neuroprotectant minocycline depresses glutamatergic neurotransmission ... · neuroprotectant...

Neuroprotectant minocycline depresses glutamatergicneurotransmission and Ca2+ signalling in hippocampalneurons

Jose Carlos Gonzalez,1,2 Javier Egea,1,2 Marıa del Carmen Godino,3 Francisco J. Fernandez-Gomez,5

Jose Sanchez-Prieto,3 Luıs Gandıa,1,2 Antonio G. Garcıa,1,2,4 Joaquın Jordan5 and Jesus M. Hernandez-Guijo1,2

1Instituto Teofilo Hernando, and2Departamento de Farmacologıa y Terapeutica, Facultad de Medicina, Universidad Autonoma de Madrid, Arzobispo Morcillo 4,E-28029 Madrid, Spain3Departamento de Bioquımica, Facultad de Veterinaria, Universidad Complutense, Madrid, Spain4Servicio de Farmacologıa Clınica, Hospital Universitario de la Princesa, Facultad de Medicina, Universidad Autonoma de Madrid,Madrid, Spain5Grupo de Neurofarmacologıa, Departamento de Ciencias Medicas, Facultad de Medicina, Universidad Castilla-La Mancha yCentro Regional de Investigaciones Biomedicas, Albacete, Spain

Keywords: glutamate release, synaptic transmission, tetracycline

Abstract

The mechanism of the neuroprotective action of the tetracycline antibiotic minocycline against various neuron insults is controversial.In an attempt to clarify this mechanism, we have studied here its effects on various electrophysiological parameters, Ca2+ signalling,and glutamate release, in primary cultures of rat hippocampal neurons, and in synaptosomes. Spontaneous excitatory postsynapticcurrents and action potential firing were drastically decreased by minocycline at concentrations known to afford neuroprotection. Thedrug also blocked whole-cell inward Na+ currents (INa) by 20%, and the whole-cell Ca2+ current (ICa) by about 30%. Minocyclineinhibited glutamate-evoked elevation of the cytosolic Ca2+ concentration ([Ca2+]c) by nearly 40%, and K+-evoked glutamate releasefrom synaptosomes by 63%. Minocycline also depressed the frequency and amplitude of spontaneous excitatory postsynapticcurrents, but did not affect the whole-cell inward current elicited by c-aminobutyric acid or glutamate. This pharmacological profilesuggests that the neuroprotective effects of minocycline might be associated with the mitigation of neuronal excitability, glutamaterelease, and Ca2+ overloading.

Introduction

In the last few years, controversial data have accumulated concerningthe neuroprotective effects of the tetracycline antibiotic minocycline(Blum et al., 2004; Domercq & Matute, 2004; Stirling et al., 2005;Jordan et al., 2007). Thus, minocycline has been shown to affordprotection against brain ischaemia (Yrjanheikki et al., 1999), excito-toxicity (Tikka et al., 2001), and spinal cord injury (Stirling et al.,2004). However, other studies have shown no effects, or evenexacerbated neurotoxicity (Yang et al., 2003; Diguet et al., 2004). Thereasons for such opposite effects remain to be determined. It isnevertheless interesting that minocycline affords neuroprotection inanimal models of focal and global cerebral ischaemia (Wang et al.,2003), Huntington’s disease (Chen et al., 2000), amyotrophic lateralsclerosis (Zhu et al., 2002), Alzheimer’s disease (Hunter et al., 2004),and Parkinson’s disease (He et al., 2001), suggesting a therapeuticneuroprotective potential of minocycline in neurodegenerativedisorders.

Several intracellular signalling pathways have been implicated inthe mechanism of the neuroprotective actions of minocycline. Forinstance, the involvement of antioxidant systems (Kraus et al.,2005), the inhibition of poly(ADP-ribose) polymerase-1, an enzymeactivated by DNA damage that promotes both cell death andinflammation (Alano et al., 2006), or the blockade of inflammatorypathways (Tomas-Camardiel et al., 2004) that involve nitric oxidesynthase (Sadowski & Steinmeyer, 2001), caspase-1 (Chen et al.,2000), matrix metalloproteinases (Mathalone et al., 2007), attenu-ation of tumour necrosis factor-a expression (Lee et al., 2004),inhibition of p38 mitogen-activated protein kinase (Lin et al., 2001),or upregulation of cyclo-oxygenase-2 and prostaglandin E2 (Atturet al., 1999). Another mechanism is linked to inhibition ofmitochondrial depolarization and blockade of permeability transitionpore opening, thus preventing the release of cytochrome c andactivation of caspase-3 (Fernandez-Gomez et al., 2005). Addition-ally, inhibition of microglial activation may represent an importantstep in the prevention of neuronal injury (Choi et al., 2005).Whether or not the neuroprotective property of minocycline has yetbeen accepted, the concentration of minocycline in the extracellularfluid of the brain that would correlate with this neuroprotectionremains unknown.

Correspondence: Dr J. M. Hernandez-Guijo, Departamento de Farmacologıa yTerapeutica, as above.E-mail: [email protected]

Received 15 March 2007, revised 3 September 2007, accepted 4 September 2007

European Journal of Neuroscience, Vol. 26, pp. 2481–2495, 2007 doi:10.1111/j.1460-9568.2007.05873.x

ª The Authors (2007). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd

Drugs that disrupt intracellular ion concentrations might result incell death or cell protection. The abnormal elevation of cytosolic Ca2+

concentration ([Ca2+]c) and the overactivation of glutamate receptorsare the two main candidates for a final common pathway in neuronaldeath (Mattson, 2003). Thus, blockers of voltage-dependent Ca2+-channels [see Kochegarov (2003) for a review] or of ionotropicglutamate receptor channels [see Danysz & Parsons (2002) for areview] can prevent neuronal Ca2+ overload and display cytoprotec-tive effects against exocytotoxic insults.In light of these data, we addressed the hypothesis that

minocycline might exert its neuroprotective actions by interactingwith receptors or ion channels involved in glutamatergic transmis-sion. Combining patch-clamp techniques with Ca2+ imaging andglutamate release, we found that minocycline decreased neuronexcitability by blocking voltage-dependent Na+-channels; thus, thiseffect caused a decrement in spontaneous action potential firing andneurotransmitter release. Furthermore, we observed that minocyclineblocked voltage-dependent Ca2+-channels, and decreased the gluta-mate-elicited [Ca2+]c elevations, which evoked a reduction inglutamate release. Hence, depression of glutamatergic neurotrans-mission may explain the neuroprotective effects of minocyclineagainst neurotoxicity. To our knowledge, this is the first electro-physiological study aimed at determining a basis for the neuropro-tective effects of minocycline.

Materials and methods

Isolation and culture of rat hippocampal neurons

All experiments were carried out in accordance with the guidelinesestablished by the National Council on Animal Care and wereapproved by the local Animal Care Committee of the UniversidadAutonoma de Madrid (Spain). Pregnant Sprague-Dawley rats werekilled by decapitation, and 18-day embryos were immediatelyremoved by caesarean section. Hippocampi were dissected rapidlyunder a stereomicroscope in cold phosphate-buffered saline [in mm:137 NaCl, 2.7 KCl, 8.1 NaH2PO4, 1.4 KH2PO4 (pH 7.4) (4 �C)]under sterile conditions. The tissue was digested with 0.5 mg ⁄ mLpapain and 0.25 mg ⁄ mL DNase. The enzymes were dissolved in aCa2+- and Mg2+-free phosphate-buffered saline solution containing1 mg ⁄ mL bovine serum albumin and 6 mm glucose at 37 �C for20 min. The papain solution was replaced with 5 mL of Dulbecco’sModified Eagle’s Medium (DMEM) supplemented with 10% fetalbovine serum. The digested tissue was then gently triturated by suctionusing a glass pipette flamed on the tip to avoid cellular damage. Thecell suspension was centrifuged for 4 min at 120 g. The supernatantwas removed, and the cells were resuspended in 5 mL of DMEM andplated at a density of 1 · 105 cells ⁄ mL on 4-cm-diameter Petri dishes(2 mL of DMEM per dish) coated with poly(d-lysine) (0.1 mg ⁄ mL).Cells were plated in DMEM (4500 mg ⁄ L glucose and 580 mg ⁄ Ll-glutamine) supplemented with 10% fetal bovine serum, 50 lg ⁄ mLstreptomycin–penicillin and 50 lg ⁄ mL gentamicin, and maintained ina 5% CO2 incubator at 37 �C. After 24 h, the medium was replacedwith fresh serum-free medium, to avoid fibroblast proliferation, butcontaining B27 supplement, essential for hippocampal neuron survivalin vitro. Under these conditions, the standard cell survival was4 weeks; the experiments were performed on neurons after11–15 days in culture.The criteria for cell selection for current recordings were adhesion

to the substrate, soma diameters of 15–30 lm, neuronal shape with noevident shrinkage or swelling, neurite extensions, and absence ofintracellular vacuoles and bright cell bodies with dark boundaries.

Cell viability studies

Coverslips containing hippocampal neurons were rinsed twice withKrebs–Hepes buffer solution [in mm: 140 NaCl, 5.9 KCl, 2.5 CaCl2,1.2 MgCl2, 10 glucose, 15 Hepes ⁄ NaOH (pH 7.4)] and exposed for20 min to glutamate (100 lm) at room temperature. Cellular deathwas determined using a fluorescein diacetate ⁄ propidium iodidedouble-staining procedure (Jordan et al., 2003). Twenty-four hoursafter exposure to glutamate, hippocampal neurons were incubated for45 s at 22–25 �C with 15 lg ⁄ mL fluorescein diacetate and4.6 lg ⁄ mL propidium iodide in phosphate-buffered saline [in mm:100 Na2HPO4, 100 NaH2PO4, 140 NaCl (pH 7.4)]. The stained cellswere examined with a standard epi-illumination fluorescence micro-scope (Axiophot, Zeiss, Germany). Cells stained with propidiumiodide represented dead cells, and cells stained with fluoresceindiacetate represented live cells. A blinded observer counted thenumber of dead and live cells in five microscopic fields (under 40·magnification), reaching approximately 300–450 cells for eachcoverslip, and the mean was considered to be the representative valuefor the coverslip. The percentage of surviving cells was determined in3–4 coverslips for each experimental condition and normalized tocontrols.

Current recordings, data acquisition and analysis

Spontaneous synaptic transmission and ionic currents were recordedusing the whole-cell configuration of the patch-clamp technique(Hamill et al., 1981); action potentials (APs) were recorded using theperforated-patch whole-cell recording technique (Korn et al., 1991).Whole-cell recordings were made with fire-polished electrodes(resistance 2–5 MW when filled with the standard intracellularsolutions) mounted on the headstage of an EPC-10 patch-clampamplifier (HEKA Electronic, Lambrecht, Germany), allowing cancel-lation of capacitative transients and compensation of series resistance.Data were acquired with a sample frequency ranging between 5 and10 kHz and filtered at 1–2 kHz. Recording traces with leak currents>100 pA or series resistance >20 MW were discarded.The perforated patch was obtained using pipettes containing

50 lg ⁄ mL amphotericin B (Sigma, Madrid, Spain) and a pipette-filling solution containing (in mm): 10 NaCl, 135 KCl, 5 Mg-ATP,0.3 Na-GTP, 14 EGTA, and 20 Hepes ⁄ KOH (pH 7.3). Amphoteri-cin B was dissolved in dimethylsulphoxide and stored at )20 �C instock aliquots of 50 mg ⁄ mL. Fresh pipette solution was preparedevery 2 h. To facilitate sealing, the pipette was first dipped in abeaker containing the internal solution and then back-filled with thesame solution containing amphotericin B. Pipettes with series resis-tance of 2–3 MW were used to form gigaseals. Recording startedwhen the access resistance decreased below 15 MW, which usuallyhappened within 10 min after sealing (Rae et al., 1991). Seriesresistance was compensated by 80% and monitored throughout theexperiment.Petri dishes containing the cells were placed on an experimental

chamber mounted on the stage of a Nikon eclipse T2000 invertedmicroscope. During the preparation of the seal with the patch pipette,the chamber was replaced with a control Tyrode solution containing(in mm): 137 NaCl, 1 MgCl2, 2 CaCl2, 4 KCl, 10 glucose, and10 Hepes ⁄ NaOH (pH 7.4). Once the patch membrane was rupturedand the whole-cell configuration of the patch-clamp technique hadbeen established, the cell being recorded was locally, rapidly andcontinuously superfused with an extracellular solution of identicalcomposition to the chamber solution (see Results for specificexperimental protocols). The neuron was constantly superfused, and

2482 J. C. Gonzalez et al.

ª The Authors (2007). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing LtdEuropean Journal of Neuroscience, 26, 2481–2495

the external solutions were rapidly exchanged through a gravitysystem consisting of a multibarrelled pipette with a single outlet andfive inlets controlled by solenoid electrovalves, the common outlet ofwhich was placed within 100 lm of the cell to be patched. The flowrate was 1 mL ⁄ min.

Currents were obtained by superfusing the whole-cell clampedpostsynaptic neuron with a Tyrode solution; tetrodotoxin (0.3 lm) wasadded, when required, to block spontaneous AP propagation; glycine(10 lm) was added to record glutamate-evoked currents. The internalsolution used for synaptic transmission recording contained (in mm):160 Cs-methanesulphonate, 10 EGTA, 10 glucose, 1 MgCl2, 5 Mg-ATP, 0.3 Na-GTP, and 10 Hepes ⁄ CsOH (pH 7.3). In someexperiments, the unselective c-aminobutyric acid (GABA) receptorantagonist bicuculline (40 lm) was added to the Tyrode solution toblock the GABAergic transmission. For voltage-dependent Na+ (INa)and Ca2+ (ICa) currents, glutamatergic transmission and GABA- orglutamate-evoked current recordings, the cells were dialysed with anintracellular solution containing (in mm): 160 CsCl, 10 EGTA,20 TEA.Cl, 10 glucose, 1 MgCl2, 5 Mg-ATP, 0.3 Na-GTP, and10 Hepes ⁄ CsOH (pH 7.3). For recordings of K+ outward currents(IK), the intracellular solution contained (in mm): 10 NaCl, 135 KCl,5 Mg-ATP, 0.3 Na-GTP, 14 EGTA, and 20 Hepes ⁄ KOH (pH 7.3).

Data acquisition was performed using PULSE programs (HEKAElektronic, Lambrecht, Germany). The data analysis was performedwith the MiniAnalysis program (Synaptosoft, Decatur, GA, USA) andPULSE programs (HEKA Elektronic, Lambrecht, Germany). Allexperiments were performed at room temperature (22–24 �C).

Measurement of the cytosolic Ca2+ concentration [Ca2+]cin single fluo-4-loaded cells

For these experiments cells, were plated at a density of2 · 105 cells ⁄ dish on 4-cm-diameter Petri dishes. Cells were loadedwith fluo-4 ⁄ AM (5 lm) for 45 min at 37 �C in Krebs–Hepes solution[in mm: 144 NaCl, 2 CaCl2, 5.9 KCl, 1.2 MgCl2, 11 glucose,10 Hepes ⁄ NaOH (pH 7.4)]. Loading with fluorescent dye wasfinished by washing the cells twice with Krebs–Hepes. Cells werekept at room temperature for 30 min before starting the experiment inan inverted fluorescence microscope (Nikon Eclipse TE300). Fluo-4was excited with light at 485 nm; emitted light was transmittedthrough a 505-nm dichroic mirror and 520-nm emission filter beforebeing detected by a CCD camera. Data were analysed using theMetafluor vs. 2.2 program (Universal Imaging, Downingtown, USA).

Synaptosomal preparation

Animal care and experimental procedures were performed followingthe guidelines set by the Animal Research Committee of ComplutenseUniversity, Madrid. Synaptosomes were purified on discontinuousPercoll gradients as described previously (Dunkley et al., 1986).Briefly, the cerebral cortex was isolated from adult male Wistar rats(2–3 months old) and homogenized in medium containing 0.32 m

sucrose (pH 7.4). The homogenate was centrifuged for 2 min at 2000 gat 4 �C, and the supernatant was spun again at 950 g for 12 min. Fromthe pellets formed, the loosely compacted white layer containing themajority of synaptosomes was gently resuspended in 8 mL of 0.32 m

sucrose (pH 7.4), and 2 mL of this suspension was then placed onto a3-mL Percoll discontinuous gradient containing: 0.32 m sucrose, 1 mm

EDTA, 0.25 mm dl-dithiothreitol, and 3%, 10% or 23% Percoll(pH 7.4). After centrifugation at 25 000 g for 10 min at 4 �C, thesynaptosomes were recovered from between the 10% and the 23%

Percoll bands, and diluted in a final volume of 30 mL of Hepes buffermedium [in mm: 140 NaCl, 5 KCl, 5 NaHCO3, 1.2 NaH2PO4,1 MgCl2, 10 glucose and 10 Hepes (pH 7.4)]. Following centrifuga-tion at 22 · 103 g for 10 min, the synaptosomes were resuspended in8 mL of Hepes buffer medium, and the protein content was determinedby the Biuret method. Finally, 1 mg of the synaptosomal suspensionwas diluted in 8 mL of Hepes buffer medium and spun at 3000 g for10 min. The supernatant was discarded, and the pellet containing thesynaptosomes was stored on ice. Under these conditions, the synap-tosomes remain fully viable for at least 4–6 h.

Glutamate release

Glutamate release was assayed by on-line fluorimetry (Nicholls et al.,1987). Synaptosomal pellets were resuspended in Hepes buffermedium (0.67 mg ⁄ mL) and preincubated at 37 �C for 1 h in thepresence of 16 mm bovine serum albumin, which served to bind anyfree fatty acids released from synaptosomes during the preincubation.After preincubation, the synaptosomes were pelleted and resuspendedin fresh incubation medium without bovine serum albumin. A 1-mLaliquot was transferred to a stirred cuvette containing 1 mm NADP+,50 U of glutamate dehydrogenase and 1.33 mm CaCl2 or ‘Ca

2+-free’solution containing 200 nm Ca2+. The fluorescence resulting fromNADPH formation was followed in a Perkin Elmer LS-50 lumines-cence spectrometer at excitation and emission wavelengths of 340 and460 nm, respectively. The traces were calibrated by adding 2 nmol ofglutamate at the end of each assay, and the data points were obtainedat 2-s intervals and corrected for Ca2+-independent release. Thus, theCa2+-dependent release was calculated by subtracting the releaseobtained during the period of depolarization with Ca2+-free mediumfrom the release obtained with 1.33 mm CaCl2.

Chemicals

Minocycline, GABA, bicuculline, glutamate, ionomycin, NADP+

glutamate dehydrogenase, fluorescein diacetate and other chemicalcomponents were obtained from Sigma (Madrid, Spain). DMEM,B-27, fetal bovine serum, penicillin–streptomycin and gentamicin werepurchased from Gibco-Invitrogen (Barcelona, Spain). Tetrodotoxinwas obtained from Tocris (Bristol, UK). Fluo-4 ⁄ AM and propidiumiodide were purchased from Molecular Probes (Barcelona, Spain).

Statistical analysis

Data were expressed as mean ± SEM for n ¼ number of cells from atleast three different cell batches. Student’s t-test was used to determinestatistical significance between group means. P-values of less than0.05 were considered significant.

Results

Minocycline inhibits glutamate-induced neuronal cell death

We used the fluorescein diacetate staining method to analyse theeffects of minocycline on glutamate-induced cytotoxicity. Glutamatetreatment produced a significant decrease in cell viability in hippo-campal cultures (about 40%, Fig. 1A). Dying neurons exhibitedseveral of the hallmarks of apoptosis, including cell shrinkage andcondensation of the cytoplasm, and chromatin aggregation andfragmentation in situ visualized with propidium iodide staining. Todetermine the effect of minocycline on glutamate-induced toxicity,

Minocycline decreases neuronal excitability 2483


hippocampal neurons were pretreated with 10–150 lm minocyclinefor 30 min before the 20-min exposure to 100 lm glutamate. By 24 hafter glutamate exposure, pretreatment with minocycline affordedcytoprotection in a concentration-dependent manner. Concentrationslower than 75 lm did not significantly modify glutamate-induced celldeath (Fig. 1A). However, other compounds, such as tetracycline(100 lm), did not show a neuroprotective effect. Figure 1B shows theconcentration–response curves for minocycline in preventing celldeath induced by glutamate exposure. The prevention of neuronaldeath amounted to 3.7 ± 6%, 2.3 ± 7%, 18.6 ± 6%, 55.8 ± 4%,

87.3 ± 2% and 87.2 ± 4% at minocycline concentrations of 10, 25,50, 75, 100 and 150 lm, respectively. Minocycline had an EC50 of79 lm. Whether or not the neuroprotective property of minocyclinehas been accepted, the concentration of minocycline in the brain thatwould correlate with this neuroprotection remains unknown.

Minocycline decreases the amplitude and the numberof spontaneous excitatory postsynaptic currents (sEPSCs)in hippocampal neurons in culture

When in culture, hippocampal neurons change their density, distribu-tion and interneuronal connections as a function of the culture age.Hence, we always used neurons after 11–15 days in culture through-out this study. When clamped at )80 mV, after blockade ofGABAergic synaptic transmission, sEPSCs, with a frequency ofabout 4.5 Hz, were recorded as inward currents. In the sample neuronshown in Fig. 2A, in the presence of 40 lm bicuculline to abolish theGABA component of such events, inward sEPSCs were recorded overseveral minutes. Note that minocycline (100 lm) reduced theiramplitude in a reversible manner.Figure 2B shows an event frequency histogram of the record shown

in Fig. 2B. Note the pronounced frequency decrement evoked byminocycline and its reversibility upon washing out the drug. The effectexerted by minocycline on the sEPSC frequency shows concentrationdependence, as shown in Fig. 2C (3.6 ± 2%, 8.6 ± 4%, 16.5 ± 4%,35.6 ± 2% and 47.8 ± 5% of the decrement induced by 3, 10, 30, 100and 300 lm minocycline, respectively, after 2 min of perfusion). TheIC50 was 64 lm. The amplitude of individual events (Fig. 2D) wasalso reversibly blocked by the drug. Averaged data from 9–21 neuronsshowed sEPSC amplitude depressions of 3.1 ± 2%, 7.7 ± 2%,18.2 ± 3%, 37.4 ± 1% and 42.5 ± 3% by the minocycline concentra-tions mentioned above. The IC50 was 42 lm (Fig. 2E).These results suggest that glutamatergic synaptic transmission in

hippocampal neurons is drastically affected by neuroprotectiveconcentrations of minocycline (30 lm and higher). This effect couldbe related to modulation of the presynaptic and ⁄ or postsynapticprocesses involved in synaptic transmission. A presynaptic effectcould be related to neuronal excitability and ⁄ or neurotransmitterrelease, whereas a postsynaptic effect could be mainly related to adirect action of minocycline on glutamatergic and ⁄ or GABAergicpostsynaptic receptors. We explored these possible targets forminocycline.

Minocycline decreases the firing of spontaneous actionpotentials

To test the possible role of minocycline as a modulator of neuronalexcitability, we recorded, under current clamp conditions, thespontaneous AP firing and how these APs could be modified byconstant neuron perfusion with minocycline. Recordings wereperformed using the whole-cell-perforated configuration of thepatch-clamp technique, under the current clamp mode.The resting membrane potential (Em) was )62.1 ± 1.9 mV in 21

cells tested. Em remained constant except for the membrane potential

Fig. 1. Neuroprotective effects of minocycline against glutamate-induced celldeath. Minocycline (10–150 lm) and tetracycline (100 lm) were added 30 minbefore a 20-min exposure to glutamate and maintained until the end of theexperiment. (A) Cell viability assayed 24 h after 100 lm glutamate exposure.(B) Averaged results of the neuroprotective effect of minocycline (ordinate) asa function of drug concentration (abscissa). Data represent the means ± SEM ofthree or four independent experiments for each drug concentration. **P < 0.01vs. control conditions in the absence of minocycline.

Fig. 2. Minocycline decreases the frequency and amplitude of spontaneous excitatory postsynaptic currents (sEPSCs). In the presence of the c-aminobutyric acidreceptor antagonist bicuculline (40 lm), sEPSCs were recorded as inward currents at )80 mV holding potential, in embryonic rat hippocampal cells kept in culturefor 11–15 days. (A) An original record, where sEPSCs are drastically modified by minocycline (100 lm) (top horizontal bar). Similar results were obtained in 13different neurons. (B) Frequency histogram for the record shown in A (bin size 4). (C) Concentration–response curve for the sEPSC frequency decrement evokedby minocycline. (D) Time course for the amplitude measured for each individual event of A. (E) Concentration–response curve for the sEPSC amplitude decrementevoked by minocycline. Data are means ± SEM from 9–21 neurons.



variations due to the autorhythmicity activity present in these cells.Under whole-cell-perforated patch-clamp in 2 mm Ca2+, most hippo-campal neurons presented autorhythmicity characterized by the firingof regular AP trains with a frequency of 3.61 ± 0.5 Hz (n ¼ 8 cells),with or without periods of electrical quiescence (Fig. 3A). When thefiring pattern was constant, the frequency of spike discharge was alsoconstant for at least 2–3 min, with a spike interval of 277 ms. TheseAPs reached peaks of 40.3 ± 2.8 mV (slope 13.9 ± 0.6 mV ⁄ ms, risetime 5.08 ± 0.5 ms, decay time 26.6 ± 2.4 ms, AP area390 ± 47.6 mV ms, and half-width 5.1 ± 0.2 ms). We next examinedthe effects of minocycline on spontaneous AP firing (Fig. 3A and B).

The concentration of 100 lm was selected because it has been provento afford maximal neuroprotection in several studies (Fernandez-Gomez et al., 2005), including ours (Fig. 1). After 1 min of perfusion,100 lm minocycline produced a progressive shortening of thedischarge (2.43 ± 0.5 spikes ⁄ s) with no significant modifications inthe kinetic parameters in eight neurons tested (AP peaks of39.9 ± 1.8 mV, slope 13.1 ± 0.6 mV ⁄ ms, rise time 5.8 ± 0.4 ms,decay time 23.3 ± 2.1 ms, AP area 328 ± 29 mV ms, half-width4.8 ± 0.6 ms). The insets in Fig. 3A show expanded AP trains before,during and after minocycline washout. Note the elevation of baseline,reflecting the cell depolarization that triggers APs; the area of such

Fig. 3. Reduction by minocycline of the rate of spontaneous action potentials. (A) An original record of spontaneous action potential firing obtained from aneuron using the current-clamp mode of the patch-clamp technique. Minocycline, 100 lm, was perfused as shown by the top horizontal bar. Insets represent anexpanded time scale for three trains of action potentials recorded under different experimental conditions. (B) The events number as a function of time for theoriginal record shown in A (bin size 5). (C) Averaged data for spike frequency under the experimental conditions shown in A. Data are means ± SEM of the numberof cells shown in parentheses. **P < 0.01.



depolarization was 88 ± 5 mV s in controls, vs. 45 ± 4 mV s in thepresence of minocycline.

Reduction by minocycline of evoked AP firing

The threshold Em for spontaneous AP firing was about 10 mV positivewith respect to the resting membrane potential. Before currentinjection to evoke AP trains, the membrane potential was held at)70 mV in order to prevent spontaneous AP firing. Short currentinjections of the order of 10 pA, applied with durations from a fewmilliseconds to several seconds, generated repetitive APs that werefollowed by prolonged afterhyperpolarizations. Their mean frequencywas 33.3 spikes ⁄ s during the 300-ms pulse; with longer currentpulses, the amplitude and frequency declined with time. The AP firingwas interrupted at the end of the pulse by a progressive decline ofspike amplitude, probably due to INa inactivation caused by sustainedmembrane depolarization. In some cases, at the beginning of the pulse,a frequency higher than 33.3 Hz, which decayed with time, wasobserved.

Figure 4A shows AP trains evoked during a 300-ms currentinjection pulse of 10 pA. The AP amplitude was 50–80 mV, and theAP duration measured midway between the threshold ()25 mV) andthe peak was 3 ms. After repolarization (5 ms after the AP peak), themembrane hyperpolarized up to 10 mV more negative than the restingEm, as shown in Fig. 4A (left). The middle trace in this panel showsthe AP trains during neuronal perfusion with minocycline. In mostcells, this drug induced an almost complete suppression of AP firing;recovery of the initial AP firing pattern was produced uponminocycline washout (right trace).

Averaged results from seven cells, corresponding to the first six APsin each train, are plotted in Fig. 4B–J. Note the augmentation of APintervals (Fig. 4B) and the diminution of amplitude (Fig. 4C) and AParea (Fig. 4D) caused by minocycline. Minocycline did not affect therise time (Fig. 4E), but it slowed down the decay time (Fig. 4F) andaugmented the halfwidth (Fig. 4G) and the afterhyperpolarization area(Fig. 4I); latency was unmodified (Fig. 4J). These changes suggest thatminocycline mitigates neuronal excitability. Thus, we decided to testits effects on the ion currents involved in AP generation andtermination.

Minocycline inhibits voltage-dependent Na+-channels

The activation of Na+-channels is necessary for AP generation andpropagation. Thus, we tested first the effects of minocycline on INausing the whole-cell configuration of the patch-clamp technique. The)80-mV clamped neurons were continuously superfused with anextracellular solution containing 137 mm NaC1 and 0 mm Ca2+ (seeMaterials and methods for further details). Figure 5A represents thecurrent–voltage (I ⁄ V) curve generated by 10-mV increasing depolar-izing steps of 8 ms duration, given from )60 mV to +40 mV at 10-sintervals. INa exhibited very fast activation kinetics followed by aninactivation phase that reached basal level in about 2.5 ms (seeoriginal traces in Fig. 5A). The activation threshold was around)55 mV, INa peaked at )45 mV, and the apparent reversal potentialwas at +40 mV (Fig. 5A). Minocycline (100 lm) induced a 20%blockade of peak current and slightly shifted the I ⁄ V relationshiptowards more hyperpolarizing potentials. The control currents wererecovered upon washout of minocycline. Averaged peak INaamounted to 3.01 ± 0.3 nA (n ¼ 20 cells). Figure 5B shows thetime course of INa elicited by 8-ms depolarizing pulses from aholding potential of )80 mV, applied at 10-s intervals. The initial

peak INa of 4.2 nA was rapidly and reversibly depressed byminocycline. Data from 9–11 cells were used to plot the concentra-tion–response curve shown in Fig. 5C, showing average blockades of3.2 ± 1%, 5.9 ± 1%, 11.0 ± 0.3%, 20.1 ± 2% and 22.3 ± 2% with 3,10, 30, 100 and 300 lm minocycline, respectively, after 2 min ofperfusion. Slight recoveries of INa were seen upon washout ofminocycline (93.5 ± 2.8%). Pooled results using the previous proto-col gave a concentration–inhibition curve with a calculated IC50 forINa blockade of 67 lm (Fig. 5C).

Effects of minocycline on K+ currents

The activation of outward K+ currents (IK) is important for the rapidtermination of APs. Therefore, modulation of IK could exert profoundeffects on the firing rate. Figure 6A shows that minocycline did notaffect voltage-dependent K+ channels. IK was recorded using anextracellular solution containing nominal zero Ca2+, so all the outwardcurrent would be expected to be due to Ca2+-independent voltage-activated K+-channels. The I ⁄ V curve was obtained in the absence andpresence of minocycline, in 15 neurons clamped at )80 mV andstimulated for 200 ms with 10-mV-increment steps at 10-s intervals.Augmentation of voltage-dependent IK (IKv) was linear at voltagesbetween )20 and +70 mV. The insets represent original IKv traces atthe indicated voltages, with (black circles) or without (black squares)minocycline. Notice the absence of effect by minocycline at allvoltages tested. Figure 6B shows the time course of the voltage-dependent K+ currents generated by the application of 200-msdepolarizing pulses to +50 mV, repeated every 10 s from a )80-mVholding potential. The current amplitude was similar before and duringminocycline perfusion (top horizontal bar) (6.3 ± 0.3 vs.6.1 ± 0.4 nA, respectively, n ¼ 25 cells).Comparison of the I ⁄ V curves in the presence and in the absence

of Ca2+ indicates that between about )30 and +50 mV (intervalvalues of potential where AP firing occurs), outward currents arepartially Ca2+-dependent. The depolarizing steps that trigger Ca2+

influx by activation of Ca2+-channels also activate Ca2+-channelsand voltage-dependent K+-channels, which can be quantified by thecharacteristic hump of the control I ⁄ V curve, which mirrors the I ⁄ Vcurve for ICa in these cells. The IKCa present in hippocampal neuronsrepresents 19.5 ± 3% at )20 mV, 28.0 ± 3% at 0 mV, 31.8 ± 2% at+10 mV and 19.7 ± 2% at +30 mV of the total IK. Figure 7Arepresents the I ⁄ V curve of IKCa obtained after subtraction of theI ⁄ V curve obtained in 0 mm Ca2+ (black circles) from that obtainedin 2 mm Ca2+ (black squares). Figure 7B shows the blockadeexerted by minocycline on IKv and IKCa during a 200-ms depolar-izing pulse to +10 mV in the absence and in the presence ofextracellular Ca2+. Minocycline (100 lm) exerted quick blockadefrom the first seconds of application and remained constant duringthe full period of minocycline perfusion. After minocycline washout,the control outward current was fully recovered. Note how thehigher blocking effects were exerted at the range of voltagedepolarizing pulses that maximally activates Ca2+-channels; thus,blockade of IKCa by minocycline amounted to 30.4 ± 4% at)20 mV, 55.9 ± 6% at 0 mV, 38.2 ± 3% at +10 mV and33.1 ± 7% at +30 mV (Fig. 7C). Figure 7D shows the concentra-tion–response curves for minocycline inhibitory effects of IKCa,measured in each individual cell at the end of the 2-min superfusionperiod with each drug concentration. The blockades exerted by 3,10, 30, 100 and 300 lm minocycline were 5.0 ± 4%, 7.3 ± 2%,12.3 ± 6%, 47.2 ± 9% and 55.7 ± 12%, respectively, in 4–6 neuronstested. The IC50 was 54 lm.



Fig. 4. Reduction by minocycline of evoked action potentials (APs). (A) Original traces obtained in response to current injections of 10 pA over 300 ms, appliedevery 10 s to hippocampal neurons under control conditions (left trace), in the presence of minocycline (middle trace), and after its washout (right trace). Note howthe sustained discharge evoked by current injection is first reduced and subsequently suppressed. AP firing resumes after minocycline washout. (B–I) Comparativeeffects of minocycline (100 lm) vs. control conditions on different parameters of APs. Data are plotted to a linear fit (B, H and I), to a Boltzmann fit (D, F and G), orto an exponential decay fit (C and E). Ordinates show the averaged kinetic parameters of individual APs; abscissae show the first six consecutive APs of a traingenerated by each current injection (E–I), except in B, where the abscissa represents the AP intervals. (J) Averaged data on latency time to the first AP. Data aremeans ± SEM of seven cells. *P < 0.05, with respect to control.



Blockade by minocycline of inward Ca2+ currents

Blockade by minocycline of ICa could affect IKCa as well as neurotrans-mitter release. To test this possibility, we studied the effects ofminocycline on ICa using 10 mm Ca2+ as charge carrier. To obtain thecurrent family traces that allowed the plotting of the I ⁄ V curve of Fig. 8A,hippocampal neurons voltage-clamped at)80 mVwere stimulatedwith50-ms depolarizing pulses of 10-mV increasing steps given at 10-sintervals, before and during cell perfusion with 100 lm minocycline.Before minocycline administration, peak ICa presented activation ataround )30 mV, peaked at 10 mV, and showed a reversal potential near+70 mV. During the 50-ms depolarizing pulses, a clear inactivation ofthe current appeared (see original traces in Fig. 8A insets). In thepresence of minocycline, no shifts of the I ⁄ V curve were observed.Minocycline inhibited ICa at all potentials tested. Tominimize variationsbetween data from different cells, ICa was normalized from eachindividual cell (ICa ⁄ ICa max.). No modifications of current activation(time constants, sact 2.8 ± 0.3 ms and 2.5 ± 0.2 ms for control andminocycline, respectively) and deactivation kinetics (time constants,sdeact 50.5 ± 14 ms and 48.5 ± 12 ms for control and minocycline,respectively) were observed. The time course of ICa peaking is shown inthe prototype cell of Fig. 8B. This cell was stimulated with 50-ms testpulses to 0 mV, from a holding potential of)80 mV.Note in Fig. 8B thatthe initial ICa amounted to 2.25 nA and was depressed in 10 s to 1.6 nAin the presence ofminocycline; the current recovered after drugwashout.Mean averaged ICa current amounted to 2.48 ± 0.2 nA in 34 cells. Theaverage pooled results for the inhibition by minocycline of peak ICa areshown in Fig. 8C. Inhibition of ICa was measured in each individualneuron at the end of the 2-min perfusion period with each drugconcentration; blockade amounted to 6.1 ± 2%, 8.3 ± 1%, 13.4 ± 2%,28.1 ± 2% and 36.3 ± 1% at minocycline concentrations of 3, 10, 30,100 and 300 lm, respectively. The IC50 for ICa blockade was 45 lm

(data from six to nine cells).

Effects of minocycline on the elevation of [Ca2+]c evokedby glutamate in fluo-4-loaded hippocampal neurons

As we found significant blockade of both INa and ICa (Figs 5 and 8)and a drastic decrease of neuronal excitability (Figs 1–3), we thoughtit of interest to follow the time course of the [Ca2+]c changes elicitedby glutamate in single fluo-4-loaded cells, and whether these changeswere modified by minocycline.Figure 9 shows the effects of minocycline on the [Ca2+]c increments

evoked by glutamate pulses. Fast perfusion of 30 lm glutamate plus10 lm glycine (5-s pulses) rapidly enhanced [Ca2+]c from 250arbitrary fluorescence units to a peak in few seconds (son 1.7 ± 0.2 s).Minocycline (100 lm) reduced by half the amplitude of these Ca2+

peaks and slowed down their rate of activation (son 3.8 ± 0.9 s)(Fig. 9A and B). Fluorescence decayed at similar rates in controls(soff 16.6 ± 1.4 s) and during minocycline perfusion (soff 17.1 ±1.8 s). In about a minute, the fluorescence reached basal levels.Figure 9B shows quantitative pooled data (peak) of the [Ca2+]ctransients elicited by repetitive glutamate pulses, and the effects ofminocycline (top horizontal bars). Minocycline (100 lm) blocked theincrement of [Ca2+]c spikes by about 30% in 10 neurons tested. Notethat in the presence of minocycline, the Ca2+ signal was half thatevoked by glutamate in control conditions, and that upon washout, the[Ca2+]c signals recovered their control values. Data from five to 10neurons tested showed average decrements of 3.9 ± 1%, 5.4 ± 1%,13.9 ± 3%, 29.6 ± 1% and 36.3 ± 2% with 3, 10, 30, 100 and 300 lm

minocycline, respectively. The concentration–response curve plottedin Fig. 9C shows an IC50 of 52 lm.

Fig. 5. Blocking effects of minocycline on voltage-dependent Na+-channels.(A) Hippocampal neurons were voltage-clamped at )80 mV. Depolarizing8-ms test pulses to different voltages were applied at 10-s intervals. Normalizedpeak inward currents are plotted as a function of the potential before and after1 min of 100 lm minocycline perfusion (n ¼ 10). Insets show original INatraces recorded in control conditions (black squares) and during minocyclineapplication (black circles) at the indicated voltage step. (B) Time course ofINa. The holding potential was )80 mV, and currents were evoked by 8-msdepolarizing pulses at 0 mV at 10-s intervals. Once they were INa stabilized,cells were continuously superfused with minocycline as indicated by thehorizontal bar. (C) Quantitative inhibition of INa by several minocyclineconcentrations. The amplitude of peak current was normalized to 100% inevery individual neuron studied (control current); then, the peak currentblockade after 2 min of exposure to the drug was expressed as percentage ofcontrol; the number of experiments for each point was 9–11. The IC50 was67 lm. Data are means ± SEM; at 30 lm or higher concentrations, theinhibition of INa by minocycline showed statistical significance (P < 0.01, withrespect to control).



Minocycline reduces the K+-evoked glutamate release but failsto alter the release induced by the Ca2+ ionophore ionomycin

In the next set of experiments (Fig. 10), we tried to ascertain whetherminocycline could contribute to the decrease in neuronal activity, notonly by blocking depolarizing ionic channels, but also by additionaldirect modulation of glutamate release.Depolarization of nerve terminals with KCl opens voltage-depen-

dent Ca2+-channels and initiates neurotransmitter release (Tibbs et al.,1989). Depolarization of nerve terminals with KCl also promotes theCa2+-independent release of glutamate that can be estimated in theabsence of Ca2+ (5 mm EGTA). This Ca2+-independent release wassubtracted from that measured in the presence of 1.33 mm Ca2+ toobtain the Ca2+-sensitive release component. In the cerebrocorticalnerve terminal preparation, the Ca2+-independent release of glutamateafter 5 min of depolarization with 30 mm KCl was 1.46 ± 0.1 nmolglutamate ⁄ mg, whereas the Ca2+-dependent release reached2.84 ± 0.1 nmol glutamate ⁄ mg (n ¼ 3). The Ca2+-dependent releasewas not altered by the addition of minocycline at 10 lm

(3.04 ± 0.2 nmol glutamate ⁄ mg, n ¼ 3). However, at 50 lm and100 lm, minocycline reduced the Ca2+-dependent release by40.5 ± 5% (n ¼ 3), and 63.3 ± 2% (n ¼ 5), respectively (Fig. 10Aand B). The Ca2+-independent release was not modified by minocy-

cline at any of the concentrations tested (1.36 ± 0.1 nmol gluta-mate ⁄ mg at 100 lm, n ¼ 3). The IC50 calculated for the reduction ofthe glutamate Ca2+-dependent release by minocycline was 52 lm.The vesicular release of glutamate can also be induced by the Ca2+

ionophore ionomycin in a manner that is independent of plasmamembrane depolarization, and therefore independent of the activity ofvoltage-dependent Ca2+-channels. Thus, ionomycin-induced releasereflects the modulation of release machinery downstream of Ca2+

influx (Sihra et al., 1992). The release of glutamate induced by 4 lm

or 6 lm ionomycin (2.16 ± 0.1 nmol ⁄ mg, n ¼ 3, and2.89 ± 0.2 nmol ⁄ mg, n ¼ 3, respectively) was not altered by mino-cycline at 100 lm (2.33 ± 0.1 nmol ⁄ mg and 3.02 ± 0.1 nmol ⁄ mg,respectively) (Fig. 10C). These results therefore suggest that theminocycline-mediated inhibition of glutamate release is not theconsequence of modified sensitivity of the release machinery to Ca2+.

Effects of minocycline on GABA receptors and glutamatereceptors

At the beginning of this study, we considered the possibility thatminocycline could modulate synaptic transmission at a postsynapticlevel. The modulatory action on synaptic transmission may involve adirect action on GABA or glutamate receptors in addition to the effectson ion channels. Therefore, we investigated the effects of minocyclineon ligand-gated ionotropic receptors involved in excitatory and

Fig. 6. Effects of minocycline on voltage-dependent K+-channels. (A) Nor-malized voltage-dependent outward K+ currents vs. membrane potential beforeand during perfusion with 100 lm minocycline in 0 mm external Ca2+

(n ¼ 15). From a holding potential of )80 mV, outward K+ currents wereevoked by 10-mV step depolarizations, applied for 200 ms at 10-s intervals.The original traces and the I ⁄ V curves show no effects of minocycline (blackcircles) on IKv (black squares). (B) Time course of IKv; notice the absence ofany effect mediated by minocycline (top horizontal bar). Filled diamondsrepresent the peak amplitude of IKv measured during 200-ms depolarizingpulses to +50 mV, repeated every 10 s. (C) Averaged quantitative inhibition ofIKv by minocycline 100 lm (after 2 min). The amplitude of peak current wasnormalized to 100% in every individual neuron (control current). Data aremeans ± SEM of 25 cells.

Fig. 7. Effects of minocycline on Ca2+ and voltage-dependent K+-channels.(A) The I ⁄ V curves reflect the IKCa (triangles) obtained after subtraction of theoutward current recorded in 0 mm Ca2+ (circles) from that recorded in 2 mm

Ca2+ (squares). Depolarizing pulses of 200 ms from )20 to +70 mV at 10-mVincrements (VH )80 mV) and 10-s intervals were given to construct these I ⁄ Vcurves. (B) Current recordings obtained in a neuron clamped at )80 mV anddepolarized to +10 mV either with or without 2 mm external Ca2+. Note howthe presence of Ca2+ induced Ca2+ influx and activates a Ca2+- and voltage-dependent current that was sensitive to blockade by minocycline. (C) Aver-aged quantitative inhibition of IKCa by minocycline (after 2 min) at theindicated voltage test pulses (abscissa). The amplitude of peak current wasnormalized to 100% in every individual neuron (control current). (D) Aver-aged results of the percentage current blockade after 2 min of minocyclineperfusion with each drug concentration (abscissa). The IC50 was 54 lm. Atconcentrations higher than 30 lm, the effect of minocycline decreased instatistical significance with respect to controls. Data are means ± SEM of thenumber of cells shown in parentheses. **P < 0.01.



inhibitory transmission in the hippocampus. The effects of minocy-cline were examined on GABA- and glutamate-induced whole-cellcurrents in primary cultures of rat hippocampal neurons bathed intetrodotoxin-containing external solution (0.3 lm) to prevent sponta-neous synaptic activity.In hippocampal neurons clamped at )80 mV, the application of 0.3

and 1 mm glutamate (100-ms pulses) induced fast inward currents of2.2 ± 0.2 nA (n ¼ 8) and 2.7 ± 0.3 nA (n ¼ 10), respectively; thisresponse was also measured in the presence of 100 lm minocycline,and no statistically different values were observed when comparedwith controls. The effect of minocycline on glutamatergic receptorswas also examined in the absence of extracellular Mg2+, where a peakof 7.5 ± 0.6 nA was evoked by 300 lm glutamate. Minocycline hadlittle effect on peak glutamate-induced currents (1.4 ± 0.8% blockadein eight neurons studied). As shown in Fig. 11A (left panel), theglutamate-dependent current was characterized by fast activationkinetics followed by a complex inactivation phase. The effect exertedby minocycline was visible only on the initial fast inactivation peakcurrent; this could be related to non-N-methyl-d-aspartate receptors,because no effect was seen in Mg2+-free external solution, where theN-methyl-d-aspartate component is more predominant. Minocycline(10 and 100 lm) did not significantly reduce the current evoked by300 lm glutamate (1.4 ± 2%, n ¼ 4, and 11.8 ± 2%, n ¼ 8, respec-tively) (see right panel in Fig. 11A) or 1 mm glutamate (0.95 ± 3%,n ¼ 4, and 14.4 ± 3%, n ¼ 10, respectively).We also examined the effects of minocycline on GABA receptors.

We recorded GABAergic-induced whole-cell currents in a hippocam-pal neuron voltage-clamped at )80 mV. Application of a 100-msGABA pulse (100 lm) evoked a chloride current that peaked at10.6 ± 1.1 nA (n ¼ 24) with characteristic fast activation kineticsfollowed by the typical slow inactivating kinetics. Minocyclineexerted no effects on GABA-induced responses in these cells.Responses induced by 100 lm GABA in these neurons wereessentially unaffected by minocycline applied for several minutes at10 lm (3.8 ± 0.6%, n ¼ 6) and 100 lm (5.5 ± 4.4%, n ¼ 16)(Fig. 11B).

Discussion

The main aim of this study was to obtain better knowledge about themechanism responsible for the neuroprotective effects of minocyclineon glutamate-induced cytotoxicity. We used hippocampal neurons inculture, which are formed by approximately 80% of glutamatergicneurons (pyramidal cells) vs. 20% of GABAergic neurons (interneu-rons and granular cells); similar rates has been described in cortexprimary culture (Millan et al., 2003). As shown in Fig. 1, theglutamate treatment produced a significant decrease in cell viability.To determine the effect of minocycline on glutamate-induced toxicity,hippocampal neurons were pretreated with a wide range of concen-trations (10–150 lm), which were observed to afford cytoprotection tothe neuronal hippocampal culture in a concentration-dependentmanner. We found in this study that neuroprotectant concentrationsof minocycline against brain ischaemia (Yrjanheikki et al., 1999),excitotoxicity (Tikka et al., 2001), spinal cord injury (Stirling et al.,2004) and 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (He et al.,2001) depressed synaptic transmission in cultured hippocampalneurons. This depression is explained by reduction of sEPSCs,recorded in the presence of the GABAA receptor blocker bicuculline.The current frequency, the amplitude and the area of individual eventsare reduced approximately by 50% (Fig. 2). We demonstrate howminocycline could contribute to decrease neuronal excitability, not

Fig. 8. Blocking effects of minocycline on voltage-dependent Ca2+-channels.(A) Normalized peak inward currents (ICa) are plotted as a function ofpotential in 10 mm Ca2+ (n ¼ 9). I ⁄ V curves were obtained before and 1 minafter perfusion of 100 lm minocycline. Neurons voltage-clamped at )80 mVwere stimulated with 50-ms depolarizing pulses at the indicated voltages(abscissa) at 10-s intervals. The insets show original current traces obtained attest potentials in control conditions (black squares) and during minocyclineapplication (black circles). (B) Time course of the inhibition by minocyclineof ICa. Depolarizing test pulses of 50 ms to 0 mV from a holding potential of)80 mV were applied at 10-s intervals in 10 mm Ca2+. Once ICa stabilized,cells were continuously perfused with minocycline (top horizontal bar).(C) Peak ICa (%) blockade after 2 min of perfusion with each drug concen-tration. The amplitude of the initial ICa peak was normalized to 100% in everyindividual neuron tested (control). The IC50 was 45 lm. Data are mean-s ± SEM of six to nine cells. The effect of minocycline at 30 lm or higherconcentrations showed statistical significance (P < 0.01) with respect tocontrols.



only by blocking depolarizing ionic channels, but also by an additionaldirect modulation of glutamate release. Hence, minocycline wasprobably reducing glutamate release, a mechanism that was directlytested in cortical synaptosomes, using high K+ as a depolarizingstimulus (Fig. 10). A postsynaptic effect explaining sEPSC depression

is discarded, considering that minocycline did not affect eitherglutamate- or GABA-induced inward currents (Fig. 11).The presynaptic action of minocycline was corroborated by

recording the spontaneous AP firing. Thus, the drug reduced thefrequency of spontaneous APs recorded in current-clamped hippo-campal neurons (Fig. 3). Furthermore, the drug diminished theamplitude and the frequency of AP trains elicited by current injection,leading to a late suppression of cell firing (Fig. 4). The reduction of APfrequency is probably due to an increase in the after-hyperpolarisationduration, a slight increment of baseline, and a drastic increment in APdecay time. These changes augment the refractory period, and thesubsequent decrease, of AP frequency. This effect of minocycline onAP generation and propagation may be explained by a combination ofeffects on various ion channel currents. For instance, minocyclinereduced INa by 20% (Fig. 5), ICa by 30% (Fig. 8), and IKCa by 50%(Fig. 7). We measured the contribution of Ca2+-channels and voltage-dependent K+-channels to the total outward K+ current at potentialsoccurring during an AP, i.e. from )20 mV to +30 mV. This impliesthat over 22% of the K+ current measured in the first millisecond of adepolarizing step is activated by Ca2+ influx. Thus, blockade byminocycline of ICa (Fig. 8) may explain its blocking effect on IKCa(Fig. 7). Additionally, the possibility of Ca2+-dependent K+ currentsuppression exerted by a direct action of minocycline on Ca2+-dependent K+-channels is completely excluded by the similar IC50

obtained in both experimental approaches, blockade of ICa and IKCa.The Ca2+ currents were recorded in 10 mm external Ca2+ however, theuse of high Ca2+ only shifted I ⁄ V 10 mV to the right (the control andthe blockade curve). We performed some control experiments,recording the blockade exerted by minocycline at two potentials (0and +10 mV) in comparison to 2 and 10 mm Ca2+, and no differencewas detected. The extracellular concentration of Ca2+ did not affect therelative blockade exerted by minocycline. The blockade obtained in10 mm Ca2+ at +10 mV is the same as that observed at 0 mV in 2 mm

Ca2+. It was interesting that blockade was higher at test potentials (i.e.0 mV), where ICa is known to be maximal; this corroborates the well-known observation that Ca2+ entering through voltage-dependentCa2+-channels rapidly activates small- and large-conductance Ca2+-dependent K+-channels. As these channels control the post-hyperpo-larization phase of the AP and hence AP firing frequency [see Stocker(2004) for a review], the halving by minocycline of IKCa (Fig. 7) mayexplain the reduction of AP frequency and even the suppression of APfiring (Figs 3 and 4).The increment in the intracellular concentration of Ca2+ during

neuronal ischaemia evoked by glutamate-derived hyperexcitabilityplays a particularly important role in the neurotoxic cascade resultingin acute neuronal cell death. Additionally, a reduction in the Ca2+

influx leads first to a decrement in cytosolic Ca2+ level to initiate theexocytotic process, and glutamate release, and second, to a decrementin the Ca2+-induced Ca2+ release responsible for maintenance of

Fig. 9. Minocycline decreases the cytosolic Ca2+ spike evoked by glutamate.(A) Time course of cytosolic Ca2+ concentration ([Ca2+]c) in Fluo-4 ⁄ AM-loaded hippocampal neurons. Cells were stimulated with 30 lm glutamate plus10 lm glycine for 5 s at 2-min intervals (black circles). Notice that glutamaterapidly raises the [Ca2+]c, which returns to basal levels in a few seconds.Minocycline (100 lm), which produced a marked decrement in [Ca2+]c, wasperfused for the time indicated by the top horizontal bar. (B) Averaged data ofpeak measured for every individual [Ca2+]c spike indicated as arbitraryfluorescence units (AFU). (C) The calculated IC50 of the [Ca2+]c peakblockade by minocycline was 52 lm. This curve was plotted with dataobtained through the normalization of the responses with respect to control, ineach individual cell. Data are means ± SEM of 5–10 neurons tested. The effectof minocycline (30 lm or higher) shows statistical differences (**P < 0.01)with respect to controls.



vesicular transport for neurotransmitter release. The partial blockadesexerted by minocycline of INa and ICa were translated into a 30–40%decrement of [Ca2+]c elevations elicited by glutamate (Fig. 9). Thismay explain the 60% blockade of K+-evoked glutamate release(Fig. 10). The K+-evoked release is not affected by either Na+-channelor K+-channel blockers, but is sensitive to inhibition by Ca2+-channelblockers (Millan et al., 2002). Nevertheless, the possibility of directinterference of minocycline with the exocytotic machinery releaseitself, downstream of Ca2+ entry, was excluded by the observation thatminocycline did not affect the ionomycin-induced release of

Fig. 10. Minocycline reduces glutamate release in a concentration-dependentmanner in cerebrocortical nerve terminals. (A) The release of glutamateevoked by 30 mm KCl in the presence of 1.33 mm Ca2+ or 5 mm EGTA wasdetermined in the presence and absence of minocycline (10, 50 and 100 lm)added 100 s before depolarization. Note the lack of effect on external Ca2+-independent glutamate release (EGTA traces). Traces are the means of three tofive experiments using two synaptosome preparations. (B) Averaged data ofglutamate release in the absence (control) and the presence of minocycline.Data are means ± SEM of the number of cells shown in parentheses.**P < 0.01, with respect to control. (C) Minocycline fails to affect ionomy-cin-induced glutamate release. The lower trace shows the basal spontaneousrelease obtained. Glutamate release was induced by 6 lm ionomycin in theabsence (control) or the presence of minocycline (100 lm), added 100 s before.Traces are the means of three to five experiments using two synaptosomepreparations.

Fig. 11. Effects of minocycline on glutamate- or c-aminobutyric acid(GABA)-evoked currents. (A) Hippocampal neurons in culture were perfusedwith Tyrode control solution and stimulated with 300 lm glutamate for 100 msat 30-s intervals in the absence and presence of minocycline (added 2 minbefore). The histogram represents the average data for glutamate-inducedresponses in both conditions. (B) A hippocampal neuron perfused with Tyrodesolution was stimulated with 100 lm GABA for 100 ms at 30-s intervalsbefore (control trace) and during minocycline perfusion. The histogramrepresents the average data for GABA-induced responses in control conditionsand in the presence of minocycline (added 2 min before). For each cell, theresponse in the presence of minocycline was calculated as a percentage of theresponse in control conditions (100%). Data are means ± SEM of number ofneurons tested (indicated in parentheses). No statistical differences were foundbetween the different experimental groups.



glutamate. Thus, these combined effects of minocycline may consid-erably reduce the neuronal Ca2+ overload evoked by excess glutamatestimulation of N-methyl-d-aspartate receptors, occurring during brainischaemic insults (Siesjo et al., 1995). Observation of the effects ofminocycline with the different experimental approaches suggested that30 lm was the threshold concentration to exert all these effects. Theeffectiveness of minocycline achieved with the same concentrationrange (see similar calculated IC50) suggests that the blockade exertedon cytosolic Ca2+ elevation, and neurotransmitter release, whicheventually led to a decrement in neuronal excitability, are mainlyevoked by a direct blockade of voltage-dependent Ca2+-channels,without discounting a small contribution of other mechanisms, such asNa+-channel blockade.We have not considered an effect of minocycline on the different

cell types and the possibility that pooling data from all cells may maskcell-type-specific effects because: (i) during the experiment performed,the hippocampal neurons recorded showed a pyramidal-like shape; (ii)the homogeneous effect recorded in every experimental approach, asshown by the low SEM, indicates a normal distribution effect, whichexcludes the possibility that different effects exerted by minocyclinecould be associated with different cell populations; and (iii) to preventvariations in the development of the primary culture, we employed11–15-day-old neurons.Minocycline exhibits neuroprotective effects against neuronal

damage in animal models of focal and global brain ischaemia(Yrjanheikki et al., 1999; Wang et al., 2003), Huntington’s disease(Chen et al., 2000; Wang et al., 2003), amyotrophic lateral sclerosis(Zhu et al., 2002), Alzheimer’s disease (Hunter et al., 2004), andParkinson’s disease (He et al., 2001). In particular, during and afterischaemic insults, there is consensus that excess glutamate release andimpairment of glutamate sequestration by astrocytes might be thecause of exacerbated neurotoxicity and neuronal death [see Blocket al. (2007) for a review]. In fact, therapeutic targets to mitigate suchneurotoxicity include Ca2+-channel blockers to reduce excess gluta-mate release (Gribkoff & Winquist, 2005) or glutamate receptorblockers (Garcıa de Arriba et al., 2006). The cell viability experimentsin which minocycline exerted a important neuroprotective effect, incontrast to the smaller effect evoked by other compounds, such astetracycline, are in accordance with other studies showing thatminocycline and doxycycline markedly reduce the size of infarctionin both focal and global transient ischaemia in the adult rat (Clarket al., 1994; Yrjanheikki et al., 1999; Xu et al., 2004); by contrast,tetracycline, which is less able to cross the blood–brain barrier to enterthe central nervous system, is not neuroprotective at the same doses(Yrjanheikki et al., 1998).The data available on the mechanism responsible for the neuropro-

tective actions of minocycline are scarce and controversial (Jordanet al., 2007). Several reports attribute the minocycline neuroprotectiveeffects to various intracellular signalling pathways, including antiox-idant systems (Kraus et al., 2005), nitric oxide synthase (Sadowski &Steinmeyer, 2001) and blockade of inflammatory responses [seeStirling et al. (2005) for a review]. Our results, however, stronglysuggest that minocycline acts at an earlier plasmalemmal step bylimiting glutamate release and the ensuing [Ca2+]c elevation in targetneurons. Minocycline may prevent the activation of this Ca2+-dependent intracellular pathway, thus preventing neuronal death.The regulatory mechanism exerted by minocycline on Ca2+ entry andmembrane potential leads to down-modulation of synaptic transmis-sion. This decrement in neuronal excitability, together with the markeddecrement in glutamate release, may explain the cytoprotectiveproperties of this drug. On the other hand, downregulation of theneuronal network activity may prevent microglial overactivation, with

a favourable effect on neurodegenerative diseases. In fact, minocyclinehas been demonstrated to inhibit microglial activation (Yrjanheikkiet al., 1999; He et al., 2001; Tikka et al., 2001), a finding that is inaccordance with its neuroprotective effects in animal models ofneurodegenerative diseases or ischaemia (Yrjanheikki et al., 1999;Chen et al., 2000; Zhu et al., 2002; Wang et al., 2003; Hunter et al.,2004).In conclusion, our observation that minocycline mitigates the

excitability of hippocampal neurons by direct blocking of ionicchannels involved in the generation and propagation of APs, anddepresses glutamate release and Ca2+ overloading by the partialblockade exerted on voltage-dependent Ca2+-channels, may explainthe well-studied neuroprotective properties of this tetracycline deriv-ative in various in vitro and in vivo models of neurotoxicity.

Acknowledgements

We acknowledge the financial support of Ministerio Educacion y Ciencia toJ. M. Hernandez-Guijo (grant BFU2004-07998 ⁄ BFI), to A. G. Garcıa (grantSAF2006-03589), to J. Sanchez-Prieto (grant BFU2004-01375), to L. Gandıa(grant SAF 2004–07307) and to J. Jordan (grant SAF2005-07919-C02-01),Mutua Madrilena to L. Gandıa, Consejerıa de Sanidad de Castilla-La Mancha(grant 04005-00) to J. Jordan, and Instituto de Salud Carlos III (grantRD06 ⁄ 0026) to J. Sanchez-Prieto. J. Carlos Gonzalez is a fellow of MinisterioEducacion y Ciencia. J. Egea is a fellow of Comunidad Autonoma Madrid. Wealso thank ‘Fundacion Teofilo Hernando’ for continued support.

Abbreviations

AP, action potential; DMEM, Dulbecco’s Modified Eagle’s Medium; GABA,c-aminobutyric acid; ICa, whole-cell Ca

2+ current; IK, outward K+ current; IKCa,

calcium-dependent outward K+ current; IKv, voltage-dependent outward K+

current; INa, whole-cell inward Na+ current; sEPSC, spontaneous excitatorypostsynaptic current.

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neuroprotectant minocycline depresses glutamatergic neurotransmission ... · neuroprotectant...

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