Excessive Na⁺/H⁺ Exchange in Disruption of Dendritic Na⁺ and Ca²⁺ Homeostasis and Mitochondrial Dysfunction following in Vitro Ischemia

Abstract

Neuronal dendrites are vulnerable to injury under diverse pathological conditions. However, the underlying mechanisms for dendritic Na⁺ overload and the selective dendritic injury remain poorly understood. Our current study demonstrates that activation of NHE-1 (Na⁺/H⁺ exchanger isoform 1) in dendrites presents a major pathway for Na⁺ overload. Neuronal dendrites exhibited higher pH_i regulation rates than soma as a result of a larger surface area/volume ratio. Following a 2-h oxygen glucose deprivation and a 1-h reoxygenation, NHE-1 activity was increased by ∼70–200% in dendrites. This elevation depended on activation of p90 ribosomal S6 kinase. Moreover, stimulation of NHE-1 caused dendritic Na⁺_i accumulation, swelling, and a concurrent loss of Ca²⁺_i homeostasis. The Ca²⁺_i overload in dendrites preceded the changes in soma. Inhibition of NHE-1 or the reverse mode of Na⁺/Ca²⁺ exchange prevented these changes. Mitochondrial membrane potential in dendrites depolarized 40 min earlier than soma following oxygen glucose deprivation/reoxygenation. Blocking NHE-1 activity not only attenuated loss of dendritic mitochondrial membrane potential and mitochondrial Ca²⁺ homeostasis but also preserved dendritic membrane integrity. Taken together, our study demonstrates that NHE-1-mediated Na⁺ entry and subsequent Na⁺/Ca²⁺ exchange activation contribute to the selective dendritic vulnerability to in vitro ischemia.

Introduction

Neuronal dendrites are vulnerable to injury under diverse pathological conditions, including cerebral ischemia, epilepsy, and Alzheimer disease (1, 2). The hallmark of dendritic injury is the formation of focal swelling or beads along the length of the dendritic arbor (3). However, the underlying mechanisms for this selective dendritic injury remain poorly understood. The initial NMDA or kainite-mediated swelling in dendrites of cultured neurons depends on intracellular accumulation of Na⁺ and Cl⁻ but not Ca²⁺ (4). On the other hand, excessive Ca²⁺ entry plays a role in the long lasting structural damage and delayed recovery in hippocampal slices in response to NMDA (4, 5). A correlation between dendritic bead formation and ATP reduction/mitochondrial dysfunction has been demonstrated in cultured hippocampal neurons following glutamate exposure (6). However, the relationship between selective dendritic damage, loss of Na⁺ and Ca²⁺ homeostasis, and mitochondrial dysfunction following ischemia remains to be defined.

NHE-1 (Na⁺/H⁺ exchanger isoform 1) is a plasma membrane protein present in virtually all mammalian cells and plays a central role in intracellular pH (pH_i) and cell volume regulation (7). NHE-1 activity is directly activated by intracellular acidification and/or by protein phosphorylation mediated by ERK-p90 ribosomal S6 kinase (p90^RSK)² in ischemic neurons (8). Excessive NHE-1 activation results in intracellular Na⁺ accumulation, which subsequently promotes Ca²⁺ entry via reversal of Na⁺/Ca²⁺ exchange (NCX_rev) and plays an important role in myocardium ischemia/reperfusion injury (9). We recently reported that NHE-1 activity in the soma of neurons and astrocytes is stimulated following ischemia, and inhibition of NHE-1 activity is neuroprotective (8, 10). In addition, inhibition of NHE-1 either pharmacologically or by genetic knockdown reduces infarction at 24 h following in vivo focal ischemia (11). However, it remains unexplored whether concurrent activation of NHE-1 and NCX_rev contributes to the selective vulnerability of postsynaptic neuronal dendrites to ischemic damage.

In the current study, we demonstrated that neurons exhibited robust NHE-1-dependent pH_i regulation in their dendrites as a result of their large surface area/volume ratio. Further, in vitro ischemia (oxygen glucose deprivation and reoxygenation, OGD/REOX) stimulated NHE-1 activity in large dendrites (Lg-dendrites). NHE-1-mediated Na⁺ entry and subsequent stimulation of NCX_rev activity contributed to selective ischemic damage of dendrites. The underlying mechanisms involved the loss of mitochondrial Ca²⁺ homeostasis and mitochondrial membrane dysfunction.

EXPERIMENTAL PROCEDURES

Materials

Hanks' balanced salt solution was from Mediatech Cellgro (Manassas, VA). Neurobasal medium, B-27 supplement, fura-2/AM, SBFI/AM, BCECF/AM, rhod-2/AM, MitoTracker Green, TMRE, calcein/AM, JC-1, Vybrant® DiO, SYTO 60, and 4-bromo-A-23187 were from Invitrogen. Saponin, tetraphenylboron, gramicidin, and monensin were purchased from Sigma. RU360 was from EMB Chemicals (Gibbstown, NJ). Pluronic F-127 was from BASF Corp. (Parsippany, NJ). HOE 642 was a kind gift from Aventis Pharma (Frankfurt, Germany). SEA0400 was a kind gift from Taisho Pharmaceutical Co. Ltd. (Omiya, Saitama, Japan). BI-D1870 was purchased from the School of Life Science, University of Dundee (Dundee, Scotland).

Pure Cortical Neuron Cultures

Pure cortical neurons from embryonic day 14–16 mouse fetuses (SV129/Black Swiss) were prepared as described previously (8). The cortices were removed from E14–16 fetuses and treated with 0.5 mg/ml trypsin at 37 °C for 25 min. The cells were centrifuged at 300 × g for 4 min. The cell pellet was diluted in B-27 supplemented neurobasal medium (2%) containing 0.5 mm l-glutamine and penicillin/streptomycin (100 units/ml and 0.1 mg/ml, respectively). The cells were seeded at a density of 1 × 10⁵ cells/cm² on glass coverslips in 6-well plastic plates coated with poly-d-lysine. The cultures were maintained in an incubator (model 3130, Thermo Forma, Waltham, MA) with 5% CO₂ and atmospheric air at 37 °C. Half of the medium was replaced twice a week. 10–15-day cultures were used in the study.

OGD Treatment

10–15-day neuronal cultures grown on coverslips in 6-well plates were rinsed with an isotonic OGD solution (pH 7.4) containing 0 mm glucose, 21 mm NaHCO₃, 120 mm NaCl, 5.36 mm KCl, 0.33 mm Na₂HPO₄, 0.44 mm KH₂PO₄, 1.27 mm CaCl₂, and 0.81 mm MgSO₄. This solution has a K⁺ concentration (∼5.8 mm) that is similar to that of the neurobasal medium (5.6 mm) used for cell cultures. The cells were incubated in 1 ml of OGD solution for 2 h in a hypoxic incubator (model 3130, Thermo Forma) containing 94% N₂, 1% O₂, and 5% CO₂. Normoxic control cells were incubated for 2 h in 5% CO₂ and atmospheric air in a buffer identical to the OGD solution except for the addition of 5.5 mm glucose. REOX was achieved by the addition of glucose (5.5 mm) and incubation at 37 °C in 5% CO₂ and atmospheric air. Alternately, REOX was performed on the microscope stage by superfusion with HCO₃⁻-EMEM at 37 °C, equilibrated with 5% CO₂ and ∼18% O₂ (monitored by an in-line oxygen electrode, model 16-730; Microelectrodes, Bedford, NH).

pH_i Measurement

pH_i measurement and prepulse treatment were performed as described previously with some modifications (8). Briefly, pure neuronal cultures grown on coverslips were incubated with 2.5–5 μm BCECF/AM for 30 min during normoxia or during the last 30 min of REOX at 37 °C. The coverslips were washed with HCO₃⁻-free HEPES-EMEM and placed in a temperature-controlled (37 °C) open bath imaging chamber (model RC24, Warner Instruments, Hamden, CT). The chamber was mounted on the stage of the TE 300 inverted epifluorescence microscope, and 1–3 neurons were visualized with a ×100 oil immersion objective. The cells were excited every 10–30 s at 440 and 490 nm, and the emission fluorescence at 535 nm was recorded. Images were collected using a Princeton Instruments MicroMax CCD camera and analyzed with MetaFluor image-processing software. Fluorescence changes in regions of interest in soma, Lg-dendrites, and small dendrites (Sm-dendrites) were determined. Lg-dendrites were defined as dendritic segments with a width of 5.3 ± 1.2 μm, whereas Sm-dendrites were ones with a width of 1.8 ± 0.4 μm. The ratio of the background-corrected fluorescence emissions (F490/F440) for each region was calibrated using the high K⁺/nigericin technique (8). pH_i values were calculated for soma, Lg-dendrites, and Sm-dendrites using the respective BCECF calibration values collected from each region.

For the prepulse treatment, cells were subjected to an acid load by a transient application (1.5 min) of a 30 mm NH₄⁺/NH₃ solution. NH₄⁺/NH₃ solutions were prepared by replacing 30 mm NaCl in the HEPES-buffered solution with an equimolar concentration of NH₄Cl. pH_i recovery rates were determined from the slope of a fitted linear regression within the first minute after NH₄⁺/NH₃ prepulse (8). To minimize differential allosteric effects of H⁺ on NHE-1 activity, pH_i recovery rates were measured at pH_i ∼6.2 throughout the study. In the Na⁺-free experiments, NaCl in the HEPES-buffered solution was replaced with an equimolar concentration of NMDG. NMDG-substituted Na⁺-free solutions (∼5 min) do not cause cell swelling in acutely isolated CA1 neurons (12).

Determination of Intrinsic Buffer Power (β_i)

β_i was determined in somata, Lg- and Sm-dendrites over a range of pH_i by subjecting the cells to progressively decreasing concentrations of NH₄⁺ in Na⁺-free HEPES-EMEM as described previously (8). The total H⁺ net efflux rate (J_H⁺ mm H⁺/min) was determined in three neuronal regions by multiplying β_i by ΔpH_i/Δt at pH_i ∼6.2. In some experiments, J_H⁺ was also calculated in the presence of HCO₃⁻. The buffering by CO₂/HCO₃⁻ was determined as β_HCO3⁻ = 2.3 × [HCO₃⁻]_i, where [HCO₃⁻]_i = S × PCO₂ × 10^{(pH_i − pK)}, where S = 0.0314, PCO₂ = 40 mm Hg, and pK = 6.12. At pH 6.2, the contribution of β_HCO3⁻ to total buffering in normoxic cells was ∼6%.

Intracellular Na⁺ Measurement

Intracellular Na⁺ concentration ([Na⁺]_i) was measured with the fluorescent dye SBFI/AM as described previously with some modifications (14). Cultured neurons grown on coverslips were loaded with 30 μm SBFI/AM plus 0.02% pluronic acid during a 45-min REOX following a 2-h OGD. The coverslips were placed in the open bath imaging chamber and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C. Using the Nikon TE 300 inverted epifluorescence microscope and a ×100 oil immersion lens, neurons were excited at 345 and 385 nm, and the emission fluorescence at 510 nm was recorded. Regions of interest (1–3 cells/area) were drawn to determine SBFI fluorescence changes in soma, Lg-dendrites, and Sm-dendrites. The 345/385 ratios were analyzed with the MetaFluor image-processing software. Absolute [Na⁺]_i was determined for each cell by performing an in situ calibration as described previously (14). Multiple time point data acquisition induced phototoxicity in neurons. Therefore, [Na⁺]_i was only determined in normoxic controls or 45-min REOX-treated neurons.

Intracellular Ca²⁺ Measurement

Neurons grown on coverslips were incubated with 5 μm fura-2 AM during a 2-h OGD. Following OGD, the cells were placed in the open bath imaging chamber and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C. Using the Nikon TE 300 inverted epifluorescence microscope and a ×100 oil immersion objective lens, neurons were excited every 5 min at 345 and 385 nm, and the emission fluorescence at 510 nm was recorded. Images were collected and analyzed with the MetaFluor image-processing software. At the end of each experiment, the cells were exposed to 1 mm MnCl₂ in Ca²⁺-free HCO₃⁻-EMEM and 5 μm 4-bromo-A-23187. The Ca²⁺-insensitive fluorescence was subtracted, and the MnCl₂-corrected 345/385 emission ratios were converted to [Ca²⁺] as described previously (14).

Measurement of Mitochondrial Ca²⁺

Neurons on coverslips were incubated at 37 °C for 60 min with 200 nm MitoTracker Green and 9 μm rhod-2/AM, which was reduced with a minimum of sodium borohydride in HCO₃⁻-EMEM containing 3 mm sodium succinate (14). Coverslips were then incubated for 2 h under either OGD or normoxia conditions. For REOX, coverslips were placed in the perfusion chamber on the stage of the Leica DMIRE2 confocal microscope and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C. Cells (1–3 in the field) were visualized with a ×100 oil immersion objective and scanned sequentially for MitoTracker Green (excitation 488 nm (argon laser line), emission 500–545 nm) and rhod-2 (excitation 543 nm (HeNe laser), emission 544–677 nm). The MitoTracker Green signal was used to maintain focus prior to each sequential scan. Sequential scans were analyzed using the Leica confocal software. Average grayscale values were collected from regions of interest around mitochondrial clusters exhibiting colocalization of MitoTracker Green and rhod-2. Ca²⁺_m levels were expressed as relative change of rhod-2 signals from the base-line values, and summarized data represent the average of the calculated values from 2–3 cells as described previously (14).

Measurement of Mitochondrial Membrane Potential (Ψ_m)

The fluorescent probe JC-1 was used to monitor Ψ_m as described previously (14). Neurons on coverslips were loaded with 9 μm JC-1 during 2 h of OGD at 37 °C. Following OGD, the cells were placed in the temperature-controlled open bath imaging chamber and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C. Cells were visualized using the Nikon TE 300 inverted epifluorescence microscope and a ×60 oil immersion objective. Cells were excited at 480 nm, and emission fluorescence images were recorded at 535 nm (the monomer) and 640 nm (JC-1 aggregates). The ratio of the aggregate to monomer fluorescence was measured in regions of interest in soma, Lg-dendrites, and Sm-dendrites (2–5 cells/area). In this study, we applied 1.0 μm FCCP for 1 min to determine the maximal loss of JC-1 signals. Ψ_m was expressed as the percentage of the maximal FCCP-induced change under normoxic controls (14). We believe that the FCCP-sensitive loss of JC-1 signals largely reflects changes in Ψ_m and is not affected by plasma membrane potentials. This is based on a study where 2.5 μm FCCP caused the immediate collapse of Ψ_m and complete depolarization of plasma membrane potential. However, a low concentration of FCCP (0.25 μm) had no effect on plasma membrane potential (15).

To further confirm that the changes in JC-1 reflect changes of Ψ_m, we also conducted some parallel experiments using the cationic membrane-permeant fluorescence probe TMRE. Neurons were loaded with 5 nm TMRE and 200 nm MitoTracker Green in a buffer supplemented with 1 μm tetraphenylboron for 30 min 37 °C. The coverslips were then placed in the perfusion chamber on the stage of the Leica DMIRE2 confocal microscope and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C supplemented with 5 nm TMRE. Cells (1–3 in the field) were visualized with a ×100 oil immersion objective and scanned sequentially for MitoTracker Green (excitation 488 nm (argon laser line), emission 500–545 nm) and TMRE (excitation 543 nm (HeNe laser), emission 544–677 nm). Sequential scans were analyzed using the Leica confocal software. The MitoTracker Green signal was used to maintain focus prior to each sequential scan and to identify mitochondrial clusters exhibiting colocalization of MitoTracker Green and TMRE. Maximal Ψ_m dissipation was induced by FCCP (1.0 μm) at the end of each experiment. At a concentration of 5 nm, TMRE behaves in the non-quench mode and decreases its fluorescence intensity when Ψ_m is reduced. Data are expressed as relative percent change in FCCP-sensitive TMRE signals.

Determination of Surface Area/Volume Ratio in Soma and Dendrites

To determine differences in the ratio of surface area to volume in soma and dendrites, neurons grown on coverslips were loaded with 0.5 μm calcein/AM (cytosol dye) and 5 μm SYTO 60 (nucleus dye) for 30 min at 37 °C. The coverslips were then placed in the perfusion chamber on the stage of a Leica DMIRE2 confocal microscope and visualized with a ×100 oil immersion objective. A 110-μm-thick image stack (300 slices at 512 × 512 pixels) was collected sequentially (excitation 488 nm (argon laser line), emission 500–545 nm; excitation 543 nm (HeNe laser), emission 544–677) and imported into ImageJ (version 1.41, National Institutes of Health). A cellular region (soma, nucleus, and Lg- or Sm-dendrites) was defined, and the surface area was calculated by summing the product of the region perimeter with the distance between each image section (0.38 μm). The volume of the region was calculated with the region area and section distance. The soma volume was corrected by subtracting the calculated volume for the nucleus. No attempt was made to correct for the intracellular volumes of endoplasmic reticulum or mitochondria.

Detection of Dendritic Beading Formation (Varicosities)

To monitor dendritic beading formation, neurons grown on coverslips were loaded with the plasma membrane dye Vybrant® DiO as per the manufacturer's instructions. Following OGD, the coverslips were placed in the open bath imaging chamber and superfused (1 ml/min) with HCO₃⁻-EMEM at 37 °C on the stage of a Leica DMIRE2 confocal microscope. A single neuron was visualized with a ×100 oil immersion objective and scanned (512 × 512, 200 Hz) with an argon laser (excitation 488 nm, emission 500–545 nm). The images were analyzed for dendrite beading with ImageJ analysis software. Beads with a diameter ∼4 times larger than the width of the corresponding dendrite were counted in a 90 × 90-μm area. Data represent the average of the calculated values from three or four experiments.

Immunoblotting

Cells were washed with ice-cold PBS and lysed with 30 s sonication at 4 °C in anti-phosphatase buffer (pH 7.4) containing 145 mm NaCl, 1.8 mm NaH₂PO₄, 8.6 mm Na₂HPO₄, 100 mm NaF, 10 mm Na₄P₂O₇, 2 mm Na₃VO₄, 2 mm EDTA, and 0.2 μm microcystin and protease inhibitors as described previously (11). Protein content was determined by the bicinchoninic acid method. Protein samples (40 μg/lane) and prestained molecular mass markers (Bio-Rad) were denatured in SDS 2× sample buffer and then electrophoretically separated on 8% SDS gels. The resolved proteins were electrophoretically transferred to a PVDF membrane (14). The blots were incubated in 7.5% nonfat dry milk in Tris-buffered saline (TBS) overnight at 4 °C and then incubated for 1 h with polyclonal anti-NHE-1 (1:500), polyclonal anti-NHE-2 (1:500) (16), polyclonal anti-NHE-3 (1:1000; Alpha Diagnostic International, San Antonio, TX), polyclonal anti-NHE-5 (1:1000). The blots were rinsed with TBS and incubated with horseradish peroxidase-conjugated secondary IgG for 1 h. Bound antibody was visualized using an enhanced chemiluminescence assay (Amersham Biosciences).

Immunofluorescence Staining

Cells grown on coverslips were fixed in 4% paraformaldehyde in PBS for 15 min. After rinsing, cells were incubated with a blocking solution for 20 min followed by application of a primary polyclonal antibody for NHE-1 (1:50; Abcam Inc., Cambridge, MA). After rinsing in PBS, cells were incubated with Alexa Fluor^TM 488 goat anti-rabbit IgG (1:200; Invitrogen) for 1 h. The coverslips were then covered with Vectashield mounting medium (Vector Laboratories, Burlingame, CA). Fluorescence images were captured by the Nikon TE 300 inverted epifluorescence microscope (×40) using a Princeton Instruments MicroMax CCD camera and MetaMorph image-processing software.

Statistics

Statistical significance was determined by Student's t test or an analysis of variance (Bonferroni post hoc test) in the case of multiple comparisons. A p value smaller than 0.05 was considered statistically significant. n values represent the number of cultures in each experiment.

RESULTS

Surface Area/Volume (A/V) Ratio in Soma and Dendrites

In order to accurately calculate ionic flux rates in soma and dendrites, we first estimated surface area to volume (A/V) ratios in these cellular regions. Fig. 1A shows a single slice two-dimensional image of cultured neurons from a confocal stack image (300 slices, 110 μm thick). The arrows in Fig. 1A illustrate the areas in the somata, Lg-dendrite, and Sm-dendrite where the A/V ratios and ionic changes were determined. Fig. 1B is a three-dimensional reconstruction of the stack of images with Metamorph software, highlighting neuronal morphology with the distinctly higher A/V ratios in Lg- and Sm-dendrites. The A/V ratio in Lg-dendrites was 3.8 times larger than somata (Fig. 1C). An ∼7 times larger A/V ratio was estimated for Sm-dendrites. Interestingly, 2 h of OGD and 1 h of REOX did not significantly change the A/V ratio either in soma or in dendrites.

FIGURE 1.

Surface area/volume ratios in soma and dendrites. A, a single two-dimensional (2-D) confocal image taken from a 300-slice image stack (110 μm thick, 512 × 512 pixels) of a neuron loaded with calcein/AM. Soma, Lg-dendrites, and Sm-dendrites are indicated (arrows). B, the 300-slice image stack was rendered into a three-dimensional (3-D) image using Metamorph software to illustrate A/V ratios determined in soma, Lg-dendrites, and Sm-dendrites (arrowheads). C, A/V ratios were calculated in three regions of normoxic control neurons or neurons subjected to 2 h of OGD and 1 h of REOX. Data are mean ± S.E. (error bars), n = 4. *, p < 0.05 versus soma. D, β_i was determined in three regions under normoxia and following 2-h OGD/1-h REOX. β_i in each region was plotted against pH_i and fit with a linear regression. The fitted slopes for each region were not significantly different under either normoxic (p = 0.97) or OGD/REOX (p = 0.474) conditions.

We then determined β_i in three regions as shown in Fig. 1D. β_i in each region was plotted against pH_i and fit with a linear regression. The slopes of the lines in the three regions were not significantly different under normoxic control or OGD/REOX conditions. These findings imply that the changes of pH_i regulation may result from altered function of H⁺ transporters, such as NHEs.

Changes of pH_i in Soma and Dendrites following OGD/REOX

Lg-or Sm-dendrites exhibited more alkaline resting pH_i values than soma under normoxic conditions (Fig. 2A). Inhibition of NHE-1 with its potent inhibitor HOE 642 (1 μm) or the newly developed NHE-1 kinase p90^RSK inhibitor fluoromethylketone (FMK; IC₅₀ of 15 nm, (17)) acidified pH_i and significantly decreased pH_i recovery rates (Fig. 2, A and B). OGD/REOX caused an alkalization of pH_i in soma (a shift from 6.96 ± 0.03 to 7.19 ± 0.05; p < 0.05). Inhibition of NHE-1 reversed the OGD/REOX-mediated increase in pH_i (Fig. 2C). Moreover, inhibition of the NHE-1 kinase p90^RSK with FMK prevented the post-OGD alkalization. OGD/REOX did not trigger additional changes in pH_i in dendrites. However, either HOE 642 or FMK significantly acidified dendrites following OGD/REOX. These data suggest that NHE-1 activation plays a role in resting pH_i maintenance and contributes to the intracellular post-OGD alkalization.

FIGURE 2.

A, resting pH_i in the soma, Lg-dendrites, and Sm-dendrites under normoxic conditions. In the drug treatment experiments, neurons were exposed to either HOE 642 (1 μm) or FMK (3 μm) for 30 min prior to the pH_i determination. B, pH_i regulation in the soma, Lg-dendrites, and Sm-dendrites under normoxic conditions. Data are mean ± S.E. (error bars), n = 3–4. #, p < 0.05 versus corresponding normoxia; *, p < 0.05 versus corresponding soma. C, changes of pH_i at 60 min of REOX following 2 h of OGD. In the drug treatment experiments, either HOE 642 (1 μm) or FMK (3 μm) was present only during the 60-min REOX. Data are mean ± S.E., n = 3–4. *, p < 0.05 versus corresponding normoxia; #, p < 0.05 versus OGD/REOX.

Increased H⁺ Efflux in Soma and Dendrites following OGD/REOX

We further determined NHE-1 activity in soma and dendrites by measuring the pH_i recovery rate following the NH₃/NH₄⁺ prepulse-induced acidification. As shown in Fig. 3A, when neurons were exposed to 30 mm NH₃/NH₄⁺, pH_i in Lg-dendrites rose rapidly as NH₃ diffused into the cell and combined with H⁺ to form NH₄⁺ (a and b) and then declined slowly (b and c). Returning cells to the standard HCO₃⁻-free HEPES-EMEM solution caused pH_i to decrease due to the rapid diffusion of NH₃, which was dissociated from the newly formed NH₄⁺, and trapping H⁺ inside the cells (c and d). Both normoxic control and OGD/REOX-treated cells were able to restore pH_i to their basal levels (Fig. 3A). However, the pH_i recovery rate increased by ∼2-fold in Lg-dendrites following OGD/REOX (1.23 ± 0.24 unit/min versus 0.57 ± 0.05 unit/min in normoxic neurons, p < 0.05).

FIGURE 3.

Increased H⁺ efflux rate in somata and dendrites following OGD/REOX. A, representative pH_i changes in Lg-dendrites subjected to NH₄⁺/NH₃ prepulse-mediated acid-loading. pH_i recovery rate was determined by fitting a slope to the pH_i values within the first minute following the prepulse in either normoxic or 2-h OGD/1-h REOX-treated neurons. pH_i recovery rates were determined at ∼6.2 to normalize for the allosteric regulation of H⁺ on NHE-1 activity. B, summary data of pH_i recovery rates under normoxic and OGD/REOX conditions. Data are mean ± S.E. (error bars), n = 3–4. *, p < 0.05 versus corresponding normoxia. #, p < 0.05 versus corresponding soma. C, pH_i recovery rates were corrected for the relative differences in surface area to volume ratios in three regions. Data are mean ± S.E., n = 3–4. *, p < 0.05 versus corresponding normoxia; #, p < 0.05 versus corresponding soma. D, proton flux (J_H⁺) was calculated at pH ∼6.2 during pH recovery following NH₄⁺/NH₃ prepulse. Data are mean ± S.E. n = 3–4. *, p < 0.05 versus corresponding normoxia; #, p < 0.05 versus corresponding soma.

pH_i recovery rates were significantly higher in the Lg-dendrites (90%) and in the Sm-dendrites (330%) than the soma under normoxic conditions (Fig. 3B). The apparent higher pH_i recovery rates in the dendrites could result from the larger A/V ratios in the dendrites. Thus, we corrected the pH_i recovery rate for A/V ratio in the three different regions. After the correction, the rates were similar in all three regions under normoxic control conditions (Fig. 3C). 2-h OGD/1-h REOX triggered a further increase in the H⁺ efflux in the soma (264%), the Lg-dendrites (218%), and the Sm-dendrites (69%; Fig. 3B). After the correction for the A/V ratio, the OGD/REOX-induced elevation in pH_i recovery rates remained significant in soma and Lg-dendrites (Fig. 3C).

This finding was further validated by calculating J_H⁺ (Fig. 3D). Dendrites exhibited smaller J_H⁺ than soma under normoxia and OGD/REOX conditions. The OGD/REOX-mediated selective stimulation of J_H⁺ persisted in soma and Lg-dendrites but not in Sm-dendrites. Similar changes of J_H⁺ were observed in the presence of HCO₃⁻ (21 mm; Fig. 3D). The lack of changes in the A/V ratios and β_i following OGD/REOX suggest that the OGD/REOX-induced stimulation of pH_i recovery rates mainly reflect J_H⁺.

Differential NHE-1 Activity in Soma and Dendrites

We directly evaluated NHE-1-dependent pH_i regulation activity in the soma and the dendrites using the NHE-1 inhibitor HOE 642 (1 μm, IC₅₀ of 0.08 μm) at a concentration that inhibits only the NHE-1 isoform (18). As shown in Fig. 4A, the OGD/REOX-mediated elevation of the H⁺ extrusion rate in the somata was nearly abolished in the presence of HOE 642. However, HOE 642 only partially blocked the elevated pH_i recovery rate in the dendrites (∼50–60%). To determine the possible role of other isoforms of NHE in neuronal processes, we examined the effects of removing extracellular Na⁺, which inhibits the function of all NHE isoforms by abolishing the inward Na⁺ driving force. In the absence of extracellular Na⁺, H⁺ extrusion was absent in soma, similar to the NHE-1 inhibition via HOE 642 (Fig. 4A). In the dendrites, the pH_i recovery rate was eliminated by ∼76–86%. Inhibiting all NHE isoforms with a general NHE inhibitor EIPA (100 μm) had a similar effect as removing extracellular Na⁺. Moreover, the residual Na⁺-independent H⁺ extrusion in the Sm-dendrites could be mediated by vacuolar H⁺-ATPase. Inhibition of vacuolar H⁺-ATPase with a specific inhibitor, bafilomycin (1 μm), abolished the residual H⁺ extrusion in both the Lg- and the Sm-dendrites (Fig. 4A). These data imply that NHE-1 is the dominant isoform in soma. However, in the dendrites, pH_i regulation is governed by NHE-1 as well as other NHE isoforms and H⁺-ATPases.

FIGURE 4.

Differential NHE-1 activity in soma and dendrites. A, 2-h OGD/1-h REOX-treated neurons exhibited different pH_i recovery rates following the NH₄⁺/NH₃ prepulse. To inhibit NHE-1 activity, 1 μm HOE 642 was present during the 60-min REOX. In some prepulse studies, Na⁺-dependent H⁺ extrusion was blocked by replacing NaCl in the HEPES-buffered solution with an equimolar concentration of NMDG. Na⁺-free HEPES-EMEM, either with or without 1 μm bafilomycin, was used to evaluate function of other NHE isoforms and vacuolar H⁺-ATPases. Inhibition of all NHE isoforms was examined with the general NHE inhibitor EIPA (100 μm). Data are mean ± S.E (error bars), n = 3–5. *, p < 0.05 versus soma under OGD/REOX; #, p < 0.05 versus corresponding OGD/REOX. B, HOE 642-sensitive pH_i regulation. OGD/REOX data are from the experiments in Fig. 3B. Data are mean ± S.E. n = 3–5. *, p < 0.05 versus soma under normoxia. #, p < 0.05 versus corresponding normoxia. C, NHE-1, NHE-2, NHE-3, or NHE-5 protein expression in cultured cortical neurons. Cerebellar tissue was used as positive control for NHE-3, and blots were probed for β-tubulin as a loading control. D, expression of NHE-1 protein in soma (arrow), Lg-dendrite (arrowhead), and Sm-dendrite (open arrowhead). Negative Control, primary antibody was omitted. Scale bar, 10 μm.

The HOE 642-sensitive portion of the H⁺ extrusion rate was obtained under normoxic control and OGD/REOX conditions (Fig. 4B). Consistently, NHE-1 activity (HOE 642-sensitive portion) in the soma and the Lg-dendrites was significantly elevated following OGD/REOX. Sm-dendrites exhibited a significantly higher basal activity of NHE-1, and OGD/REOX did not cause additional activation. Fig. 4C demonstrates that NHE-1 is the dominant form in cortical neurons, whereas the NHE-3 isoform is restricted to the cerebellum (19). Localization of NHE-1 protein in soma and Lg- and Sm-dendrites was also shown in Fig. 4D.

NHE-1-mediated Na⁺ Entry in Soma and Dendrites following OGD/REOX

The robust NHE-1 activation in the soma and the dendrites following REOX led us to speculate that NHE-1 plays a role in the dendritic Na⁺_i dysregulation following in vitro ischemia. [Na⁺]_i in the soma and the dendrites was monitored under normoxic controls and at 45 min REOX. There were no significant differences in the base-line [Na⁺]_i between the soma and the dendrites (Fig. 5A, arrowhead). A 45-min REOX following 2 h OGD led to an increase in [Na⁺]_i throughout the neuron. Localized increases in [Na⁺]_i were detected in the dendrites (Fig. 5A, arrow). Summary data show that [Na⁺]_i increased from a resting level of 12.5 ± 0.3 to 44.4 ± 2.6 mm in the soma (p < 0.05; Fig. 5B). The Sm-dendrites exhibited the largest Na⁺ accumulation (56.5 ± 4.1 mm, p < 0.05). Inhibition of NHE-1 with HOE 642 during REOX abolished the OGD/REOX-induced Na⁺ overload in both the soma and the dendrites. The rise in HOE 642-sensitive changes in [Na⁺]_i was shown in Fig. 5C. These data imply that OGD/REOX-mediated accumulation of [Na⁺]_i is largely mediated via NHE-1 activation.

FIGURE 5.

NHE-1-mediated Na⁺ entry in soma and dendrites following OGD/REOX. A, representative SBFI pseudocolored images of changes in [Na⁺]_i in normoxic and OGD/REOX-treated neurons. Arrowhead, low levels of [Na⁺]_i; arrow, localized increases in [Na⁺]_i. B, summary data of [Na⁺]_i in soma, Lg-dendrites, and Sm-dendrites of neurons under normoxic conditions and 2-h OGD/45-min REOX. Data are mean ± S.E. (error bars), n = 3–5. *, p < 0.05 versus normoxia; #, p < 0.05 versus OGD/REOX; α, p < 0.05 versus soma OGD/REOX. C, HOE 642-sensitive change in [Na⁺]_i. Data are mean ± S.E., n = 3–5. *, p < 0.05 versus corresponding normoxia; #, p < 0.05 versus soma under OGD/REOX. D, pH_i recovery rates were determined. Normoxia and OGD/REOX data are from the experiments in Fig. 3B. BI-D1870 (1 μm), FMK (3 μm), or HOE 642 (1 μm) was present only during REOX. Data are mean ± S.E., n = 3. *, p < 0.05 versus normoxia; #, p < 0.05 versus OGD/REOX; α, p < 0.05 versus soma OGD/REOX; β, p < 0.05 versus Sm-dendrite normoxia. E, pH_i recovery rates were determined in the presence of 21 mm HCO₃⁻. HOE 642 (1 μm) was present only during REOX. Data are mean ± S.D., n = 4–8 cells. *, p < 0.05 versus normoxia; #, p < 0.05 versus OGD/REOX.

We recently reported that stimulation of NHE-1 depends on activation of the ERK-p90^RSK signal transduction pathways and phosphorylation of NHE-1 (8). In the current study, we examined whether direct inhibition of the NHE-1 kinase p90^RSK with its potent inhibitor BI-D1870 (BI-D, IC₅₀ of 10–30 nm (20)) could reduce OGD/REOX-mediated NHE-1 activation. The OGD/REOX-induced Na⁺_i loading in Lg- and Sm-dendrites was abolished by BI-D1870 (Fig. 5B). This effect is similar to the one mediated by the NHE-1 inhibitor HOE 642. Thus, p90^RSK function is largely responsible for NHE-1 activation and Na⁺ entry. These data also imply that the initial increased intracellular acidification associated with OGD (8) is not sufficient to drive NHE-1 activity but requires altered phosphorylation of the transporter.

We further compared the effects of BI-D along with FMK on inhibition of NHE-1 activation. Both BI-D and FMK abolished the OGD/REOX-mediated stimulation of pH_i recovery in the soma and the dendrites (Fig. 5D). Especially in the Sm-dendrites, BI-D or FMK profoundly suppressed the pH_i recovery rate, which was only ∼25% of the normoxic basal levels (p < 0.05; Fig. 5D). These data suggest that p90^RSK pathways play a more dominant role in pH_i regulation in the dendrites than in the soma.

Last, the pH_i recovery rates in all three regions were determined in the presence of 21 mm bicarbonate under normoxia and OGD/REOX conditions (Fig. 5E). The results are similar to those in the absence of physiological bicarbonate. These data suggest that the role for bicarbonate-dependent ion transporters in regulation of pH_i is negligible under these conditions.

To further test NHE-1-mediated Na⁺ overload, we examined whether there was a differential Na⁺ overload in soma and Sm-dendrites when Na⁺/K⁺-ATPase was blocked by ouabain (0.1 mm) during REOX. As shown in supplemental Fig. 1A, Na⁺_i clearly loaded faster in Sm-dendrites than in soma under these conditions. Moreover, at 30 min of REOX, blocking NHE-1 activity decreased the Na⁺ influx in Sm-dendrites (supplemental Fig. 1B). A similar trend was also observed in Lg-dendrites (supplemental Table 1).

Delayed Dysregulation of Ca²⁺ Depends on a Concurrent Activation of NHE-1 and NCX_rev

One consequence of Na⁺_i overload is to trigger NCX_rev and Ca²⁺ entry. To investigate the possible concerted activation of NHE-1 and NCX_rev, we first monitored changes of local [Ca²⁺]_i following REOX. The somata [Ca²⁺]_i was 68 ± 9 nm under normoxic control conditions and increased modestly to 104 ± 8 nm after 2 h of OGD (p < 0.05), which remained unchanged over the initial 35-min REOX (Fig. 6, A and B). In contrast, 2 h of OGD caused a slightly higher elevation in the dendritic [Ca²⁺]_i (157 ± 21 nm, p < 0.05). Twenty min of REOX triggered a secondary rise in the dendritic [Ca²⁺]_i that initiated from local “hot spots” and then spread toward the soma over time (Fig. 6, A and C). The amplitude of the dendritic Ca²⁺ dysregulation was significantly higher than the soma. By 45 min of REOX, the dendritic [Ca²⁺]_i rose dramatically to 1206 ± 440 nm and spread to the soma, which exhibited slightly lower levels (766 ± 300 nm). Sustained elevation in [Ca²⁺]_i during REOX (60–100 min) resulted in cell death, as reflected by a sudden loss of the dye (data not shown).

FIGURE 6.

Changes in [Ca²⁺]_i in soma and dendrites following OGD/REOX. A, representative fura-2 pseudocolored images of changes in [Ca²⁺]_i in neurons at 0, 20, 30, 40, and 45 min REOX. Arrowhead, low levels of [Ca²⁺]_i. Arrow, localized increases in [Ca²⁺]_i. B and C, summarized changes of [Ca²⁺]_i in soma (B) or Sm-dendrites (C). In the HOE 642 or SEA0400 studies, the drugs (1 μm) were present only during 1 h of REOX. Data are mean ± S.E. (error bars), n = 4. *, p < 0.05 versus 0 min REOX; #, p < 0.05 HOE versus non-treated OGD/REOX; α, p < 0.05 HOE or SEA versus non-treated OGD/REOX.

We speculated that NCX_rev contributes to this Ca²⁺ dysregulation as a result of the robust NHE-1 activation and Na⁺ overload. First, we investigated whether inhibition of NHE-1 activity with HOE 642 could block the delayed rise in [Ca²⁺]_i. As shown in Fig. 6 (B and C), when HOE 642 was present only during 60 min of REOX, no secondary rise in [Ca²⁺]_i occurred in the soma and the dendrites following REOX. These data imply that NHE-1 activation is involved in the secondary loss of Ca²⁺_i homeostasis during REOX. To establish whether this Ca²⁺ rise results from activation of NCX_rev, we conducted the experiments in the presence of SEA0400 (1 μm), a potent inhibitor of NCX_rev. As shown in Fig. 6, B and C, REOX failed to elicit the secondary elevation in [Ca²⁺]_i in both the soma and the dendrites. The results were similar to those of the HOE 642-treated cells. This led us to conclude that concerted activation of NHE-1 and NCX_rev contributed to the delayed Ca²⁺ dysregulation in soma and dendrites following in vitro ischemia. However, these inhibitors did not affect basal levels of [Na⁺]_i and [Ca²⁺]_i under normoxic control conditions (supplemental Table 2).

Changes of Ca²⁺_m and Ψ_m in Soma and Dendrites following OGD/REOX

Excessive dendritic Na⁺_i and Ca²⁺_i overload will affect mitochondrial Ca²⁺ homeostasis and mitochondrial function. To investigate this, we first monitored changes of Ca²⁺_m in the soma and the dendrites during 0–60 min REOX. There was a slow and progressive elevation in the somata Ca²⁺_m starting by 10 min REOX and reaching a plateau value (∼2.5-fold control) by 40 min REOX (Fig. 7A). Interestingly, the somata Ψ_m did not decrease significantly, whereas Ca²⁺_m was increasing during early REOX. However, Ψ_m depolarized significantly after Ca²⁺_m reached its plateau levels, and it was reduced to 47 ± 6% of control at 60 min of REOX (Fig. 7A).

FIGURE 7.

Changes in Ca²⁺_m and Ψ_m in soma and dendrites following OGD/REOX. Changes in Ca²⁺_m and Ψ_m were monitored in soma (A), Lg-dendrites (B), and Sm-dendrites (C). Blue lines, Ψ_m data determined with JC-1 and expressed as the percentage of the maximal FCCP-induced change in the JC-1 ratio of normoxic controls. Red lines, Ca²⁺_m determined by the relative change in rhod-2 fluorescence. In the HOE study, 1 μm HOE 642 was present only during 0–1 h REOX. Data are mean ± S.E. (error bars), n = 3–4. *, p < 0.05 versus 0 min REOX; #, p < 0.05 versus OGD/REOX.

By 10 min of REOX, Ca²⁺_m levels had increased significantly in the Sm-dendrites but not in the Lg-dendrites (Fig. 7, B and C). The rate of Ca²⁺_m increase in the Sm-dendrites during early REOX was significantly faster than in the soma (0.018 versus 0.003 relative change/min, p < 0.05). At 60 min REOX, Ca²⁺_m was increased by ∼4.5-fold in the Sm-dendrites and ∼3.2-fold in the Lg-dendrites. Ca²⁺ accumulation in the mitochondria remained elevated throughout the neuron until ∼100 min, when a sudden loss in the rhod-2 dye signal occurred, probably due to a collapse of mitochondrial function (data not shown).

The earlier rise in Ca²⁺_m in the dendrites was accompanied by a faster decrease of Ψ_m (Fig. 7, B and C). Ψ_m in the dendrites (particularly in the Sm-dendrites) depolarized at a rate twice that of the somata (1.2 versus 0.7%/min). Ψ_m in the Sm-dendrites dropped to the lowest level (17 ± 4% of control) by 40 min of REOX. The kinetics of the Ψ_m collapse in the dendrites exhibited a significant negative correlation with the dendritic Ca²⁺_m accumulation (Pearson product moment correlation coefficient = −0.964, p < 0.001). Thus, compared with the soma, the dendrites show two characteristics: earlier onset time and larger magnitude in the loss of mitochondrial Ca²⁺ homeostasis and Ψ_m. This demonstrates that the dendritic mitochondria are more sensitive to OGD/REOX damage than the soma. These changes are consistent with the earlier loss of Na⁺ and Ca²⁺ homeostasis in the dendrites.

Interestingly, inhibition of NHE-1 activity with 1 μm HOE 642 prevented the REOX-mediated changes of Ca²⁺_m and Ψ_m in soma. In the presence of 1 μm HOE 642, there were no significant increases in Ca²⁺_m in the Lg-dendrites and the somata. A slow accumulation in Ca²⁺_m was detected in the Sm-dendrites, which was not statistically significant from 0 min REOX. Moreover, the delayed depolarization of the somata Ψ_m during 50–60 min of REOX was absent with the HOE 642 treatment (Fig. 7A). Strong protective effects of HOE 642 on Ψ_m were also found in the Lg- and the Sm-dendrites. The small dendritic Ψ_m depolarized to ∼44% of control (instead of 17% of control) at 60 min of REOX when NHE-1 activity was inhibited (p < 0.05). Thus, preservation of Ψ_m by NHE-1 inhibition may result from decreased mitochondrial Ca²⁺ loading.

To determine the role of NHE-1 in mitochondrial dysfunction following OGD/REOX, we examined whether FMK (3 μm) would prevent mitochondrial damage in the soma and the dendrites. As shown in Fig. 8, A and B, inhibition of the NHE kinase p90^RSK during REOX attenuated loss of Ψ_m following OGD/REOX in the soma and the Lg-dendrites. In the Sm-dendrites, FMK effectively prevented depolarization of Ψ_m as early as 10 min of REOX (Fig. 8C). Interestingly, these effects were similar to the direct inhibition of NHE-1 mediated by HOE 642. Taken together, we can firmly conclude that blocking either NHE-1 or p90^RSK significantly preserves mitochondrial function in ischemic neurons.

FIGURE 8.

Effects of FMK on changes of Ψ_m in soma and dendrites following OGD/REOX. Changes in Ψ_m were determined with JC-1 and expressed as the percentage of the maximal FCCP-induced change in the JC-1 ratio of normoxic controls in soma (A), Lg-dendrites (B), and Sm-dendrites (C). In the FMK study, 3 μm FMK was present during 0–1 h REOX. Data are mean ± S.E. (error bars), n = 4. *, p < 0.05 versus 0 min REOX; #, p < 0.05 versus OGD/REOX.

In a parallel study, in order to confirm the reliability of the Ψ_m determination with JC-1, we also used the cationic dye TMRE to monitor changes of Ψ_m at 0 and 60 min of REOX (supplemental Fig. 2, A and B). Both JC-1 and TMRE measurements indicated that the REOX-induced decrease in Ψ_m was more profound in the Lg- and the Sm-dendrites. Additionally, inhibition of NHE-1 with HOE 642 during REOX significantly attenuated the OGD/REOX-induced decrease in Ψ_m in all areas of the cells (p < 0.05; supplemental Fig. 2B).

Role of Mitochondrial Uniporter in Mitochondrial Ca²⁺ Accumulation

To investigate the role of the uniporter in mitochondrial Ca²⁺ loading following OGD/REOX, we determined changes of Ca²⁺_m when the mitochondrial uniporter was inhibited with 10 μm RU360 during 60 min of REOX. Inhibition of the uniporter prevented accumulation of Ca²⁺_m in the mitochondria in all regions of the neuron (Fig. 9).

FIGURE 9.

Effects of mitochondrial uniporter inhibitor RU360 on Ca²⁺_m in soma and dendrites following OGD/REOX. Ca²⁺_m was assessed by the relative change in rhod-2 fluorescence when the mitochondrial uniporter inhibitor, RU360 (10 μm), was present during 60-min REOX in soma (A), Lg-dendrites (B), and Sm-dendrites (C). Data were plotted against Ca²⁺_m data from Fig. 7 for OGD/REOX and OGD/REOX + HOE. Data are mean ± S.E. (error bars), n = 3–4. *, p < 0.05 versus 0 min REOX; #, p < 0.05 versus OGD/REOX.

Dendritic Damage Is Reduced by Inhibition of NHE-1 following OGD/REOX

We further investigated dendritic damage by monitoring the dendritic beading (varicosities) and membrane integrity changes following OGD/REOX. As shown in Fig. 10, A and B, 2 h of OGD caused some swelling in the dendrites without formations of varicosities (arrow). Varicosities developed in the dendrites over 30–60 min REOX (Fig. 10C, arrowhead). At 60 min of REOX, the bead density increased by >13 times compared with the normoxic neurons (Fig. 10, C and E). In contrast, in the presence of 1 μm HOE 642 during REOX, the bead formation increased only by ∼3 times (Fig. 10, D and E, arrow; p < 0.05). These data suggest that inhibition of NHE-1 activity with HOE 642 not only reduces the dendritic ionic dysregulation but also decreases dendritic swelling following OGD/REOX.

FIGURE 10.

Dendritic beading in neurons following OGD/REOX. A–D, dendritic beading formation in DiO-loaded cells was detected at 0, 30, 45, and 60 min of REOX. In the HOE studies, 1 μm HOE 642 was present only during 0–1 h of REOX. Arrow, swelling in dendrites without formations of varicosities; arrowhead, varicosities. E, summary data of dendritic beading. The data of normoxia control and normoxia plus HOE 642 (red) groups overlap. Data are mean ± S.E. (error bars), n = 3–4. *, p < 0.05 versus 0 min REOX. #, p < 0.05 versus OGD/REOX.

DISCUSSION

Robust NHE-1 Activity in Dendrites

In the current study, we characterized NHE-1 activity in the somata and the Lg- and the Sm-dendrites under normoxic and OGD/REOX conditions. We observed that pH_i regulation in the Lg- and the Sm-dendrites was ∼90–300% faster than in the somata under normoxic conditions. The differential pH_i regulation rates between the dendrites and the soma were abolished when they were corrected by the differences in the A/V ratios. Therefore, the data illustrate that the dendrites can change pH_i more rapidly than the soma due to the small cytosolic volume compared with its surface area. Robust NHE activity has previously been detected in the hippocampal nerve terminals following intracellular acidification under normoxic conditions (21). In our study, following OGD/REOX, the Na⁺-dependent H⁺ extrusion activity was further elevated in the soma (264%) and the Lg-dendrites (218%), whereas the A/V ratios remained unchanged.

The H⁺ extrusion mechanisms in the soma and the dendrites have not been well defined. In this study, we concluded that the somata pH_i regulation under HCO₃⁻-free conditions is exclusively mediated by NHE-1 activity, which is consistent with our previous findings using both HOE 642-mediated pharmacological inhibition and NHE-1 genetic knockdown approaches (11). Moreover, the significance of NHE-1 in neuronal ionic regulation is further highlighted by the abundant expression of NHE-1 compared with NHE-2, NHE-3, or NHE-5 in the neurons. This is consistent with the earlier reports on the preeminence of both NHE-1 mRNA and protein expression in brains over the isoforms NHE-2 to -4 and the abundance of NHE-3 in the cerebellum (19, 22).

We report here that NHE-1 plays a dominant role in the regulation of dendritic pH_i (60%). The remaining pH_i regulation in the dendrites depended on the functions of the less abundant NHE isoforms (NHE-2 and NHE-5) and vacuolar H⁺-ATPases. The H⁺ pumps are highly expressed in the vesicles of synaptic terminals and responsible for acid loading and accumulation of neurotransmitters (23). Although the H⁺ pumps are typically expressed in membranes of organelles, they have been detected in the plasma membrane of hippocampal astrocytes and are active in pH_i regulation under Na⁺- and HCO₃⁻-free conditions (24, 25). Our findings of H⁺ pump activity in the dendritic plasma membrane suggest that dendrites are equipped with multiple H⁺ extrusion mechanisms to counteract the robust Ca²⁺-dependent intracellular acidification during synchronous neural activity (26).

Recently, spatial nonuniformity in pH_i has been reported in the proximal and distal dendrites of oligodendrocytes (27). The alkaline microdomains in the perikaryon and proximal dendrites of the oligodendrocytes were attributed to localized increases in NHE activity, whereas the acidic pH_i in the distal dendrites may be the result of Na⁺/HCO₃⁻ cotransporter-mediated HCO₃⁻ extrusion (27). Thus, the pH_i microdomain and regulatory mechanisms in the oligodendrocyte distal dendrites appear to be different from those in the Sm-dendrites of neurons. These findings suggest that different cell types express different pH_i-regulating mechanisms in regulating microdomain pH_i.

Lack of NHE-1 Activation in Sm-dendrites following OGD/REOX

The basal level of NHE-1-mediated H⁺ extrusion was high in the Sm-dendrites compared with soma. OGD/REOX did not further stimulate it. This suggests that, given their large A/V ratio and their higher basal J_H⁺, Sm-dendrites are able to maintain pH_i without significant further elevation of NHE activity. This may also be a reflection of the sophisticated pH_i regulatory mechanisms expressed in Sm-dendrites, preventing overstimulation of H⁺ extrusion. On the other hand, this Sm-dendritic J_H⁺ phenomenon may be unique under the circumstances of culture models in plastic plates but not characteristic of dendrites under in vivo conditions. Indeed, we have observed much higher A/V ratios and faster pH_i recovery rates in neurites grown in microfluidic devices, a model that mimics the slow diffusion and convection of the in vivo microenvironment (28). It remains to be investigated whether OGD/REOX affects NHE-1 function differently in the Sm-dendrites using the microfluidic device model.

p90^RSK-mediated Stimulation of NHE-1 Activity following OGD/REOX

In the current study, OGD/REOX-mediated stimulation of NHE-1 activity was abolished in both the soma and the dendrites when NHE-1 kinase p90^RSK was inhibited by its potent inhibitors BI-D1870 and FMK. Activation of p90^RSK and NHE-1 phosphorylation is downstream of the ERK1/2 signaling pathways (8). FMK is a novel, specific inhibitor for p90^RSK isoforms 1 and 2 (17). FMK blocks the α₁-adrenocepter-mediated NHE-1 phosphorylation and stimulation in rat ventricular myocytes (29). On the other hand, BI-D1870 has been shown to be a potent ATP-competitive inhibitor of all p90^RSK isoforms (20). In this study, we found that FMK and BI-D1870 were equally effective in inhibiting OGD/REOX-mediated NHE-1 activation, implying a role for p90^RSK isoforms 1 and 2. Moreover, the p90^RSK inhibitors reduced the NHE-1 activity to below the base-line levels. This suggests that p90^RSK is also involved in the regulation of basal NHE-1 activity in all three regions.

NHE-1-mediated Na⁺ Entry following OGD/REOX

Disruption of dendritic ionic homeostasis occurs during early cerebral ischemia and may play a role in irreversible dendritic damage. Excessive Na⁺ influx via ionotropic glutamate receptors or tetrodotoxin-sensitive Na⁺ channels leads to neuronal death under excitotoxic or hypoxic conditions (4, 30, 31). However, subsequent studies have suggested that hypoxia-induced Na⁺ influx could be through pathways other than ionotropic glutamate receptors or tetrodotoxin-sensitive channels (32). In cultured hippocampal neurons, Na⁺ entry immediately after anoxia results from activation of NHE and a Gd³⁺-sensitive pathway (33). Recent reports demonstrate that dendritic damage following brief in vivo ischemia (34) or axonal morphological changes following in vitro hypoxia (35) are independent of ionotropic glutamate receptor activation. In the present study, we observed that OGD/REOX triggered an ∼3-fold increase in [Na⁺]_i (∼50 mm). The Na⁺ accumulation was eliminated when NHE-1 activity was inhibited by its inhibitor HOE 642 or the p90^RSK kinase inhibitor BI-D1870. Thus, the elevated NHE activity in the dendrites not only accelerates pH_i recovery after OGD/REOX but also intensifies disruption of Na⁺ ionic homeostasis and causes dendritic vulnerability to ischemic damage. These findings also suggest that Na⁺/K⁺-ATPase function is not sufficient to maintain Na⁺_i homeostasis following OGD/REOX when there is an increase in Na⁺ influx. Blocking NHE-1 activity would decrease the need for Na⁺ extrusion via Na⁺/K⁺-ATPase and preserve cellular ATP levels (Fig. 11). This imbalance between Na⁺ extrusion via Na⁺/K⁺-ATPase and NHE-1-mediated Na⁺ influx can also have a significant impact on [Na⁺]_i in Sm-dendrites, particularly because of their large A/V ratio and high basal J_H⁺, even without further elevation of NHE-1 activity following OGD/REOX. Taken together, we conclude that the OGD/REOX-mediated stimulation of NHE-1 plays a dominant role in dendritic Na⁺ overload.

FIGURE 11.

Illustration of dendritic ionic disruption and mitochondrial dysregulation in ischemic neurons. Following ischemia, activation of NHE-1 causes an increase in dendritic [Na⁺]_i, which triggers NCX_rev and leads to increases in [Ca²⁺]_i. The [Na⁺]_i overload also causes increased consumption of ATP by Na⁺/K⁺-ATPase to maintain dendritic ionic homeostasis. On the other hand, the [Ca²⁺]_i overload stimulates Ca²⁺_m uptake by the uniporter (UP) and formation of the mitochondrial permeability transition pore (PTP). Blocking NHE-1 and NCX_rev would reduce disruption of Na⁺ and Ca²⁺ homeostasis and preserve mitochondrial bioenergetics and dendritic membrane integrity.

We failed to detect elevation of NHE-1-mediated H⁺ extrusion in the Sm-dendrites after OGD/REOX. The causes for the discrepancy between the NHE-1-mediated H⁺ extrusion and Na⁺ overload in the Sm-dendrites are not apparent. One possible explanation is that a subtle increase in NHE-1 activity may not be detected with the instantaneous measurement of H⁺ extrusion (dpH_i/dt), whereas its impact on Na⁺ overload over time (at a steady-state level) can be revealed. This speculation is also supported by HOE 642-sensitive effects on mitochondrial dysfunction and Ca²⁺ dysregulation. Future study is needed to further address this issue.

NHE-1-dependent Changes in Dendritic Ca²⁺ following OGD/REOX

Activation of NHE activity in hippocampal nerve terminals following intracellular acidification is accompanied with an elevation in [Na⁺]_i and [Ca²⁺]_i as well as increased postsynaptic currents (21). The authors attribute these changes to the concurrent activation of NHE and NCX_rev in the nerve terminals (21). These findings suggest that NHE and NCX_rev could play a coordinating role in the regulation of dendritic Na⁺ and Ca²⁺ homeostasis and affect Ca²⁺-dependent release of neurotransmitters. In dendrites and dendritic spines close to postsynaptic localities, all three isoforms of NCX (NCX-1 to -3) are preferentially expressed, suggesting a role for NCX in Ca²⁺ signaling at the excitatory postsynaptic sites (36).

In this study, 2-h OGD triggered a moderate elevation in dendritic Ca²⁺_i, but, during 60 min of REOX, a delayed accelerated Ca²⁺_i rise occurred. The OGD/REOX-induced Ca²⁺ deregulation was initiated in the dendrites and then propagated to the soma. Interestingly, the secondary Ca²⁺_i deregulation can be prevented when either NHE-1 activity was inhibited by HOE 642 or NCX_rev was blocked by SEA0400. These findings imply that a coupled NHE-1 and NCX_rev function is a major contributor to Ca²⁺_i deregulation in the dendrites of cultured cortical neurons. Moreover, we believe that NCX-1 is the dominant NCX isoform in this study because SEA0400 has a high affinity against the reverse mode function of NCX-1 (IC₅₀ ∼50 nm) as compared with NCX-2 (IC₅₀ ∼1 μm) and is ineffective against either NCX-3 or NCKX-2 (37).

The role of NCX_rev in dendritic Ca²⁺_i deregulation has also been examined during sustained NMDA exposure. Delayed Ca²⁺ deregulation in CA1 neurons of acute hippocampal slices depended on Na⁺ loading but was not prevented by the nonspecific NCX inhibitor KB-R7943 (38). However, when Na⁺ loading is potentiated with low levels of ouabain (30 μm), NMDA can trigger secondary Ca²⁺ deregulation in dendrites, which is completely blocked by KB-R7943 (39). These findings further suggest that activation of NCX_rev requires excessive Na⁺ loading. Our previous thermodynamic analysis predicts that NCX_rev occurs when [Na⁺]_i is elevated to ∼20 mm in cortical neurons at a resting membrane potential of −60 mV (40).

The vulnerability of neuronal dendrites is characterized by the initial membrane depolarization, mitochondrial structure collapse, and dendritic beading in the dendrites and the subsequent propagation toward the soma during hypoxia and activation of NMDA receptors (1, 6). Our current study illustrates that blocking of NHE-1 activity attenuated many similar changes in the dendrites following OGD/REOX. Therefore, concerted activation of NHE-1 and NCX_rev may also play a role in dendritic injury in conditions such as glutamate-mediated neurotoxicity, epilepsy, etc.

Changes of Dendritic Ψ_m and Ca²⁺_m following OGD/REOX

In the current study, depolarization of Ψ_m in the Sm-dendrites occurred 40 min earlier than the soma following OGD/REOX. The loss of Ψ_m in the dendrites closely correlated with the Ca²⁺_m accumulation. Mitochondria are capable of sequestering large amounts of Ca²⁺ under various pathological conditions (41). Increases in free mitochondrial Ca²⁺ would occur when Ca²⁺ entry into the mitochondria exceeds the capacity of mitochondrial Ca²⁺ extrusion and the mitochondrial robust phosphate buffering system (42). The Ca²⁺_m accumulation reported in this study probably reflects Ca²⁺ entry via a voltage-dependent Ca²⁺ uniporter before the collapse of Ψ_m (41). Small increases in Ca²⁺_m stimulate Ca²⁺-dependent dehydrogenases and mitochondrial metabolism, but massive Ca²⁺ loading of mitochondria leads to depolarization of Ψ_m (6, 42). Sustained loss of Ψ_m will eventually trigger the opening of the permeability transition pore and release of Ca²⁺_m and collapse of mitochondria bioenergetics (6). In the current study, most of the Ca²⁺_m accumulation occurred before Ψ_m decreased below 50% and remained at a sustained level when Ψ_m was ∼45% of control in the soma and ∼20% of control in the Sm-dendrites. This implies that a residual level of Ψ_m (20%) for a short period (∼40 min) is sufficient to maintain high mitochondrial Ca²⁺ levels. However, when low Ψ_m and high Ca²⁺_m were extended past 60 min, there was a sudden loss of residual Ψ_m that coincided with the loss of the rhod-2 signal, suggesting a release of Ca²⁺_m from the permeability transition pore under these conditions.

Interestingly, when either NHE-1, p90^RSK, or NCX_rev was inhibited during 60-min REOX, loss of Ψ_m and Ca²⁺_m accumulation in dendrites was significantly reduced. This finding is consistent with the earlier reports on NHE-1 inhibition-induced attenuation of the mitochondrial Ca²⁺ overload and mitochondrial permeability transition pore opening in cardiomyocytes and in ischemic/reperfused rat hearts (43–45). Taken together, these studies demonstrate a conserved role of the NHE-1 signaling mechanism in ischemic reperfusion injury among multiple cell types. Moreover, it has been suggested that NHE-1 inhibitors, including HOE 642, have a direct effect on reactive oxygen species production and mitochondrial permeability transition pore formation (13). In the current study, it is unknown whether any protective effects mediated by HOE 642 result from its direct actions on mitochondria. We speculate that such a possibility is low in light of the similar protective effects offered by inhibition of p90^RSK or NCX_rev in this study as well as the protection observed in NHE-1 transgenic knock-out neurons (11). Moreover, in general, mild acidosis can inhibit neurotransmission, whereas alkaline pH_i stimulates excitability. Thus, we cannot rule out that HOE 642 may protect neurons in part via directly correcting NHE-1-mediated alkalinization.

In summary (Fig. 11), the current study reports that NHE-1-mediated Na⁺ entry and subsequent stimulation of NCX_rev activity contribute to the selective vulnerability of dendrites following in vitro ischemia. A newly emerging hypothesis speculates that dendritic Na⁺ overload and the subsequent activation of Na⁺/K⁺-ATPase would consume more ATP and further collapse mitochondrial biogenesis (6). However, to date, the mechanisms underlying the excessive Na⁺ influx and mitochondrial dysfunction are not well defined. Our current study demonstrates that activation of NHE-1 in dendrites presents a major pathway for Na⁺ overload. NHE-1 inhibition prevents Na⁺ accumulation, which is required for dendritic beading. Blocking NHE-1 function also attenuates loss of the dendritic Ψ_m and Ca²⁺_m homeostasis and preserves mitochondrial bioenergetics and dendritic membrane integrity.