• chloride;
  • macrophage;
  • mice;
  • potassium;
  • proton;
  • α-synuclein.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Brain mononuclear phagocyte (perivascular macrophage and microglia, MG) inflammatory neurotoxins play a principal role in the pathogenesis of Parkinson’s disease; chief among these are reactive oxygen species (ROS). We posit that aggregated, misfolded and oxidized α-synuclein (a major constituent of Lewy bodies), released or secreted from dying dopaminergic neurons, induces microglial ROS production that is regulated by ion channels and as such affects disease progression. To address this hypothesis, we performed patch clamp recordings of outward ionic currents in murine microglia and characterized their links to ROS production during α-synuclein stimulation. Aggregated nitrated α-synuclein induced ROS production in a dose-dependent manner that was inhibited by voltage-gated potassium current blockade, and to a more limited degree, by chloride current blockade. Interestingly, ROS produced in MG primed with tumor necrosis factor alpha and activated with phorbol myristate acetate was attenuated by voltage-gated potassium current blockade and more completely by chloride current blockade. In contrast, amyloid beta or cell membrane extract failed to induce microglial ROS production. Similar results were obtained using bone marrow-derived macrophages. The association of ROS production with specific plasma membrane ion currents provides a link between regulation of microglial ion transport and oxygen free radical production. Understanding these linkages may lead to novel therapeutics for Parkinson’s disease where modulation of redox-related stress may slow disease progression.

Abbreviations used



bone marrow-derived macrophages


buthionine sulfoximine




Dulbecco’s Modified Eagles Media


flufenamic acid


Hanks’ balanced salt solution


delayed rectifier K+ channel


inward-rectifier K+ current


macrophage colony stimulating factor




mononuclear phagocyte


niflumic acid


5-nitro-2-(3-phenylpropylamino) benzoic acid


Parkinson’s disease


phorbol myristate acetate


reactive oxygen species


substantia nigra pars compacta


tumor necrosis factor alpha

Parkinson’s disease (PD) is a devastating neurodegenerative disorder that manifests clinically as motor and gait disturbances including rigidity, resting tremor, slowness of voluntary movement and postural instability, and in some cases, dementia (Dauer and Przedborski, 2003). Pathologically, PD is characterized by the progressive loss of midbrain dopaminergic neurons in the substantia nigra pars compacta (SNpc) along with the loss of their terminals in the dorsal striatum. Of emerging importance, dopaminergic neuronal injury is associated with microglial activation and neuroinflammation (McGeer et al. 1988). Under steady-state conditions, MG serve a sentry role, responding to injury with a stereotypic response that includes phagocytosis of cellular debris, presentation of antigen, and secretion of a plethora of immune factors that orchestrate the innate inflammatory response and recruit leukocytes to injured or diseased brain areas. Moreover, microglial activation occurs throughout the disease course, suggesting that it contributes to nigrostriatal degeneration (Vila et al. 2001; Teismann et al. 2003; McGeer and McGeer 2004).

While the inciting events (environmental toxins, aging and genetics) for idiopathic PD are unknown, oxidative stress is likely a central component in the disease process (Jenner 2003; Przedborski et al. 2003; Andersen 2004). The relationship between redox state and neurodegeneration is underscored by the fact that following microglial activation, glutathione (GSH) levels are decreased by 40–50% in the SNpc of PD patients (Sofic et al. 1992; Sian et al. 1994). This decrease in GSH, an important redox buffer in mammalian cells (Dringen 2000), is an early indicator of oxidative stress in PD (Nakamura et al. 1997).

A major histopathological hallmark of sporadic PD is the presence of Lewy bodies, intracellular inclusions consisting primarily of aggregated proteins and lipids, in degenerating neurons (Goedert et al. 1998; Duda et al. 2000; Hurtig et al. 2000; Galvin et al. 2001; Shults 2006). The major constituent of these inclusions is α-synuclein, a protein normally concentrated in pre-synaptic terminals (Jakes et al. 1994). While the conditions that lead to the abnormal aggregation of the α-synuclein in sporadic PD are debated, a role for oxidative stress has been posited that involves the oxidative nitration of the molecule, forming covalently bonded dimers which lead to accelerated aggregation of the protein (Souza et al. 2000; Paxinou et al. 2001; Krishnan et al. 2003). In fact, antibodies directed against nitrated α-synuclein clearly demonstrate staining of Lewy bodies and other inclusions in PD and other synucleinopathies (Giasson et al. 2000).

Microglial activation and phagocytic activity is accompanied by the production of reactive oxygen species (ROS) such as superoxide anion (O2) and hydrogen peroxide (H2O2) (Colton and Gilbert 1987). ROS production in mononuclear phagocytes (MP; macrophages, dendritic cells and MG) is mediated by the membrane bound electrogenic enzyme complex, NADPH-oxidase (Babior 1999, 2004). Many factors, including pro-inflammatory cytokines that are elevated during neuroinflammatory processes, influence the activity of NADPH-oxidase.

In bone marrow-derived MP, the phorbol ester-stimulated production of superoxide is greatly enhanced by pre-incubation with tumor necrosis factor alpha (TNF-α) (Phillips and Hamilton 1989). The generation of ROS by MP is accompanied by plasma membrane depolarization and cytosolic acidification (DeCoursey 2004). Production of ROS thus requires both a compensatory movement of charge across the plasma membrane and buffering or removal of the acid load, raising the question of which ion channel species are required for sustained production of ROS and subsequent increased oxidative stress during neuroinflammatory events. The few studies performed that address the role of ion channels in ROS production in MG support a role for voltage- or Ca2+-activated potassium channels (Spranger et al. 1998; Khanna et al. 2001). However, a comprehensive study of ionic currents involved in mediation and regulation of MP ROS production has not been previously performed despite the importance of these cells in the pathogenesis of neurodegenerative diseases and the role of oxidative stress in disease progression. In the current work, we examined the roles of plasma membrane ionic currents in the generation of ROS in activated murine MP. We found that aggregated α-synuclein could induce ROS production in murine MP whereas, other potential phagocytic stimuli were not effective. We compared the ion channel sensitivity of ROS production induced by aggregated α-synuclein with that induced by TNF-α priming and phorbol ester stimulation, the latter a model for generalized activation of MG in the course of a neuroinflammatory response. We show that inhibition of specific ionic currents affects ROS production in murine MP, including novel evidence consistent with a role for a plasma membrane chloride conductance. However, the relative importance of potassium and chloride currents varied depending on the activating stimuli. Understanding the mechanisms that regulate the activity of these ion channels could provide therapeutic tools for altering the progression of neurodegeneration by modulating ROS production. In addition, characterizing the response of the glutathione redox buffer system to factors that affect ROS production may ultimately provide insights into the mechanisms that regulate the balance between production and removal of ROS in the course of PD.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Macrophage and microglia cultures

Bone marrow-derived macrophages (BMM) and microglia (MG) were prepared from C57BL/6 adult male (6–12 weeks old) and neonatal mice (1–2 days old), respectively (Charles River Laboratories Inc., Wilmington, MA, USA). All tissue harvest procedures followed National Institute of Health (NIH)-established guidelines and were approved by the University of Nebraska Medical Center Institutional Animal Care and Use Committee. The animals were anesthetized with isoflurane and killed by decapitation. Bone marrow cells were obtained from femurs after the marrow cavities were flushed with Hanks’ balanced salt solution (HBSS). After dissociation of red cells with ACK lysis buffer (0.15 mol/L NH4Cl, 10 mmol KHCO3, 0.1 mmol Na2EDTA, pH 7.2), the cell suspension was passed through a 40 µm filter, spun at 93 × g for 5 min, and the pellet resuspended in complete medium (Dulbecco’s modified Eagles media (DMEM) supplemented with 10% fetal bovine serum, 2 mmol/L-glutamine, and 1% penicillin/streptomycin) containing 2 µg/mL macrophage colony stimulating factor (MCSF), a generous gift from Wyeth Pharmaceuticals (Cambridge, MA, USA). The bone marrow cells were cultured in complete medium in a 5% CO2/ 37°C incubator. Non-adherent cells were removed from flasks at 1, 4, and 7 days by successive DMEM washes. The cells were replated for experiments following 7–14 days in culture.

Microglia were prepared using previously described techniques (Dobrenis 1998). Brains were removed and placed in HBSS at 4°C. The tissue was dissociated using a 10-mL plastic pipette and then incubated in 0.25% trypsin at 37°C for 30 min. After adding cold, heat-inactivated fetal bovine serum, the tissue was washed several times with cold HBSS. The tissue was then triturated by pipetting through a series of sterile Pasteur pipettes with reduced bores and then filtered through a 40 µm filter. These mixed glial cells were cultured in complete medium containing 2 µg/mL MCSF. To obtain highly purified MG, beginning 7 days after harvest, the culture flasks were gently shaken and the supernatants containing floating MG were transferred to new flasks. The flasks were incubated for 30 min to allow the MG to adhere, and loose cells were removed by washing with DMEM. These MG cells were cultured in complete medium containing MCSF for 7–14 days and then replated for experiments.

Whole-cell patch clamp recording

Bone marrow-derived macrophages and MG were removed from flasks by scraping, suspended in complete media with MCSF, and plated onto sterile glass cover slips within 12-well culture plates. BMM were plated at a density of 106 cells/well and MG were plated at a density of 105 cells/well. Electrophysiological recordings were performed 1-7 days following replating. Cover slips were transferred one at a time to a recording chamber (RC-13, Warner Instrument Corp., Hamden, CT, USA) mounted on the fixed stage of a Nikon Eclipse E600FN microscope. Cells were continuously perfused via gravity-fed reservoirs. Whole-cell patch clamp recordings were made using an Axopatch 200B patch clamp amplifier (Axon Instruments, Foster City, CA, USA) interfaced with a Digidata 1322A hardware data acquisition system (Axon Instruments) and controlled with pClamp Version 8.1 software (Axon Instruments) on a Dell microcomputer. Currents were filtered at 1 kHz with a four-pole low-pass Bessel filter and digitized at a sampling interval of 100 µs. Patch pipettes were fabricated from borosilicate glass capillary tubing (WPI, Sarasota, FL, USA) using a Sutter P97 microelectrode puller, with tip resistances of 3–6 MΩ. Immediately following break-in to whole-cell mode, pipette access resistance, membrane resistance and membrane capacitance were measured and recorded. Following experimental recordings, passive membrane properties were remeasured to assure that the quality of the recording was not altered during data acquisition.

To measure voltage-dependent potassium and proton currents, whole-cell currents were recorded in response to a series of voltage steps from −160 mV to +100 mV in 20 mV steps for 1000 ms at 10 s intervals, from a holding potential of −60 mV. For these experiments, the pipette solution contained (in mmol) 130 K+ gluconate, 10 EGTA, 1 CaCl2, 1 MgCl2, and 10 HEPES (pH 7.2, 280 mOsm). The bathing solution contained (in mmol) 135 Na+ gluconate, 5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose (pH 7.4, 290 mOsm). To measure stretch-activated chloride currents, whole-cell currents were recorded in response to a series of voltage steps from 0 mV to +40 mV in 10 mV steps at 10 s intervals, from a holding potential of −10 mV. For these experiments, the pipette solution contained (in mmol) 130 KCl, 1 EGTA, 0.3 CaCl2, 2 Mg ATP, 0.2 Tris GTP, and 10 HEPES (pH 7.2, 280 mOsm). The bathing solution was the same as above except NaCl (135 mmol) replaced Na+ gluconate. To activate chloride current, the cells were perfused with a hypotonic solution with NaCl reduced to 115 mmol for a final osmolarity of 250 mOsm (85% of normal).

Measurement of membrane currents and drug effects

The inward rectifier potassium current was measured as the peak current evoked at the step to −140 mV. To estimate the magnitude of the transient outward potassium current, the peak current evoked during the step to +40 mV was subtracted from the current magnitude at the end of the 1000-ms step. To estimate the magnitude of the slow outward proton current, the peak current evoked at the end of the 1000-ms step to +100 mV was subtracted from the current magnitude at the beginning of the step. The stretch-activated chloride current was measured as the difference in current magnitude evoked at +40 mV and the holding current measured at −10 mV, after 5 min perfusion in hyposmotic solution. To account for initial leakage current, this current was then subtracted from the same differential current in control solution. Finally, changing the chloride concentration in the bathing solution from 144 mmol to 124 mmol (in the hyposmotic solution) would shift the chloride equilibrium potentia (ECl) by −3.8 mV. To account for the resulting 7.6% increase in driving force, the stretch-activated current was multiplied by 0.924 to compensate for this effect.

To calculate the effects of channel blocking agents on the voltage- and stretch-evoked currents, the current magnitudes evoked in the presence of the agents were measured and expressed as a percentage of the magnitudes measured in control solution. For statistical analysis, paired Student’s t-tests were used with statistical significance at p < 0.05.

Production, nitration, and aggregation of recombinant mouse α-synuclein

For α-synuclein production, a BL21 bacterial clone transformed with the his-tag containing expression vector, pMSYN, was obtained (kindly provided by Dr Eric Benner). A kanamycin plate was streaked overnight, and a single colony picked the next day and grown in 5 mL Luria Broth (LB) + 50 µg/mL kanamycin to confluence. Five microliter of culture was used to inoculate 500 mL aliquots of LB and grown until the culture measured an optical density at 600 nm (OD600) of 0.55. Cultures were induced for 4 h with 1 mmol isopropyl-beta-D-thiogalacto pyranoside (IPTG) and the bacteria centrifuged and lysed using BugBuster lysis buffer (Novagen, Madison, WI, USA). The his-tag containing mouse α-synuclein was then purified using NTA-binding columns (Qiagen, Valencia, CA, USA). The his-tag was removed using a thrombin-based capture kit (Novagen, Madison, WI, USA) and then incubated with polymyxin-coated beads (Sigma, St. Louis, MD, USA) to remove residual endotoxin. Samples were then dialyzed and lyophilized. To prepare nitrated α-synuclein, protein samples were rehydrated in phosphate-buffered saline to 2 mg/mL and peroxynitirite (100 µL/mg protein) was added dropwise while mixing. The nitrated protein was dialyzed, lyophilized, and stored at −80°C. To aggregate the protein, samples were rehydrated in a glycine-based buffer (80 mmol NaCl, 50 mmol glycine at pH 2.9) and then incubated in a hot plate at 54°C for 5 days. Protein quality was assessed at each step using either polyacrylamide gel electrophoresis or Western blotting methods. For visualization of the unaggregated and aggregated protein via atomic force microscopy, samples were deposited on mica, glued to a glass slide and dried under argon gas flow. The image was taken in air, in height, amplitude and phase modes using a Molecular Force Probe 3D controller (Asylum Research Inc., Santa Barbara, CA, USA).

Measurement of extracellular H2O2 production

Bone marrow-derived macrophages and MG were removed from flasks by scraping, suspended in complete media with MCSF and plated into 96-well culture plates at 200 μL/well. To measure ROS production, BMM and MG were cultivated at a density of 105 cells/well. H2O2 production was measured using the fluorescent dye, Amplex Red (Molecular Probes; Eugene, OR, USA). This dye is non-fluorescent in the reduced state and fluoresces with a peak at 590 nm when oxidized by H2O2 in the presence of horseradish peroxidase. Culture media was removed from the wells and replaced with 100 µL reaction mixture (0.1 U/mL horseradish peroxidase and 50 µmol Amplex Red in HBSS containing 1.0 mmol CaCl2 and 1.0 mmol MgCl2; final pH 7.4) with/without activators (as described below) and with/without channel blockers. Fluorescence measurements were taken using a SpectraMax Gemini fluorometer (Molecular Devices, Menlo Park, CA, USA) with Ex/Em = 530/590 nm at 37°C. Readings were taken immediately after adding reaction mixture (time = 0) and repeated at 15, 45 and 90 min. The plates were returned to the incubator in between measurements. The dye fluorescence was calibrated to [H2O2] by taking fluorescence measurements of reaction mixtures containing H2O2 concentrations from 0 to 10 µmol and generating a regression line relating [H2O2] to raw fluorescence values.

To measure α-synuclein-activated ROS production, aggregated or unaggregated protein was added to the reaction mixture from 100 µg/mL stock solutions. The pH of the reaction mixture was tested and was not altered significantly by the addition of aggregated protein stock at the highest concentration used (500 nmol). To measure phorbol myristate acetate (PMA)-stimulated ROS production, 48 h prior to the experiment the culture media in one half of the wells (48 wells) was replaced with complete media containing 200 ng/mL TNF-α and no MCSF (200 µL/well). The other 48 wells received fresh media without MCSF. ROS production was stimulated by adding 250 nmol/L PMA to the reaction mixture. For all experiments, 6 or 8 wells were used for each experimental group. The effects of channel blockers were expressed as a percentage of α-synuclein- or TNF-α + PMA-stimulated experimental groups. For statistical analysis of channel blocker effects, raw fluorescence readings were converted to log values and analyzed using a Mixed Model anova with statistical significance at p < 0.05. Experiments were performed at least thrice for each experimental group using tissue harvests from different animals.

Measures of oxidized and reduced glutathione

To investigate the effect of TNF-α on glutathione levels in MP, the culture medium was replaced with MCSF-free medium containing TNF-α (200 ng/mL final concentration) 24 h or 48 h prior to harvesting. Glutathione concentration was measured as described previously (Reed et al. 1980; Mosharov et al. 2000). In brief, the medium was aspirated and the cells washed with ice-cold saline. Then, 100 μL of cold phosphate-buffered saline was added to the plate, the cells harvested by scraping and the suspension homogenized with a pipette. One aliquot of the suspension was used to determine protein concentration using the Bradford reagent (Bio-Rad Laboratories, Hercules, CA, USA). The other aliquot was mixed with an equal volume of metaphosphoric acid solution (containing16.8 mg/mL HPO3, 2 mg/mL EDTA, and 9 mg/mL NaCl), incubated on ice for several minutes and centrifuged. The supernatant was treated with iodoacetate to block free sulfhydryl groups followed by derivatization of the amines with 2,4-dinitrofluorobenzene, which adds a chromophore. Reduced (GSH) and oxidized (GSSG) glutathione were separated by HPLC using a μ-Bondapak-NH2 300 × 3.9 mm column (Waters, Mildford, MA, USA) and eluted as described previously (Mosharov et al. 2000). The eluent was monitored at 355 nmol. The concentrations of GSH and GSSG were determined by comparison with a calibration curve generated with authentic samples of the same.

Buffers and solutions

All recording buffer and pipette solution constituents were purchased from Sigma. HBSS, DMEM and all culture media constituents were purchased from Invitrogen (Carlsbad, CA, USA). Amplex Red was purchased from Molecular Probes. PMA, buthionine sulfoximine (BSO) and all channel blocking agents were purchased from Sigma except charybdotoxin and kaliotoxin, which were purchased from Alomone Labs Ltd (Jerusalem, Israel). Channel blockers were prepared from stock solutions and dissolved in bathing solution or reaction mixture for electrophysiological recordings and ROS experiments, respectively.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

To study the role of membrane ion channels in MP ROS production, we induced murine MP activation using two paradigms that reproduce different aspects of the neuroinflammatory context that could occur in the progression of PD. We first examined the effects of specific ion channel blockade on ROS production following activation of MG with aggregated nitrated α-synuclein, likely encountered by MG surrounding degenerating dopaminergic neurons. We then compared this profile of ion channel sensitivity with that observed during ROS production in MG or BMM primed with the pro-inflammatory cytokine, TNF-α, and activated with the phorbol ester, PMA.

To address the linkages between plasma membrane ion currents and ROS production, we studied the expression patterns of several currents that have been identified in murine MP and their sensitivity to specific blocking agents. We focused on voltage-dependent and stretch-activated currents that could compensate for charge movement during a respiratory burst.

Voltage-dependent whole-cell currents

To study the voltage-dependent currents, whole-cell patch clamp recordings were performed from a total of 104 cells (BMM, n = 44; MG, n = 60). The mean (± SEM) resting membrane potential was −52 ± 18 mV (n = 18) for BMM and −65 ± 7 mV (n = 21) for MG. The mean membrane resistance was 1.0 ± 0.4 GΩ (n = 41) for BMM and 1.2 ± 0.5 GΩ (n = 54) for MG.

We measured whole-cell currents evoked by a series of voltage commands from −160 mV to +100 mV, from a holding potential of −60 mV. The most consistent voltage-dependent current observed in MP recordings was an inward current evoked at hyperpolarized membrane potentials. This current, evoked at membrane potentials more negative than the K+ equilibrium potential, showed properties consistent with an inward-rectifier K+ current (KIR) (Eder 1998; Walz and Bekar 2001). Thus, the inward conductance increased in magnitude, activated more rapidly, and exhibited inactivation, with increasing levels of hyperpolarization. The mean peak amplitude measured at −160 mV and normalized to cell capacitance was −14.9 ± 8.9 pA/pF (n = 39) for BMM and −20.9 ± 12.0 pA/pF (n = 51) for MG. The mean amplitude of KIR was significantly larger in MG cells compared to BMM (p < 0.01), which likely accounts for the larger resting membrane potential observed in MG cells (Eder 1998). The magnitude of KIR varied widely, ranging from 3.8 to 45.2 pA/pF for BMM and from 3.8 to 71.8 pA/pF for MG cells.

A second ion current was observed at depolarized membrane potentials that showed the properties of an inactivating delayed rectifier K+ channel (KDR) previously characterized in MPs (Eder 1998; Walz and Bekar 2001) and identified as a Kv1.3 type potassium channel (Khanna et al. 2001; Fordyce et al. 2005). This current was activated at membrane potentials above −40 mV and showed a partial inactivation during the prolonged depolarizing pulse (shown for MG cells in Fig. 1). This transient outward current was observed only with depolarizations from a −60 mV holding potential, showing steady-state inactivation when evoked from more depolarized holding potentials (data not shown). Finally, the transient current was blocked by 1 mmol 4-aminopyridine (4-AP) (0.5% of control, n = 5; p < 0.05) and 30 nmol carybdotoxin (CTX) (2.3% of control, n = 4; p < 0.01) (Fig. 1), consistent with this type of KDR current. The percentage of cells that expressed KDR current was significantly greater in microglial cultures when compared with BMM cultures [seven of 44 BM cells (16%) vs. 29 of 60 MG cells (48%), Fisher’s Exact Test, p < 0.001]. The KDR current, when present, showed a wide variation in magnitude, and in most cells where the current was distinctly present (as determined by kinetic properties), the magnitude was small. In MG cells, the peak current magnitude measured at +40 mV averaged 153 ± 77 pA and ranged from 48 pA to 340 pA (normalized to cell capacitance, 4.6 pA/pF, range 1.5-12.1 pA/pF).


Figure 1.  KDR channel currents expressed in bone marrow macrophage and microglia. Representative recordings from two different microglial cells before (control) and during perfusion with bathing solution containing (a) 1 mmol 4-aminopyridine (4-AP) or (b) 30 nmol charybdotoxin. Current traces show responses to 1000 ms steps from −60 mV to −40 mV through +40 mV at 20 mV intervals. Calibration bar: 500 ms, 300 pA (a), 100 pA (b). Histograms at right show cumulative data. The outward KDR current was completely inhibited by both drugs (4-AP, p < 0.05, n = 5 experiments; charybdotoxin, p < 0.01, n = 4 experiments).

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A third voltage-dependent current observed in MP was a slowly developing outward current evoked at strong depolarizing potentials. This current activated slowly over the course of the 1000 ms duration depolarization at membrane potentials of +60 mV and greater (shown for BM cells in Fig. 2). The current measured at +100 mV was reversibly inhibited by 100 µmol/L LaCl3 (54.9% of control, n = 5; p < 0.05) and completely and reversibly blocked by 100 µmol/L ZnCl2 (1.2% of control, n = 5; p < 0.01) (Fig. 2). The degree of blockade of this current by both cations was voltage-sensitive, with maximum effects observed at more depolarized potentials. The voltage sensitivity, kinetics, and ionic blocker sensitivity of this current are consistent with a proton-dependent current that has been described previously in brain macrophages (Eder 1998). This current was observed in 10 of 44 BMM (23%) and four of 60 MG cells (7%). The mean amplitude of the slow outward current, measured at +100 mV and normalized to cell capacitance, was 1.8 ± 1.7 pA/pF for BMM (n = 10) and 1.8 ± 1.3 pA/pF for MG cells (n = 4). The amplitude varied considerably across cells that expressed the current, ranging from 0.3 to 5.7 pA/pF for BMM and from 0.5 to 3.1 pA/pF for MG cells.


Figure 2.  H+ channel currents expressed in bone marrow macrophages and microglia. Representative recordings from two different bone marrow macrophages before (control), during, and following (wash) perfusion with bathing solution containing (a) 100 µmol ZnCl2 or (b) 100 µmol LaCl3. Current traces show responses to 1000 ms steps from −60 mV to +40 mV through +100 mV at 20 mV intervals. Calibration bar: 500 ms, 200 pA (a), 300 pA (b). Histograms at right show cumulative data. The slow outward current was completely abolished by ZnCl2 (p < 0.01, n = 4 experiments) and inhibited by LaCl3 (mean = 55% of control, p < 0.05, n = 4 experiments).

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Stretch-activated whole-cell currents

To study stretch-activated currents, we recorded microglial cells during perfusion with a hyposmotic bath solution, which led to cell swelling and plasma membrane stretching. Whole-cell patch clamp recordings were performed from 30 microglial cells with a mean membrane resistance of 1.2 ± 0.7 GΩ. For these experiments, the recordings were performed at a holding potential of −10 mV (near the predicted chloride equilibrium potential), to inactivate any transient outward K+ current that might be present and to eliminate steady shifts in chloride ion during the course of the experiment. Under these conditions, outward current evoked by depolarizing commands was minimal in normosmotic bath solution (Fig. 3a, control). Perfusion with a hyposmotic solution (nominally 250 mOsm, 85% of normal) resulted in the development of an outwardly rectifying current (Fig. 3a, hyposmotic). This outward current increased slowly during a 5 min perfusion in hyposmotic solution, and persisted for at least 5 min after returning to control solution (Fig. 3a, 5 min normal) when recordings were performed with pipettes containing 2 mmol ATP. In preliminary recordings performed using pipettes without ATP, stretch-activated outward currents decayed within several minutes (data not shown). Thus, all experiments characterizing channel blocker effects were performed using pipettes containing ATP.


Figure 3.  Stretch-activated Cl channel currents expressed in microglia. (a) Representative recordings from a microglial cell before (control) and 5 min following perfusion with a hyposmotic bathing solution (253 mOsm) showing activation of an outwardly rectifying current. The right-most trace (5 min normal) shows recordings made following return to control solution for 5 min. Current traces show responses to 1000 ms steps from −10 mV to 0 mV through +40 mV at 10 mV intervals. Calibration bar: 500 ms, 500 pA. The outward current was inhibited by (b) 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) (mean 3.5% of control, p < 0.01, n = 6 experiments), (c) niflumic acid (NFA) (mean 30.3% of control, p < 0.05, n = 5 experiments) and (d) flufenamic acid (FNA) (200 µmol, mean 44.3% of control, p < 0.01, n = 5 experiments; 500 µmol, mean 2.4% of control, p < 0.001, n = 5 experiments). Histograms at right show cumulative data.

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Figure 5.  Aggregated α-synuclein stimulates reactive oxygen species (ROS) generation in mononuclear phagocyte cultures. (a) Incubation of microglial cultures for 90 min with aggregated α-synuclein causes a dose-dependent increase in accumulation of extracellular H2O2. Histogram shows mean values ± SEM for 50 nmol (n = 4; p < 0.001), 100 nmol (n = 6; p < 0.001), and 500 nmol (n = 3; p < 0.001) aggregated α-synuclein. ROS production in microglial cultures incubated with unaggregated α-synuclein was not significantly different from controls. Incubation of microglial cultures for 90 min with (b) unaggregated (UA) or aggregated (AA) amyloid-β protein over a range of 1-30 µmol (n = 2 each condition) or (c) membrane homogenate over a range of 1-50 µg/mL (n = 3 each condition) did not result in a significant increase in ROS production in microglial cultures. For both amyloid-β protein and membrane homogenate experiments, microglial cultures produced significant ROS in response to 250 nmol phorbol myristate acetate following incubation in 200 ng/mL tumor necrosis factor-α for 48 h.

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Stretch-activated outward currents were present in all microglial cell recordings (n = 30). This was the most robust whole-cell current recorded in murine MP, with an average magnitude of 1141 ± 480 pA and a range from 400 to 2388 pA. The outward current was inhibited by several chloride transport inhibitors. Thus, 200 µmol 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) reduced the stretch-activated current to 3.5% of control (Fig. 3b, n = 6, p < 0.01) and 200 µmol niflumic acid (NFA) reduced the current to 30.3% of control (Fig. 3c, n = 5, p < 0.05). Flufenamic acid (FNA) reduced the current to 44.3% of control at 200 µmol (Fig. 3d, n = 5, p < 0.01) and to 2.4% of control at 500 µmol (Fig. 3d, n = 5, p < 0.001). The activation characteristics and drug sensitivities of this outward current are consistent with a stretch-activated chloride current that has been described previously in rodent microglial cells (Schlichter et al. 1996; Eder et al. 1998).

Reactive oxygen species production and ion channel blockade

We determined whether acute blockade of specific ionic currents affected α-synuclein or PMA-stimulated ROS production. As generation of ROS is accompanied by plasma membrane depolarization and cytosolic acidification (DeCoursey 2003a), blockade of depolarization-activated currents might inhibit the sustained production of ROS by abolishing the compensatory repolarization of the membrane that would occur when these conductances are activated. Additionally, a proton conductance is activated by cytosolic acidification (Eder 1998) and its inhibition could thus lead to uncompensated pH changes. Activation of MP is also accompanied by elevations in cytosolic Ca2+ concentration. In human macrophages, a chloride conductance activated by cytosolic Ca2+ has been observed (Holevinsky et al. 1994), and chloride channels are known to modulate store-operated Ca2+ influx in human MG (McLarnon et al. 2001). In rodent MP, chloride channels activated by membrane stretching have been identified (Schlichter et al. 1996; Eder et al. 1998; Eder 2005). The currents mediated by these channels can be inhibited by several agents including NPPB and FNA (Schlichter et al. 1996). Thus, we also examined the effects of chloride channel inhibitors on ROS production. For all experiments measuring ROS accumulation, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assays were performed at the end of the experiment and no toxicity was observed in the cultures following exposure to any channel blocker at the concentrations used.

α-Synuclein stimulated reactive oxygen species production and effects of ion channel blockade

We examined ROS production in response to stimulation of MG with aggregated α-synuclein, the major protein constituent of Lewy bodies that are observed frequently in degenerating dopaminergic neurons in PD brain. Other authors (Zhang et al. 2005) and ourselves posit that MG are exposed to this protein from dead or dying dopaminergic neurons and may serve as an activating stimulus that drives the production of ROS, leads to increased oxidative stress and damages healthy neurons, thus hastening the progression of PD.

Purified mouse recombinant α-synuclein was first nitrated and then aggregated for 5 days as described in Materials and methods. Figures 4(a)–(c) show examples of nitrated protein, unaggregated and aggregated, visualized by atomic force microscopy. Analysis of protein aggregation was also assessed semi-quantitatively: particle heights were measured and categorized as either spherical oligomers (2–6 nm in height), protofibrils (1.5–3 nm in height), or fibrils (5–8 nm in height) (Apetri et al. 2006). Samples of the nitrated protein consisted only of oligomers (100%; 27 total particles) prior to aggregation (Figs 4a and d). Following 5 days of aggregation, the percentage of oligomers decreased (to 70%; 47 total particles) and protofibrils appeared (25%; 15 total particles), along with a small percentage of fibrils (3 particles) (Figs 4b–d). Thus, a shift in particle distribution occurred during aggregation, indicating that the protein applied to the microglial cultures consisted predominantly of oligomeric species along with some protofibrils. Western blot analysis demonstrated that the protein contained nitrotyrosine residues in dimeric and larger aggregates following aggregation for 5 days (Fig. 4e). Further, protein samples subjected to nitration contained larger molecular weight aggregates following 5 days of aggregation and less monomeric protein (Fig. 4f).


Figure 4.  Nitrated α-synuclein forms aggregates following 5 days at low pH and high temperature. (a) Atomic force micrograph of unaggregated nitrated mouse α-synuclein. (b) Atomic force micrograph of nitrated mouse α-synuclein following 5 days at 54°C, and at higher magnification (c). Samples in (a–c) were dissolved in a pH 2.7 glycine buffer at a concentration of 100 µg/mL. (d) Distribution of particle type prior to and following 5 days aggregation. (e) Western blot using anti-nitrotyrosine antibody of nitrated protein sample following 5 days aggregation as in (b). (f) Western blot using anti-α-synuclein antibody of nitrated protein sample (left) and native protein sample (right) following 5 days aggregation as in (b).

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Application of aggregated α-synuclein to microglial cultures for 90 min caused a dose-dependent increase in the accumulation of H2O2 (Fig. 5a; 50 nmol: 141% of control, n = 4, p < 0.001; 100 nmol: 153% of control, n = 6, p < 0.001; 500 nmol: 143% of control, n = 3, p < 0.001). In contrast, ROS production in microglial cultures incubated with unaggregated α-synuclein was not significantly different from controls (Fig. 5a). We compared α-synuclein stimulation of ROS production with amyloid-β, another protein known to cause activation of MG. Incubation of microglial cultures for 90 min with unaggregated or aggregated amyloid-β protein over a range of 1-30 µmol (n = 2) did not result in a significant increase in ROS production over control cultures (Fig. 5b). Further, incubation of microglial cultures with brain membrane homogenates (to simulate phagocytosis of neuronal debris) over a range of 1-50 µg/mL (n = 3) also did not result in a significant increase in ROS production in microglial cultures (Fig. 5c). For both amyloid-β protein and membrane homogenate experiments, microglial cultures produced significant ROS in response to 250 nmol PMA following incubation in 200 ng/mL TNF-α for 48 h (Figs 5b and c).

The generation of H2O2 in response to aggregated α-synuclein was partially inhibited by blockade of chloride, potassium, or proton dependent ion currents (Fig. 6). Co-application of the chloride channel blockers NPPB (200 µmol) or NFA (200 µmol) with 100 nmol aggregated α-synuclein resulted in a 16% and 18% decrease in accumulation of H2O2, respectively, relative to protein alone (Fig. 6). The effects of potassium channel blockade were somewhat larger; thus, co-application of margatoxin (10 nmol) or CTX (30 nmol) with 100 nmol aggregated α-synuclein resulted in a 27% and 32% decrease in accumulation of H2O2, respectively, relative to protein alone (Fig. 6). Finally, co-application of the proton channel inhibitor ZnCl2 (100 µmol) with 100 nmol aggregated α-synuclein resulted in a 24% decrease in accumulation of H2O2 relative to protein alone (Fig. 6).


Figure 6.  Blockade of ion channels inhibits α-synuclein-induced reactive oxygen species (ROS) production. In microglial cultures, H2O2 accumulation was inhibited by co-application of 100 nmol aggregated α-synuclein and the chloride channel blockers, 5-nitro-2-(3-phenylpropylamino) benzoic acid (200 µmol; n = 3) or niflumic acid (200 µmol, n = 3), the potassium channels blockers, margatoxin (10 nmol, n = 3) or charybdotoxin (30 nmol, n = 3), and the proton channel blocker, ZnCl2 (100 µmol, n = 3).

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α-Synuclein and mononuclear phagocyte ion channel expression

To determine whether the differences in ion channel blocker sensitivities observed between PMA- and α-synuclein-stimulated ROS production were because of effects of these agents on ion channel expression, we performed patch clamp recordings from MG that were acutely exposed to aggregated α-synuclein protein (90 min to 3 h). However, this prior exposure did not significantly alter the expression levels of the ionic currents observed in these cells. Thus, 33% (4/12) of microglial cells acutely exposed to 100 nmol aggregated α-synuclein expressed the voltage-dependent KDR current compared with 38% (3/8) of control cells from the same cultures, and 11% (3/27) of cells expressed a proton current compared with 7% (4/60) control cells (Table 1). Acute application of aggregated α-synuclein also had no significant effect on the expression (Table 1) or magnitude of the stretch-activated chloride current. Thus, all cells exposed acutely to 100 nmol aggregated α-synuclein expressed the chloride current, with an average amplitude of 965 ± 402 pA (n = 12) compared with 1141 ± 480 pA (n = 30) for cells incubated in normal media (Student’s t-test, p = 0.27).

Table 1.   Tumor necrosis factor-α (TNF-α) and mononuclear phagocyte ion channel expression
CurrentControlTNF-α (24 h)TNF-α (48 h)
(a) BMM: TNF-α
KDR7/44 (16%)4/10 (40%)2/10 (20%)
H+10/44 (23%)5/10 (50%)7/10 (70%)*
CurrentControlTNF-α (24 h)TNF-α (48 h)
CurrentControlTNF-α (24 h)TNF-α (48 h)
(b) Microglia: TNF-α
KDR29/60 (48%)7/15 (47%)8/32 (25%)
H+4/60 (7%)8/15 (53%)**7/32 (22%)**
Cl30/30 (100%) 6/6 (100%)
  1. (a) Cumulative data for bone marrow macrophage recordings showing numbers of cells expressing each current type in control recordings and following 24 h or 48 h incubation with 200 ng/mL TNF-α. Proton current expression was significantly increased following 48 h TNF-α incubation (Fisher’s Exact Test, p < 0.05)*. (b) Cumulative data for microglial recordings. Proton current expression was significantly increased following both 24 and 48 h TNF-α incubation (Fisher’s Exact Test, p < 0.05)**. (c) Cumulative data for microglial recordings following acute incubation with 100 nmol aggregated α-synuclein for 90 min to 3 h.MP, mononuclear phagocyte; BMM, bone marrow-derived macrophages.

(c) Microglia: α-synuclein
KDR3/8 (38%)4/12 (33%)
H+4/60 (7%)3/27 (11%)
Cl30/30 (100%)12/12 (100%)

Phorbol myristate acetate-stimulated reactive oxygen species production in tumor necrosis factor-α-primed mononuclear phagocyte

Bone marrow-derived MP (Phillips and Hamilton 1989) and MG (Colton et al. 1992) can be stimulated to produce ROS with PMA, which activates NADPH-oxidase. In isolated cultures of murine bone marrow-derived MP, PMA-stimulated production of superoxide is enhanced by preincubation with the pro-inflammatory cytokine, TNF-α (Phillips and Hamilton 1989). We measured the accumulation of H2O2 for 90 min in control cultures of BMM and MG stimulated with PMA compared with cultures that had been pre-incubated with TNF-α for 48 h (Fig. 7). As reported previously (Phillips and Hamilton 1989), PMA-stimulated ROS production was greatly enhanced by TNF-α pre-incubation in BMM cultures, as shown in a representative experiment in Fig. 7(a). The absolute values of H2O2 accumulation after 90 min varied considerably, averaging 6.8 ± 0.7 µmol/well and ranging from 1.9 to 12.7 µmol/well for PMA-stimulated, TNF-α pre-incubated BMM cultures. The effect of TNF-α pre-incubation on ROS production was also observed in microglial cultures (shown for a representative experiment in Fig. 7b), demonstrating that brain-derived MP can also be induced to produce ROS under conditions observed during an active inflammatory state. The accumulation of H2O2 in microglial cultures was similar to that observed in BMM cultures, averaging 7.0 ± 0.8 µmol/well and ranging from 1.9 to 10.8 µmol/well for PMA-stimulated, TNF-α pre-incubated MG cultures after 90 min.


Figure 7.  Tumor necrosis factor-α enhances phorbol myristate acetate-stimulated mononuclear phagocyte reactive oxygen species production. Graphs show representative experiments measuring H2O2 production in bone marrow macrophage (a) and microglia (b). At the initiation of the experiment, culture media was removed from 96-well plates and replaced with reaction media (control) or reaction media containing 250 nmol phorbol myristate acetate. Half of the wells were incubated in 200 ng/mL tumor necrosis factor-α for 48 h. Data points represent mean ± SEM for n = 6 wells per group for each experiment. These graphs are representative of control groups used for all experiments using ion channel blockers; for each channel blocker group at least three plates from three different cell harvests were used (n = 6 wells per group) for both bone marrow macrophage and microglial cultures.

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The NADPH-oxidase inhibitor, diphenylene iodonium (10 µmol), completely blocked H2O2 production (Fig. 8), demonstrating that the generation of ROS in murine macrophages is entirely dependent on the activity of this membrane-bound enzyme complex. However, incubation of BMM cultures with BSO, which inhibits glutathione synthesis, led to increased production of ROS (data not shown). Thus, co-incubation with TNF-α and 10 μmol or 30 μmol BSO, 48 h prior to PMA stimulation, increased H2O2 production to 117% and 133% of controls, respectively.


Figure 8.  Blockade of potassium, chloride and proton currents inhibits phorbol myristate acetate-stimulated mononuclear phagocyte reactive oxygen species production. Histograms show inhibition of reactive oxygen species production in (a) bone marrow macrophage cultures and (b) microglial cultures. Values are expressed as mean percent inhibition relative to control values for TNF-α incubated, phorbol myristate acetate-stimulated groups. Channel blockers used were: 100 µmol LaCl3; 100 µmol ZnCl2 (proton current inhibitors); 30 nmol charybdotoxin (CTX); 43 nmol kaliotoxin (KTX) (KDR current inhibitors); 1 mmol 4-aminopyridine (4-AP); 200 µmol niflumic acid (NFA); 200 µmol 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB); 200 µmol and 500 µmol flufenamic acid (FNA) (Cl current inhibitors). The NADPH-oxidase inhibitor, diphenylene iodonium (DPI, 10 µmol) completely blocked H2O2 production (n = 2). All drug groups in bone marrow macrophage experiments differed from controls (Mixed Model ANOVA: LaCl3p < 0.05; ZnCl2, p < 0.05; CTX, p < 0.001; KTX, p < 0.001; 4-AP, p < 0.05; NFA, p < 0.0001; NPPB, p < 0.05). For the microglial experiments, ZnCl2 (p < 0.05), KTX (p < 0.01), NFA (p < 0.001), NPPB (p < 0.001), and FNA (200 µmol and 500 µmol, p < 0.001) groups were significantly different from controls. ( ) indicate number of different plates per group, n = 6 wells/group; error bars indicate ± SEM. (c) ROS production is inhibited in low chloride solutions. NaCl was replaced with Na Gluconate in the reaction buffer to reduce chloride to 50% (75 mmol Na gluconate and 70 mmol NaCl) or 5% (145 mmol Na gluconate and no NaCl) of normal. Histogram shows cumulative H2O2 production (mean ± SEM) normalized to control (100% chloride) for three separate experiments.

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Ion channel blockade and phorbol myristate acetate-stimulated reactive oxygen species production

Blockade of the KDR conductance led to moderate, but significant, inhibition of PMA-stimulated ROS production in BMM cultures (Fig. 8a). Thus, 30 nmol CTX and 1 mmol 4-AP reduced H2O2 accumulation by 35% (n = 7) and 36% (n = 8), respectively. Additionally, 43 nmol kaliotoxin, which also blocks KDR channels, reduced H2O2 accumulation by 49% (n = 4). The inhibition of proton conductance also led to a significant inhibition of PMA-stimulated ROS production by BMM. Thus, 100 µmol LaCl3 and 100 µmol ZnCl2 reduced H2O2 accumulation by 29% (n = 6) and 20% (n = 10), respectively. However, blockade of chloride conductance had the greatest effect on PMA-stimulated ROS production in BMM. Thus, 200 µmol NPPB and 200 µmol NFA reduced H2O2 accumulation by 88% (n = 4) and 66% (n = 4), respectively.

Similar effects of the channel blockers were obtained in experiments measuring PMA-stimulated ROS production in microglial cultures (Fig. 8b). Thus, inhibition of KDR with 30 nmol CTX, 1 mmol 4-AP, and 43 nmol kaliotoxin reduced H2O2 accumulation by 20% (n = 5), 10% (n = 3), and 32% (n = 5), respectively. However, of the KDR blockers, only the effect of kaliotoxin was statistically different from controls. Inhibition of proton conductance with 100 µmol LaCl3 and 100 µmol ZnCl2 reduced H2O2 accumulation by 12% (n = 5) and 22% (n = 7), respectively, with only the ZnCl2 inhibition reaching statistical significance. As with BMM, the effects of chloride channel blockers had the most pronounced effects on PMA-stimulated ROS production in MG cultures. NPPB (200 µmol) significantly reduced H2O2 accumulation by 79% (n = 4), NFA (200 µmol) significantly reduced H2O2 accumulation by 68% (n = 6), and FNA significantly reduced H2O2 accumulation by 29% at 200 µmol (n = 4) and 85% at 500 µmol (n = 4).

Since blockade of chloride conductance had the largest effect on ROS production in TNF-α-primed MP, we performed additional experiments where the extracellular concentration of chloride ions in the reaction buffer was reduced. If chloride flux is necessary for sustained ROS production, this manipulation should also result in decreased ROS accumulation. Indeed, when the chloride concentration was lowered to 50% or 5% of normal, ROS production was reduced to 63% and 62% of controls, respectively (Fig. 8c; n = 3, p < 0.05).

TNF-α and mononuclear phagocyte ion channel expression

As we utilized TNF-α incubation to prime the production of ROS in our MP cultures, it was important to establish what effects, if any, this cytokine has on ion current expression patterns. As TNF-α has been reported to transiently up-regulate the expression of KDR (McLarnon et al. 2001; Vicente et al. 2003), we recorded from cells incubated in TNF-α for 24 or 48 h. The percentage of BMM cells expressing KDR current increased transiently after 24 h of TNF-α incubation, and returned to control levels after 48 hr TNF-α exposure (Table 1); however, the increase was not statistically significant. This transient up-regulation was not observed in microglial cells, where the percentage of cells expressing KDR was higher in control cultures (Table 1).

TNF-α incubation significantly increased the expression of the proton current in both BMM and MG cells (Table 1). The proton current was present in 23% (10/44) of control BMM cells, increasing to 50% (5/10) and 70% (7/10) of cells following 24 and 48 h in TNF-α, respectively (Table 1). Thus, a sustained increase in expression of the proton current was observed in BMM cells following TNF-α exposure. The expression of the proton current was also increased in MG following TNF-α exposure. The current was present in 7% (4/60) of control cells, 53% (8/15) of cells exposed to TNF-α for 24 h, and 22% (7/32) of cells exposed to TNF-α for 48 h (Table 1). Thus, in microglial cells, proton current expression increased transiently after 24 h of TNF-α exposure, but still remained elevated following 48 h exposure relative to unstimulated cells.

TNF-α pre-incubation had no significant effect on the expression (Table 1) or magnitude of the stretch-activated chloride current. Thus, all cells exposed for 48 h to TNF-α still expressed the chloride current, with an average amplitude of 1081 ± 241 pA (n = 6) compared with 1141 ± 480 pA (n = 30) for cells incubated in normal media (Student’s t test, p = 0.76).

TNF-α and intracellular glutathione

The average glutathione concentration in BMM cells incubated in the presence of MCSF was 41 ± 2.6 μmol/g of protein (mean± SEM, n = 11; ‘control’ in Fig. 8). The glutathione concentration increased slightly 24 h after incubating cells in medium without MCSF, and returned to the initial value after 48 h (Fig. 9). In contrast, in the presence of TNF-α, glutathione levels significantly increased to ∼170% after 24 h compared with untreated controls (93.2 ± 12.3 vs. 54.8 ± 3.2 μmol/g of protein) and to 235% after 48 h (95.3 ± 11.9 vs. 40.5 ± 3.5 μmol/g of protein; Fig. 9). PMA stimulation of MP incubated for 48 h in medium with or without TNF-α elicited a small (20-30%) increase in glutathione concentration within 30–60 min. This was followed by a slow decrease in glutathione concentration (data not shown).


Figure 9.  Tumor necrosis factor alpha (TNF-α) increases mononuclear phagocyte glutathione production. Comparison of intracellular glutathione concentrations in bone marrow-derived macrophages treated with 200 ng/mL TNF-α for 24 or 48 h vs. untreated controls. Values are mean ± SEM, n = 11 for control cells cultured with macrophage colony stimulating factor, n = 14 at 24 h and n = 7 at 48 h for untreated cells, n = 10 at 24 h and n = 7 at 48 h for TNF-α treated cells. The results for untreated and TNF-α treated cells were significantly different with p < 0.002.

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  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Oxidative stress plays a major role in the progressive destruction of SNpc dopaminergic neurons in PD. Growing evidence suggests that MG may contribute to redox stress in disease by producing ROS during phagocytosis of debris from dying and degenerating neurons (Green et al. 2001; Block et al. 2004; Zhang et al. 2005). As compensatory ionic fluxes are required to sustain ROS generation, we characterized the relative roles of plasma membrane ion currents in this process, with the long-term goal of abrogating neuronal damage by modulating these fluxes during PD-associated microglial activation. We reasoned that dopaminergic neuronal degeneration is accompanied by microglial activation in response to material, especially α-synuclein, released from dead and dying cells (Thomas et al. 2006). Thus, initial neuronal insult could lead to a positive feedback cycle of oxidative damage and accelerated progression of the disease. This hypothesis is supported by a wealth of published data. First, monomeric and aggregated α-synuclein can be released from cells in culture, and this release is elevated with proteasomal and mitochondrial dysfunction (Lee et al. 2005). Second, oligomeric α-synuclein levels are increased in the blood of PD patients (El-Agnaf et al., 2006). Third, extracellular Lewy bodies are observed in PD tissue surrounded by activated MG (McGeer et al. 1988; Yamada et al. 1992). Fourth, aggregated α-synuclein can activate rodent MG resulting in the production of ROS and a neurotoxic phenotype (Zhang et al. 2005).

In the current work we characterized the relative roles of several ionic currents in the generation of ROS induced by aggregated α-synuclein in murine bone marrow macrophage and MG. It is conceivable that, depending on the neuroinflammatory context within which microglial cells are found, different ionic conductances may play varying roles that depend on unique signaling pathways. Thus, we compared the ion channel sensitivity of ROS production induced by aggregated α-synuclein with ROS production induced by direct activation of NADPH-oxidase. Direct activation of the oxidase with PMA following priming with TNF-α served as a control to elicit a more generalized inflammatory cascade. In microglial cultures stimulated with aggregated α-synuclein, ROS production was moderately reduced by blockade of voltage-activated K+ or proton currents, whereas blockade of chloride currents played a smaller role. Whereas, the most significant effects observed for PMA-stimulated ROS production resulted from blockade of chloride currents, while blockade of either voltage-activated K+ or proton current resulted in moderate effects on ROS generation. Overall, our results are consistent with a role for several plasma membrane ion channels in mediating compensatory ion fluxes during ROS generation. The relative roles of these currents appear to depend on the activation stimulus, suggesting that different signaling pathways involving ionic conductances may be engaged in the course of neuroinflammatory responses. The links between ROS production, microglial activation and specific plasma membrane ion channel activities provide a step towards understanding how ion transport in MP lineage cells can modulate oxygen-free radical production induced during inflammation and neurodegenerative processes.

Reactive oxygen species production by MP is accompanied by plasma membrane depolarization and cytosolic acidification, regardless of whether translocation of negative charge is directed into phagosomes or across the plasma membrane into the extracellular space (DeCoursey 2004). In fact, uncompensated electron translocation by NADPH-oxidase would result in a rapid depolarization of the membrane to levels (+200 mV) that would prevent further electron translocation and ROS production (DeCoursey 2004). Thus, sustained ROS production requires a continuous compensatory movement of positive charge outward across the plasma membrane. Prior studies linking ROS production and plasma membrane ion currents primarily used human neutrophils or eosinophils, as they are capable of large respiratory bursts during the process of engulfing and killing microbial targets (Ahluwalia et al. 2004; DeCoursey 2004). It is likely that the voltage- and pH-dependent proton conductance present in all phagocytes plays a significant role in ROS generation (Eder and DeCoursey 2001; DeCoursey 2003b; Eder 2005). A role for other ion currents including potassium, chloride, or non-specific cations has been hypothesized by others (Holevinsky et al. 1994; Holevinsky and Nelson 1995; Schmid-Antomarchi et al. 1997; Ahluwalia et al. 2004). The few studies performed using murine MG appear to support a role for voltage- or Ca2+-activated potassium channels (Spranger et al. 1998; Khanna et al. 2001; Fordyce et al. 2005). The definitive establishment of roles for ion currents may be complicated by the fact that, in addition to charge compensation, another function of these ionic conductances may be to compensate or regulate pH changes or volume changes that accompany the respiratory burst (Eder 2005).

While a proton-dependent plasma membrane conductance appears ubiquitous in MP (Eder and DeCoursey 2001; DeCoursey 2003b), the expression levels of this conductance vary widely. This may be related to the relative potential of different MP phenotypes to produce ROS. It is, therefore, interesting that we observed an increase in expression of the proton-dependent current following TNF-α-induced MP activation, which significantly increased the potential of the cells to generate ROS. This is consistent with a role for this conductance in compensating for the ion flux accompanying ROS production. The evidence that micromolar concentrations of La3+ and Zn2+, agents that inhibit the proton current, lead to decreased ROS production support this idea. The fact that we did not observe a more complete inhibition of ROS production by these cations might be partly explained by the voltage-dependent nature of their blockade. Thus, the membrane potential may not reach the levels of depolarization required for a complete blockade of the current by these cations during a respiratory burst. In fact, the primary stimulus for activating this current during ROS generation may be a decrease in pH, which shifts the threshold for activation of this conductance to more hyperpolarized levels (Eder 1998). Alternatively, the proton conductance may not play as important a role in rodent MP as has been hypothesized for human leukocytes (DeCoursey 2004).

Previous studies have provided evidence that support a role for voltage- or Ca2+-activated potassium channels in ROS production in murine MP (Spranger et al. 1998; Khanna et al. 2001; Fordyce et al. 2005). Consistent with our results, Khanna et al. (2001), using a single-cell fluorescence assay, demonstrated that nanomolar concentrations of CTX inhibited ROS production in rat MG. However, the level of inhibition by CTX was greater than observed in our study. More recently, a specific role for the Kv1.3 type potassium channel in ROS production by rat microglial cells was demonstrated by the same laboratory (Fordyce et al. 2005). It is possible that the greater effects of potassium channel blockade in these studies compared to our study are because of differences in channel expression. The proportion of cells expressing transient outward current, and the mean magnitude of this current, appear to be greater in unstimulated rat MG (Fordyce et al. 2005) compared with murine cells. Another possibility that may account for the differences is that rat MG may express higher levels of BK-type Ca2+-activated potassium channels than murine cells (Eder 1998). These channels are also sensitive to blockade by CTX. In any case, the differences are quantitative and results from both studies support a role for the transient outward KDR in ROS production. However, it will be of great interest to determine the expression levels of this channel in brain MP in the context of specific neurodegenerative disorders, to further evaluate its role in ROS production in different diseases.

Our results are consistent with a role for plasma membrane chloride current in maintenance of ROS production in murine MP, especially following activation by pro-inflammatory agents. Our data show a robust expression of chloride current in murine MG, and the effects of several chloride channel inhibitors on the stretch-activated current, closely parallel their effects on ROS production. Thus, 200 µmol NPPB and 500 µmol FNA abolished the chloride current and strongly inhibited ROS production, while 200 µmol NFA had more moderate effects on both the ion current and ROS production. Further, decreasing extracellular chloride led to decreases in ROS accumulation, consistent with a role for chloride ion flux in compensation for membrane potential changes during ROS generation.

There is some indirect evidence supporting a role for Cl conductances in ROS production. In human neutrophils, a sustained Cl efflux is causally related to the respiratory burst stimulated by TNF-α (Menegazzi et al. 1999). Since activation of MP is accompanied by elevations in intracellular Ca2+, it is interesting to note that Cl channels modulate store-operated Ca2+ influx in human MG (McLarnon et al. 2000). A Cl current activated by cytosolic Ca2+ was observed in human monocyte-derived macrophages (Holevinsky et al. 1994) and a chloride channel activated by NADPH-oxidase-derived H2O2 has been recently identified in a liver-derived cell line (Varela et al. 2004). As observed in our recordings, a chloride conductance activated by membrane stretching was identified in murine (Eder et al. 1998) and rat (Schlichter et al. 1996) microglial cells. Thus, in rodent MG, one likely stimulus for the Cl current is cell swelling induced by the acidosis that follows activation of a respiratory burst (Morihata et al. 2000). It will be interesting to determine what factors, besides membrane stretching, regulate Cl conductances present in rodent MP. The different effects of chloride channel blockade observed between PMA-stimulated and α-synuclein-stimulated ROS production may reflect differential activation of this current as the expression of the channel itself does not appear to be changed by these alternate activation pathways. Our data suggests that ROS generation induced by α-synuclein-stimulated activity does not engage plasma membrane chloride conductances as robustly as that observed following PMA-induced activation of the NADPH-oxidase. Nevertheless, we demonstrate here for the first time a role for membrane chloride conductance in murine MP ROS production.

TNF-α induces an increase in glutathione concentration in rat hepatocytes and in human hepatoma cells by transcriptional activation of the catalytic subunit of γ-glutamylcysteine ligase, the rate-limiting enzyme in glutathione synthesis (Morales et al. 1997; Zou and Banerjee 2003). TNF-α was found to elicit a similar effect in MP, where intracellular glutathione levels were increased ˜twofold after 24 h and remained elevated after 48 h. The increased intracellular glutathione concentration in cells exposed to TNF-α correlated with increased PMA-stimulated generation of ROS production and enhanced channel expression. It will be of interest to determine if the enhanced antioxidant capacity of MP in TNF-α-treated cells confers an increased capacity for ROS generation with a concomitant increase in expression of channels that serve to compensate for the accompanying charge flux. Furthermore, if changes in the ambient redox potential, viz. under pathophysiological conditions, can modulate the magnitude of the respiratory burst, these changes could be linked to the etiology of neuronal dysfunction.

The balance between ROS generation and buffering in cellular compartments dictates the overall redox state in a brain region, a balance which is tipped chronically towards oxidative stress in the course of PD. During an inflammatory process, MP undergo changes that increase their capacity to both produce and buffer ROS, rendering them more capable of causing neuronal injury or death. In future studies it will be important to distinguish between mechanisms that are essential to the signaling functions of redox reactive species, and those that are up-regulated during MP activation and mediate neurotoxic functions. Inhibition of the latter, while leaving the former mechanisms intact, would then form a rational therapeutic approach for PD and other neurodegenerative disorders.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

This work was supported in part by grants from the NIH (P20 RR17675 to MPT, R37 NS036126, P01 NS31492, R01 NS034239, P20 RR15635, and P01 NS043985 to HEG and DK64959 to RB). The authors would like to thank Drs Lee Mosley and Pawel Ciborowski and Ms. Robin Taylor for critically reading the manuscript.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
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