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Research ArticleNeural Control

Hydrogen peroxide inhibits neurons in the paraventricular nucleus of the hypothalamus via potassium channel activation

Published Online:26 Jun 2019https://doi.org/10.1152/ajpregu.00054.2019

Abstract

The paraventricular nucleus (PVN) of the hypothalamus is an important homeostatic and reflex center for neuroendocrine, respiratory, and autonomic regulation, including during hypoxic stressor challenges. Such challenges increase reactive oxygen species (ROS) to modulate synaptic, neuronal, and ion channel activity. Previously, in the nucleus tractus solitarius, another cardiorespiratory nucleus, we showed that the ROS H2O2 induced membrane hyperpolarization and reduced action potential discharge via increased K+ conductance at the resting potential. Here, we sought to determine the homogeneity of influence and mechanism of action of H2O2 on PVN neurons. We recorded PVN neurons in isolation and in an acute slice preparation, which leaves neurons in their semi-intact network. Regardless of preparation, H2O2 hyperpolarized PVN neurons and decreased action potential discharge. In the slice preparation, H2O2 also decreased spontaneous excitatory postsynaptic current frequency, but not amplitude. To examine potential mechanisms, we investigated the influence of the K+ channel blockers Ba2+, Cs+, and glibenclamide on membrane potential, as well as the ionic currents active at resting potential and outward K+ currents (IK) upon depolarization. The H2O2 hyperpolarization was blocked by K+ channel blockers. H2O2 did not alter currents between −50 and −110 mV. However, H2O2 induced an outward IK at −50 mV yet, at potentials more positive to 0 mV H2O2, decreased IK. Elevated intracellular antioxidant catalase eliminated H2O2 effects. These data indicate that H2O2 alters synaptic and neuronal properties of PVN neurons likely via membrane hyperpolarization and alteration of IK, which may ultimately alter cardiorespiratory reflexes.

INTRODUCTION

Reactive oxygen species (ROS) have diverse cellular functions (16), including a role as signaling molecules to modulate neuronal circuits (25, 55). The ROS H2O2 is produced by dismutation of superoxide. Because of its relative stability and ability to freely cross the plasma membrane, H2O2 has been shown to be important in a variety of neurophysiological processes. For instance, H2O2 alters neurotransmission, ion channel function, and cellular activity in several central nuclei (22, 32). In the nucleus tractus solitarius (nTS), the initial site of chemoafferent termination and hypoxic reflex modulation, we have demonstrated that H2O2 produces complex responses. H2O2 reversibly hyperpolarized membrane potential (Vm), reduced membrane resistance, and shifted action potential (AP) threshold in a hyperpolarized direction; changes were due to activation of Ba2+-sensitive K+ currents (IK) (42). Upon H2O2 washout, nTS neurons became more excitable, in part, due to enhancement of Ca2+ currents (41). H2O2 signaling can be terminated by enzymatic breakdown of H2O2 by catalase, glutathione peroxidase, and peroxidoxins (63).

The nTS reciprocally projects to the paraventricular nucleus (PVN) of the hypothalamus (19, 50), which is critical in eliciting the full cardiorespiratory and neuroendocrine response to a variety of stimuli and stressors, including the peripheral chemoreflex (24, 40, 46, 51). In the PVN, H2O2 is a potent modulator that contributes to sympathetic nerve overactivity of the cardiac sympathetic afferent reflex, as well as renovascular and salt-sensitive hypertension (64–66). However, H2O2 also reduces the effectiveness of exogenous glutamate in the PVN on sympathetic activation (7). These results suggest that H2O2 may contribute to the balance of inhibitory and excitatory drives within the PVN, their efferent projections, and the physiological consequences. Given the influence of H2O2 on nTS activity, the reciprocal connection between the nTS and PVN, the contribution of the PVN to peripheral reflexes, and previous work demonstrating that H2O2 modulates PVN influence on cardiovascular function, we sought to examine the influence of H2O2 on PVN parvocellular synaptic and neuronal activity. Our present studies show that H2O2 hyperpolarizes Vm due to activation of voltage-gated IK to ultimately limit glutamatergic synaptic drive.

METHODS

Animals and Ethical Approval

Animal protocols were approved by the Animal Care and Use Committees of the University of Missouri and handled according to National Institutes of Health guidelines. Three- to 6-wk-old male Sprague-Dawley rats (ENVIGO, Indianapolis, IN) were housed in an in-house animal facility at 22°C with a 12:12-h light-dark cycle and food and water available ad libitum.

Immunohistochemistry and Expansion Microscopy

Isoflurane-anesthetized rats were transcardially perfused with cold 0.01 M phosphate-buffered saline (PBS, ~200 ml, pH 7.4), followed by cold 4% paraformaldehyde (PFA, 200 ml, pH 7.4; Sigma-Aldrich). Brains were postfixed for 2 h in 4% PFA and then rinsed in 0.01 M PBS. Forebrains were removed, and coronal sections of the area containing the PVN were sectioned at 100 μm using a vibrating microtome (model VT 1000S, Leica) in 0.01 M PBS. Sections were rinsed (3 times for 15 min each) with 0.01 M PBS and then permeabilized in 0.1% Triton X-100-PBS containing 3% normal donkey serum (NDS; Millipore) for 6 h at 4°C. Tissue sections were incubated with primary antibodies against the antioxidant catalase [rabbit anti-catalase, 1:300 dilution; catalog no. ab1877, Abcam; research resource identifier (RRID): AB_302649] and NeuN (mouse anti-NeuN, 1:500 dilution; catalog no. MAB377, Millipore; RRID: AB_2298772) in 0.1% Triton X-100-PBS and 3% NDS for 24 h at 4°C. Specificity of these antibodies has been previously established (42). Slices were rinsed with cold 0.01 M PBS (4 times for 30 min each) at 4°C and incubated for 24 h at 4°C in appropriate fluorescent secondary antibody (Jackson ImmunoResearch) in 0.1% Triton X-100-PBS and 3% NDS. On the following day, slices were washed (3 times for 20 min each) with room-temperature PBS.

Expansion microscopy was used to improve visualization of small structures, as described by others (8, 10). After catalase and NeuN antibody binding, slices were incubated with 1 mM methacrylic acid-N-hydroxysuccinimide ester (catalog no. 730300, Sigma-Aldrich) in PBS for 1 h at 22°C, rinsed with PBS (3 times for 20 min each at 22°C), and then incubated in a 24-well dish for 45 min at 4°C in monomer solution containing 1× PBS, 2 M NaCl, 2.5% (wt/wt) acrylamide (catalog no. A4058, Sigma-Aldrich), 0.15% (wt/wt) N,N′-methylenebisacrylamide (catalog no. M7279, Sigma-Aldrich), and 8.625% (wt/wt) sodium acrylate (catalog no. 408220, Sigma-Aldrich). Slices were placed in a custom-made gel chamber (8, 10), and gelatin solution (~100 µl) was added. The gelatin solution, consisting of monomer solution (188 µl), accelerator solution [4 µl, 0.2% from 10% stock N,N,N′,N′-tetramethylethylenediamine (catalog no. T7024, Sigma Aldrich)], and inhibitory solution [4 µl, 0.01% from 0.5% stock solution (4-hydroxy-TEMPO, catalog no. 176141, Sigma-Aldrich)], was mixed on ice in low light. The initiator solution [4 µl, ammonium persulfate, 0.2% (wt/wt) from 10% stock solution (catalog no. A3678, Sigma-Aldrich)] was added immediately before the gelatin solution to prevent premature gelation. Slices in the gelation chamber were incubated at 37°C for 2.5 h. Subsequently, excess gel was removed from around the samples, and the tissue was placed in a six-well dish containing digestion buffer [1× Tris base-acetic acid-EDTA buffer, 0.5% Triton X-100, 0.8 M guanidine hydrochloride (catalog no. G3272, Sigma-Aldrich), and proteinase K (final concentration 8 U/ml; catalog no. P8107S, New England Biolabs)] overnight (12 h) at 22°C. Tissue was then transferred to another six-well dish and washed with excess (1–2 ml) deionized water (3–5 times for 15 min each). The amount of water was gradually increased with each wash to prevent folding of the tissue. Slices were removed from the dish and placed on a slide, a coverslip was mounted using Vectashield (catalog no. H-1000, Vector Laboratories), and the slide was sealed with nail polish.

Images were acquired using a Nikon A1 HD25 confocal microscope mounted on a Ti2 inverted microscope controlled by NIS-Elements AR software. Immunoreactivity was visualized at 488- and 561-nm wavelengths via a Plan Apo λ ×4 (2, 048 × 2,048) or Apo ×40 WI λS digital interference contrast N2 (1,024 × 1,024) objective. Images (×40 magnification) were acquired through the z direction in 0.2-μm steps and deconvolved via NIS-Elements AR software. Catalase immunoreactivity that colocalized with NeuN immunoreactivity was quantified in a 200 × 200-μm region of the PVN from a maximal-intensity projection of ×40 images and presented as the percentage of NeuN neurons that express catalase (i.e., catalase + NeuN/NeuN ×100). FIJI ImageJ (RRID: SCR_002285) was used to quantify immunoreactivity, adjust brightness and contrast only, and calculate plot profiles.

PVN Slice Generation, Electrophysiology, and Protocols

PVN slices were prepared from rats that were anesthetized with 5% isoflurane and decapitated. The forebrain was removed and placed in ice-cold N-methyl-d-glucamine (NMDG)-artificial cerebrospinal fluid (aCSF) cutting solution (in mM: 93 NMDG, 93 HCl, 2.5 KCl, 1.2 NaH2PO4, 10 MgSO4, 30 NaHCO3, 20 HEPES, 25 d-glucose, 5 l-ascorbic acid, 2 thiourea, 3 sodium pyruvate, and 0.5 CaCl2) aerated with 95% O2-5% CO2 (pH 7.4, 300–310 mosM). Slices (~290 µm) were cut with a vibrating microtome (model VT 1000S, Leica), allowed to rest in NMDG solution for 12 min at 32°C, and placed in recording aCSF (see below) until used in experiments. The submerged sections were secured with nylon threads attached to a stainless steel harp and superfused at a flow rate of 3–4 ml/min with standard recording aCSF (in mM: 124 NaCl, 3 KCl, 1.2 NaH2PO4, 1.2 MgSO4, 25 NaHCO3, 11 d-glucose, and 2 CaCl2) saturated with 95% O2-5% CO2 (pH 7.4, 300 mosM) at 31–33°C. Neurons were visualized using a fixed-stage upright microscope (model BX51WI, Olympus) with ×40 magnification, differential interface contrast, and an infrared-sensitive camera. Glass recording electrodes (3.5–5.0 MΩ; 8250, King Precision Glass, Claremont, CA) were filled with a solution containing (in mM) 10 NaCl, 130 K-gluconate, 11 EGTA, 1 CaCl2, 10 HEPES, 1 MgCl2, 2 MgATP, and 0.2 NaGTP (pH 7.3, 295–300 mosM). In some experiments, the filling solution also contained 0.5% Neurobiotin or 1 mg/ml Alexa Fluor 594 hydrazide (28). The pipette was guided using a piezoelectric micromanipulator (ThorLabs, Newton, NJ). Data were recorded using an amplifier (MultiClamp 700B, Molecular Devices), filtered at 2 kHz, and sampled at 10 kHz using Clampex v9 and v10 (Molecular Devices, San Jose, CA).

After recording and cell filling were completed, slices were fixed with 4% PFA for 20 min and then placed in PBS. Neurobiotin-containing cells were exposed to streptavidin-Cy3 and then washed with PBS. The tissue was subsequently cleared with Visikol HISTO solution according to the manufacturer’s instructions and mounted on an indented slide, and a coverslip was applied. Three-dimensional images were acquired using a conventional fluorescence microscope equipped with an ORCA-ER camera (Hamamatsu) and spinning-disk confocal unit (Olympus). Appropriate filter sets and excitation wavelengths were used to visualize the fluorophores. z Stacks (0.5-μm separation) were taken and neurons were traced using the Simple Neurite Tracer plugin of FIJI ImageJ (34).

Voltage-clamp protocols.

Neurons were clamped at −60 mV, near the Cl− reversal potential for GABAA receptors under our recording conditions; thus the recorded synaptic currents are likely glutamatergic excitatory postsynaptic currents [spontaneous EPSCs (sEPSCs)]. sEPSCs were monitored in gap-free mode. Holding currents (Ihold) and the root mean square (RMS) of the current were measured during periods that lack synaptic currents. EPSCs, Ihold, and RMS were measured during the last 30 s of a given treatment (37). IK were evoked between −100 and +40 mV (10 mV per step, 125 ms) from a −90-mV prepulse and normalized to the cell’s initial capacitance (pA/pF) (27). Current amplitude was measured for 2 ms at the end of the 125-ms pulse (late) and immediately after the initial capacitive current (early). For fitting of IK to the Boltzmann equation, the current was normalized to its maximum current (I/Imax).

Current-clamp protocols.

Resting Vm and spontaneous AP discharge (APd) were measured under current-clamp conditions with zero Ihold (I = 0). Spontaneous APd (Hz) was measured during 1-min epochs. Current-evoked APs were initiated through ramp (−20 to 120 pA, 1-s ramp) or step (−20 to 120 pA, 20-pA steps, 100 ms) depolarization from the cell’s resting potential. Phase plane plots (time derivative of voltage vs. voltage) were generated to identify AP threshold, identified as the point at which slope exceeded 10 mV/ms, and to examine the rate of voltage change due to ionic currents (5) in aCSF and H2O2. AP amplitude, afterhyperpolarization, half-width, rise and decay time, and slope were determined from initial threshold values (29).

PVN Neuron Dissociation, Electrophysiology, and Protocols

Neurons were dissociated to examine the direct effect of H2O2 on recorded neurons, independent of pharmacological synaptic blockers or diffusion of H2O2 through antioxidant-containing tissue layers in the slice. Similar to our previous studies in the nTS (41), the forebrain of isoflurane-anesthetized (VetOne, Boise, ID) rats was quickly removed and chilled in ice-cold low-Ca2+/high-Mg2+ aCSF (in mM: 124 NaCl, 3 KCl, 1.2 NaH2PO4, 1.2 MgSO4, 25 NaHCO3, 11 d-glucose, 1 CaCl2, and 2 MgCl2) saturated with 95% O2-5% CO2 (pH 7.4, ~300 mosM). Horizontal slices (~400 µm) containing the PVN were cut using a vibrating microtome (model VT 1000S, Leica). Subsequently, the PVN was excised and incubated in Ca2+/Mg2+-free Hanks’ balanced salt solution (Gibco, Big Cabin, OK) with 10 U/ml papain (Worthington, Lakewood, NJ). After 30 min in the incubator (model MCO-18AIC, Sanyo) at 37°C and 5% CO2, the papain-Hanks’ balanced salt solution was replaced with ~1 ml of high-glucose Dulbecco's modified Eagle’s medium (Gibco) containing 10% horse serum (Gibco) to stop enzymatic activity. The tissue was then triturated using progressively smaller fire-polished glass pipettes until cells were visibly dispersed. Cells were concentrated using a centrifuge (700 rpm for 5 min; model 5804 R, Eppendorf), and the supernatant was replaced with ~0.3 ml of Neurobasal-A (Gibco) + B-27 supplement (2%; Gibco), GlutaMAX (1%; Gibco), insulin-transferrin (1%; Gibco), and penicillin-streptomycin (1%; Gibco). PVN cells were plated on poly-d-lysine (100 µg/ml)-coated glass coverslips and allowed to settle and attach for 2 h in the incubator. Subsequently, dishes were flooded with Neurobasal-A as culture medium.

Recordings were made within 3 days of isolation. Neurons were visualized on an inverted microscope (model IMT2, Olympus) with phase optics. Recording electrodes were pulled from borosilicate glass (catalog no. B150-86-7.5, Sutter Instruments) and positioned using a hydraulic micromanipulator (model MX6600R, Siskiyou). Neurons were bathed in a normal physiological external solution (i.e., aCSF) consisting of (in mM) 137 NaCl, 3 KCl, 10 d-glucose, 10 HEPES, 2 CaCl2, and 1 MgCl2 (pH adjusted to 7.4 with NaOH, ~315 mosM). To record Vm and IK in whole cell mode, electrodes (3–6 MΩ) were filled with a solution containing (in mM) 10 NaCl, 130 K-gluconate, 11 EGTA, 1 CaCl2, 10 HEPES, 1 MgCl2, 2 MgATP, and 0.2 NaGTP (pH 7.3, 295–300 mosM). In some cells, to block IK, electrodes contained 124 mM CsCl, 11 mM EGTA, 1 mM CaCl2, 10 mM HEPES, 1 mM MgCl2, 4 mM MgATP, 0.2 mM NaGTP, 20 mM phosphocreatine, 0.02 mM leupeptin, and 35 U/ml phosphocreatine kinase (pH adjusted to 7.2 with CsOH, ~300 mosM). Signals were filtered at 2 kHz and acquired at 10 kHz using an Axopatch 200B amplifier and Clampex 10 software controlled by a Digidata 1320 interface (Molecular Devices, Sunnyvale, CA).

Voltage-clamp protocols.

Neurons were initially clamped at −60 mV. Because H2O2 has been shown to target inward-rectifier and twin-pore K+ (KIR and K2P, respectively) channels in other tissue (23, 31), membrane voltage ramps (−110 to −40 mV, 1 s) were used to identify possible changes in K+ conductance through these channels (2, 4, 42). Slope conductance (Gslope) was measured on ramps between −110 and −50 mV. The subtracted H2O2-sensitive currents were identified by subtracting the current obtained during H2O2 exposure from the current obtained immediately prior in aCSF alone (the control). In addition, we evoked IK between −110 and +40 mV (10 mV per step, 140 ms). Currents (pA/pF) were measured and fit to the Boltzmann equation, as described above for slice experiments.

Current-clamp protocols.

Vm and spontaneous APd were measured under the I = 0 condition. Evoked APd was elicited by step depolarization (−5 to 45 pA, +10- to 20-pA steps, 100-ms duration). To characterize PVN phenotype, a hyperpolarization to −100 mV preceded depolarization in some cells (35). The current varied depending on the excitability of the particular cell. AP properties were recorded and measured as described above for our slice protocols.

Drugs

In both slice and isolated cell protocols, physiologically relevant concentrations of H2O2 [100–400 µM (41, 42, 52)] were freshly prepared from stock (American Chemical Society certified 30%, Fisher Scientific) and added to the recording solution immediately before use. In brain slices, we recorded neurons one to two cell layers deep and perfused H2O2 for 5 min to allow penetration into the slice and to compensate for dead space in the perfusion tubing. H2O2 application for isolated cells matched that for the slice. BaCl2 (Ba2+, 100 μM, a general K+ channel blocker) or glibenclamide [3 μM, an ATP-sensitive K+ (KATP) channel blocker] was individually added to the extracellular solution for 5 min before exposure to H2O2 in subsets of experiments. Diazoxide (100 μM, a KATP channel activator) was also added before H2O2 in some experiments. CsCl (Cs+, as described above) or catalase (500 U, a general antioxidant) was included in the intracellular solution. Parameters during application of antagonists were considered the baseline for each H2O2 application. All drugs were obtained from Tocris (Bio-Techne, Minneapolis, MN) or Sigma-Aldrich (St. Louis, MO).

Statistics

Data were analyzed by Clampfit (Molecular Devices), OriginPro (Origin Laboratories, Northampton, MA), and Microsoft Excel. Statistical analysis was performed with GraphPad Prism 8.0 (GraphPad Software, La Jolla, CA), Excel with the Real Statistics Using Excel add-in (http://www.real-statistics.com/), or OriginPro software. Data were tested for normality by the Shapiro-Wilk test and subsequently by Student’s paired t-test or 1- or 2-way repeated-measures (RM) ANOVA where appropriate. Fisher’s least significant difference post hoc test identified individual differences. Fitting to the Boltzmann equation was done in OriginPro. All data are presented as means ± SE. Results were considered significantly different at P < 0.05.

RESULTS

H2O2 Hyperpolarizes PVN Neurons and Reduces Synaptic Currents

Recordings targeted neurons in the dorsal and medial parvocellular subnuclei of the PVN. Overall, PVN neurons had a Vm of −53.0 ± 2.0 mV, capacitance of 28.1 ± 1.9 pF, and initial input resistance of 738.7 ± 93.2 MΩ (n = 29). These properties are consistent with previous reports of PVN parvocellular neurons (9, 56). In addition, neurons were generally classified as type II based on their lack of large A-type transient IK, transient low-threshold spikes in some neurons, and lack of bursting discharge (35, 56, 61). Representative images of slice, recording, and neuronal morphology are shown in Fig. 1, A and B.

Fig. 1.

Fig. 1.H2O2 hyperpolarizes paraventricular nucleus (PVN) neurons. A: an in vitro PVN slice with the recording electrode on the neuron (inset). F, fornix; OT, optic tract; 3V, third ventricle. Dashed line represents PVN region; smaller inset shows area of recording. B: morphology of 4 neurons reconstructed following recording. Scale bars = 20 μm. C: representative trace of spontaneous synaptic activity and holding current in artificial cerebrospinal fluid (aCSF) and H2O2. Note outward current in response to H2O2. Cell was voltage-clamped at −60 mV. D: group data showing holding current (Ihold) in aCSF, H2O2, and wash. Ihold was significantly greater in H2O2 than aCSF (baseline). E: H2O2 did not alter root mean square (RMS) noise. F: membrane potential (Vm), measured in current (I) clamp (I = 0), reversibly hyperpolarized following H2O2 application. *P < 0.05 vs. aCSF (by 1-way repeated-measures ANOVA with Fisher’s least significant difference test).

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In PVN slices, bath application of H2O2 altered Vm and synaptic neurotransmission. As shown in Fig. 1C, H2O2 (150 μM, 5 min) induced an outward hyperpolarizing current. The shift in Ihold is quantified in Fig. 1D and demonstrates a significant reversible outward Ihold following H2O2 (−17.8 ± 5.4 and +28.5 ± 15.4 pA in aCSF and H2O2, respectively, n = 23, P = 0.006 vs. aCSF by 1-way RM ANOVA). Of all neurons studied, the majority (18 of 23) increased Ihold >20%. This hyperpolarizing current by H2O2 occurred in the absence of alterations in current RMS noise (Fig. 1E; n = 23, P = 0.915 vs. aCSF by 1-way RM ANOVA) and was also observed as Vm hyperpolarization (Fig. 1F). Vm hyperpolarized to −70.8 ± 4.2 mV from an initial level of −53.4 ± 2.7 mV (n = 19, P = 0.0001 vs. aCSF by 1-way RM ANOVA). A similar H2O2 effect was observed with 300 μM H2O2 (Ihold: −3.9 ± 6.2 and +38.9 ± 16.7 pA in aCSF and H2O2, respectively, P = 0.01 by paired t-test; RMS: 3.0 ± 0.6 and 3.7 ± 0.8 Hz in aCSF and H2O2, respectively, P = 0.04 by paired t-test, n = 6), yet its response was not reversible.

In addition to membrane hyperpolarization, H2O2 reduced sEPSCs (Fig. 2A). As shown in Fig. 2A and quantified in Fig. 2, B and C, H2O2 did not consistently alter sEPSC amplitude (14.2 ± 1.1 and 12.7 ± 1.4 pA in aCSF and H2O2, respectively, P = 0.098 vs. aCSF by 1-way RM ANOVA) but significantly attenuated sEPSC frequency (8.1 ± 1.2 and 5.2 ± 0.9 Hz in aCSF and H2O2, respectively, n = 23, P = 0.0008 vs. aCSF by 1-way RM ANOVA). During aCSF and H2O2 application, sEPSC rise time from 10 to 90% of amplitude was comparable (1.42 ± 0.09 and 1.58 ± 0.13 ms, P = 0.22 by 1-way RM ANOVA) as was decay time from 90 to 10% of amplitude (3.70 ± 0.18 and 3.94 ± 0.49 ms, P = 0.61 by 1-way RM ANOVA). A similar H2O2 effect on sEPSC frequency was observed with 300 μM H2O2 (9.0 ± 2.0 and 5.6 ± 2.2 Hz in aCSF and H2O2, respectively, n = 6, P = 0.08 by paired t-test).

Fig. 2.

Fig. 2.Exogenous H2O2 decreases synaptic currents in paraventricular nucleus (PVN) neurons. A: zoomed traces of spontaneous synaptic activity from Fig. 1 showing reversible H2O2-induced decrease in spontaneous excitatory postsynaptic current (sEPSC) frequency. Cells were voltage-clamped at −60 mV. Note change in holding current (Ihold). B and C: group data showing sEPSC amplitude (amp) and frequency (freq) in artificial cerebrospinal fluid (aCSF), H2O2, and wash. Note decrease in frequency, but no change in amplitude, with H2O2. *P < 0.05 vs. aCSF (by 1-way repeated-measures ANOVA with Fisher’s least significant difference test).

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APd and AP Intrinsic Properties After H2O2

Consistent with H2O2-induced hyperpolarization of Vm, there was also an attenuating effect on APd (Fig. 3A). Under aCSF baseline, 44% (7 of 16) of PVN neurons fired spontaneously; after H2O2 application, 19% (3 of 16) of PVN neurons fired spontaneously (P = 0.25 by Fisher’s exact test). In those spontaneously active neurons, APd frequency was 2.1 ± 0.9 Hz at aCSF baseline and 0.05 ± 0.03 Hz following H2O2 application (P = 0.07 by paired t-test; Fig. 3B).

Fig. 3.

Fig. 3.H2O2-induced hyperpolarization reduces discharge and alters action potential (AP) properties. A: membrane potential in artificial cerebrospinal fluid (aCSF) and H2O2; H2O2 hyperpolarized membrane potential and reduced spontaneous discharge. Membrane potential is shown adjacent to traces. B: mean decrease in AP discharge (APd) by H2O2. One cell did not fire spontaneously during aCSF control but did fire during H2O2 treatment. Spont, spontaneous. C: injection of ramp current (inset) induced discharge, and H2O2 reduced this discharge and AP overshoot. THR, threshold. D: AP phase plane plots of the first evoked event of data in C. Note reduced upward slope and overshoot. Mem pot, membrane potential. Full parameters are presented in Table 1.

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We subsequently utilized depolarizing current ramps to further examine AP properties. As shown in Fig. 3C, current-induced membrane depolarization evoked one or more APs at baseline aCSF following H2O2 application. The H2O2-induced hyperpolarization eliminated discharge in 6 of the 14 neurons examined with ramp depolarization; in the remaining 8 neurons, H2O2 decreased the overall number of APs evoked during the ramp (12.1 ± 2.9 and 2.6 ± 1.5 in aCSF and H2O2, respectively, P < 0.05 by paired t-test). Phase plane plots examined the rate of Vm change per millisecond versus Vm (mV/ms vs. mV) of the first evoked AP. H2O2 significantly decreased AP overshoot (Fig. 3D), and nonsignificant decreases were seen in overall amplitude, decay slope, and rheobase. AP properties are described in Table 1.

Table 1. Action potential characteristics in brain slice preparation

aCSFH2O2 (n = 7)P
Peak amplitude from THR, mV62.33 ± 2.6355.36 ± 3.820.102
Overshoot from 0 mV, mV19.47 ± 3.369.45 ± 2.870.009*
AHP amplitude from THR, mV−17.05 ± 2.32−18.21 ± 2.180.516
Half-width, ms1.02 ± 0.111.14 ± 0.190.416
Threshold, mV−42.86 ± 3.12−45.91 ± 1.410.370
Rise time from 10 to 90%, ms0.62 ± 0.090.68 ± 0.110.552
Rise slope from 10 to 90%, mV/ms86.04 ± 11.8575.75 ± 12.210.234
Decay time from 90 to 10%, ms0.81 ± 0.100.90 ± 0.160.470
Decay slope from 90 to 10%, mV/ms−69.86 ± 12.85−60.97 ± 11.460.178
Rheobase44.29 ± 9.85178.786 ± 21.10.107

Values are means ± SE; n equals number of cells. aCSF, artificial cerebrospinal fluid; THR, threshold; AHP, afterhyperpolarization. *Statistically significant.

H2O2 Produces Biphasic Responses in Voltage-Gated IK

Outward IK were investigated to examine their role in H2O2 alterations on neuronal properties. Step depolarization from −100 to +40 mV from a brief prepulse of −90 mV elicited IK in PVN neurons, and H2O2 reduced IK (Fig. 4A). The resultant (H2O2-sensitive) current was obtained by subtraction of the H2O2 current from the aCSF current. The current-voltage relationship in aCSF and H2O2 is shown in Fig. 4, B and C. The early- and late-current amplitudes were similar, with H2O2 decreasing both. The H2O2-sensitive current occurred at voltages more positive than −30 to −20 mV (Fig. 4, B and C). H2O2 moderately increased IK in the late response at voltages near the resting potential between −60 and −50 mV, up to −30 mV (Fig. 4C, inset). The current at −50 mV increased from 0.09 ± 0.3 pA/pF in aCSF to 2.4 ± 1.5 pA/pF in H2O2 (P = 0.02 by 2-way RM ANOVA). Early and late currents at a given voltage were compared with their maximal currents (I/Imax) in aCSF and H2O2 and were fit with a Boltzmann equation. H2O2 decreased the late-current half-activation (6.5 ± 0.8 and 0.5 ± 0.9 mV in aCSF and H2O2, respectively, P = 0.001 by paired t-test) and increased its slope (16.7 ± 0.52 vs. 20.9 ± 1.2, P = 0.002 by paired t-test). Early-current half-activation also decreased with H2O2 (−0.4 ± 1.5 and −3.6 ± 1.6 mV in aCSF and H2O2, respectively, P = 0.01 by paired t-test) but had no effect on its slope (17.1 ± 0.35 and 17.7 ± 0.8, P = 0.44 by paired t-test).

Fig. 4.

Fig. 4.Biphasic effects of H2O2 on K+ currents. A: depolarization evoked K+ currents (IK) during artificial cerebrospinal fluid (aCSF) and H2O2 treatment. H2O2 decreased outward currents at higher voltage commands. H2O2-sensitive current is current in aCSF − current in H2O2. Inset: voltage command. Peak inward Na+ currents are truncated for clarity. B: early current immediately following pipette capacitive current (B in panel A) during aCSF and H2O2 treatment. C: late steady-state current at the end of the current trace (C in panel A) during aCSF and H2O2 treatment. H2O2-sen, H2O2-sensitive current. Note H2O2-induced decrease (*P < 0.05) in IK at voltages >0 mV. At lower voltages (−50 to −30 mV), H2O2 increased IK (^P < 0.05). Inset: current-voltage relationship at lower voltages. Significance was determined by 2-way repeated-measures ANOVA with Fisher’s least significant difference test. D: H2O2-induced decrease in IK tends to reduce action potential (AP) interevent interval (IEI) in response to current injection. Because of hyperpolarization by H2O2, more current was needed to evoke similar AP discharge (APd). E: quantification of IEI in aCSF and H2O2.

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To examine if the H2O2-induced decrease in IK alters neuronal discharge, we depolarized PVN neurons via current steps (100 ms, 20-pA steps). As described above, H2O2 hyperpolarized PVN neurons, yet upon further depolarization, multiple APs were evoked. The interevent interval (IEI) of two consecutive minimally evoked APs was determined as an indicator of IK influence on firing rate in aCSF and H2O2. As shown in the representative traces (Fig. 4D), H2O2 decreased the AP IEI, which is quantified in Fig. 4E (P = 0.07, aCSF vs. H2O2, n = 8).

Together, these data demonstrate that H2O2 hyperpolarizes PVN neurons and reduces excitatory synaptic currents. H2O2 also increases IK at voltages near the resting potential yet decreases currents at more depolarized potentials to alter APd.

PVN Neurons in Isolation Have Intrinsic Properties Similar to Those in the Slice

PVN neurons were mechanically isolated to further identify the cellular mechanism(s) of the action of H2O2 and examine the direct action of H2O2 on somas. Neurons were dissociated and recorded within 2.5 ± 0.2 days of isolation. Across all cells (n = 63), PVN neurons had a Vm of −50.9 ± 1.2 mV, capacitance of 18.8 ± 0.92 pF, and initial input resistance of 1,525.6 ± 90.9 MΩ. Vm in isolation was comparable to that observed in the slice (P = 0.384 by unpaired t-test). However, capacitance was significantly less (P < 0.001 by unpaired t-test), while input resistance was greater (P < 0.001 by unpaired t-test), in isolated cells. Such differences are not unexpected, as cells in isolation lack many of the processes observed in their native network and, thus, are electrically compact. Similar to our slice data, neurons were generally classified as type II based on their lack of large A-type IK and the existence of transient low-threshold depolarization in some cells. A representative example of our isolated cells and their characterization are shown in Fig. 5, A–C. Figure 5B illustrates Vm in response to hyperpolarization to −100 mV followed by depolarization to −35 mV (35). The representative cell exhibited APd followed by subsequent depolarization and lack of large A-type IK (Fig. 5C).

Fig. 5.

Fig. 5.H2O2 effects on neurons independent of neuronal network. A: representative image of isolated paraventricular nucleus (PVN) neurons with attached recording pipette. B: membrane potential in response to brief hyperpolarization to −100 mV followed by depolarization to −35 mV illustrating resulting action potential discharge and subsequent depolarizing potential characterizing parvocellular PVN neurons. Inset: current protocol. C: K+ current (IK) in response to depolarizing voltage steps with a −100-mV prepulse (top) and resultant currents evoked by subtraction of IK without a preceding prepulse from IK with a −100-mV prepulse (bottom). Note the lack of large A-type IK. D: H2O2 (200 μM)-induced hyperpolarization of membrane potential in isolated PVN neurons. Initial resting potential of the cell is shown at the beginning of the trace. E and F: group data of PVN neurons exposed to H2O2 and vehicle. H2O2 hyperpolarized membrane potential (mem pot) at minutes 3 and 5. Vehicle did not alter membrane potential. aCSF, artificial cerebrospinal fluid. *P < 0.05 vs. aCSF baseline (by 1-way repeated-measures ANOVA with Fisher’s least significant difference test). Cells were recorded at 0 holding current (I = 0).

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H2O2 Hyperpolarizes Neurons Independent of Surrounding Network

Figure 5D shows effects of H2O2 on Vm in isolated cells. Compared with aCSF baseline, H2O2 (5 min, 200 μM) hyperpolarized Vm and decreased APd. Hyperpolarization occurred over 5 min of H2O2 application and was not readily reversible after 8 min of aCSF wash. Vm data are quantified in Fig. 5E and demonstrate a significant hyperpolarization within 3 min of H2O2 application (P = 0.008 by 1-way RM ANOVA) that persisted after 5 min (−58.5 ± 2.1 and −66.1 ± 2.6 mV in aCSF and H2O2 at 5 min, respectively, n = 17, P = 0.002 by 1-way RM ANOVA). By contrast, as shown in Fig. 5F, aCSF alone (i.e., vehicle) did not alter Vm (−62.5 ± 2.8 and −64.8 ± 3.6 mV in aCSF and aCSF at 5 min, respectively, n = 7, P = 0.260 by 1-way RM ANOVA), which remained unchanged following an additional 8 min of aCSF wash. We also examined Vm alterations by 100 and 400 μM H2O2. Vm was also hyperpolarized with H2O2 at 100 μM (−60.0 ± 3.8 and −62.1 ± 3.7 mV in aCSF and H2O2 at 5 min, respectively, n = 6, P = 0.01 by paired t-test) and 400 μM (−48.5 ± 4.3 and −51.8 ± 5.2 mV in aCSF and H2O2 at 5 min, respectively, n = 6, P = 0.02 by paired t-test). Because the magnitude of response was greatest at 200 μM (data not shown), this concentration was used for the remainder of the isolated cell study.

H2O2 Reduces AP Overshoot and Slows Decay in Isolated PVN Neurons

In isolation, few (4 of 10) neurons spontaneously fired APs. Of those that did exhibit spontaneous discharge, the number of APs during 5 min at aCSF baseline and 5 min in H2O2 was 7.8 ± 5.5 and 0.75 ± 0.75, respectively (n = 4, P = 0.3 by paired t-test). We subsequently utilized depolarizing current steps (up to 55 pA, 100 ms) to examine AP properties. Cells were maintained at −60 mV during aCSF control and in H2O2 with a bias current to examine properties at similar channel activation profiles. As shown in Fig. 6A, current-induced membrane depolarization typically evoked one AP over the 100-ms current step during aCSF baseline and in H2O2, which was not different between groups (1.1 ± 0.1 and 1.0 ± 0.1 APs/100 ms in aCSF and H2O2, respectively, P = 0.33 by paired t-test). aCSF alone (i.e., vehicle) also did not alter the number of APs over time (1.0 ± 0.0 and 1.0 ± 0.0 APs/100 ms in aCSF and 5 min of aCSF, respectively). Figure 6B, a phase plane plot of the APs in Fig. 6A, was used, in part, to determine AP properties. As illustrated in the phase plane plot and similar to slice APs, H2O2 decreased AP overshoot and tended to decrease overall amplitude and decay slope. Full AP properties for aCSF control and following application of either H2O2 or aCSF vehicle are shown in Table 2.

Fig. 6.

Fig. 6.Action potential (AP) characteristics in H2O2. A: representative current-evoked APs in artificial cerebrospinal fluid (aCSF) and H2O2. Cells were maintained at −60 mV with bias current. H2O2 reduced AP amplitude. B: phase plane plot of APs in A demonstrating H2O2-induced reduction in rise and decay slope, as well as amplitude. Mem pot, membrane potential. Full AP details are presented in Table 2.

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Table 2. Action potential characteristics in isolation

aCSFH2O2 (n = 9)PaCSFVehicle (n = 5)PaCSFH2O2 + Cat (n = 6)P
Peak amplitude from THR, mV74.31 ± 6.6768.29 ± 9.230.11563.14 ± 10.2860.21 ± 13.280.58764.53 ± 9.5164.80 ± 10.800.893
Overshoot from 0 mV, mV48.24 ± 6.0541.26 ± 7.390.023*35.75 ± 9.0133.79 ± 10.520.63937.84 ± 7.3636.26 ± 7.090.372
AHP amplitude from THR, mV−26.41 ± 2.52−26.54 ± 2.710.891−22.39 ± 3.21−23.64 ± 3.680.212−26.30 ± 2.06−25.82 ± 2.750.540
Half-width, ms1.73 ± 0.111.88 ± 0.170.2612.35 ± 0.552.34 ± 0.520.8261.46 ± 0.261.48 ± 0.250.534
THR, mV−26.08 ± 1.14−27.03 ± 2.930.694−27.40 ± 3.39−26.42 ± 4.420.490−26.69 ± 2.78−28.53 ± 4.100.347
Rise time from 10 to 90%, ms0.96 ± 0.071.02 ± 0.110.2731.07 ± 0.191.08 ± 0.170.7050.89 ± 0.080.91 ± 0.090.161
Rise slope from 10 to 90%, mV/ms69.73 ± 10.3663.42 ± 12.200.11460.96 ± 19.1257.11 ± 19.690.34663.71 ± 15.5867.08 ± 19.060.439
Decay time from 90 to 10%, ms1.40 ± 0.111.58 ± 0.160.3432.25 ± 0.512.04 ± 0.430.1331.10 ± 0.321.10 ± 0.280.896
Decay slope from 90 to 10%, mV/ms−44.49 ± 4.80−35.51 ± 4.210.086−35.89 ± 15.49−34.25 ± 13.890.571−61.77 ± 13.88−58.51 ± 13.630.170
Rheobase19.44 ± 3.6724.44 ± 5.160.13529.00 ± 7.4833 ± 9.170.37419.17 ± 4.5520 ± 4.830.771

Values are means ± SE; n equals number of cells. aCSF, artificial cerebrospinal fluid; Cat, catalase; THR, threshold; AHP, afterhyperpolarization. *Statistically significant.

We previously demonstrated that the exogenous H2O2 antioxidant catalase in the recording pipette eliminated H2O2 effects in the nTS (42). In PVN neurons, H2O2 in the presence of exogenous intracellular catalase (500 U) did not alter the number of evoked APs during current-evoked depolarization (1.2 ± 0.2 and 1.0 ± 0.0 in aCSF and H2O2, respectively, P = 0.36 by paired t-test) or the properties of the first AP (Table 2). Notably, the reductions in peak amplitude, overshoot, and rise and decay slope by H2O2 alone were eliminated by addition of catalase.

H2O2-Induced Hyperpolarization Is Blocked by K+ Channel Blockers and Catalase

H2O2 hyperpolarizes neurons near their resting potential (Fig. 5). This may suggest activation by H2O2 by one or more channels at these potentials. K+ channels are known to set the resting potential and contribute to APd rate. Thus we examined the potential K+ channel mechanism(s) by which H2O2 may modulate Vm. In isolated cells, we applied H2O2 in the bath in the presence of an individual K+ channel blocker, including those that block KIR or K2P channels that are activated by H2O2. In addition, H2O2 was applied to dissociated cells, which were recorded with pipettes containing the intracellular K+ inhibitor Cs+ or the antioxidant catalase.

As shown in Fig. 7A, inclusion of catalase in the pipette eliminated the H2O2-induced hyperpolarization. Quantitative data showing the overall change in Vm in response to 5 min of H2O2 exposure compared with its baseline control, as well as the change in Vm in response to H2O2 in the presence of catalase or K+ channel blockers, are shown in Fig. 7B. As mentioned previously, H2O2 alone produced a significant hyperpolarization. In contrast, H2O2 did not affect membrane polarization when catalase was present in the pipette (P = 0.73 by paired t-test, n = 6), and this response was significantly less than the response to H2O2 alone (P = 0.047 by 1-way ANOVA; Fig. 7B). H2O2 in the presence of extracellular Ba2+ (100 μM, a general K+ channel blocker, n = 10) produced a varied response. Overall, Vm after H2O2 in Ba2+ was not different from Vm after Ba2+ alone (P = 0.36 by paired t-test); however, this response did not reach significance compared with the response to H2O2 alone (P = 0.12 by 1-way ANOVA; Fig. 7B). Similarly, in the presence of glibenclamide (3 μM, a KATP channel blocker, n = 11), H2O2 did not change Vm compared with glibenclamide alone (P = 0.20 by paired t-test), although the H2O2-induced change in Vm was not different from the change induced by H2O2 alone (P = 0.24 by 1-way ANOVA; Fig. 7B). The lack of hyperpolarization by H2O2 in the presence of glibenclamide may suggest a potential role for KATP channels. To further examine these channels, we activated KATP channels with diazoxide (100 μM), reasoning that if KATP channels contribute to H2O2-induced hyperpolarization, then their activation would mimic the response to H2O2 and occlude any further Vm change with additional H2O2. Diazoxide alone did not alter Vm (−60.8 ± 6.1 and −60.2 ± 6.5 mV in aCSF and diazoxide, respectively, P = 0.61 by paired t-test). H2O2 in the presence of diazoxide did not further alter Vm (P = 0.067 vs. diazoxide alone, n = 5), which was significantly reduced compared with Vm after application of H2O2 alone (P = 0.04 by 1-way ANOVA; Fig. 7B).

Fig. 7.

Fig. 7.H2O2-induced hyperpolarization is blocked by exogenous catalase (Cat) and K+ channel blockers. A: membrane potential (Vm) during exposure to artificial cerebrospinal fluid (aCSF) and H2O2 in the presence of exogenous catalase (500 U, in the pipette). Vm following H2O2 with catalase eliminated hyperpolarization. B: group data showing change in Vm with H2O2 alone and in the presence of blockers. Catalase blocked H2O2-induced hyperpolarization, as did Ba2+, glibenclamide (Glib), and diazoxide (Diaz). BSL, baseline; Cntl, control. *P < 0.05 vs. preceding aCSF or blocker alone (by paired t-test). The magnitude of change compared with H2O2 alone was significantly reduced by catalase and diazoxide. #P < 0.05 vs. H2O2 alone (by 1-way ANOVA with Fisher’s least significant difference test).

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H2O2 Does Not Alter KIR /K2P Currents but Enhances Voltage-Gated Currents at Negative Potentials

Previous work by our group and others showed that H2O2 modulates Vm via activation of KIR/K2P channels (42). Moreover, the reduction of H2O2-induced hyperpolarization by Ba2+ in Fig. 7 suggests contribution of one or more K+ channels, including KIR/K2P channels. To further examine the contribution of these channels to the H2O2 responses, we compared currents in response to voltage ramps (−110 to −40 mV, 1 s) within the conducting range of KIR and K2P channels (4, 15) at baseline and with H2O2 perfusion in isolated cells. Before application of the voltage ramp, H2O2 produced a general positive (outward) shift in Ihold, consistent with hyperpolarization of resting membrane potential (Fig. 8A). The increase in Ihold after H2O2 (P = 0.05, n = 9) is illustrated in Fig. 8B. Moreover, compared with the prior control period (paired t-test), there was no increase in Ihold after H2O2 in the presence of intracellular catalase (n = 6, P = 0.84) as well as extracellular Ba2+ (n = 10, P = 0.41), glibenclamide (n = 11, P = 0.72), and diazoxide (n = 6, P = 0.62). Across the treatments, the change in Ihold compared with the change induced by H2O2 alone was significantly reduced by catalase and diazoxide, with a near-significant elimination of Ihold with glibenclamide (P = 0.06 by 1-way ANOVA; Fig. 8B). Replacement of Cs+ with intracellular K-gluconate to block K+ channels also eliminated the H2O2-induced changes in Ihold (H2O2 + Cs+ vs. Cs+ alone, P = 0.55 by paired t-test, n = 9), which was reduced compared with H2O2 alone (Fig. 8B).

Fig. 8.

Fig. 8.H2O2 increases K+ currents (IK) at voltages near resting potential. A: voltage ramps in the absence and presence of H2O2 (red). H2O2 induced an upward shift in holding current (Ihold; B) and did not alter slope conductance (Gslope; C) but enhanced transient outward IK (D-E). Subtracted current is shown below traces. B–E: quantification of parameters highlighted in A during exposure to H2O2 alone and following 1 or more blockers. B: magnitude of change in Ihold before voltage ramps was reduced compared with H2O2 alone by exogenous catalase (Cat), diazoxide (Diaz), and Cs+-based electrodes. C: magnitude of change in Gslope during voltage ramps, as measured between −110 and −50 mV, was not altered by H2O2 alone or with various treatments. D and E: H2O2 increases an outward IK, enhancing the peak amplitude (D) and area (E) between −50 and −40 mV. Outward IK was eliminated by catalase and K+ channel modifiers. For C–E, 1 = no change in H2O2 from its baseline (BSL). *P < 0.05, H2O2 vs. preceding individual baseline (by paired t-test); #P < 0.05, H2O2 in treatment vs. H2O2 only (by 1-way ANOVA with Fisher’s least significant difference test).

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A voltage ramp between −110 and −40 mV elicited a modest inward current in PVN neurons, and application of H2O2 did not alter this current (Fig. 8A). This is readily evident in the subtracted current, which shows, specifically, the lack of change in current with H2O2. Quantitatively, the voltage ramp during aCSF elicited an average Gslope of 0.79 ± 0.41 pA/mV (nS, measured between −110 and −50 mV, n = 9). Addition of H2O2 did not alter Gslope (0.70 ± 0.36 nS, n = 9, P = 0.26 by paired t-test), suggesting little activation of KIR /K2P channels. Figure 8C shows the magnitude of change in Gslope between the control period (aCSF or blocker alone) and the addition of H2O2 to the K+ blocker. The individual addition of K+ blockers with H2O2 did not alter Gslope (Fig. 8C; 1-way ANOVA).

Interestingly, upon ramp depolarization to approximately −50 mV, following H2O2 an evoked outward current that was not observed under aCSF conditions became readily evident (Fig. 8A). This current was reminiscent of the H2O2 increase in IK observed in slices (Fig. 4) at similar voltages. This H2O2-evoked current averaged 24.6 ± 10.5 pA at its peak, with an area of 1,549.4 ± 728.2 pA × ms (between −50 and −40 mV, P = 0.047, H2O2 vs. aCSF). Exogenous intracellular catalase eliminated the H2O2-evoked increase in peak outward current, as did Ba2+, glibenclamide, and diazoxide, as well as an intracellular Cs+-based solution, compared with their individual blocker alone (P < 0.05 by paired t-test). The magnitude of the increase in current in H2O2 from its baseline control (e.g., H2O2/aCSF) on peak amplitude and area and the attenuation by K+ channel blockers are shown in Fig. 8, D and E (by 1-way ANOVA with Fisher’s least significant difference test).

H2O2 Increases IK at Negative Potentials but Attenuates IK at Positive Potentials in Isolated Cells

As in our above slice preparation, we also monitored IK across several step voltages and the influence of H2O2. Consistent with results obtained in the slice (Fig. 4) and our isolated cell voltage ramps (Fig. 8), at negative potentials H2O2 increased IK (Fig. 9B, inset). H2O2 again decreased the early and late IK at the most depolarized potentials (Fig. 9). The reduced H2O2-sensitive current occurred at voltages more positive than −20 mV. I/Imax values during aCSF and H2O2 were fit with a Boltzmann equation. H2O2 decreased the late-current half-activation (1.3 ± 1.0 and −1.4 ± 0.7 mV with aCSF and H2O2, respectively, n = 5, P = 0.04 by paired t-test) and increased slope (14.8 ± 0.37 vs. 16.0 ± 0.32, P = 0.001 by paired t-test). Early-current half-activation did not change significantly (2.8 ± 0.9 and 0.8 ± 0.59 mV with aCSF and H2O2, respectively, P = 0.20 by paired t-test), and H2O2 had no effect on its slope (15.1 ± 0.52 and 15.4 ± 0.34, P = 0.44 by paired t-test).

Fig. 9.

Fig. 9.H2O2 has a biphasic influence on K+ current (IK) in isolated cells. A: early current immediately following the pipette capacitive current during exposure to artificial cerebrospinal fluid (aCSF) and H2O2 and the H2O2-sensitive (H2O2-sen) current. B: late steady-state current at the end of the current trace during aCSF and H2O2 exposure and the H2O2-sensitive current. Inset: current-voltage relationship at lower voltages. Note decrease (*P < 0.05) in current at voltages >0 mV; at lower voltages, H2O2 increased IK (^P < 0.05). Statistical significance was determined by 2-way repeated-measures ANOVA with Fisher’s least significant difference test.

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The Antioxidant Catalase Is Located Throughout the PVN

H2O2 modulated PVN neuronal activity, which was ablated by elevated catalase in the pipette. Consistent with our nTS studies (42), the antioxidant enzyme catalase was found throughout the PVN, including in NeuN-identified neurons (Fig. 10). Catalase was localized to 60 ± 2% (n = 3) of NeuN-identified neurons. Catalase immunoreactivity was located perinuclearly.

Fig. 10.

Fig. 10.Catalase is present within paraventricular nucleus (PVN) neurons. A: overview of catalase (green) antioxidant and NeuN (red) immunoreactivity across 3 representative levels of the PVN (outlined). Approximate bregma level (mm) of section is shown at bottom left of each image. Scale bars = 500 μm. 3V, 3rd ventricle; OT, optic tract; F, fornix. B: magnified image of catalase in yellow box shown in caudal PVN in A. Note catalase expression perinuclear to NeuN-positive neurons throughout the PVN. Scale bar = 50 μm. C: neuron from B demonstrating catalase surrounding nuclei. Scale bar = 10 μm. D: plot profile of catalase (green) and NeuN (red) from the dashed line in C. Catalase is not expressed in the plasma membrane but, rather, in the cytosol several micrometers from the membrane and distinct from nuclei.

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DISCUSSION

In the present study we examined the role and potential mechanisms of the ROS H2O2 in synaptic and neuronal activity in PVN neurons in the in vitro brain slice and cells in isolation. Our data demonstrate that H2O2 hyperpolarizes PVN neurons to limit spontaneous EPSC activity and APd. Hyperpolarization is due to activation of a K+ conductance near the resting Vm of the cell. These ROS-mediated effects can be eliminated by exogenous antioxidants, suggesting an intracellular site of action by H2O2. Together, these data suggest that H2O2 limits PVN activity via ion channel modulation.

The PVN is a critical nucleus in homeostasis and reflex responses of the autonomic nervous, endocrine, and respiratory systems. The PVN receives innervation from the nTS, which is activated via carotid body inputs during hypoxic stimulus (26). Short- and long-term exposure to intermittent hypoxia (IH) activates central neural substrates to augment breathing and sympathetic nervous system output to help maintain physiological function but also leads to overexcitation of these systems. Repeated IH and its episodic reoxygenation induce ROS production (44). H2O2 is a stable ROS that is increased in IH (20). The PVN has been suggested to play a prominent role in the chemoreflex responses (51), including the IH physiological (mal)adaptations (36, 54). For instance, IH induces an increase in NMDA receptors, a decrease in nitric oxide production (12), and an elevation of the transcription factor ΔFosB, suggesting elevated neuronal activity (54). IH increases PVN ROS, while central infusion of a ROS scavenger decreases IH-induced blood pressure, as well as PVN Fos and ROS production (33). Intravenous antioxidants decrease the activity of PVN neurons, in parallel with a reduction in sympathetic activity (62). Exogenous catalase in the PVN reduces elevated sympathetic nerve activity and blood pressure in response to cardiac sympathetic afferent reflex (65), renovascular hypertension (64), and salt-induced hypertension (66). These studies suggest H2O2 contributes to the sympathoexcitation in several conditions; however, not all studies agree. Cardoso et al. reported that increasing H2O2 via blockade of catalase blunts the sympathoexcitation induced by microinjection of glutamate (7), suggesting that H2O2 limits glutmatergic signaling. Our results showing that H2O2 hyperpolarizes neurons and reduces EPSCs in the PVN align with those of Cardoso et al. (7). It is tempting to speculate that increasing H2O2 via catalase inhibition would produce similar neuronal hyperpolarization and synaptic reduction in the slice, although perhaps to a lesser degree, where basal neuronal activity may contribute to endogenous ROS production.

We recorded neurons within or dissociated from the dorsal and medial subnuclei that encompass the parvocellular neurons that serve neuroendocrine and autonomic functions (58). The PVN region we examined receives dense inputs from the nTS (14), and acute hypoxia activates Fos in neurons within this region (11). In addition, neurons in this region project to the nTS and are activated (FOS) during hypoxia (49). Although we do not know the site to which our recorded neurons project, the neurochemical released, or whether they contribute to chemo-, baro-, or neuroendocrine responses to a hypoxic challenge and, thus, represent a heterogeneous cell population, the majority of neurons responded to H2O2. The differing magnitude of response to H2O2 may be due to the phenotype or the amount of antioxidants within each neuron. The latter interpretation is supported by the observation that catalase was not observed to the same degree in every PVN neuron. Nevertheless, H2O2 induced hyperpolarization in neurons recorded in the slice and in isolation. Each preparation has its limitations: the brain slice does not recapitulate the possible influence of active afferent projections on PVN neurons in vivo, while cells in isolation lack the synaptic contacts and dendritic processes that may contribute to the H2O2 response in vivo and in the slice. The redundancy of response, however, confirms that PVN neurons are intrinsically sensitive to ROS.

The H2O2 concentrations examined (<200 µM) encompass those expected during endogenous ROS production, considering the predicted 10- to 100-fold decrease with cell entry (48). Given the moderate and reversible response, our data suggest that H2O2 did not produce permanent oxidative damage but, rather, altered neuronal activity via one or more mechanisms. Hyperpolarization was observed following H2O2 application in neurons in the slice and in isolation, suggesting that H2O2 may influence somal activity in neurons independent of network synaptic activity. Membrane hyperpolarization was likely responsible for the decrease in APd, as evident by the increase in rheobase. These data are consistent with our previous work in the PVN-projecting nTS (42) and other central nuclei (3, 18, 39, 43, 53). The hyperpolarization-induced decrease in discharge was most evident in the slice, where ongoing synaptic input likely contributes to background Vm, bringing neurons closer to their threshold. Local excitatory circuits contribute to background glutamatergic signaling in the PVN (6). Thus the hyperpolarization of Vm by H2O2 may likely also reduce EPSC frequency. However, whether sEPSCs originate from these local glutamatergic circuits or other central nuclei that impinge on the recorded neuron is not known. While the Vm of the synaptic terminals adjacent to the neurons is also not known, if H2O2 hyperpolarizes their Vm similar to PVN somas, it is likely that H2O2 also reduces sEPSC frequency via reduction in terminal neurotransmitter release. EPSC amplitude, rise, and decay were not altered by H2O2, suggesting that glutamate receptor conductance and released quantal content were likely not altered. These results are consistent with reduced nTS neurotransmission (42) and hippocampal long-term potentiation (1), although increases in excitatory synaptic events have also been observed with small changes in amplitude (39). Whether H2O2 influences GABAerigc signaling in the PVN, as in other nuclei (39, 60), to potentially influence sympathetic activity (64–66) requires further examination.

While overall discharge is reduced, H2O2 had a modest influence on AP properties. In our brain slice recordings, where APs were evoked from the cell’s Vm, AP rheobase likely increased due to H2O2-induced membrane hyperpolarization. We previously observed a similar phenomenon in nTS slices (42). When bias current was applied to maintain Vm in dissociated cells, rheobase again increased, although not to the extent observed in the slice preparations. This would suggest an alteration in the ion channel responsible for evoking discharge. However, the most prominent effect of H2O2 was the decrease in AP overshoot and slowing of its decay, in accordance with reduction in the initial depolarizing inward voltage-dependent Na+ current and outward hyperpolarizing IK, respectively (5). Consistent with this interpretation, a reduction in voltage-dependent outward IK by H2O2 was observed at potentials more positive than −10 mV. This reduction was observed in both the initial and steady-state current, similar to that observed in the hippocampus (21) but not consistent with our previous studies in the nTS (41, 42). The decrease in IK, however, permitted a modest reduction in AP IEI, suggesting that H2O2 may increase discharge rate despite its effects on AP kinetics. While the molecular identity of these H2O2-modulated IKs in the PVN is unknown, several voltage-gated channels are expressed throughout the PVN (17), and future studies are required to fully delineate the channel subtype responsible.

The hyperpolarization by H2O2 occurred whether the cells were held at −60 mV or allowed to remain at their resting Vm, suggesting activation of one or more ion channels at these negative potentials. K2P and KIR channels, including the ATP-sensitive KATP channel, modulate Vm and overall excitability, are influenced by H2O2 (3, 42), and are expressed in the PVN (17). To determine the contribution of these channels, we examined Gslope between −100 and −50 mV, within the range of the initial Vm of our recorded cells. However, contrary to the nTS (42) and our initial expectations, H2O2 did not alter Gslope, suggesting that these channels did not appreciably contribute to currents within these voltages. Thus the H2O2-induced effects occur via distinct mechanisms in the nTS and PVN. Use of exogenous Ba2+ to block KIR/K2P channels (4) also did not alter conductance. Glibenclamide, to block KATP, and diazoxide, to activate KATP, which we predicted would mimic H2O2 and occlude any further effect if these channels contribute, also did not alter Gslope. Together, these studies suggest that K2P, KIR, and KATP channels are not directly responsible for the H2O2 hyperpolarization under our recording conditions. Regardless of the current effects, in the presence of Ba2+, glibenclamide, and diazoxide, H2O2 failed to hyperpolarize Vm. It is possible that these compounds influenced additional targets. For instance, Ba2+ is a prototypical blocker of a number of K2P/KIR channels but also inhibits a number of voltage-gated K+ channels (30, 59). Glibenclamide is an inhibitor of the KATP channel but also blocks voltage-gated channels (13, 47). Thus, Ba2+ and glibenclamide may have influenced K+ conductance through nontraditional pathways. Interestingly, our data show that H2O2 increases a voltage-gated IK at negative voltages near the resting potential, and Ba2+, glibenclamide, and internal Cs+ blocked the increase in this evoked current. These data suggest that Ba2+ and glibenclamide may influence voltage-gated channels and that such an increase in voltage-gated IK at negative resting potentials may be one mechanism by which H2O2 induces hyperpolarization.

The production and elimination of H2O2 occur through endogenous enzymes (57). Catalase is an endogenous antioxidant that has been located in peroxisomes and mitochondria (38, 45). In the nTS, when we elevated intracellular catalase via its placement in the recording pipette, the influence of H2O2 on Vm was eliminated (42). In the present study we confirm these results and demonstrate that catalase is located throughout the PVN in the majority of NeuN-identified neurons. It may be that catalase was below our level of detection in the minority of NeuN cells that lacked catalase, and perhaps other antioxidants are more greatly expressed in these cells. Regardless, exogenous catalase application via pipette placement eliminated the H2O2 hyperpolarization and ionic current changes. These results suggest that H2O2, when exogenously applied, is the primary ROS modulating this effect. It was reasoned that if H2O2 is broken down to a hydroxyl radical (·OH) to mediate membrane hyperpolarization, then exogenous catalase would not prevent this response; this did not occur. Together, these findings suggest that H2O2 likely alters neuronal intracellular, rather than extracellular, sites.

Perspectives and Significance

The present results suggest that H2O2 hyperpolarizes PVN Vm via modulation of K+ channels. This inhibition limits neuronal discharge, which may help explain the role of H2O2 in cardiorespiratory homeostasis and visceral reflexes. Future studies are needed to further determine the influence of this ROS on neuronal phenotypes and projections from the PVN that influence cardiovascular, respiratory, and endocrine function.

GRANTS

This work was funded by National Heart, Lung, and Blood Institute Grants RO1 HL-128454 and HL-098602 (to D. D. Kline).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

D.D.K. conceived and designed research; H.A.D., M.P.M., D.M., and D.D.K. performed experiments; H.A.D., M.P.M., and D.D.K. analyzed data; H.A.D., M.P.M., D.M., and D.D.K. edited and revised manuscript; H.A.D., M.P.M., D.M., and D.D.K. approved final version of manuscript; M.P.M. and D.D.K. interpreted results of experiments; D.D.K. prepared figures; D.D.K. drafted manuscript.

ACKNOWLEDGMENTS

We thank Dr. Eileen M. Hasser for helpful discussion and comments on the manuscript.

Present address of M. P. Matott, Dept. of Neuroscience, Cell Biology, and Physiology, Wright State University, Dayton, OH 45435.

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AUTHOR NOTES

  • * Co-first authors, H. A. Dantzler and M. P. Matott, contributed equally to this work.

  • Address for reprint requests and other correspondence: D. D. Kline, University of Missouri Dalton Cardiovascular Research Center, 134 Research Park Dr., Columbia, MO 65211 (e-mail: ).
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