Summary
Neuromodulatory control by oxytocin is essential to a wide range of social
1,2
, parental
3
and stress-related behaviors
4
. Autism spectrum disorders (ASD) are associated with deficiencies in oxytocin levels
5
and with genetic alterations of the oxytocin receptor (OXTR)
6
. Thirty years ago, Muhlethaler et al.
7
found that oxytocin increases the firing of inhibitory hippocampal neurons, but it
remains unclear how elevated inhibition could account for the ability of oxytocin
to improve information processing in the brain. Here, we describe a simple yet powerful
mechanism by which oxytocin enhances cortical information transfer
8
while simultaneously lowering background activity, thus greatly improving signal-to-noise.
Increased fast-spiking interneuron (FSI) activity not only suppresses spontaneous
pyramidal cell firing, but also enhances the fidelity of spike transmission and sharpens
spike timing. Use-dependent depression at the FSI-pyramidal cell synapse is both necessary
and sufficient for the enhanced spike throughput. Notably, we show the generality
of this novel circuit mechanism by activation of FSIs with cholecystokinin, or with
channelrhodopsin-2. This provides insight into how a diffusely delivered neuromodulator
can improve the performance of neural circuitry that requires synapse specificity
and millisecond precision.
Results
The CA1 region of hippocampus receives potent excitatory input from neighboring area
CA3 through the Schaffer Collateral (SC) pathway. Activation of SC axons evokes a
monosynaptic excitatory post-synaptic potential (EPSP) onto CA1 pyramidal cells, as
well as exciting a variety of CA1 interneurons. These interneurons then deliver a
millisecond-delayed inhibitory postsynaptic potential (IPSP), termed feed-forward
inhibition. Thus, both the stimulation threshold and the timing of spikes evoked in
CA1 pyramidal cells by SC activation are dictated by a finely tuned balance of monosynaptic
excitatory and disynaptic inhibitory inputs
8,9
.
In agreement with previous results
8
, we found that stimulation of the SC pathway in acute rat hippocampal slices evoked
spikes with a short latency and moderate jitter (Fig 1a). Strikingly, bath application
of TGOT (Thr4,Gly7-Oxytocin, 200 nM), a specific agonist for oxytocin receptors, dramatically
increased the probability of evoking a spike in the postsynaptic neuron from 0.50
to 0.82, while simultaneously suppressing the spontaneous activity of CA1 pyramidal
cells by 57% from 1.4 Hz to 0.6 Hz (Fig 1a–d). The combination of increased evoked
spike probability (signal) and reduced spontaneous activity (noise) resulted in an
enhanced signal-to-noise. TGOT also reduced the latency and increased the temporal
precision of evoked spikes (Fig 1e,f).
In agreement with previous work
7,10
, TGOT increased the rate and amplitude of spontaneous inhibitory postsynaptic currents
(IPSCs) onto CA1 pyramidal cells (Fig 1g, S1). Blockade by 10 µM bicuculline or by
100 nM tetrodotoxin indicated that these events were mediated by GABAA receptors and
likely required an increase in interneuron firing rather than a change in spontaneous
presynaptic release. The specific oxytocin receptor antagonist OTA ((d(CH2)5
1,Tyr(Me)2,Thr4,Orn8,des-Gly-NH2
9)-Vasotocin, 1 µM) blocked the TGOT-induced effects, suggesting that these actions
were solely mediated by the oxytocin receptor
10
. The TGOT-induced increase in spontaneous IPSCs was also abolished by the potent
P/Q-type calcium channel blocker ω-Agatoxin IVA, but unaffected by the N-type calcium
channel antagonist ω-Conotoxin GVIA, suggesting that these events primarily arise
from FSIs with little contribution from RS interneurons (Fig 1g, S1)
11,12
.
To test more directly whether TGOT precisely targeted FS interneuron subtypes, we
used whole cell recordings in CA1 strata oriens and pyramidale, stratum radiatum interneurons
being unresponsive to TGOT
10
and lacking OXTR expression
13
. We found a clear distinction: FSIs were responsive to TGOT, whereas RS interneurons
were not (Fig 2a). FSIs displayed robust responses upon application of 20 and 200
nM TGOT (Fig S2a–c), the latter producing a near-saturated effect. Dividing the increase
in IPSCs onto pyramidal cells (27.3 Hz, Fig 1g) by the increase in FSI firing rate
(8.8 Hz per FSI, Fig 2a), we calculate that on average each pyramidal cell receives
input from at least ~3.1 TGOT-responsive FSIs in our slices.
To clarify mechanisms by which TGOT depolarizes FSIs, we voltage clamped FS perisomatic-targeting
(basket and axo-axonic) and RS basket cells at −65 mV. TGOT induced a large inward
current in FSIs (Fig 2b, S2g), but as expected had no effect on the RS cells (data
not shown). TGOT also increased the rate of spontaneous IPSCs onto the FSI (Fig S2d–f),
as predicted from the FSI-FSI connectivity that may serve to regulate the distribution
and extent of inhibition.
To test whether the TGOT-induced inward current arises from G-protein signaling within
the FSI itself, we replaced the GTP in the intracellular recording solution with 1
mM GTPγS, a non-hydrolysable GTP analog that renders G-proteins constitutively active.
The GTPγS-induced current largely occluded the TGOT-induced current (Fig 2b, S2g),
verifying that the TGOT effects involve G-protein signaling within the recorded neuron.
The amplitude and kinetics of the TGOT-induced current were unaffected by intracellular
BAPTA, indicating that the intracellular signaling mechanism is likely not Ca2+-dependent
14
. In voltage ramp recordings from FSIs, the TGOT-induced current reversed at −3.1±3.4
mV (Fig 2c, S2h), suggesting that the currents were generated by a non-selective cation
channel. Partial replacement of external sodium by NMDG (50 mM) shifted the reversal
potential to more negative values (−13.8±3.7 mV, P<0.05 unpaired two-tailed t-test,
data not shown), pointing to Na+ as the predominant charge carrier of the TGOT-induced
inward current.
To investigate the mechanisms of the enhanced fidelity of spike transmission, we obtained
whole cell current clamp recordings from CA1 pyramidal cells and elicited spikes synaptically
or by current injection on interleaved trials (Fig 2d,e, S3, S4a). TGOT increased
the fidelity of synaptically evoked spikes in whole cell mode, paralleling its effect
in cell-attached recordings (Fig 1), but reduced the probability of evoking spikes
by whole cell current injection. This apparent reduction in pyramidal cell excitability
was coupled to a hyperpolarization of the cell membrane (Fig S4b). As TGOT had no
effect on the holding current or membrane resistance in voltage clamp recordings of
pyramidal neurons in the presence of bicuculline (Fig S4c), we concluded that the
reduction in spontaneous activity and excitability was wholly attributable to enhanced
inhibitory tone. This increase in inhibitory tone, however, made the enhanced EPSP-spike
coupling all the more surprising.
We hypothesized that the enhanced EPSP-spike coupling might arise from a shift in
the synaptic excitatory-inhibitory balance. Indeed, the disynaptic IPSP was reduced
by TGOT (Fig S4g,h), and bicuculline abolished the TGOT-induced increase in evoked
spike probability in cell-attached recordings (Fig 2f). To most rigorously isolate
inhibitory inputs, we stimulated the Schaffer Collaterals while holding the cell at
0 mV under voltage clamp, and found that the evoked disynaptic IPSC was reduced by
TGOT (Fig 2g,h). In contrast, the evoked excitatory postsynaptic current (EPSC), isolated
at −65 mV in the presence of bicuculline, was unaffected. This selective reduction
of the evoked IPSC while sparing the EPSC, shifts the excitatory-inhibitory (E-I)
balance and could account for the increase in evoked spike probability. This reduction
in feed-forward inhibition could arise from either a reduction in excitatory to inhibitory
(E→I) transmission, causing fewer interneurons to be activated, or a reduction in
inhibitory to excitatory (I→E) transmission, causing each interneuron to be less effective.
We recorded from FSIs while stimulating the SC pathway but found no effect of TGOT
on E→I transmission (Fig S4i,j). In contrast, stimulating the stratum pyramidale while
blocking excitatory transmission with NBQX and AP5 revealed a TGOT-induced suppression
of I→E transmission onto pyramidal cells (Fig S4k,l).
Using paired whole cell recordings, we investigated how TGOT reduces the evoked IPSC
at the I→E synapse. TGOT increased the spontaneous firing of presynaptic FSIs and
also diminished the FSI-pyramidal cell unitary IPSC, without affecting RS interneurons
(Fig 3a,b). When the TGOT-induced depolarization of the presynaptic FSI was countered
with a hyperpolarizing bias current, however, the spontaneous firing in the presynaptic
cell ceased and the unitary evoked IPSC was maintained at its pre-TGOT amplitude.
This rescue suggests that TGOT induces a use-dependent depression of the IPSC
15,16
, and that the increase in spontaneous FSI firing is necessary for the reduction in
the evoked feed-forward IPSC.
To test whether the TGOT-induced increase in the FSI firing rate was sufficient to
account for the observed synaptic depression, we drove 10 s trains of action potentials
in the absence of TGOT (Fig 3c). The frequency dependence of the residual IPSC following
a 10 s train in control ACSF (Fig 3d, colored diamonds) matched closely with that
of the residual IPSC in TGOT (Fig 3d, black symbols). Thus, the TGOT-mediated increase
in FSI spontaneous firing is not only necessary (Fig 3a,b) but also sufficient (Fig
3c,d) to account for the observed decrease in evoked IPSC amplitude (Fig 2g,h), and
enhancement of EPSP-spike coupling (Fig 1). Recovery of the IPSC was nearly complete
by 4.5 s following the 50 Hz train, consistent with a rapid switching of the FS synapses
between baseline and depressed states
16
(Fig 3e). We also observed a modest, frequency-dependent increase in the spike width
over the 10 s trains (Fig S5) that would be expected, if anything, to increase presynaptic
release, contrary to the use-dependent depression that was observed.
The specificity of TGOT for FSIs suggested that this mechanism may be a general property
of this network, and that any peptide, network state, or signal that increases the
spontaneous activity of FSIs will also increase the fidelity of spike transmission.
We tested this hypothesis using two independent approaches, first stimulating FSIs
with the peptide cholecystokinin (CCK), and second targeting this population with
the light-activated ion channel channelrhodopsin-2 (ChR2).
CCK activates FS basket cells
11
, transiently increasing their firing rate in a manner reminiscent of TGOT. In close
agreement with our TGOT results, CCK enhanced inhibitory tone and suppressed the evoked
feed-forward IPSC without affecting the evoked EPSC (Fig 4a,b, S6). In cell-attached
recordings, CCK increased the probability of evoking spikes in CA1 pyramidal cells
by SC stimulation, while simultaneously suppressing the spontaneous firing of these
cells (Fig 4c,d). Furthermore, both the latency and the jitter of the evoked spikes
were reduced by CCK (Fig S6f), just as they were with TGOT (Fig 1f).
We then used ChR2 to selectively activate FSIs in acute hippocampal slices from PV-cre
BAC transgenic mice. Immunostaining confirmed that the ChR2 was efficiently targeted
to the parvalbumin-expressing (PV+) FSIs (Fig S7). Optogenetic activation of FSIs
induced IPSCs in CA1 pyramidal cells that showed a strong synaptic depression (Fig
4e), consistent with our paired recording data (Fig 3), and with previous reports
15,16
. In agreement with our TGOT and CCK results, driving FSIs with a brief train of blue
light pulses preceding an electrical stimulus to the SC pathway modestly increased
the probability of eliciting a spike in a postsynaptic pyramidal cell, relative to
interleaved control trials in which the blue light was omitted (Fig S8a). Examination
of the spike latency and jitter, however, revealed a high probability that monosynaptic
inhibition contaminated a subset of these recordings. When the recordings with the
shortest latency and lowest jitter were excluded (Fig S8d–f, see methods), the remaining
neurons all exhibited a pronounced increase in evoked spike probability following
ChR2 stimulation (Fig 4f,g). Taken together, three interventions, TGOT, CCK and ChR2,
therefore all converge on a single surprising conclusion: that activation of FSIs
enhances the fidelity of spike transmission in the hippocampus. TGOT also increased
the evoked population spike amplitude in the presence of kainate-induced gamma rhythms
(Fig S9), thus confirming that the TGOT-induced enhancement in EPSP-spike coupling
is robust under more in vivo-like conditions.
We constructed a minimal computational model to investigate the mechanisms linking
FSI activation to the evoked spike probability, latency and jitter. We mimicked the
enhanced FSI activity by increasing the rate and amplitude of spontaneous IPSCs. The
synaptic depression at the FSI-pyramidal synapse was simulated by reducing the evoked
IPSC to 60% of its basal value. In agreement with our experimental results, these
changes reduced the simulated evoked IPSP, increased the simulated evoked spike probability,
and sharpened the evoked spike timing (Fig S10).
We then asked why a decrease in feed-forward inhibition shrinks evoked spike latency
and jitter, in apparent conflict with the idea that feed-forward inhibition enforces
sharp spike timing
8,9
. Resolution is achieved by considering how the EPSC and IPSC conductances (gEPSC
and gIPSC) regulate membrane voltage near the spike-firing threshold. A reduction
in gIPSC allows an unaltered gEPSC to push the membrane potential up to the spike
firing threshold more reliably and more quickly and precisely (Fig 4h,i). In contrast,
if gEPSC is reduced to nearly the same degree as gIPSC in order to clamp the likelihood
of spike firing
8
, the latency and jitter are increased.
Finally, we probed the functional consequences of the strikingly incomplete depression
of the FS synapses (Fig 3c) and the effects of varying the latency between the onset
of gEPSC and gIPSC. Fidelity of spike transmission (defined as the fraction of sweeps
containing precisely one postsynaptic spike) is maximal when IPSCs are depressed by
approximately 40% (Fig 4j), the value we observed experimentally in response to either
TGOT or CCK application (Fig 2g,h, 4a,b). Likewise, a residual IPSC of 50–60% was
optimal in considerations of global spike jitter (Fig S10h). Thus, the empirically
observed TGOT response in FSIs seems well suited in multiple respects to the efficient
retuning of overall circuit performance.
Discussion
Our experiments reveal a generalized mechanism through which oxytocin improves the
fidelity and temporal precision of information transfer through brain networks. Oxytocin
enhanced circuit performance in three ways: increasing throughput of output spikes,
sharpening submillisecond spike timing, and suppressing background firing. Each of
these improvements in circuit signal-to-noise could be traced to the action of oxytocin
on FSIs and reproduced through quantitative simulations, as well as through other
interventions that specifically activate FSIs.
The rapid onset and recovery of FSI use-dependent synaptic depression is well suited
to shift circuit dynamics rapidly yet stably in response to oxytocin, whether delivered
quickly and focally, as in synaptic release
4
, or presented diffusely at low doses, as in volume transmission. The partial depression
of FSI synapses (residual, ~35%) and the sparing of RS interneurons ensures that modulation
by oxytocin avoids the dangers associated with a complete loss of inhibition such
as dramatically impaired spike timing precision
8
(Fig S10, S11) and epileptogenesis.
Our experiments provide a circuit mechanism linking three disparate aspects of ASD
17
. Oxytocin signaling has been implicated in ASD by genetic analysis and pharmacological
studies
18,19
. PV-positive FSIs are important in autism etiology
20
, presumably due to their role in excitation-inhibition balance and neuronal oscillations,
both of which are likely impaired in ASD. Deficiencies in signal-to-noise, observed
as unreliable cortical evoked potentials in ASD
21
, offer a valuable endophenotype, but have not yet been linked to a circuit defect
or a therapeutic strategy. Our finding that FSIs are direct targets of oxytocin
18,19
and can potently modulate circuit signal-to-noise, shows these cells may be uniquely
poised to counteract deficits in rapid information processing in psychiatric disorders
21
. In healthy individuals, oxytocin signaling through FSIs may provide a salience cue,
capable of transiently enhancing cognitive performance
1,3,22
. Indeed, increasing PV+ interneuron activity was sufficient to recover hippocampal-dependent
behavioral deficits in a mouse model of Alzheimer’s Disease
23
. There may be parallels in the visual cortex as well, where optogenetic activation
of PV+ interneurons operates like a salience cue and sharpens orientation tuning
24
.
What is the functional logic for oxytocin selectively targeting FSIs amidst the wide
variety of interneuron types? One rationale is that the FSIs engaged by oxytocin are
physiologically and functionally distinct from RS interneurons, which themselves are
regulated by endocannabinoids
25
. Thus, by targeting select interneuron populations, neuromodulators are able to regulate
different forms of inhibition, with oxytocin affecting feed-forward inhibition and
endocannabinoids tuning feed-back inhibition.
Another modulator, noradrenaline (NA) enhances circuit signal-to-noise in slice and
in vivo through a variety of mechanisms across multiple brain regions including the
hippocampus
26,27
and auditory system
28
. In auditory brainstem, Kuo and Trussell elegantly described how NA suppresses cartwheel
inhibitory neuron spiking, relieving their output synapses from tonic depression
28
. Although their mechanism differs from ours in direction of change and functional
outcome, an emergent general principle is that modulation of inhibitory neuron tonic
firing and variation in use-dependent synaptic depression can regulate signal-to-noise.
In the hippocampus, Madison, Nicoll and colleagues delineated several monoamine responses
across excitatory and inhibitory neurons that enhance circuit signal-to-noise
26,29,30
. Although oxytocin and NA both enhance signal-to-noise of CA1 pyramidal neurons,
the widespread effects of NA contrast sharply with the exquisitely focused mechanism
we uncovered. Oxytocin accomplishes both the enhanced fidelity of spike transmission
and the suppression of background activity by selectively targeting a single locus:
FSI activity. Furthermore, FSI synaptic depression in hippocampal CA1 (Fig 4) is representative
of that in dentate gyrus
16
, cortex
15
, and elsewhere, suggesting similar modulation of signal-to-noise by FSI activity
may be essential in many brain regions.
Methods
Slice preparation
Rat hippocampal slices (350 µm thick) were prepared using a Leica VT 1000S vibratome
from p21-p28 Sprague-Dawley rats of either sex in ice-cold sucrose slicing solution
containing (in mM) 206 Sucrose, 11 D-Glucose, 2.5 KCl, 1 NaH2PO4, 10 MgCl2, 2 CaCl2
and 26 NaHCO3. Rats were anaesthetized with isofluorane inhalation before decapitation
and dissecting out of the hippocampus. Mouse transverse hippocampal slices (300µm)
were prepared using a Vibratome 1000 plus (Vibratome, St. Louis, MO). Mice were deeply
anaesthetized with intraperitoneal injection of pentobarbital (100 mg/kg body weight)
and then transcardially perfused with ~30 ml ice-cold sucrose-ACSF solution containing
(in mM) 252 sucrose, 24 NaHCO3, 1.25 NaH2PO4, 3 KCl, 2 MgSO4, 10 D-Glucose and 0.5
CaCl2. All slices from rats and mice were allowed to recover submerged in artificial
cerebro-spinal fluid (ACSF) for 1 hr at 34°C, and then maintained at room temperature
until recording. For recordings from rat tissue, ACSF contained (in mM) 122 NaCl,
3 KCl, 10 D-Glucose, 1.25 NaH2PO4, 2 CaCl2, 1.3 MgCl2, 26 NaHCO3, 3 Na-Pyruvate, 2
Na-Ascorbate and 5 L-Glutamine. For mouse recordings, ACSF contained (mM) 124 NaCl,
26 NaHCO3, 2.5 KCl, 1.25 NaH2PO4, 2 CaCl2, 2 MgSO4, 5 L-Glutamine, and 10 D-Glucose.
All slice preparation and recording solutions were oxygenated with carbogen gas (95%
O2/5% CO2, pH 7.4).
Electrophysiological recordings
Recordings were performed in a submerged chamber at 32–34°C with constant bath perfusion
of ACSF at ~5 mL/minute for rats, ~2 mL/minute for mice. Slices were allowed 15–45
min to equilibrate before recording. Because the GABAB blocker CGP52432 (2 µM) did
not affect the TGOT enhancement of evoked spike probability, recordings were pooled
from control ACSF (N=7 cells) and CGP52432 (N=8 cells) conditions to measure spike
probability, suppression of spontaneous firing, and evoked spike timing (Fig 1). For
cell-attached measurement of TGOT influence on spontaneous activity, results were
pooled from recordings in control ACSF (N=15 cells) and in the presence of CGP52432
at 2 µM (N=8 cells). To prevent ictal activity, the CA3 region of each slice was removed
before recordings in bicuculline. Recordings were made using glass pipettes with a
tip resistance of 2–4 MΩ. For cell-attached recordings, pipettes were filled with
ACSF and the amplifier was set in voltage clamp mode. Slices were visualized with
an upright microscope (Zeiss Axioskop 2 FS plus) using infrared differential interference
contrast (IR-DIC) optics. Data were recorded with a MultiClamp 700B amplifier (Axon
Instruments, Union City, CA), filtered at 10 kHz using a Bessel filter and digitized
at 20 kHz with a Digidata 1322A analog-digital interface (Axon Instruments). For whole
cell recordings, experiments were discarded if the series resistance changed significantly
or reached 20 MΩ. Spontaneous IPSCs onto pyramidal cells were detected in voltage
clamp using a 50 mM Cl- internal solution containing (in mM) 70 CsMeSO3, 35 CsCl,
15 TEA-Cl, 1 MgCl2, 0.2 CaCl2, 10 HEPES, 0.3 EGTA, 10 Tris-Phosphocreatine, 4 Mg-ATP,
and 0.3 Na-GTP. For evoked IPSC and EPSC recordings, the internal solution contained
(in mM) 130 CsMeSO3, 8 CsCl, 1 MgCl2, 10 HEPES, 0.3 EGTA, 10 Tris-Phosphocreatine,
4 Mg-ATP, and 0.3 Na-GTP. Bicuculline (10 µM), TTX (100 nM) and OTA (1 µM) were delivered
as indicated in the bathing solution throughout the recording (Fig 1, S1). Calcium
channel blockers ω-agatoxin IVA at 0.5 µM (AgaIVA) or ω-conotoxin GVIA at 1 µM (GVIA)
were delivered by pre-treating the slice for 30 min in an interface chamber before
recording in control ACSF. AgaIVA and GVIA recordings were performed in separate slices
from the same experimental animal.
Synaptic events were evoked using a tungsten bipolar stimulating electrode placed
in the Schaffer Collateral excitatory afferents from area CA3 to deliver stimuli 100
µs in duration. With the exception of Figure S4k,l, the stimulating electrode was
placed far from the recorded cell (~400 µm to ~800 µm) to minimize monosynaptically
evoked IPSCs. In Figure S4k,l, monosynaptic IPSCs were evoked using sub-maximal stimulation
by placing the stimulating electrode in the pyramidal cell layer close to the recorded
cell (~100 µM), and including 10 µM NBQX and 50 µM AP5 in the bath to block excitatory
transmission. For evoked IPSP measurement, data were pooled from evoked spike successes
and failures and from recordings in the presence (N=5 cells) or the absence (N=1 cell)
of the GABAB antagonist CGP52432 (2 µM). Evoked disynaptic feed-forward IPSCs (Fig
2g,h; Fig 4a,b) were recorded as outward currents at a holding potential of 0 mV in
control ACSF. Evoked EPSCs were isolated by including 10 µM bicuculline in the bath
and holding the cell at −65 mV. Two out of 14 recordings in Figs 4c,d and S6e,f were
performed in the continuous presence of AM-251 (2 µM) to confirm the persistence of
the CCK-induced enhancement of EPSP-spike coupling even when endocannabinoid signaling
was blocked.
For current clamp recordings, and all interneuron recordings except for the voltage
ramp experiments, the intracellular solution contained (in mM) 130 K-Gluconate, 1
MgCl2, 10 HEPES, 0.3 EGTA, 10 Tris-Phosphocreatine, 4 Mg-ATP, and 0.3 Na-GTP. For
interneuron recordings this solution was supplemented with 0.1% biocytin. GTP was
omitted in experiments featuring GTPγS. For voltage clamp recordings of TGOT-induced
currents in FSIs, traces were divided into 10 s segments, with the mean value of each
segment plotted as a function of time to exclude synaptic events. See Fig S2d for
exemplar raw trace. All recordings were baseline-subtracted to adjust for the leak
current measured during the first 2 min before the onset of the GTPγS-induced current.
Traces were time-aligned to the wash-in of TGOT (red bar). For one cell in the GTPγS
data set in which baseline recording period was 10 min rather than 15 min, the pre-TGOT
period was aligned to the start of the other recordings, and the remainder of the
trace starting with TGOT wash-in was aligned to the TGOT wash-in of the other traces.
FSI Voltage Ramp Recordings
For voltage ramp recordings, the internal solution contained (in mM) 50 K-Gluconate,
70 CsMeSO3, 10 TEA-Cl, 1 MgCl2, 10 HEPES, 0.3 EGTA, 10 Tris-Phosphocreatine, 4 Mg-ATP,
and 0.3 Na-GTP. The pipette reference potential was set to zero and a junction potential
of −15.1 mV (calculated using pClamp) was corrected post hoc. An additional, empirically
measured correction factor of 3.3 mV was applied to correct for a change in the junction
potential introduced by partial replacement of sodium with NMDG in the voltage ramp
ACSF. Apart from the voltage ramp recordings, other membrane potentials reported are
not corrected for liquid junction potentials. After obtaining a whole cell recording
from a putative interneuron, the fast-spiking phenotype was verified as described
below. The amplifier was then switched to voltage clamp mode and the bath solution
was substituted for Voltage Ramp ACSF containing (in mM) 112 NaCl, 10 D-Glucose, 3
KCl, 1.25 NaH2PO4, 10 TEA-Cl, 1.3 MgCl2, 2 CaCl2,26 NaHCO3, 5 4-Aminopyridine, 0.1
CdCl2 and 0.001 TTX. Voltage ramps ~1 s in duration between −91 and +29 mV were applied
once every 10 s until the current at each potential reached a steady state for >2
min, at which point TGOT was applied. In 3 out of 13 recordings the voltage ramp-activated
current (1) became more negative at all potentials shortly after TGOT application,
and (2) failed to return to baseline after washout of the drug. It was assumed that
this global shift was caused by a change in the space clamp or access resistance and
these recordings were excluded from further analysis.
Drugs and reagents
All salts and buffers for intracellular and extracellular solutions, as well as ATP,
GTP, GTPγS, phosphocreatine and biocytin were purchased from Sigma (St. Louis, MO).
TGOT ((Thr4,Gly7)-Oxytocin), OTA ((d(CH2)5
1,Tyr(Me)2,Thr4,Orn8, des-Gly-NH2
9)-Vasotocin) and CCK (cholecystokinin octapeptide) peptides were purchased from Bachem
(Torrance, CA), dissolved at 1 mM in ddH2O and stored at −20°C until use within 6
months of purchase. Bicuculline, TTX, NBQX, and D-AP5 were purchased from Ascent Scientific
(Princeton, NJ). ω-conotoxin GVIA and ω-agatoxin IVA were purchased from Peptides
International (Louisville, KY). Stock solutions were prepared and stored according
to manufacturer specifications.
Interneuron labeling and classification
Physiological classification of interneuron subtypes was based on established criteria
11,25,31
. Fast-spiking cells were defined as those including (1) peak firing rates >200 Hz
with little firing rate accommodation, (2) characteristic FS action potential waveform,
and (3) minimal hyperpolarization-induced sag current due to Ih. Following interneuron
recordings, slices were transferred to a fixative solution containing 4% paraformaldehyde,
0.2 % picric acid and 1× phosphate buffered saline for 24–72 h before being stained
with 3,3’-diaminobenzidine tetrahydrochloride (0.015%) using a standard ABC kit (Vector).
Neuronal cell types were identified based on morphology of axonal and dendritic arbors
and electrophysiological properties of the cell. The FS perisomatic-targeting set
includes both basket cells (shown), and axo-axonic cells (not shown). Because of technical
challenges of discriminating FS basket and axo-axonic cells unequivocally, both cell
types were pooled into a single group of FS perisomatic-targeting cells. When analyzed
separately, both putative types were equivalently responsive to TGOT.
Analysis of cell-attached and intracellular recording data
Analysis of spikes, evoked synaptic currents, and synaptic potentials were performed
offline using custom written routines in MATLAB (Mathworks). Spontaneous IPSCs were
detected using a modified version of the detectPSPs script by Phil Larimer (http://www.mathworks.com/matlabcentral/fileexchange).
Spike jitter histograms were calculated by subtracting the latency of each spike from
the average latency of spikes evoked in that cell. The average latency and jitter
were calculated separately for control and TGOT/CCK conditions in each cell. To measure
the spike width, raw data was oversampled to 133 kHz using the MATLAB spline function.
Time course of spontaneous activity in pyramidal cell attached recordings was calculated
by averaging over all cells and smoothing in time with a boxcar filter (width=7 sweeps).
Optical stimulation of channelrhodopsin-2
Photostimuli were produced by three Luxeon Rebel LEDs (470 nm, Philips Lumileds, San
Jose, CA) driven by a custom-built controller. The LEDs were placed below the recording
chamber for full slice illumination once stable recording conditions were reached.
Light pulses were 5 ms in duration with a power of approximately 0.5 mW/mm2. ChR2-evoked
IPSCs were recorded from CA1 pyramidal neurons (n=10 cells, N=4 animals). Six of these
neurons were recorded in the same region of the same slice as neurons recorded in
the cell-attached data set.
Data analysis for cell-attached recordings involving blue light stimulation
In the full data set, blue light stimulation increased the spike probability in 13
of 16 neurons (Fig S8a, 12% increase in spike probability including all neurons, P<0.05,
two-tailed t-test). In recordings from rat neurons the average increase in spike firing
probability with TGOT or CCK was not correlated with spike latency, whereas in the
mouse data we found a strong correlation between control spike latency and the ChR2-induced
increase in spike probability (Fig S8d,e). In the mouse data set, the shortest latency
spikes showed the weakest increase in spike firing probability. Plotting the latency
against the jitter of spikes elicited under control conditions, we found a clear separation
between two groups of cells, in which evoked spikes from one set of cells occurred
with very short latency and little jitter and spikes from another set of cells occurred
at longer latency and with more jitter (Fig S8f).
Because of the smaller size of the mouse brain, we found our slice angle to be less
reliably transverse than in the rat preparation. As a result, the stimulating and
recording electrodes were placed closer to one another in the mouse slice in order
for the stimulating electrode to recruit a sufficient number of excitatory Schaffer
Collateral fibers to drive an action potential in the postsynaptic CA1 pyramidal cell.
This change in recording configuration unfortunately increases the probability of
directly activating inhibitory fibers with the stimulating electrode and generating
a monosynaptic IPSC. A well-documented set of physiological parameters, including
synaptic kinetics and cell excitability
25
ensure that the physiologically relevant disynaptic IPSC arises mostly from FSIs.
The monosynaptically activated IPSC, however, will arise from a less targeted subset
of neurons, and therefore be less susceptible to modulation by interventions that
selectively target FSIs.
In the cell-attached recording configuration, it was impossible to determine directly
the relative monosynaptic and disynaptic contributions to the feed-forward IPSC. The
monosynaptic IPSC relies only on a single GABAergic synapse, however, whereas the
disynaptic IPSC relies on three sequential steps: (1) a glutamatergic synapse onto
the interneuron, (2) the subsequent action potential in the interneuron, and finally
(3) the GABAergic transmission onto the postsynaptic pyramidal cell. The monosynaptic
IPSC will therefore occur with a shorter latency and less jitter than the disynaptically
evoked IPSC. As a result, spikes in pyramidal cells in which the feed-forward IPSC
is dominated by a monosynaptic component will be expected to occur with a shorter
latency and less jitter than spikes in cells experiencing a more physiological disynaptic
feed-forward IPSC. We therefore excluded the tightly clustered group of neurons with
very short latency and low jitter spikes from the mouse data set (N=9 cells) and analyzed
only the neurons in which spikes occurred with a longer latency and more jitter, consistent
with disynaptic feed-forward inhibition (N=7 cells). All of these remaining cells
demonstrated an increase in spike firing probability following blue light stimulation
(7 out of 7 cells, 28% increase in spike probability; P<0.01 paired two-tailed t-test).
In the complete data set (N = 16 cells) we observed a modest increase in spike latency
following blue light stimulation of PV interneurons across all 16 neurons (Fig S8b,c).
However, in the 5 out of 7 cells from the restricted data set that fired at least
5 spikes in both the control and blue light stimulation conditions, light activation
of PV interneurons reduced the latency (Fig S8g, 10.35 ms in control, 10.07 ms following
light stimulation; P = 0.73 paired two-tailed t-test) and jitter (Fig S8h, 16.58 ms2
control; 11.78 ms2 light stimulation; P = 0.23 paired two-tailed t-test) of spikes.
Although this reduction in latency and jitter did not reach statistical significance,
the trend is consistent with our TGOT and CCK results.
Immunohistochemistry
At the end of each ChR2 recording session, slices were fixed overnight with 4% paraformaldehyde
(PFA)/phosphate buffered saline (PBS) solution and cryoprotected by immersion in 30%
sucrose/PBS solution overnight at 4°C. Tissues were embedded in Tissue Tek, frozen
on dry ice, and cryosectioned at 20 µm thickness. Sections for were processed using
1.5% normal goat serum (NGS) and 0.1% Triton X-100 in all procedures except washing
steps, where only PBS was used. Sections were incubated in blocking solution for 1
hr, followed by incubation with the primary antibodies overnight at 4°C. Cryostat
tissue sections were stained with the primary antibodies: mouse anti-Parvalbumin (1:1000,
Sigma) and rabbit anti-DsRed (1:500, Chemicon). Secondary antibodies conjugated with
Alexa fluoro dyes 488, 594 (Molecular Probes) raised from the same host used for blocking
serum were applied for 1 hr at room temperature. Nuclear counterstaining was performed
with 100 ng/ml 4,6-diamidino-2-phenylindole (DAPI) solution in PBS for 5 min. Fluorescent
images were captured using a cooled-CCD camera (Princeton Scientific Instruments,
NJ) using Metamorph software (Universal imaging, Downingtown, Pennsylvania).
Virus injection
Adeno-associated virus carrying ChR2 fused to the fluorescent marker mCherry AAV2/1.EF1.dflox.hChR2(H134R)-mCherry.WPRE.hGH,
(University of Pennsylvania Gene Therapy Program Vector Core) was injected bilaterally
into dorsal hippocampal CA1 region of Pvalb-cre (PV-Cre) transgenic mice
32
(aged between postnatal days 15–19) at three sites: 2.2, 1.8 and 1.6 mm posterior
from bregma, 2.4, 2.1, 1.7 mm from midline, and 1.2, 1.1, and 1 mm below cortical
surface, respectively. Animals were anesthetized with isoflurane, mounted in a stereotactic
apparatus and kept under isoflurane anesthesia during surgery. We injected 100 nL
of virus at each location over a 2 min period using a glass micropipette (tip diameter
~20 µm) attached to a Nanoliter 2000 pressure injection apparatus (World Precision
Instruments). The pipette was held in place for 3 min following each injection before
being completely retracted from the brain. Mice were returned to their home cage for
2–3 weeks before acute slice preparation to allow for virus expression.
Computational model of EPSP-spike coupling
The computater modeling was performed using NEURON and automated using MATLAB. A simplified
pyramidal cell, consisting of a soma, a single axon and a single dendrite was initialized
to starting parameters before each stimulus. Background and voltage-gated conductances
were based on reported models
33,34
. Small adjustments were made to improve agreement of parameters such as cell excitability
and action potential waveform between the model and experimental observations. Each
sweep consisted of (1) a “monosynaptic” EPSC onto the dendrite, (2) a “disyanptic”
feed-forward IPSC onto the soma and dendrite 2 ms after the evoked EPSC (unless otherwise
specified), and (3) multiple “spontaneous” IPSCs onto the soma with randomly distributed
amplitudes and timing. To isolate the role of the feed-forward IPSC from changes in
inhibitory-tone, spontaneous IPSCs were omitted in the simulation used to generate
Fig 4h,i. At the outset of each set of sweeps, the “evoked” EPSC-IPSC amplitudes were
set empirically by increasing the EPSC and IPSC conductances together with a fixed
ratio of 6:1 until ~50% chance of spike propagation was reached. Experimental measurement
of IPSC/EPSC ratio ranged from 2.62 to 5.20 (mean±S.E.M. of 3.65±0.28). This experimentally
measured range is presumed to be an underestimate of the true ratio due to imperfect
isolation of the IPSC reversal potential, causing a presumed GABAergic contribution
to the measured EPSC in some cells. In the model, IPSC/EPSC ratios from 4:1 up to
6:1 showed a pronounced TGOT-induced increase in evoked spike probability, with 6:1
supporting the strongest influence of TGOT on spike timing and jitter. Variability
was introduced by using pseudo-random number generation to vary independently (1)
the evoked EPSC conductance, (2) the evoked IPSC conductance and (3) the spontaneous
IPSC timing and amplitudes. Evoked EPSC and IPSC conductances were varied independently
on each sweep according to a normal distribution centered on the empirically determined
mean value, with a standard deviation that was 5% of the mean. TGOT was simulated
by (1) reducing the evoked somatic IPSC conductance to 60% of “baseline”, while sparing
the evoked EPSC and the dendritic IPSC, (2) doubling the spontaneous IPSC amplitude,
and (3) increasing the spontaneous IPSC rate from 5 Hz to 35 Hz. The IPSC reversal
potential was set at −110 mV for Fig S10b–c, consistent with the calculated GABAA
reversal potential in our whole cell recording conditions. For the rest of the simulations,
the IPSC reversal potential was set to −90 mV, consistent with cell-attached recording
conditions. The increase in evoked spike probability was robust as the GABAA reversal
potential was varied from −80 mV to −120 mV (Fig S12), while the reduction in latency
and latency jitter were decreased in magnitude but remained statistically significant.
Supplementary Material
1