrained to have a resistance between 4 and 8 MΩ, when filled with intracellular solution containing (in mM unless indicated): potassium gluconate 140, potassium chloride 4, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (Hepes) 10, magnesium chloride 2, adenosine-5′-triphosphate dipotassium (ATP) salt 4, guanosine-5′-triphosphate (GTP) lithium salt 0.4, ethylene glycol tetraacetic acid (EGTA) 0.5 and in some experiments, 0.01% dimethyl sulfoxide (DMSO) and 1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) tetrapotassium salt 10 or 5. The osmolarity of the intracellular solution was balanced to 285–295 mOsm with distilled water and pH adjusted to 7.4 with concentrated potassium hydroxide.

The recording system was specifically designed to make dynamic-clamp and visual protocols, and is fully described elsewhere [28]. Briefly, current clamp recordings under bridge mode were made with a Dagan Instruments BVC-700 intracellular amplifier. The membrane voltage and current output were low-pass filtered at 10 kHz and acquired at 40 kHz. Experiment control, data acquisition and data analysis were accomplished with in-house software using the LabView programming environment (National Instruments). Visual stimuli were generated using the VisionEgg software library [29].

Electrode cortical insertion

Whole-cell patch electrodes were introduced into the cortex using a motorized micromanipulator (Narishige SM-21), with the electrode impedance monitored continuously with a 50% duty cycle, 10 Hz current pulses alternating between 0 and −1.11 nA (this value was chosen to allow rapid decade reduction of current amplitude). The manipulator was typically adjusted so that the pipette entered the cortex at an angle between 20 and 30 degrees off the perpendicular. For the initial descent, contact with the cortical surface and penetration, the electrode was advanced in continuous mode (∼750 µm/second), with a strong positive pressure of 100–300 mmHg applied to the interior of the pipette. After penetration of the cortical surface was detected by a transient deflection of the measured voltage, the electrode advance was continued until a predetermined depth (100–2000 microns) was reached. The electrode was then immediately retracted 100–200 microns in continuous mode, and the positive pressure quickly reduced to 40–60 mmHg (the step pressure Pstep) for advancing stepwise into the tissue. The electrode resistance was then determined by visual inspection and fully compensated by the bridge circuit of the amplifier, and voltage offset adjusted according to an estimated tip offset potential (−14 mV with the K-gluconate based solution used here).

Whole-cell access

Subsequent manipulations to achieve whole-cell access mode (Figure 1) followed either the classical blind technique described in the literature (first described by Blanton and colleagues [4]), which we refer to “Wash and Suction” (WS), or with the Touch and Zap protocol. In either case the electrode was advanced in 3–4 micron steps through the cortex until a small deflection of the voltage response was observed (5–10% of the unbalanced response to the −1.11 nA pulses thus, given our electrodes, on the order of several hundred microvolts corresponding to a resistance increase of a few hundred kΩ), indicating contact with the cell membrane. If no change of the electrode resistance was detected after stepping several hundred microns, Pstep was reduced by 5–10 mmHg, and the descent continued. This adjustment was repeated as necessary, unless the attempt was abandoned and the electrode replaced. Thus, the range of Pstep values reported here represent an upper bound for a minimum value of this parameter, therefore even lower values, preferable to reduce the tissue perturbation, may be applicable for the Touch and Zap method (for example 18–26 mmHg used by Margrie and colleagues [17] for blind patching, or 5–15 mmHg for two-photon visually-guided patching in vivo (A. Schramm and P. Kara, and L.J. Graham, unpublished data).

Figure 1. Summary flowchart for the whole-cell access methods described in the text.

The orange diamonds highlight an explicit decision point by the experimenter (e.g., the first orange diamond depends on the choice to employ the WS method or a Touch and Zap variant), whereas the gray diamonds depend on the observed recording properties.

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A key subtlety at this step was to distinguish between an increase in electrode resistance due to contact with a cell, and that due to other causes. In particular, it was critical that the electrode tip be clear before contact with the target cell, assuming that the surface of the electrode tip must be uncontaminated to allow the molecular-level interaction between glass and lipid membrane that underlies the gigaohm-seal. For example, and in contrast with other reports (for example see [13], [17], [30]), in our experience a cardiac or respiratory artifact, possibly due to a blocked electrode or excessive tissue movement, either during the descent or in on-cell configuration (with gigaohm-seal or not), was associated with a poor intracellular recording. In general if we observed fluctuations of the electrode resistance on the order of 20%, the recording attempt was abandoned and a new electrode was used.

Once the electrode reached the target cell, the next step differentiated between the WS and Touch and Zap methods (Figure 1). For the standard WS method the positive pipette pressure was maintained on the order of ten seconds in order to “wash” the membrane. In our experience, a stable electrode resistance during this period (after the increase on contact), that can be easily modulated with small pressure changes (±10–20 mmHg), is correlated with successful seal formation and whole-cell access. After the “wash” the pressure was removed, and a giga-seal formed typically with the aid of light suction applied to the pipette. During seal formation the amplitude of the current steps was progressively reduced from −1.11 nA to −110 pA to −10 pA to avoid large voltage changes (e.g. >150 mV). When a stable gigaohm-seal was achieved, punctuate suction pulses were applied by mouth to achieve whole-cell access. If strong suction was required in order to obtain a gigaohm-seal, subsequent whole-cell access was more difficult and usually had a large access resistance.

The Touch and Zap method specifically avoids the “wash” step of the standard WS method. Thus, the pipette pressure was removed immediately on cell contact – the “Touch” – and under good conditions a seal began to form spontaneously. Also in distinction to the WS method, no (or very little) mouth suction was applied to the pipette a priori at this point. In fact, given the normal intracranial pressure of between 5 and 10 mmHg [31], [32], versus the pressure of the pipette interior, the released of the applied pipette pressure likely results in a small but significant negative pressure gradient across the pipette tip, thus an “automatic” suction.

In contrast to the WS approach, during seal formation the hyperpolarizing current pulses (initially used to monitor the electrode resistance) were maintained at −1.11 nA, which had two effects. First, because seal formation is facilitated by hyperpolarized membrane potentials [17], [33] a positive feedback was established, since voltage deflections became increasingly hyperpolarizing as the seal resistance increased. Second, given the magnitude of the resistance increase, the voltage responses to −1.11 nA could reach the breakdown voltage for the cell membrane within a few seconds, and whole-cell access was achieved by automatic electroporation – the “zap”. In about 25% of the recordings the access resistance seen by the electrode after the zap was close to the final value; in the remainder a smaller second zap followed within a few seconds (typically between at a potential between 100–150 mV less hyperpolarized than the first zap) which reduced the resistance further, again close to the final configuration. When whole-cell access was stable, we defined this endpoint as “pure Touch and Zap” (TZ, see Figure 1 and Results for examples). In some cells the seal quickly reformed after the first or second zap to a giga-seal configuration, and in that case the seal was re-broken with mouth suction. If stable access was achieved, we refer to this endpoint as “Touch, Zap and Suction” (TZS, see Figure 1 and Results; although not used here, we may note that in principle, re-establishing whole-cell access could be attempted by another zap). Finally, if a seal failed to form spontaneously within few seconds after the initial release of pressure (i.e. no zap) we then applied a constant hyper-polarizing current and/or pressure manipulations, as in the standard method. Then, if these procedures resulted in the establishment of a true giga-seal, whole-cell access was achieved by suction, similar to the WS method. If a stable recording was reached we referred to the endpoint as “Touch and Suction” (TS, see Figure 1 and Results). Interestingly, we found that if suction was needed for seal formation at any point, it was rare to see a successful zap.

The achieved endpoint out of the three possibilities following the touch step was in part automatic, and in part dependent on the experimentalist. TZ access was the most straightforward, being almost completely automatic, since no intervention has to be done by the experimentalist between the pressure released right after the touch and the whole cell access. Likewise, the TZS access was essentially imposed by the cell in question – either there was a resealing after the initial zap, or not. Using the TS approach for a given recording was more subjective, specifically deciding when to abandon spontaneous seal formation and resort to more classical manipulations. For our work, we typically chose TS access when there was only a slow increase in resistance, for example no electroporation many seconds after releasing the pipette pressure. Because this decision to follow the TS protocol was essentially arbitrary, the number of TZ and TZS recordings reported here is a lower bound on the number of cells which would have been electroporated if the zap had been tried for longer.

Electrode compensation

Once stable whole-cell access was achieved, the amplifier capacitance compensation was adjusted to accelerate the fast electrode component as much as possible without oscillations during voltage responses to −100 to +300 pA subthreshold 10 Hz current steps. On-line Ra compensation using the amplifier bridge circuit was then adjusted using the same stimulus by the standard method of visually estimating the inflection point of the response that distinguishes the fast electrode artifact component from the (normally slower) passive membrane charging curve.

At this stage, application of a constant small positive pressure (10–20 mmHg) to the pipette often reduced Ra, although in our experience this tended to shorten the recording duration. Ra was typically estimated and compensated several times over the first minutes of the recording, with subsequent estimates made at intervals of several minutes.

Electrophysiological and recording parameters, and data analysis

For off-line analysis we obtained the linear cell parameters with a heuristic inspired by the standard on-line visual method for bridge correction mentioned above, including resting potential Vrest, the cell membrane time constant τ0, the uncompensated Ra and the cell input resistance Rin. An often unappreciated aspect of the visual approach is that the fast time constant of the electrode (and bridge compensation circuit) is implicitly ignored, in contrast with direct methods that model the electrode, such as fitting a two exponential expression to the entire current step response [34], [35]. The intuition is not altogether misplaced: Not only does the short duration of the artifact compromise fit confidence, in any event it is the electrode resistance, not time constant, which is essential for balancing the bridge.

We considered 4 to 10 current step responses (0.5 second hyperpolarizing or depolarizing current) for each cell, where the steady-state response was limited to about ±10 mV from rest, without strong synaptic activity nor action potentials or other obvious non-linearities, for example sags. We assumed that the fastest time constant of the response was due to the electrode, and that the upper bounds of this and the cell time constant were a few hundreds of microseconds and tens of milliseconds, respectively. Assuming that the electrode time constant was one or two orders of magnitude smaller than the cell allowed the cell response fit to discount the electrode artifact by simply ignoring the first part of the response. Assuming an upper bound on the cell time constant allowed an independent estimation of the steady-state voltage, and permitted the estimation of cell time constant to focus on the actual transient. Finally, we assumed the step response could be reasonably approximated by a single exponential. This last assumption was made for simplicity, and improving the method with higher order fits will be considered in future work.

Vrest was first estimated by averaging periods of the membrane voltage prior to the application of stimulus current, rejecting periods that showed spontaneous spiking activity. The steady-state response of each trace, Vss, was then estimated as the average voltage over 100 milliseconds, starting 100 milliseconds after stimulus onset. The voltage just prior to the current step, VO, was estimated by a third order polynomial fit (to account for rapid membrane fluctuations) to the voltage over the two milliseconds just before stimulus onset. The trace voltage V was shifted to obtain V′ = (Vss−V) for depolarizing stimuli or V′ = (V−Vss) for hyperpolarizing stimuli. A first estimate of τ0 was then obtained by linear regression of the logarithm of V′ from 0.5 milliseconds after stimulus onset (thus after much of the electrode artifact had decayed), until V′ reached 0.3(Vss−Vrest) for depolarizing stimuli or 0.3(Vrest−Vss) for hyperpolarizing stimuli. A second iteration was made to refine the estimate by better isolation of the cell component, with the start of the V′ fit now set to stimulus onset plus 20% of the first estimate of τ0. The response fit was then extrapolated to stimulus onset and compared to VO. This voltage difference, divided by the stimulus amplitude, gave the on-line Ra compensation error (about 4 MΩ on average for all cells in this study), and the trace voltage was adjusted as necessary. Rin was obtained for each trace by the difference between Vrest and Vss adjusted by the Ra error, divided by the amplitude of the current step. The linear parameters of the traces were then averaged to obtain the values for the cell. Mean Ra for a recording, and the Ra slope, ΔRa/Δt, corresponding to the evolution of Ra during the recording, and thus stability, were obtained by a linear regression of Ra estimates at various time points of the recording.

For all cells an initial IV protocol was made to establish basic intrinsic biophysical properties, including input impedance and membrane time constant as described above, as well as non-linear responses to establish cell type. The latter classification was based entirely on voltage responses to 500 millisecond current steps; cells with classical current-evoked action potentials of at least 50 mV in amplitude (measured from the resting potential) were classified as neurons, specifically into four physiological types as described by Nowak et al (2003) [42], thus, regular-spiking, fast-spiking, intrinsic bursting and chattering cells. Cells with no obvious fast voltage-dependent properties (time constant <5 ms) were classified as glial cells. The remaining recordings were not considered further in this study. This physiologically-based criteria, thus spiking or not, is standard for distinguishing glial cells and neurons, but since the detailed in vivo electrophysiology of glial cells is less well known we consider this classification as putative. A subset of the neurons and glial cells reported here were tested for visual responses, typically with stimulus sets composed of full field or circumscribed moving sinusoidal gratings (described in [28]).

Recording durations were determined by extrinsic and intrinsic factors. The former was defined by the specific experimental protocol, the shortest being a simple measurement of the voltage response to imposed current steps as described above (minimum recording duration of about one minute), and the longest aimed at measuring and manipulating visual responses. For glial cells, only a subset of cells were tested for visual properties, and of those typically only one to five visual protocols were made, each with a duration of about one minute. Intrinsic factors included the stability of the cellular and electrode properties. First, if there was a large Ra (typically greater than 50 MΩ) following whole-cell access, then the recording was abandoned after a single IV protocol. For stable recordings, a recording was usually terminated if the value of Ra increased more than 50% of its initial value (the most likely cause), or if the resting potential strayed more than ∼20 mV from its initial value, or if there were large (many millivolt) movement artifacts in the voltage trace. If Ra was not verified systematically during the longer visual response protocols, recording duration took into account stable visual responses, including spike heights for neurons within 20 mV of that measured at the beginning of the recording.

The recording depth was taken as the distance of the electrode trajectory between the cortical surface (indicated by a sharp deflection of the electrode voltage during the initial, fast descent) and the recorded cell. This represents an upper bound of the true trajectory length because of dimpling of the cortical surface due to the electrode insertion. The actual cortical depth of the recordings was not measured systematically, but can be estimated taking into account the angle of the electrode relative to the cortical surface, as described above.

For the TZ and TZS protocols, we measured the delay or “zap delay” (Tzap) between the release of the pipette pressure and the abrupt reduction of the resistance seen by the electrode. We also report the peak hyperpolarization or “zap voltage” (Vzap) achieved by the −1.11 nA pulses.

When comparing between whole-cell access methods, unless otherwise indicated, significant differences in the mean among pair-wise data sets were assessed using MANOVA. Statistical significance in the difference between parameter means taken over all neurons and all glial cells was established by a two-tail Student T test, and significant differences between proportions of neurons versus glial cells for the WS and the Touch and Zap methods was evaluated using the two proportion z test. Unless indicated, results are presented as sample average ± the standard deviation, with the sample population in parentheses.

Results

Results were obtained from 372 whole-cell patch-clamp recordings (out of approximately 1800 attempts, with one attempt per electrode). Quantitative analysis was made on a subset of these cells for which at least one current clamp protocol (for example firing rate versus input current curve) have been performed. Two glial cells with very high input resistances (>500 GΩ) were excluded from further analysis. For the remaining cells (N = 201), 33 were recorded with the WS method, and 168 with the Touch and Zap method including 87 recordings with the TZ endpoint, 60 with the TZS endpoint and 21 with the TS endpoint (separated by species and cell type in Table 1). While the WS method has been used extensively by the corresponding author [36], [37], the WS cohort presented here was obtained at an early stage of this study for comparative purposes, and only from rat recordings; the Touch and Zap method was developed over the course of this study on both rat and cat. In general, there were no significant differences in measured properties as function of species; unless mentioned neuron and glial cell recordings are pooled across species.

Relative Frequency of Touch Variants

For the protocols that commenced with the touch step, the first observable concerns the frequency among the possible TZ, TZS and TS endpoints. Interestingly, the ranking of the variants was the same for neurons and glial cells. TZ whole-cell access was most frequent, occurring in 48% of the neuron and 61% of glial cell recordings. The TZS variant was the next most frequent, with 40% of neuron and 24% of glial cell recordings, and TS recordings were obtained in 12% of the neurons and 15% of the glial cells. From this result, it can be seen that fast and spontaneous seal formation, with whole-cell access initiated by electroporation (i.e. TZ and TZS) was observed in most cells subsequent to the touch step, thus 89% of the neuron and 73% of the glial cells. Because the proportion of TS recordings was small and, as explained above, depended on experimenter judgment, unless mentioned the following results consider only the TZ and TZS endpoints of the Touch and Zap method. Since the relative numbers of recordings of neurons versus glial cells with the TZ and TZS endpoints were similar between rat and cat (Table 1), this supported grouping results for the two species unless otherwise mentioned.

The second observation regards how the recording approaches influence the proportion of neuron versus glial cells in blind patch protocols. Our results suggest that protocols starting with the touch step increase the chances of recording glial cells: the proportion of glial cell recordings following the touch step (46 out of 168 recordings, or 27%), was significantly greater than recordings using the WS method (5 out of 28 recordings, or 15%).

Relative Proportions of Neuron Physiological Types and Glial Cells

In Table 2 we report the numbers and relative proportions of physiological neuron types and glia cells recorded over all the Touch and Zap variants (thus, TZ, TZS and TS) in the rat and in the cat. For comparison, this table includes the relative percentages of corresponding physiological neuron types as reported by Nowak et al (2003) [42], from adult cat visual cortex in vivo using sharp microelectrodes. Of note is that the relative populations of cat neurons in this study are consistent with the study by Nowak et al (2003) [42] (no statistical difference using Z test for 2 population proportions). In addition, we did not identify chattering cells in our rat recordings, a negative finding which has been noted by previous workers (ref. Nowak et al, 2003 [42]). Indeed, the fact that the cat and rat protocols reported here were essentially identical and contemporaneous argues against the existence of chattering cells in rodent cortex.

Example Recordings with the Touch and Zap Method

We now present several examples of in vivo recordings with the different methods in the cat, including regular-spiking (Figure 2), bursting (Figure 3) and fast-spiking neurons (Figure 4), and three glial cells (Figures 5–7), illustrating the whole-cell access variants (Figures 2–6), basic measurement of intrinsic properties based on voltage responses to current steps, and visual responses. The examples, which show the TZ, TZS and TS endpoints, were chosen to highlight the diversity of recordings using the Touch and Zap method. To facilitate comparison between the endpoints, the beginning of each recording is plotted on the same scale (panels A and B of Figures 2–6). The voltage and current logs illustrated at the top of these figures were generally started when the stepwise descent into the cortex began, which gives an idea of the fairly short delay, usually less than one minute, between electrode insertion and obtaining a recording. In particular, the characteristic voltage envelopes for the different Touch and Zap endpoints can be appreciated. The envelope for the TZ endpoint (panel B in Figures 2, 3 and 5) is essentially monophasic, quickly growing as the seal forms, and then shrinking in one or two abrupt steps as electroporation achieves whole-cell access. The voltage envelope for the TZS endpoint (panel B in Figure 6) adds a second phase, with a slower increase as the cell re-seals on its own, followed again by an abrupt reduction as whole-cell access is re-achieved by mouth suction. Finally, the voltage envelope for the TS endpoint (panel B in Figure 4) develops much more slowly than the other two, finished by an abrupt reduction as whole-cell access is achieved by suction.

Figure 2. Regular-spiking neuron in cat visual cortex in vivo recorded with the TZ method.

(A) Voltage and current recordings under current clamp mode during the approach to the neuron, detection of contact (“touch”; at ∼6 seconds), release of pressure with immediate seal formation, electroporation of the membrane and establishment of whole-cell access (“zap”), and initial measures of the voltage response to current steps. (B–C), expanded traces from (A) highlighting the access to the whole-cell configuration. (B) corresponds to the region marked by the orange square in (A), and (C) corresponds to the region marked by the grey square in (B). The “touch” is characterized by a small increase in the electrode resistance (C). The positive pressure in the pipette is released within a second after the touch, followed by an immediate increase in the electrode resistance as the seal spontaneously forms. A first “zap” occurs within about one second (B), followed by a second “zap” which results in the whole-cell configuration, evidenced by the much smaller voltage deflections, and the hyperpolarization of the voltage envelope as the cell's resting potential establishes a DC bias. At this point the current pulses are typically set to 100 pA, switching polarity as necessary to estimate and compensate for Ra. (D) Voltage responses to current steps (−100, 0, 150 and 250 pA), with the response to 250 pA showing a regular-spiking firing pattern. (E) Left – Examples of preferred (180°) and non-preferred (90°) responses (10 trials overlaid) to a moving sinusoidal grating (4 Hz, 100% contrast, 0.5 second duration). Right - Total spike count over the 10 trials as a function of stimulus direction, showing that this neuron is strongly tuned to stimulus orientation and modestly tuned to stimulus direction. The on-line corrected Ra for this recording was 24 MΩ, which was subsequently corrected off-line to 21 MΩ. The input resistance was 102 MΩ, giving a relative Ra of 0.21.

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Figure 3. Bursting neuron in cat visual cortex in vivo recorded with the TZ method.

(A–B). As in Figure 2, voltage and current recordings during the approach to the neuron, the “touch” (at ∼16 seconds), release of pressure with immediate seal formation, the first “zap” about two seconds later, followed by a second “zap” and establishment of whole-cell access, and initial measures of the voltage response to current steps. (C) Voltage responses to current steps (−50, 150 and 250 pA), showing a bursting firing pattern in response to the 250 pA step. (D) Left – Examples of preferred (0°) and non-preferred (90°) responses (10 trials overlaid) to a moving sinusoidal grating (4 Hz, 100% contrast, 1 second duration). Right – Total spike count over the 10 trials as a function of stimulus direction, showing the strong orientation tuning of this neuron. The on-line corrected Ra was 44 MΩ, which was subsequently corrected off-line to 61 MΩ. The input resistance was 70 MΩ, giving a relative Ra of 0.87.

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Figure 4. Fast-spiking neuron in cat visual cortex in vivo recorded with the TS method.

(A–B). As in Figure 2, voltage and current recordings during the approach to the neuron, with the “touch” at ∼36 seconds. As can be appreciated in (B), and in distinction to the TZ endpoints, here there is only a slow increase in electrode resistance after the release of the pipette pressure. At approximately 51 seconds, the “zap” attempt is abandoned, and the standard methods of suction and hyperpolarization for obtaining a gigaseal were used, with the whole-cell access achieved by mouth suction at ∼70 seconds. (C) Voltage responses to current steps (0 and 800 pA), showing a fast-spiking firing pattern to the suprathreshold stimulus. (D) Left – Examples of preferred (230°) and non-preferred (50°) responses (10 trials overlaid) to a moving sinusoidal grating (4 Hz, 100% contrast, 1 second duration). Right – Total spike count over the 10 trials as a function of stimulus direction, showing that this neuron is mainly tuned to stimulus direction. The on-line corrected Ra was 37 MΩ, which was validated off-line. The input resistance was 26 MΩ, giving a relative Ra of 1.4.

https://doi.org/10.1371/journal.pone.0097310.g004

Figure 5. Glial cell in cat visual cortex in vivo recorded with the TZ method.

(A–B). As in Figure 2, voltage and current recordings during the approach to the cell, the “touch” (at ∼33 seconds), release of pressure with immediate seal formation, the first “zap” about two seconds later. The current was reduced to 100 pA about one second later, and final whole-cell access was achieved by a small suction. As for neurons (e.g. Figures 2 and 3), the “zap” is distinguished by the sudden reduction in resistance while the strong hyperpolarizing current pulses are applied. (C) Voltage responses to current steps (100 nA increments from −100 to 700 nA), showing the lack of spikes and a fast membrane time constant that are characteristic of glial cells. (D) Left - Preferred (45°) and non-preferred (135°) current clamp responses (average of 10 trials) to a moving sinusoidal grating (4 Hz, 100% contrast, 2 second duration). Right – Average standard deviation of the visual responses (over the 10 trials) as a function of stimulus direction, showing that the membrane voltage fluctuations of this cell are tuned to stimulus direction. The Ra estimated on-line was 45 MΩ (validated off-line), and the cell input resistance was 105 MΩ, thus a relative Ra of 0.43.

https://doi.org/10.1371/journal.pone.0097310.g005

Figure 6. Glial cell in cat visual cortex in vivo recorded with the TZS method.

(A–B) As in Figure 2, voltage and current recordings during the approach to the cell and subsequent whole-cell access. Here, an initial spontaneous seal with a subsequent “zap” was accomplished, but in distinction to the TZ endpoints illustrated in Figures 2, 3 and 5, the membrane resealed (between 24 and 27 seconds), and whole-cell access was re-established by suction at ∼46 seconds (not visible in (B)). (C) Voltage responses to current steps (100 nA increments from −200 to 700 nA). (D) Left – Preferred (270°) and non-preferred (0°) voltage responses (average of 10 trials) to a moving sinusoidal grating (4 Hz, 100% contrast, 5 second duration). Right – Average standard deviation of the visual responses (over the 10 trials) as a function of stimulus direction, showing that this cell is tuned to stimulus direction, with a small bias for stimulus orientation. The on-line corrected Ra was 40 MΩ, which was validated off-line. The input resistance was 99 MΩ, giving a relative Ra of 0.40.

https://doi.org/10.1371/journal.pone.0097310.g006

Figure 7. Gial cell in cat visual cortex in vivo recorded with the WS method.

(A) Voltage responses to current steps (100 nA increments from −100 to 700 nA). (B) Left – Preferred (135°) and non-preferred (45°) voltage responses (average of 10 trials) to a moving sinusoidal grating (4 Hz, 100% contrast, 5 second duration). Right – Average standard deviation of the visual responses (over the 10 trials) as a function of stimulus direction, showing that this cell is tuned to stimulus orientation, with a small bias for stimulus direction. The on-line corrected Ra was 48 MΩ, which was validated off-line. The input resistance was 100 MΩ, giving a relative Ra of 0.48.

https://doi.org/10.1371/journal.pone.0097310.g007

To illustrate the high quality recordings obtained with the Touch and Zap method each example also shows the intrinsic response to injected current steps (panel D in Figure 2 and panel C in Figure 3–6) and functional visual responses (panel E in Figure 2 and panel D in Figure 3–6), specifically the canonical selectivity of cells in primary visual cortex to the orientation or direction of a visual feature. We emphasize that the Touch and Zap method is not necessary to obtain whole-cell recordings from glial cells in vivo; Figure 7 shows the intrinsic (panel A) and visual responses (panel B) from a glial cell recorded with the WS method (but see the specific advantages of Touch and Zap below). Overall of particular note is the relatively fast dynamics of the glial visual responses in these examples, showing the ability of these cells to follow stimuli up to at least 4 Hz.

Recording duration

For the cells analyzed here, the mean recording duration was 12±17 minutes (N = 150) for all neurons, and 2.7±2.9 minutes (N = 51) for all glial cells, with a maximum duration of approximately 80 minutes and 15 minutes for neurons and glia, respectively (Figure 8).

Recording and cell parameters

To further characterize the Touch and Zap method we now compare recording protocol parameters across methods, such as the electrode resistance, the pressure applied to the pipette during cell approach, the recording depth, the delay and minimum voltage of the zap step (Table 3). Then, to assess the impact of the method on cell physiology, we next compare cellular properties obtained using the Touch and Zap (pooled TZ and TZS populations) and WS methods, including the cell input resistance and membrane time constant, an estimate of the cell size based on input resistance, the resting potential, characteristics of action potentials for neurons, the absolute and relative access resistance, and the stability of the access resistance (Table 4).

Electrode resistance.

On average, the electrode resistance, Re, was significantly larger for glial cells than for neuron recordings, but in practical terms the difference of 0.4 MΩ was small (Table 3). Across whole-cell access methods, comparing the three pooled Touch and Zap endpoints with the WS approach, there was no significant difference in Re for either glial cells or neurons.

Step pressure and recording depth.

For the ensemble of Touch and Zap recordings, there was no significant difference in the average Pstep between glial cells and neurons. (Table 3). The broad distribution of Pstep for these recordings implies that successful whole-cell access did not depend critically on this parameter. Pstep was not systematically recorded for the WS protocols, which precluded quantitative comparison with the Touch and Zap method, but the same range (40–60 mmHg) was used.

Zap delay and voltage.

The average zap delay of the pooled TZ and TZS recordings, Tzap, was significantly longer for glial cells than neurons (Table 3). This parameter was susceptible to the experimenter judgment: while standard practice was to abandon the zap attempt after a few seconds, occasionally the large current pulse was maintained for much longer. The distributions of Tzap for both cell types were therefore skewed, and thus the medians (3.2 and 2.1 seconds for glial cells and neurons, respectively) are more indicative of what to expect in practice. These values reflect the fast seal formation that is characteristic of this method, as compared to the standard WS protocol where the seal is typically formed over tens of seconds.

The average zap voltage for the pooled TZ and TZS recordings, Vzap, was significantly lower for glial cells compared to neurons, with the overall values for both cell types consistent with reported values of the lipid bi-layer membrane breakdown voltage [39], [40] (Table 3).

Rin and τ0.

There was no significant difference in either Rin or τ0 for neurons or glial cells as a function of access method (pooled TZ and TZS endpoints, compared with WS; Table 4

Abstract

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Keywords: Neuroscience, Issue 91, electrophysiology, acute slice, whole-cell patch-clamp recording, neuronal morphology, immunocytochemistry, parvalbumin, hippocampus, inhibition, GABAergic interneurons, synaptic transmission, IPSC, GABA-B receptor

Introduction

Hippocampal neuronal circuits have long been the subject of intense scrutiny, with respect to both anatomy and physiology, due to their essential role in learning and memory as well as spatial navigation in both humans and rodents. Equally, the prominent, but simple laminar organization of the hippocampus makes this region a favored subject of studies addressing structural and functional properties of cortical networks.

Hippocampal circuits are comprised of excitatory principal cells (>80%) and a smaller (10-20%), but highly diverse cohort of inhibitory interneurons1-3. Interneurons release γ-aminobutyric acid (GABA) from their axon terminals which acts at fast ionotropic GABAA receptors (GABAARs) and slow metabotropic GABAB receptors (GABABRs)4. These inhibitory mechanisms counterbalance excitation and regulate the excitability of principal cells, and thus their timing and pattern of discharge. However, GABA released from interneurons acts not only on principal cells, but also on the interneurons themselves5,6. Pre and postsynaptic receptors mediate feedback regulation and inhibitory mutual interactions among the various types of interneuron. These inhibitory mechanisms in interneuron networks are believed to be central to the generation and shaping of population activity patterns, in particular oscillations at different frequencies7.

Whole-cell patch-clamp recording is a well-established method for the examination of intrinsic properties and synaptic interactions of neurons. However, due to the high diversity of interneuron types, investigation of inhibitory interneurons requires rigorous identification of the recorded cells. As hippocampal interneuron types are characterized by distinct morphological features and neurochemical marker expression, combined anatomical and immunocytochemical examination can provide a means to determine precise interneuron identity6,8,9.

In the present paper we describe an experimental approach in which whole-cell patch-clamp recordings from single neurons or synaptically-coupled pairs are combined with intracellular labeling, followed by post-hoc morphological and immunocytochemical analysis, allowing for the characterization of slow GABAB receptor mediated inhibitory effects in identified interneurons. As an example, we focus on one major type of interneuron, a subset of the so called “basket cells” (BC), which innervates the soma and proximal dendrites of its postsynaptic targets and is characterized by a “fast spiking” (FS) discharge pattern, an axon densely covering the cell body layer, and expression of the calcium-binding protein parvalbumin (PV)10,11. These interneurons display large postsynaptic inhibitory currents, as well as prominent presynaptic modulation of their synaptic output, in response to GABABR activation12. The combination of techniques described here can be applied equally well to investigate intrinsic or synaptic mechanisms in a variety of other identified neuron types.

Protocol

Ethics Statement: All procedures and animal maintenance were performed in accordance with Institutional guidelines, the German Animal Welfare Act, the European Council Directive 86/609/EEC regarding the protection of animals, and guidelines from local authorities (Berlin, T-0215/11)

1. Preparation of Acute-hippocampal Slices

  1. Take a transgenic rat (17 to 24 day old), expressing the fluorescent Venus/YFP protein under the vGAT promoter, which labels the majority of cortical inhibitory interneurons13. Decapitate the rat. Rapidly dissect the brain (<40 sec) into semifrozen, carbogenated (95% O2/5% CO2) sucrose-based artificial cerebrospinal fluid (sucrose-ACSF, Figure 1A).

  2. Assess the dissected rat brain for Venus/YFP fluorescence with a 505 nm LED lamp and 515 emission filter, mounted on a pair of goggles.

  3. Remove the frontal third of the cortex and cerebellum; then separate the hemispheres, all with a scalpel. Remove the dorsal surface of the cortex to provide a flat surface to glue the brain down, as previously described14.

  4. Cut transverse slices (300 μm) of the hippocampal formation on a vibratome, the hemispheres should be surrounded with semifrozen, carbogenated sucrose-ACSF (Figure 1B)14. Remove additional regions of rostral cortex, midbrain and brainstem. Transfer each slice to a submerged holding chamber containing sucrose-ACSF, which is carbogenated and warmed to 35 ºC.

  5. Leave the slices to recover at 35 ºC for 30 min from the time of the last slice entering the warmed ACSF. Do this in order to reactivate metabolic processes and facilitate the resealing of cut neuronal processes. Then transfer to room temperature for storage (Figure 1C).

2. Fabrication and Filling of Recording Pipettes

  1. Pull patch pipettes from glass capillaries, so that a pipette resistance of 2-4 MΩ is achieved when filled with filtered (syringe filter, pore size: 0.2 μm) intracellular solution containing 0.1% biocytin (for intracellular labeling). Keep the intracellular solution chilled on ice to prevent degradation of its constituents.

  2. Fill patch pipettes for identification of postsynaptic currents with a solution containing a physiologically relevant low Cl- concentration (ER(Cl-)= -61 mV; see solution list).

  3. For paired recordings to identify the presynaptic receptor mediated responses, fill patch pipettes with intracellular solution with low Ca2+ buffer capacity to prevent interference with transmitter release presynaptically, as well as 4-fold higher Cl- concentration (ER(Cl-)= -20 mV) to improve signal-to-noise of observed IPSCs5 allowing accurate assessment of pharmacological responsiveness. Note that changing Cl- concentration can alter IPSC kinetics15.

3. Whole Cell Patch-clamp Recording from FS-INs

  1. Carbogenate the ACSF and feed through the perfusion system to the recording chamber, by means of a peristaltic pump (which also removes ACSF from the recording chamber through a suction line, Figure 2A). Turn on all equipment on the setup in preparation for recording.

  2. Transfer a slice to the recording chamber and hold in place with a platinum ring strung with single fibers of silk. Position the slice so that the stratum (str.) pyramidale of CA1 runs vertically through the field of view, allowing access with 2 pipettes to both the str. radiatum and str. oriens simultaneously (Figures 2C and 4A).

  3. Place the chamber into the setup and start perfusion of carbogenated and warmed (32-34 ºC) recording ACSF at a flow rate of 5-10 ml/min.

  4. Assess slice quality under IR-DIC optics at 40X objective magnification, and visualize with a CCD camera viewed on a display. Assume good slice quality if a large number of round, moderately contrasted CA1 pyramidal cells (CA1 PC) can be seen in str. pyramidale at depths of 20-30 μm below a smooth and lightly cratered surface (Figure 2C). Poor quality slices contain large numbers of highly contrasted, shrunken or swollen cells, with an uneven slice surface.

  5. Identify putative FS interneurons under epifluorescence illumination as those expressing Venus/YFP (Figure 2B), with large multipolar somata in or near the str. pyramidale. Select cells reasonably deep within the slice (50-100 μm, Figure 2C) in order to better preserve their morphological integrity.

  6. Mount the recording electrode in the pipette holder on the headstage; then apply a low, positive pressure (20-30 mBar) through the tube line. Lower the pipette to the surface of the slice, slightly offset to the center of the selected neuron.

  7. Obtain whole-cell recording configuration as described previously14,16 and see also Figures 2D and 2E:
    1. Target a cell: Increase the pressure to 70-80 mBar and rapidly lower the pipette through the slice to just above the soma of the selected cell (Figure 2D, top).

    2. Approach the cell: Press the pipette against the cell membrane to produce a “dimple” on it (Figure 2D, top). Perform this step swiftly, in order to prevent biocytin labeling of neighboring cells.

    3. Create a giga-ohm seal: Release the pressure and simultaneously apply a 20 mV voltage command to the pipette. A giga-ohm seal (1-50 GΩ; Figure 2D, bottom and Figure 2E middle) typically develops rapidly. Once sealed, apply the expected resting membrane potential (typically between -70 and -60 mV) as a voltage command.

    4. Break through the patch: Once sealed, rupture the membrane patch with a short pulse of negative pressure; thereby achieving the whole-cell configuration (Figure 2E, bottom).

  8. Compensate whole-cell capacitance and series resistance (RS). Rs is normally 5-20 MΩ and stable for up to 120 min. Abandon cells if membrane potential (VM) on break-through is more depolarized than -50 mV; RS is initially greater than 30 MΩ; or RS changes by more than 20% over the course of the recording.

  9. Identify FS-INs by their response (in current-clamp mode) to a family of hyper- to depolarizing current pulses (-250 to +250 pA, Figure 2F, top). FS-INs have relatively depolarized VM (typically -50 to -60 mV), short membrane time-constant (<20 msec) and respond to a 500 pA depolarizing current injection with a train of action potentials (APs) at frequencies >100 Hz 11 (Figure 2F, bottom), which are markedly different from those in CA1 PCs (Figure 2F, middle).

4. Extracellular Electrical Stimulation to Evoke GABABR-mediated Responses

  1. To observe synaptically evoked responses, position an extracellular stimulation electrode (a patch pipette filled with 2 M NaCl; Resistance: 0.1-0.3 MΩ) in the slice at the border of str. radiatum and str. lacunosum-moleculare. Position the electrode 200-300 μm lateral to the soma to prevent direct electrical stimulation of the cell and minimize stimulation artifacts (Figure 3A).

  2. Once the stimulation electrode is positioned, obtain whole-cell recording of the chosen cell and assess the physiological phenotype in current-clamp mode as in section 3.9 (Figure 3B).

  3. With the neuron recorded in voltage-clamp (VM -65 mV), deliver electrical stimulation of presynaptic axons at 50 V (~500 μA effective stimulus) every 20 sec, using an isolated constant-voltage stimulator. Use single stimuli (100 μsec duration, Figure 3C, top) to observe GABABR mediated IPSCs, and interleave with trains of multiple stimuli (at 200 Hz) to produce greater transmitter release.

  4. Bath apply ionotropic glutamate receptor antagonists (AMPA receptor: DNQX [10 μM]; NMDA receptor: d-AP5 [50 μM]) to reveal the isolated monosynaptic IPSC (Figure 3C, middle upper). Further isolate the GABABR-mediated IPSC with application of a GABAAR blocker (gabazine [10 μM]; Figure 3C middle lower).

  5. Confirm the resultant slow-outward current (Figure 3C lower, expanded) as being GABABR-mediated by the subsequent application of CGP-55,845 [5 μM] (Figure 3C lower, underlain in grey)

5. Paired Recordings of Synaptically Coupled FS-IN and CA1 PCs

  1. Assess GABABR-mediated presynaptic control of inhibitory synaptic transmission with simultaneous recordings, performed between synaptically-coupled IN and PC pairs as described below.

  2. First, establish a whole-cell recording of a presynaptic interneuron (as in section 3) and confirm the FS phenotype (Figure 4A).

  3. Then patch a neighboring CA1 PC (20-100 μm distance, Figure 4A) and apply brief suprathreshold depolarizing current pulses (1 msec duration, 1-5 nA amplitude) to the presynaptic IN (held in current-clamp mode) to elicit APs. If a synaptic connection is present, APs in the IN result in IPSCs in the CA1 PC, held in voltage-clamp (compensate RS to about 80%).

  4. If necessary, fill a new recording electrode and record from further CA1 PCs until a connection is found.

  5. Once a connection is established, elicit pairs of APs in the presynaptic FS-IN to assess both the unitary synaptic response and dynamic behavior. Use a typical paired-pulse protocol of 2 depolarizing stimuli with a 50 msec interval (Figure 4B).

  6. Collect control traces in baseline conditions. Then, apply the selective GABABR agonist baclofen (10 μM) to the perfusing ACSF, thus activating GABABRs, followed by the antagonist CGP-55,845 (5 μM), to fully block the receptor mediated effects. Collect ~50 traces during steady state of each drug condition (Figure 4B and C).

  7. Once the recording is complete, seal the somatic membrane by forming an outside-out patch: Slowly withdraw the pipette from the cell body in V-clamp and as the RS increases, reduce the VM to -40 mV. Do this to facilitate the formation of the outside-out patch; then remove the pipette from the bath.

6. Analysis of Electrophysiological Properties

  1. NOTE: A multitude of different software packages are available for the acquisition of electrophysiological data. Here, WinWCP, a Windows program in the free Strathclyde Electrophysiology Software package is used, which allows recording of up to 16 analog input channels and output of 10 digital signals.

  2. Low-pass filter all data at 5-10 kHz and sample at 20 kHz.

  3. Analyze physiological data with an off-line analysis suite. NOTE: Stimfit, an open source software package which includes a Python shell, is used in this instance; however other alternatives can easily be used instead.
    1. Analyze passive membrane properties of recorded neurons, acquired in current clamp, from resting membrane potential.

    2. Measure the mean resting membrane potential from the baseline of recorded responses from the beginning of the recording.

    3. Calculate input resistance, using Ohm’s law, from the voltage response to the smallest hyperpolarizing current pulses (≤ -50 pA). To improve signal to noise ratio, average multiple traces. Note: our examples are typically averages of 10-50 individual sweeps.

  4. Estimate the apparent membrane time constant by fitting a monoexponential curve to the decay of the responses to the smallest hyperpolarizing current pulses.

  5. Analyze action potential waveform to determine threshold, amplitude (threshold to peak) and duration (width measured at half height) elicited by threshold level depolarizing current pulses.

  6. Analyze GABABR mediated IPSCs from voltage clamp recordings. Filter traces off-line at 500 Hz (Gaussian filter) and assess the peak amplitude and latency of the GABABR mediated response (in averages of at least 10 traces).

  7. Detect the effect of GABABRs on the inhibitory output of INs as a change in peak amplitude of the GABAAR-mediated IPSCs measured between peak and preceding baseline. Calculate the mean amplitude from ≥50 traces for the control period and the steady-state of all pharmacological epochs.

7. Visualization and Immunocytochemistry of FS-Ins

  1. Following the recordings, fix the slices by immersion in 4% paraformaldehyde with 0.1 M phosphate buffer (PB, pH=7.35) O/N at 4 °C.

  2. If necessary slices can be transferred to PB and stored for up to ~1 week before processing.

  3. Wash slices liberally in fresh PB and subsequently in 0.025 M PB with 0.9% NaCl (PBS, pH=7.35).

  4. To reduce non-specific antibody binding, block the slices for 1 hr at RT in a solution containing 10% normal goat serum (NGS), 0.3% Triton-X100 (a detergent to permeabilize membranes) and 0.05% NaN3, made up in PBS.

  5. To label for PV expression, use an anti-PV monoclonal mouse antibody diluted in a solution containing 5% NGS, 0.3% Triton-X100, 0.05% NaN3, in PBS. Incubate primary antibodies for 2-3 days at 4 °C12. Rinse slices thoroughly in PBS.

  6. Apply fluorescent anti-mouse secondary antibodies (e.g. Alexafluor-546) along with the biotin binding-protein streptavidin, conjugated to a fluorochrome (e.g., Alexafluor-647); and incubate in a solution containing 3% NGS, 0.1% Triton-X100, 0.05% NaN3, diluted in PBS and incubate O/N at 4 °C.

  7. Liberally rinse slices 2-3x with PBS followed by 2-3 rinses in PB. Mount the slices on glass slides. Use a 300 μm agar spacer to prevent the slice from collapsing. Cover-slip slices with a fluorescent mounting medium and seal with nail-varnish.

8. Imaging and Reconstruction of Visualized FS-Ins

  1. Visualize the slices using a scanning confocal microscope, with the fluorochome reporter excited with the appropriate laser line (diode laser 635 nm for Alexafluor-647; Helium-Neon 543 nm for Alexafluor-546 labeling for PV and Argon 488 or 515 nm for Venus/YFP).

  2. Take images at an appropriate Z-resolution (typically 0.5-1 μm steps, using a 20X objective) to produce a Z-stack of the whole cell. Multiple stacks are normally required to image the whole cell, which can be digitally stitched off-line using FIJI/ImageJ software (Figure 5A).

  3. Reconstruct the cell from the stitched image stack using a semi-automatic tracing method (Simple Neurite Tracer plugin in FIJI/ImageJ software package17, Figure C).

  4. Finally, assess the PV-immunoreactivity of the interneuron with a high numerical aperture objective lens (60X silicon-immersion, N.A.=1.3). Make images of the soma, proximal dendrites and proximal axon, or alternatively of axon terminals if somatic washout of PV is too strong. Cells are deemed immunoreactive for PV if immuno-labelling is seen to align with the biocytin-labeled structures (Figure 5B).

Representative Results

Provided that slice quality is appreciably good, recording from both CA1 PCs and FS-INs can be achieved with minimal difficulty. The transgenic rat line expressing Venus / YFP under the vGAT promoter13 does not unequivocally identify FS-INs, or indeed BCs. However recordings from INs in and around str. pyramidale, where the density of FS-INs is typically high1, results in a high probability of selecting FS-INs (Figure 2B). FS-INs can be distinguished by their characteristic physiological properties different from those of both CA1 PCs and RS-INs. They have a relatively depolarized resting membrane potential (-58.9 ± 1.5 mV, 15 cells, vs. -62.6 ± 1.1 mV in CA1 PCs, 26 cells), low input resistance (92 ± 12 MΩ vs. 103 ± 14 MΩ in PCs) and fast apparent membrane time constant (15.4 ± 2.6 msec vs. 22.0 ± 2.7 msec in PCs), resulting in rapid voltage response to hyperpolarizing current pulses (Figure 2F). FS-INs discharged very brief action potentials (0.38 ± 0.01 msec) of relatively low amplitude (82.8 ± 1.0 mV) followed by very prominent fast afterhyperpolarization (mean amplitude 22.6 ± 2.9 mV). In response to large depolarizing current pulses (250 pA), FS-INs fired at high frequencies (82 ± 10 Hz; Range: 34-128 Hz; Figure 2D, left).

To assess GABABR-mediated postsynaptic currents in FS-INs, pharmacologically isolated synaptic responses were elicited by extracellular stimulation of inhibitory fibers at the str. radiatum/lacunosum-moleculare border. Figure 3 shows an example of a FS-IN, in which sequential blockade of components of the compound synaptic response isolates the monosynaptic GABABR-mediated slow IPSC. Synaptic responses were elicited by single stimuli or trains of 3-5 stimuli (50 V intensity, 0.1 msec duration, delivered at 200 Hz) to the str. lacunosum-moleculare/radiatum border (Figure 3A). The initial compound response including both excitatory and inhibitory components (Figure 3C, top) was strongly reduced by bath application of AMPA and NMDA receptor antagonists (DNQX, 10 μM and AP-5, 50 μM, respectively: Figure 3C, middle). The residual monosynaptic IPSC comprised an early fast inward current (a putative inward Cl- mediated current at the holding potential -65 mV, with a reversal potential of approximately -60 mV, in these experiments) and a slower outward current (mediated by a putative K+ conductance with a reversal potential close to 100 mV). Application of the selective GABAAR antagonist gabazine (SR-95531, 10 μM) abolished the fast IPSC, leaving an isolated the slow GABABR-mediated IPSC (Figure 3C, bottom, black trace); which was observed in response to single stimulation, but more clearly in response to short trains of stimuli. This response was confirmed as a GABABR-activated K+ conductance, as it was blocked by the potent and selective antagonist CGP-55,845 (CGP, 5 μM, Figure 3C, bottom, gray trace). FS BCs typically show large amplitude GABABR-mediated IPSCs, such as shown in our recent publication12.

To assess presynaptic regulation of FS-IN inhibitory output by GABABRs, paired recordings were performed from synaptically-coupled FS-IN and CA1 PCs. Synaptic connectivity between FS BCs and CA1 PCs is relatively high, as shown previously1,3. In these recordings, as shown previously, coupling probability is over 50% between closely located cell pairs (≤50 μm; Figure 4A)5. However, this depends strongly on the interneuron type examined. Connectivity was tested with either a long depolarizing current injection (100 msec, ≥ 500 pA) or a train of short depolarizing pulses (1 msec duration, 1-5 nA, up to 10 pulses delivered at 20 Hz) eliciting a single AP each. APs in the presynaptic interneurons elicited fast GABAAR-mediated IPSCs with short latency, rapid rise and decay in synaptically-coupled PCs. When paired-pulses (2 pulses at 20 Hz) were applied, the synapse showed short-term depression (Paired-pulse ratio < 1)1. In the example cell shown, bath application of the GABABR agonist baclofen (2-10 μM) resulted in a substantial reduction in the amplitude of the first IPSC (Figures 4B and 4C). Subsequent bath application of the antagonist CGP-55,845 invariably resulted in a recovery of the IPSC (5 out of 5 cells12; Figures 4B and 4C).

Once a recording had been completed, an outside-out patch successfully formed, the slices were fixed overnight and subsequently processed to visualize and analyze the morphology of the recorded cells and determine their neurochemical marker content. Figure 5A illustrates the morphology of one representative FS BC in a projection of combined image stacks obtained on a confocal microscope. The cell’s expression of PV was confirmed in immunocytochemical labeling that gave a clear immuno-signal over the cell body and proximal dendrites of the recorded and biocytin-labeled cell (Figure 5B). Three-dimensional reconstruction of the cell was performed from stitched image stacks using the Simple Neurite Tracer plugin in FIJI software (Figure 5C)17. The axon showed a high density of collaterals in and near the str. pyramidale with “baskets” formed around the somata of CA1 PCs (Figure 5A, inset), putatively identifying this cell as a BC. Furthermore, the localization of the soma near to str. pyramidale and the radially-oriented dendrites spanning all layers of the CA1 correspond well to the typical morphological feature of FS BCs10.

In summary, in whole-cell recordings obtained from identified FS PV+ BCs, we have demonstrated that these cells express large amplitude GABABR-mediated slow IPSCs and their synaptic output is also markedly inhibited by the activation of presynaptic GABABRs .

Figure 1. Preparation of acute hippocampal slices.(A) A freshly dissected rat brain. Note the ice surrounding the brain, maintaining the temperature of the whole brain near 0 °C. (B) Cutting of 300 μm hippocampal slices on a Leica VT1200s vibratome. The hemispheres are aligned so that the cortical surface is cut first. Note the large amount of slushy ice surrounding the hemispheres and the constant carbogenation of the icy ACSF (arrow). (C) After cutting the brain slices are moved to warmed sucrose-ACSF. The slices were turned over and trimmed to remove the forebrain and midbrain, leaving only the hippocampus and overlying cortex.

Figure 2. Whole-cell patch-clamp recording from interneurons. (A) The recording configuration around the chamber. Note the inflow and outflow are on opposing sides of the chamber to achieve a close to laminar flow. Also visible are the recording electrodes, mounted on the headstages, and the ground electrodes on the two sides, as well as the objective (40X, water-immersion) in the middle. (B) Low-power epifluorescent image of the vGAT-Venus/YFP signal in the CA1 of the hippocampus in a slice. (C) IR-DIC imaging of the same area. The arrows in B and C indicate a Venus/YFP positive interneuron in the cell body layer. (D) High-power IR-DIC images of the soma of the neuron indicated in panel B, with an approaching patch pipette forming the dimple on the surface (top) and, subsequently, the cell in the whole-cell configuration (bottom). (E) Schematic illustration of the major stages of establishing a whole-cell patch-clamp recording (left side) with cartoon responses to a test voltage-pulse monitoring the resistance at the pipette tip (right side). Top row: Pipette in the bath away from the cell; Upper middle: Dimple formation, accompanied by a reduction in the pulse amplitude, indicating a moderate increase in resistance. Lower middle: Giga-ohm seal formation. Note that the current pulse amplitude is dramatically reduced. Only the fast capacitive currents are visible at the beginning and end of the pulse before pipette capacitance is compensated. Bottom: Whole-cell recording configuration is achieved by breaking through the membrane patch under the pipette tip. Note that the large but relatively slow capacitive currents at the beginning and end of the pulse before series resistance compensation is applied. (F) Representative traces of a FS-IN (top) and CA1 PC (bottom), elicited by a family of hyper- to depolarizing current pulses (protocol shown above). Note the very high frequency discharge of the FS-IN compared to the much slower firing of the CA1 PC. Insets on the right: Comparison of the AP waveform from the CA1 PC (top) and the FS-IN (bottom, in grey, overlaid on the PC AP in black), illustrating the difference in the waveform of the AP and the AHP.

Figure 3. Pharmacological dissection of the compound synaptic response, isolating slow GABABR-mediated IPSCs in a FS-IN. (A) IR-DIC image of the slice in the recording chamber, with the recording electrode (Patch) placed at the border of the str. oriens (Ori.) and pyramidale (Pyr.) and the simulation electrode (Stim) at the border of the str. radiatum (Rad.) and lacunosum-moleculare (L-M). (B) Left: Schematic illustration of the recording configuration. Right: Superimposed voltage responses in the FS-IN elicited by a family of hyper- to depolarizing current injections. (C) Representative traces recorded from the FS-IN in response to extracellular stimulation. Top: The compound synaptic response produced in normal ACSF elicited by a single stimulus (50 V intensity, 0.1 msec duration). Middle upper: Isolated monosynaptic IPSC after bath application of DNQX (10 μM) and d-AP-5 (50 μM). Middle lower: The fast monosynaptic IPSCs is blocked following the addition of gabazine (10 μM) to the bath. Bottom: A train of 5 stimuli (at 200 Hz) reveals a slow GABABR mediated IPSC (black trace) which is blocked by subsequent application of CGP (5 μM) to the bath (superimposed grey trace).

Figure 4. Analysis of presynaptic modulation of inhibitory synaptic transmission in a paired recording from a coupled FS-IN and CA1 PC pair. (A) Left panel: IR-DIC image of the two recording pipettes, the left one patched onto the FS-IN at the border of the str. oriens (Ori.) and pyramidale (Pyr.) and the right patched onto a CA1 PC closer to the str. radiatum (Rad.). Right panel: A schematic of the recording configuration, with representative voltage responses from the FS-IN (left) and CA1 PC (right) to a family of current pulses. (B) Representative traces showing the APs elicited in the presynaptic FS-IN (top) and the short-latency unitary IPSC in the CA1 PC (bottom) under control conditions (traces on the left), after bath application of baclofen (10 μM, middle), and subsequent application of CGP (5 μM, right). Note that baclofen reduced the IPSC amplitude by ~50%, whereas CGP resulted in an almost full recovery of the IPSC amplitude. Control traces are shown underlain. (C) Time course plot of the IPSC amplitude shows the effect of baclofen and CGP. IPSCs were recorded at 10 sec intervals.

Figure 5. Visualization, reconstruction and immunocytochemical identification of a biocytin-labeled recorded FS-BC. (A) A projection of stitched stacks of images of a FS-IN imaged with a 20X objective on a confocal microscope. The somata is located at the border of the str. radiatum (Rad.) and pyramidale (Pyr.) of the CA1 area, the dendrites run radially and span all layers, whilst the majority of axon is found in and around the cell body layer; as typical for BCs. Scale bar:100 μm. Inset, top right: high magnification projection of a 20 layers, showing typical baskets of axon forming around putative CA1 PC somata (orange asterisks). Scale bar: 10 μm. (B) The biocytin-filled cell body of the IN (white pseudocolour, lower panel, arrow) shows immunoreactivity for PV (in green, upper panel). Scale bar: 20 μm. (C) Projection of the 3-dimensional reconstruction of the cell. The soma and dendrites are in black and the axon colored red. Layers of CA1 are delineated in blue. Abbreviations: Ori, str. oriens; L-M, str. lacunosum-moleculare. Scale bar: 100 μm.

Table 1. Solutions list.

Discussion

We describe a method which combines electrophysiological and neuroanatomical techniques to functionally characterize morphologically- and neurochemically-identified neurons in vitro; in particular the diverse types of cortical inhibitory INs. Key aspects of the procedure are: (1) pre-selection of potential INs; (2) intracellular recording and neuron visualization; and finally (3) morphological and immunocytochemical analysis of recorded INs. Although this study has addressed PV-INs in particular, the described protocol can be used for similar recordings from any interneuron or other neuronal types, with minimal alteration.

The relatively low number of interneurons in cortical areas makes random selection and recording from these cells highly inefficient. Divergent localization and morphological features have enabled researchers to distinguish and routinely record from some interneuron types. However in slices, identification of interneurons in and near the cell body layers remains difficult, such as with FS-INs. The advent of transgenic mouse lines, expressing the fluorescent proteins in specific interneuron populations offers an elegant solution which makes pre-selection and recording of these neurons much more efficient2. Now many transgenic lines, mostly mice, but increasingly also rats13 are available, facilitating the investigation of interneurons. When using transgenic lines, however, it is essential to establish the extent and the specificity of the reporter expression.

Quality of cellular labeling and morphology, as well as the electrophysiological recording, critically depend on viable, high quality slices; for which the brain needs to be rapidly dissected (ideally 20-40 sec), handled very carefully and continuously chilled. We find that the use of sucrose-ACSF, whereby the overall sodium concentration and thus excitability of neurons is reduced, dramatically improves the quality of the slices and interneuron survival18. However, it should be highlighted that many researchers have made use of standard recording ACSF for slice preparation, to great effect10,11,15. Morphological integrity depends heavily on the angle at which slices are cut13; which varies by region and cell type. However, many interneurons can be reliably recorded from transverse, coronal or sagittal slices. To further minimize severance of dendrites and axon, cells deeper in the tissue should be targeted, albeit sacrificing IR-DIC quality and reliability of giga-ohm seal formation. Under these circumstances recordings from both CA1 pyramidal cells and FS-INs can be reliably obtained.

Labeling and visualization of the neurons is achieved following a typical recording session lasting 30 min or longer. If recordings last less than 30 min, it is necessary to allow this time prior to fixation to enable biocytin to diffuse into distal dendritic and axonal processes, guaranteeing complete filling and thus post-hoc visualization of cells.

In whole-cell recordings, maintaining low and stable series resistances is imperative to accurate physiological measurement of synaptic and intrinsic properties, as well as thorough labeling of the neurons. However low-resistance electrodes allow rapid dialysis of the cytoplasm with the intracellular solution contained within the pipette. Supplementing the intracellular solution with ATP, GTP and phosphocreatine certainly helps to maintain the energy supplies and recording quality. However, dialysis of neurochemicals and protein can lead to diminished responses, reduced plasticity19 and can also hamper neurochemical identification. For instance PV is consistently washed out of neurons over the course of longer experiments. Therefore, examination of distal dendritic or axonal processes may be required to determine immunoreactivity of the recorded neuron. In such cases where such dialysis is a concern, recordings using perforated-patch configuration can be utilized to preserve the intracellular environment 20, however the patch must be broken at the end of recordings or the neuron subsequently “repatched” in the whole-cell configuration to allow biocytin filling.

Pharmacological investigation of recorded neurons is a useful method to assess the functional significance of divergent neuromodulatory mechanisms in shaping the intrinsic and synaptic activity of INs. In the methods described above, pharmacological isolation and paired recordings are used to assess post and presynaptic GABABR-mediated effects, respectively; however one can easily modify the combination of receptor agonists and antagonists to assess the role of alternative receptor families, for example the cannabinoid system21.

Histological and immunocytochemical processing of recorded cells is highly reliable, once protocols are established. The immunocytochemistry protocol described here has been tuned with respect to the combination of antibodies used, slice thickness and requirement to identify the neurochemical content. We utilize normal goat serum as all secondary antibodies used are raised in goat. As alternatives, it is also possible to use bovine serum albumin (BSA) or milk powder as blocking agents. It should be noted that this protocol uses phosphate buffered saline (PBS) for antibody incubation, however alternatively one could simply use 0.1 M PB in lieu of PBS. Using a relatively high concentration of detergent (0.3% TritonX-100), does not diminish apparent antigenicity, while allowing penetration of antibodies into the first 100 μm surface layer of the slice, where neurons are routinely recorded. If deeper cells are recorded, or the slices are thicker, it is advisable to resection the slices, using a cryostat or a vibratome, and perform the immunolabeling on the thin (40-70 µm) sections. For 300 μm slices, a long incubation of the antibodies (at 4 ºC to preserve structural integrity of the slices) produces highly reliable immunocytochemical results.

Identification of the neurons relies on the combined information obtained in the recording, the post-hoc morphological and the immunocytochemical analysis. In the hippocampus, due to the strict laminar organization, axonal distribution is a good indicator of the postsynaptic targets of interneurons. Severed axon or dendrites, however, can make the identification difficult or impossible. Furthermore, some ambiguity remains, for example in differentiating PV+ BCs and axo-axonic interneurons, due to similarity in the overall axonal localization. A final definitive identification would require electron microscopic analysis of postsynaptic targets1,3 or further immunocytochemical dissection22.

Disclosures

The authors declare that they have no competing financial interests.

Acknowledgments

The authors wish to thank Ina Wolter for her excellent technical assistance. VGAT-Venus transgenic rats were generated by Drs. Y. Yanagawa, M. Hirabayashi and Y. Kawaguchi in National Institute for Physiological Sciences, Okazaki, Japan, using pCS2-Venus provided by Dr. A. Miyawaki.

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