Abstract
Many proteins in the immune system are also expressed in the brain. One such class of immune proteins are T-cell receptors (TCRs), whose functions in T lymphocytes in adaptive immunity are well characterized. In the brain, TCRs are confined to neocortical neurons, but their functional role has not been determined. In mouse layer 1 neocortical neurons, TCR activation inhibited α7 nicotinic currents. TCRs modulated α7 currents via tyrosine phosphorylation of α7 nicotinic receptors (nAChRs) through src tyrosine kinases because eliminating lck kinase expression, coexpressing fyn kinase dead, or mutating tyrosine to alanine in α7 blocked the effect of TCR activation. We found that TCR stimulation decreased surface α7 nAChRs and reduced single-channel conductance. These results reveal that TCRs play a major role in the modulation of cholinergic neurotransmission in the brain mediated by α7 nAChRs and that this has a profound effect on regulating neuronal excitability.
Introduction
The CNS has been regarded as an immune privileged organ because of the blood–brain barrier and an immunosuppressive microenvironment (Sallusto et al., 2012). However, there is evidence that immune proteins, such as MHC, cytokines, and T-cell receptor (TCR) subunits, originally thought to have only immune function, are also expressed in CNS neurons (Boulanger et al., 2001; Syken and Shatz, 2003). TCRs are known for their role in adaptive immunity on T lymphocytes (Irvine et al., 2002), but this restrictive role in only immune system function has recently been challenged. TCR expression in the adult brain is restricted to the neocortex (Syken and Shatz, 2003). The TCR is an octameric complex of subunits where the αβ subunits bind to the peptide/MHC complex (Huang and Wange, 2004). T-cell antigen receptor β (TCRβ) along with MHC class I and CD3ζ are expressed in neurons, with the latter two proteins known to play a role in the induction of synaptic plasticity (Baudouin et al., 2008; Escande-Beillard et al., 2010). However, there is no reported evidence that TCRs can modify activity of any ligand-gated ion channel in the CNS. The earliest activated kinases after TCR stimulation are downstream src family kinases (Weiss and Littman, 1994).
α7 nicotinic acetylcholine receptors (nAChRs) are members of the pentameric cys loop family of ligand-gated ion channels. These receptors constitute the high-affinity α-bungarotoxin (α-BTX) binding sites in the CNS and are the second most prevalent nAChR subtype in the CNS after α4β2 (Chen and Patrick, 1997). α7 nAChRs play an important role in cognition, such as attention and memory (Levin et al., 2006; Young et al., 2007). These receptors are expressed most abundantly in the hippocampus and neocortex (Freedman et al., 1993; Christophe et al., 2002). The long intracellular loop between the third and fourth membrane-spanning regions of each α7 subunit contains putative protein kinase phosphorylation sites, including at least one tyrosine phosphorylation site (Charpantier et al., 2005). Tyrosine phosphorylation of α7 nicotinic receptors is known to modulate their activity (Charpantier et al., 2005; Cho et al., 2005).
Because α7 nAChRs and TCRs are both highly expressed in the cerebral cortex and TCRs can signal via tyrosine kinases, this opens the possibility that TCRs can have downstream effects on α7 nAChRs. We examined whether TCR activation can modulate α7 nAChRs in CNS neurons. We provide evidence that TCR activation inhibits α7 currents in cortical neurons. The negative modulatory effect of TCRs on α7 nAChR activity is mediated through activation of fyn and lck tyrosine kinases and the subsequent tyrosine phosphorylation of the cytoplasmic loop of α7. The TCR-negative regulation of α7 receptors was due to a loss of surface α7 receptors and a decrease in α7 single-channel conductance. Furthermore, we found that TCR activation decreased the excitability of neurons. Together, our results reveal a novel mechanism of modulation of neuronal excitability by altering ion channel function through phosphorylation mediated by activation of an immune receptor.
Materials and Methods
cDNA constructs.
Mouse α7 nAChR and human RIC-3 cDNA plasmids were kindly provided by Dr. Jerry Stitzel (University of Colorado Boulder, Boulder, CO) and Dr. Neil Millar (University of London, London, UK), respectively. RIC-3 is a chaperone protein that is a requirement for the functional expression of α7 nAChRs in many mammalian cell lines (Lansdell et al., 2005). Venus fluorescent protein was generously provided by Atsushi Miyawaki (Riken Brain Science Institute, Wako-shi, Saitama, Japan) (Nagai et al., 2002). We constructed a cDNA construct in which Venus fluorescent protein and hemagglutinin epitope tag were fused to α7 nAChR in the M3-M4 cytoplasmic loop (α7-Venus) and functions normally in every respect (Dau et al., 2013).
According to ProSite analysis, there is a single putative tyrosine kinase phosphorylation site in the M3-M4 cytoplasmic region of α7 at Y442. Using gene synthesis approach (Bioscience), the wild-type (WT) tyrosine 442 codon (TAC) was mutated to the alanine codon (GCT) in both α7 (α7(Y442A)) and α7-Venus (α7(Y442A)Venus).
Expression vectors for constitutively active fyn kinase (FKA) and fyn kinase dead construct (FKD) were kindly provided by Dr. Todd Holmes (New York University, New York, NY) (Nitabach et al., 2002).
Cell culture and transfection.
In this study, we cultured Jurkat cells (clone E6–1, catalog #TIB-152, ATCC). Jurkat cells are a clonal T lymphocyte cell line, which natively expresses TCRs. Other Jurkat cells used in the study include the Jurkat TCR β subunit knock-out (KO), in which there is loss of expression of the β subunit of the TCR (clone J.RT3-T3.5, catalog #TIB-153, ATCC) and the Jurkat lck KO cells, in which the cells are deficient in lck tyrosine kinase activity (JCaM 1.6, catalog #CRL-2063, ATCC). Jurkat cells were maintained in RPMI 1640 medium supplemented with 10% FBS, penicillin (100 U/ml), streptomycin (100 μg/ml), and 5% glutamine in a humidified CO2 incubator at 37°C.
We transiently transfected Jurkat cells using electroporation. On the day of electroporation, inside 35 mm Petri dishes, we coated 5-mm-diameter round glass coverslips (catalog #64–0700, Warner) with rat collagen Type 1 (0.05 mg/ml, catalog #92590, Millipore) for 3 h and washed twice with PBS, pH 7.4. In an electroporation cuvette (catalog #165–2088, Bio-Rad), 1 × 107 Jurkat cells were incubated with 6 μg of α7 nAChR cDNA, 6 μg of RIC-3 cDNA, and 0.6 μg of soluble Venus fluorescent protein cDNA, in 300 μl of incomplete RPMI 1640 medium and subjected to electroporation with a Gene Pulser Xcell (Bio-Rad) at 250 V and 960 μF. RPMI-1640 incomplete medium was identical to RPMI-1640 complete medium, except that FBS was omitted. Electroporated cells were then plated into the 35 mm dishes each containing 2.5 ml complete RPMI media. Whole-cell patch-clamp recordings were performed 2 d after electroporation. For experiments involving confocal imaging of cells, the Jurkat cells were plated onto collagen Type 1-coated coverslip bottom 35 mm dishes (catalog #P35G-0–14-C, MatTek).
HEK293T cells were also used in the study. They were grown in DMEM high glucose medium supplemented with 10% FBS, 2 mm l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin (DMEM complete medium) and maintained in a CO2 incubator at 37°C. Three to 6 d before electrophysiological recordings, the cells were incubated with trypsin for 3 min, mechanically dissociated, and seeded onto poly-dl-lysine (1 mg/ml, catalog #P9011, Sigma) coated 5 mm glass coverslips (catalog #64–0700, Warner) placed inside 35-mm-diameter Petri dishes. Cells were transfected at 60–70% confluency using Fugene Transfection Reagent (catalog #PRE2311, Promega). To each dish, 2 μg of α7 nAChR cDNA, 2 μg of RIC-3 cDNA, and 0.2 μg of soluble Venus fluorescent protein cDNA were mixed with 3 μl of Fugene transfection reagent and 250 μl of incomplete DMEM medium, which was identical to the complete DMEM, except lacking FBS. Transfection was performed according to the manufacturer's protocol. Electrophysiological recordings were performed 2–3 d after transfection.
Whole-cell patch-clamp recordings from cultured cells.
Cells were visualized with differential interference contrast illumination using an upright microscope (Nikon FN1) equipped with a CFI APO 40× W NIR objective (0.80 numerical aperture, 3.5 mm working distance). Transfected cells were identified with Venus fluorescent protein under fluorescence illumination with a mercury lamp. Standard whole-cell recordings were performed using a Multiclamp 700B amplifier (Molecular Devices) low pass filtered at 4 kHz and digitized at 10 kHz with a Digidata 1440A (Molecular Devices). Whole-cell patch-clamp recordings were performed using extracellular solution containing the following (in mm): 150 NaCl, 4 KCl, 2 CaCl2, 2 MgCl2, 10 HEPES, and 10 d-glucose adjusted to pH 7.4. Micropipette recording electrodes were pulled from borosilicate glass of 1.5 mm OD and 1.0 mm ID (catalog #1B150F-4, WPI) on a P-97 Flaming/Brown micropipette puller (Sutter Instruments). Patch electrodes (7–9 mΩ) were filled with pipette solution containing the following (in mm): 108 KH2PO4, 4.5 MgCl2, 0.9 EGTA, 9 HEPES, 0.4 CaCl2, 14 creatine phosphate (Tris salt), 4 Mg-ATP, 0.3 GTP (Tris salt), pH 7.4 with KOH. Atropine (100 nm) was present in the bath throughout all recordings to block muscarinic acetylcholine receptor responses. Series resistance was compensated 50%, and the membrane potential was held at −60 mV. Holding potentials were corrected for the liquid junction potential. ACh was delivered for 1 s duration using the two-barrel glass θ tube valve driven drug applicator, which was positioned ∼300 μm from the recorded cell (Komal et al., 2011). Solution exchange rates measured from open tip junction potential changes during application with 10% extracellular solution were typically <500 μs (10–90% peak time). The timing of agonist delivery and recordings was controlled using pCLAMP 10.2 acquisition software (Molecular Devices).
Whole-cell patch-clamp recordings from brain slices.
All experiments on mice were performed in accordance of the Canadian Council of Animal Care and approved by the Animal Care Committee of the University of Victoria. WT C57BL/6J mice, TCR β subunit KO mice (strain B6.129P2-Tcrbtm1Mom/J, stock no. 002118, The Jackson Laboratory) or α7 nAChR KO mice (strain B6.129S7-Chrna7tm1Bay/J, stock no. 003232, The Jackson Laboratory) at 10 to 15 d postnatal of either sex were anesthetized with isofluorane and decapitated. Subsequently, the brain was rapidly removed, kept for a minute in slicing solution, and sectioned coronally into 320 μm thick slices in oxygenated slicing solution with a vibratome (Leica 1000S). Slicing solution comprised of (in mm): 250 sucrose, 2.5 KCl, 1.2 NaH2PO4, 1.3 MgCl2, 2.4 CaCl2, 26 NaHCO3, and 11 d-glucose. Slices were transferred and incubated in a 37°C water bath for 1 h. Concanavilin A (ConA) (75 μg/ml) incubation time of 30 min was used for all our experiments. Slices were continuously bubbled in 95% O2 and 5% CO2 during incubation and recording.
Using infra-red video-assisted differential interference contrast illumination in combination with an upright microscope (Nikon FN1) equipped with a CFI APO 40× W NIR objective (0.80 numerical aperture, 3.5 mm working distance), whole-cell patch-clamp recordings were performed on layer 1 medial prefrontal cortical neurons from C57BL/6J mice (WT) and TCR β KO mice. Data were acquired using a Multiclamp 700B amplifier (Molecular Devices), low-pass filtered at 4 kHz, and digitized at 10 kHz with a Digidata 1440A A/D converter. Brain slices were perfused continuously with extracellular solution comprised of the following (in mm): 125 NaCl, 2.5 KCl, 1.2 NaH2PO4, 1.3 MgCl2, 2.4 CaCl2, 26 NaHCO3, and 11 d-glucose. Neurons were visualized with an upright microscope (Nikon FN1) equipped with a CFI APO 40× W NIR objective (0.80 numerical aperture, 3.5 mm working distance) using infra-red differential interference contrast and a video camera (IR-1000, Dage MTI). The patch electrodes had resistances between 9 and 11 mΩ when filled with pipette solution containing the following (in mm): 130 potassium gluconate, 5 EGTA, 0.5 CaCl2, 2 MgCl2, 10 HEPES, 3 Mg-ATP, 0.2 GTP, and 5 phosphocreatine Tris, pH adjusted to 7.4 with KOH, osmolarity adjusted to 300 mOsm with sucrose. Whole-cell voltage-clamp recordings were performed at room temperature with a MultiClamp 700B amplifier (Molecular Devices) and pCLAMP 10.2 software (Molecular Devices). Data were filtered at 4 kHz and sampled at 10 kHz with a Digidata 1440A data acquisition system (Molecular Devices). The membrane potential was corrected for liquid junction potential, and series resistance was corrected 50%. Neurons were held at −60 mV. The θ tube of the valve-driven drug applicator was positioned ∼600 μm from the recorded cell, and 100 μm PHA543613 hydrochloride (catalog #3092, Tocris Bioscience), a specific agonist for α7 nicotinic receptor, was applied for 1 s.
The firing frequency of layer 1 cortical neurons from brain slices of WT mice and TCR β KO mice for untreated and ConA treatment (75 μg/ml, 30 min) was measured in current-clamp mode of whole-cell configuration, with bridge balance correction. Methylycaconitine (MLA, 10 nm) was used in control experiments to identify the contribution by α7 nicotinic receptors toward the firing rate of layer 1 neurons. Current-clamp steps ranging from 0 pA to 200 pA (500 ms duration) in 20 pA increments were used to induce action potentials in cortical neurons. Firing frequency was calculated by dividing the number of action potentials by the 500 ms duration of each depolarizing step current. All data acquisition and analyses were performed using pClamp 10.2 software. Cells were not used for analysis if resting membrane potential (Vm) was more depolarized than −40 mV, access resistance (Ra) > 35 mΩ, or input resistance (Rinput) < 100 mΩ.
Current fluctuation analysis to estimate single-channel conductance.
Estimation of single-channel conductance of α7 nicotinic receptor was done by fluctuation analysis as previously described (Sigworth, 1980; Lambert et al., 1989; Gill et al., 1995; Brown et al., 1998). We used Clampfit 10.2 software (Molecular Devices) to conduct fluctuation analysis on the whole-cell current traces. Briefly, repeated whole-cell α7 currents evoked using 100 μm PHA543613 hydrochloride from brain slices of mice held at −60 mV were obtained. The variance of the current at each sample point of each trace was plotted against the mean current of the averaged traces at the same sample point in time. Then a linear fit was performed through the sampled points. The slope of the fit estimated the unitary current, I, of the nicotinic ion channel. The single-channel conductance was calculated using the equation γ = i/(Vh − Erev) where, Vh is the holding potential (−60 mV) and Erev is the reversal potential for α7 receptors, determined experimentally as (−0.7 mV).
Single-channel recordings.
Single-channel recordings in the cell-attached patch-clamp configuration were performed at room temperature on WT Jurkat cells electroporated with mouse α7 nAChR receptor and RIC-3 cDNA. The bath and pipette solutions contained the following (in mm): 150 NaCl, 4 KCl, 2 CaCl2, 2 MgCl2, 10 HEPES, 10 d-glucose, and 10−4 atropine adjusted to pH 7.4, with 100 μm ACh dissolved in the pipette solution. Micropipette recording electrodes were 1.5 mm OD and 1.0 mm ID borosilicate glass (catalog #1B150F-4, WPI) that were pulled on a P-97 Flaming/Brown micropipette puller (Sutter Instruments). The pulled pipettes were coated with Sylgard #184 (Dow Corning) and fire polished. Recording micropipettes had resistances between 6 and 15 mΩ. A pipette holding potential (Vp) of 60 mV was used throughout the recordings. Because recordings were made in the cell-attached configuration, this meant that the membrane potential underneath the patch of membrane was hyperpolarized by −60 mV in addition to the resting membrane potential. Single-channel currents were acquired using a Multiclamp 700B amplifier (Molecular Devices), low-pass filtered at 4 kHz, digitized at 50 kHz with a Digidata 1440A A/D converter (Molecular Devices), and collected with pClamp 10.2 software.
Clampfit 10.2 software (Molecular Devices) was used to analyze single-channel recordings to determine single-channel amplitudes and gating kinetics. Single-channel recordings were notch-filtered at 60 Hz followed by 4 kHz Gaussian filter. The “Event Detection–Single-Channel Search” feature in Clampfit was used to detect the open and closed channel events and analyzed for single-channel amplitudes and closed and open durations. Single-channel current amplitudes were analyzed by plotting an amplitude histogram fitted with two Gaussian functions: one Gaussian corresponded to the closed channel current level and the other corresponded to the open channel current level. Single-channel conductance was calculated in two ways. One way was dividing the single-channel amplitude by the driving force γ = i/(Vh − Erev) where, Vh was estimated to be −108 mV by taking an average of the resting membrane potentials recorded from whole-cell recordings from Jurkat cells (−48 mV) minus the pipette potential (60 mV) (Vh = Vm − Vp). Single-channel conductance was also verified by calculating the slopes of current–voltage relationships of single-channel recordings stepped at various pipette potentials. To assess gating kinetics, open and closed duration histograms of single-channel events were graphed on a semilog plot and fit with functions with multiple.
Immunoprecipitation and Western blot analysis.
Jurkat cells (1 × 107 cells/ml) were electroporated with 6 μg of α7-Venus, 6 μg of mouse RIC-3 in 300 μl of incomplete RPMI media. α7-Venus contains both a Venus fluorescent protein and an upstream HA epitope tag inserted into the M3-M4 cytoplasmic loop of the α7 nAChR subunit. Electroporated cells were then plated onto 35 mm dishes each containing 2.5 ml complete RPMI media. On the second day after transfection, WT α7-Venus receptor-containing Jurkat cells were treated with control solution (RPMI solution) or ConA for 30 min at 37°C. Before immunoprecipitation, cells were washed with ice-cold PBS (pH 7.4) at 4°C and resuspended in 1 ml freshly prepared immunoprecipitation buffer containing 50 mm Tris-HCl pH 8, 150 mm NaCl, 0.5% NP40, 0.5% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 1 mm EDTA, 2 mm sodium orthovanadate, 1 μg/ml pepstatin, 1 μg/ml leupeptin, 1 μg/ml aprotinin. The cells were lysed on ice for 10 min and clarified by centrifugation at 18,000 × g for 5 min. α7 receptors were immunoprecipitated using 2 μl of anti-HA antibody (clone 12ca5, gift from Ivan Sadowski, University of British Columbia) and incubated with lysate for 1 h before addition of 20 μl of BSA blocked protein A/G agarose beads (200 μg BSA in 1 ml immunoprecipitation buffer with beads) (catalog #20422, Pierce) for an additional hour with rotation. Beads were washed 3 times in 1 ml immunoprecipitation buffer, and immune complexes were released by briefly boiling in 25 μl SDS loading buffer. Proteins were resolved by SDS-PAGE and transferred to nitrocellulose membrane. Western blotting was performed using anti-HA antibody (clone 12ca5) at 1:5000 dilution to detect total cellular α7 nicotinic receptors. Tyrosine phosphorylation of α7 receptors was detected with anti-phosphotyrosine antibodies (1:2000 dilution, clone 4G10, Millipore). HRP-conjugated anti-mouse secondary antibody (catalog #NXA931, GE Healthcare) was used at 1:5000. Proteins were detected by chemiluminescence (GE Healthcare).
Surface α-BTX labeling and spectral confocal microscopy.
Jurkat cells were transfected with 6 μg of mouse α7-Venus or α7(Y442A)-Venus nAChR cDNA and 6 μg of mouse RIC-3. For cell surface receptor expression assays, 48 h after transfection cells were incubated on ice for 1 h to ensure that the temperature was <4°C. ConA was incubated (75 μg/ml) for 30 min at 37°C. Cells were then fixed with 4% paraformaldehyde in PBS for 10 min on ice and washed twice with ice-cold PBS. Surface α7 receptors were then labeled with Alexa Fluor 647 α-BTX (1:200, Fl-BTx, catalog #B35450, Invitrogen) under nonpermeabilized conditions for 30 min and washed twice with ice-cold PBS. Then the cells were plated on rat tail collagen Type I-coated dishes before imaging (0.05 mg/ml, catalog #92590, Millipore). Surface expression of Fl-BTx labeled WT α7-Venus receptors versus mutant α7(Y442A)-Venus receptors was examined with a Nikon C1si spectral confocal microscope system using a Plan Apo VC 60× 1.4 NA oil-immersion objective (0.13 mm working distance). A λ stack of X-Y images were collected simultaneously with one laser sweep onto an array of 32 photomultiplier tubes. Jurkat cells containing α7-Venus were imaged from 496.5 to 696.5 nm at 5 nm wavelength separation. Images were acquired at 512 pixels × 512 pixels, at 25 μm × 25 μm field of view and 12 bit intensity resolution. The pixel dwell time was set at 5.52 μs, and the pinhole was set to medium (60 μm diameter). Fl-BTx was excited with a 638 nm laser line at 15% maximum intensity, and emission was measured at the emission peak channel (665 nm). Images were analyzed for mean signal intensity using ImageJ v1.43r software. Using JACoP (just another colocalization plugin) in ImageJ, we also calculated the Mander's coefficients, M1 and M2, which quantify the degree of overlap between α7-Venus and FI-Bgt (Manders et al., 1993). M1 measures the percentage of pixels in the red channel that overlaps with the signal in the green channel, whereas M2 measures the percentage of pixels in the green channel that overlaps with signal in the red channel. M1 and M2 coefficients were calculated for both control and ConA-treated Jurkat cells.
Statistical analysis.
Values are reported as mean ± SE. Significant differences (p < 0.05) between two groups of data were determined using a t test for continuous data, meeting parametric assumptions of equal variances and normality. Otherwise, a Wilcoxon rank sum test was performed for nonparametric data. Comparison between three or more groups was analyzed using an ANOVA for parametric data followed by post hoc multiple pairwise analysis using a Tukey's HSD tests. For nonparametric data involving comparison of three or more groups of data, a Kruskal–Wallis rank sum test was performed followed by pairwise analyses using Wilcoxon rank sum tests. All statistical analyses were performed using the R statistical computing language (www.R-project.org).
Results
TCR activation decreases α7 nAChR responses in Jurkat cells
Because the TCR complex is involved in downstream transmembrane signaling through src family tyrosine kinases, and α7 nicotinic receptors are negatively regulated by tyrosine kinases, we examined whether TCR activation could modulate the function of α7 receptors.
To test our hypothesis, we used Jurkat cells, which are a human clonal T lymphocyte cell line that natively expresses TCRs. We activated endogenously expressed TCRs with 75 μg/ml concanavilin A (ConA), a lectin and exogenous agonist of TCRs (Palacios, 1982). We found that Jurkat cells transiently transfected with α7 nicotinic receptors and incubated for 30 min with ConA resulted in a decrease in nicotinic current responses elicited by rapid application of ACh (1 mm for 1 s) compared with Jurkat cells incubated with control solution (Fig. 1A). There was a significant decrease in the mean α7 peak current response in ConA-treated Jurkat cells (185 ± 34 pA, n = 14) compared with control (720 ± 113 pA, n = 14) (p < 0.0001, Wilcoxon rank sum test) (Fig. 1B).
TCR activation decreases α7 nAChR responses in Jurkat cells. A–D, Whole-cell recordings showing representative traces of ACh (1 mm, 1 s) induced α7 receptor currents in WT Jurkat cells (A) and TCR β KO Jurkat cells (C). ConA incubation (75 μg/ml, 30 min) significantly decreased α7 nAChR currents in WT Jurkat cells (n = 14) compared with control treatment (n = 14) (p < 0.0001, Wilcoxon rank sum test) (A,B), whereas ConA (n = 12) had no effect in TCR β KO Jurkat cells compared with control treatment (n = 12) (p = 0.4, Wilcoxon rank sum test) (C,D). E, F, Whole-cell recordings done in HEK293T cells transfected with α7 nicotinic receptor show that ConA incubation (75 μg/ml for 30 min) (n = 6) has no effect on ACh-mediated current responses compared with control treatment (n = 6) (p = 0.1, t test). Each circle superimposed on bar chart represent the average α7 nicotinic receptor response from each individual cell; bars represent mean ± SE of responses averaged across all measured cells.
To verify the role of TCR-mediated decrease in α7 nicotinic responses, we used a TCR β subunit KO Jurkat cell line. In this cell line, the absence of the β subunit renders the TCR receptor complex nonfunctional (Ohashi et al., 1985). Accordingly, the exogenous ligand ConA should not be able to activate the incomplete TCR complex. Indeed, we found no significant difference in the ACh-mediated α7 nicotinic currents between control (590 ± 147 pA, n = 12) and ConA (470 ± 128 pA, n = 12) treated TCR β subunit KO Jurkat cells (p = 0.4, Wilcoxon rank sum test) (Fig. 1D).
To further rule out the possibility that the effect of ConA was the result of a direct interaction with α7 nicotinic receptors, we used HEK293T cells, which are devoid of TCRs (Shaw et al., 2002). Similarly, we found no significant difference in α7 nicotinic currents in HEK293T cells treated with ConA (606 ± 114 pA, n = 6) compared with control treatment (407 ± 77 pA, n = 6) (p = 0.1, t test) (Fig. 1F).
Together, these results demonstrate that the decrease in α7 nicotinic current responses in Jurkat cells is a TCR activation-dependent event.
TCR activation decreases α7 nicotinic currents in layer 1 prefrontal cortical neurons
We next asked whether the phenomenon of TCR regulation of α7 receptors we observed in Jurkat cells could be found in CNS neurons. TCR β subunits are localized throughout all layers of the neocortex (Syken and Shatz, 2003), and α7 nicotinic receptors are present in layer 1 neocortical interneurons (Christophe et al., 2002) in addition to neurons in other cortical layers (Poorthuis et al., 2013). Thus, there is a high probability that TCRs and α7 nicotinic receptors are localized on the same neurons. We restricted our experiments to layer 1 of the medial frontal and prefrontal cortex, which is a simplified cortical layer essentially consisting almost entirely of GABAergic interneurons (Winer and Larue, 1989) and therefore is less heterogeneous than other layers, which contain pyramidal neurons and many subtypes of interneurons. To examine the functional interaction between TCRs and α7 receptors, we performed whole-cell patch-clamp electrophysiology on layer 1 cortical neurons of the medial prefrontal cortex
In WT mouse brain slices incubated in ConA, using the α7 selective agonist PHA543613 (100 μm for 1 s), we found that TCR activation significantly reduced α7-mediated currents (36 ± 6 pA, n = 10) compared with control-treated brain slices (80 ± 14 pA, n = 12) (p = 0.003, Wilcoxon rank sum test) (Fig. 2A). To isolate nicotinic responses, recordings were voltage-clamped at −60 mV and performed in the presence of TTX, CNQX, and atropine to inhibit action potential firing, glutamateric and muscarinic neurotransmission, respectively.
Activating TCRs decrease α7 nicotinic currents in layer 1 prefrontal cortical interneurons. A, C, Whole-cell recorded α7 nicotinic current traces elicited by the α7 specific agonist PHA543613 (PHA, 100 μm for 1 s) showed a significantly diminished α7 nicotinic current in brain slices from WT C57BL/6J mice treated with ConA (75 μg/ml for 30 min) (n = 10) compared with control treatment (n = 12) (p = 0.003, Wilcoxon rank sum test). Recordings were performed in the presence of TTX (200 nm) and CNQX (10 μm) to isolate nicotinic currents. B, C, No difference was observed in the amplitude of α7 nicotinic currents in brain slices of TCR β KO mice treated with ConA (n = 10) compared with control treatment (n = 10) (p = 0.7, Wilcoxon rank sum test). C, There was a significant increase in the mean baseline α7 nicotinic current amplitudes from neurons of TCR β subunit null mice compared with α7 nicotinic currents from neurons of WT mice (p = 0.02, Wilcoxon rank sum test).
To confirm that the ConA-mediated reduction in peak α7 nicotinic currents in brain slices was a TCR-mediated event, we repeated the recordings in brain slices from TCR β subunit KO mice (TCR β KO). We found no significant difference in α7 nAChR peak current responses recorded from layer 1 prefrontal cortical interneurons between ConA (156 ± 50 pA, n = 10) and control treatments (136 ± 41 pA, n = 10) (p = 0.7, Wilcoxon rank sum test) (Fig. 2B). These findings are consistent with our results in WT and TCR β subunit KO Jurkat cells (Fig. 1). Interestingly, we also found a significant elevation in the baseline α7 nicotinic responses from layer 1 cortical neurons in brain slices from TCR β KO mice (136 ± 41 pA, n = 12) compared with WT mice (80 ± 14 pA, n = 12) (p = 0.0004, Kruskal–Wallis rank sum test) (p = 0.02, Wilcoxon rank sum test post hoc analysis) (Fig. 2C). This result confirmed that TCRs, even in the absence of the exogenous compound ConA, had a basal activity that was sufficient to reduce α7 nicotinic currents in WT mice.
These data show that TCRs, which have traditionally been known to play an important role in adaptive immunity, also have a neuronal function in the CNS, which involves the negative regulation of function of α7 nicotinic receptors.
TCR activation inhibits α7 nicotinic currents via src family tyrosine kinases
The earliest signaling events downstream of TCR stimulation are the activation of protein tyrosine kinases and the subsequent tyrosine phosphorylation of multiple intracellular proteins (Zhang et al., 1998). One of the earliest activated kinases upon TCR activation are the src family tyrosine kinases (Parsons and Parsons, 2004). Previous work done by Charpantier et al. (2005) and Cho et al. (2005) has shown that α7 nicotinic receptors are prone to modulation by tyrosine kinases. Therefore, we asked whether two src family kinases involved in TCR signaling, fyn and lck, play a role in the modulation of α7 nicotinic responses after TCR activation.
To investigate TCR regulation of α7 nicotinic receptors involves tyrosine kinase action, we compared α7 peak current responses to ConA in the presence and absence of the broad-spectrum tyrosine kinase inhibitor genistein. Jurkat cells were transfected with α7 nAChRs, and whole-cell electrophysiology was performed to compare the peak current responses between four different treatment groups: (1) control treatment, (2) ConA treatment (75 μg/ml, 30 min), (3) genistein preincubation (10 μm, 20 min) followed by ConA (75 μg/ml, 30 min), and (4) genistein treated cells (10 μm, 20 min) (Fig. 3A). When we inhibited tyrosine kinases with genistein alone, α7 nicotinic currents increased significantly (2128 ± 335 pA, n = 14) compared with control (882 ± 149 pA, n = 20) (p = 0.0005, Wilcoxon rank sum test post hoc analysis) (Fig. 3B). Genistein when preincubated for 20 min before ConA stimulation significantly decreased the ACh-mediated responses (902 ± 175 pA, n = 15) relative to genistein alone (2128 ± 335 pA, n = 14) (p = 0.0009, Wilcoxon rank sum test). However, with genistein plus ConA stimulation α7 nicotinic responses (902 ± 175 pA, n = 15) were not significantly different from α7 responses for control treatment (882 ± 149 pA, n = 20) (p = 0.8, Wilcoxon rank sum test post hoc analysis). ConA alone (398 ± 62 pA, n = 14) had significantly attenuated α7 responses (p < 0.0001, Kruskal–Wallis rank sum test) (p = 0.02, Wilcoxon rank sum test post hoc analysis) relative to control treatment (882 ± 149 pA, n = 20). These results are consistent with previously published work showing genistein-mediated potentiation of α7 nicotinic receptor currents (Charpantier et al., 2005; Cho et al., 2005).
TCRs inhibit α7 nicotinic responses through activation of src family tyrosine kinases. Whole-cell patch-clamp recordings from Jurkat cells transfected with α7 nAChRs. A, B, ACh (1 mm for 1 s) induced α7 nAChR currents in Jurkat cells show that TCR activation by ConA (75 μg/ml for 30 min) (n = 14) significantly decreased α7 nAChRs responses compared with control treatment (n = 20) (p = 0.02, Wilcoxon rank sum test). Preincubation with the tyrosine kinase inhibitor genistein (10 μm for 20 min) (n = 15) before ConA stimulation blocked the effect of ConA-mediated decrease in α7 nicotinic current. Genistein alone (n = 14) resulted in a significant enhancement of α7 nicotinic currents compared with control treatment (n = 20) (p = 0.0005, Wilcoxon rank sum test). The genistein alone treatment had significantly enhanced α7 nicotinic currents compared with ConA preincubated with genistein (p = 0.0009, Wilcoxon rank sum test). C, D, ConA stimulation (n = 22) significantly reduced α7 nicotinic currents compared with control treatment (n = 39) (p = 0.01, Wilcoxon rank sum test). α7 receptor-mediated currents were significantly enhanced in cells cotransfected with a kinase dead form of fyn kinase (FKD) (n = 34) compared with control cells without FKD (n = 39) (p < 0.0001, Wilcoxon rank sum test), whereas cells cotransfected with FKD and treated with ConA (n = 7) still had augmented α7 currents compared with control (p = 0.01, Wilcoxon rank sum test). Cells cotransfected with a constitutively active fyn kinase (FKA) (n = 16) expressed significantly reduced α7 nicotinic responses compared with control cells (p < 0.0001, Wilcoxon rank sum test). E, F, α7 nAChR current traces from WT Jurkat cells and lck KO Jurkat cells. A significant decrease in α7 nAChR currents was observed between control-treated (n = 10) and ConA-treated (n = 13) WT Jurkat cells (p = 0.002, Wilcoxon rank sum test). However, there was no significant difference in α7 nAChR currents between control-treated (n = 13) and ConA-treated (n = 12) (p = 0.8, Wilcoxon rank sum test). These results suggest that both fyn and lck contribute to TCR-mediated decrease in α7 nAChR currents.
Because the lck and fyn kinases are the primary kinases activated upon TCR stimulation (Parsons and Parsons, 2004), we next asked whether these tyrosine kinases regulate the activity of α7 nicotinic receptors. To examine the role of fyn kinase signaling in TCR modulation of α7 receptors, we cotransfected Jurkat cells with α7 nAChRs and either the gain of function (FKA) or the loss of function (FKD) Fyn kinase expression vector. Whole-cell recordings were then performed to monitor ACh-induced α7 nicotinic currents (Fig. 3C). We observed that α7 peak currents increased robustly and significantly in Jurkat cells cotransfected with FKD (2286 ± 241 pA, n = 34) compared with that of control (970 ± 163 pA, n = 39) (p < 0.0001, Kruskal–Wallis rank sum test) (p < 0.0001, Wilcoxon rank sum test post hoc analysis), whereas ConA treatment alone reduced α7 current responses (501 ± 87 pA, n = 22) relative to control (p = 0.02, Wilcoxon rank sum test post hoc analysis) (Fig. 3D). However, in cells cotransfected with FKD, ConA stimulation in cells expressing FKD (2706 ± 631 pA, n = 7) did not lead to a significant change in the peak α7 current amplitude compared with control-treated FKD cotransfected cells (2286 ± 241 pA, n = 34) (p = 0.4, Wilcoxon rank sum test post hoc analysis). The abolition in TCR-mediated α7 current inhibition implicates the active role of fyn kinase in negatively modulating α7 nicotinic responses after TCR activation (Fig. 3C,D). When we cotransfected Jurkat cells with FKA and control treatment, it led to a significant decrease in the ACh-mediated α7 nicotinic responses (235 ± 39 pA, n = 16) compared with control treatment alone (p < 0.0001, Wilcoxon rank sum test post hoc analysis) (Fig. 3C,D). The reciprocal effect of FKA and FKD suggests that fyn kinase modulates α7 nicotinic receptor regulation and that fyn kinase is one key downstream effector after TCR activation.
Because lck kinase also plays a critical role in mediating phosphorylation of ITAM residues and downstream TCR-mediated signaling (Weiss et al., 1992), we tested the possibility for the involvement of lck kinase in TCR-mediated α7 nicotinic receptor regulation. We used Jurkat cells devoid of lck kinase (lck KO). There were four experimental groups of cells transfected with α7 nAChRs: (1) WT Jurkat cells control treatment, (2) WT Jurkat cells ConA-treated, (3) lck KO Jurkat cells control treatment, and (4) lck KO Jurkat cells ConA-treated. Whole-cell recorded ACh evoked α7 nicotinic receptor responses showed significant reduction in WT Jurkat cells upon ConA stimulation (261 ± 76 pA, n = 13) compared with control (1036 ± 308 pA, n = 10) (p < 0.001, Kruskal–Wallis rank sum test) (p = 0.002, Wilcoxon rank sum test post hoc analysis), consistent with previous data (Figs. 1 and 2). However, we observed no significant difference in α7 nicotinic currents between lck KO controls (1628 ± 420 pA, n = 13) and ConA-treated lck KO cells (842 ± 179 pA, n = 12) (p = 0.2, Wilcoxon rank sum test post hoc analysis) (Fig. 3F). Furthermore, neither lck KO control-treated cells (1628 ± 420 pA, n = 13) (p = 0.4, Wilcoxon rank sum test post hoc analysis) nor lck KO ConA-treated cells (842 ± 179 pA, n = 12) differed significantly (p = 0.8, Wilcoxon rank sum test post hoc analysis) in α7 nicotinic currents compared with control-treated WT Jurkat cells (1036 ± 308 pA, n = 10).
Together, these data strongly suggest that both lck and fyn kinases are required for TCR-mediated negative regulation of α7 nicotinic receptor function. Furthermore, these data suggest that fyn and lck are lined up in series and are not in parallel in the same TCR signaling pathway, either TCR-lck-fyn-α7 or TCR-fyn-lck-α7. Therefore, fyn and lck are each not redundant in function. However, there is one potentially paradoxical result in Figure 3B. Although ConA with genistein preincubation abolished the ConA-mediated decrease in α7 current and was not significantly different from control, ConA preincubated with genistein was still effective in decreasing the α7 current relative to genistein alone. This suggests that ConA was still effective in activating fyn and lck even when genistein preincubation, a general tyrosine kinase inhibitor, should inhibit these enzymes. The reason could be explained by the fact that genistein reversibly binds to src family of tyrosine kinases (Cho et al., 2005). Furthermore, unlike genistein, which must diffuse to its target and can also diffuse away, TCRs are structurally closely associated with fyn and lck kinases. Fyn coimmunoprecipitates with the TCR complex (Samelson et al., 1990), whereas lck physically interacts with the TCR coreceptor CD4 (Shaw et al., 1989). This may explain why TCR activation with ConA still attenuated α7 responses even with preincubation with genistein because the TCR has tighter association to fyn and lck than genistein.
Tyrosine 442 in the M3-M4 cytoplasmic loop of α7 nicotinic receptor is targeted by TCR activation
One mode by which TCR activation and fyn/lck kinases could affect α7 nAChRs is via receptor tyrosine phosphorylation. To test this, we monitored total tyrosine phosphorylation of α7 by immunoprecipitation and Western blots. Whole-cell extracts from Jurkat cells expressing α7-Venus-HA nictonic receptors, which have both a Venus fluorescent protein and an HA epitope tag, were generated (Dau et al., 2013), and α7 receptors were captured with anti-HA epitope antibody coated beads. Total α7 and tyrosine-phosphorylated α7 levels were monitored by Western blots using anti-HA and anti-4G10 antibodies, respectively (Fig. 4A). Activation of TCRs with ConA (30 min) showed immunoprecipitated α7 nAChRs with higher tyrosine phosphorylation signal compared with that of the control-treated cells (Fig. 4A,B). Similarly, α7 nAChRs in cells cotransfected with constitutively active fyn kinase showed greater tyrosine phosphorylation than control cells (Fig. 4A). We calculated a tyrosine phosphorylation index by dividing the α7 tyrosine phosphorylation signal by the total α7 nicotinic receptor expression and found that there was a significant increase in tyrosine phosphorylation of α7 nAChRs in ConA-treated cells compared with control treatment (Fig. 4B) (p = 0.02, Wilcoxon rank sum test). The bands analyzed were specific for α7 nAChRs as control immunoprecipitations with IgG-coated Sepharose beads were devoid of signal with either anti-HA or anti-phosphotyrosine antibodies (Fig. 4A). Thus, both TCR activation and fyn can potentiate tyrosine phosphorylation of α7 nAChRs.
TCR activation phosphorylates tyrosine 442 of α7 nicotinic receptors to decrease nAChR function. A, An immunoprecipitation and Western blot experiment demonstrates that TCR activation results in an increase in tyrosine phosphorylation of α7-Venus nAChRs, which also include an HA epitope tag. Anti-HA antibody-coated Sepharose beads are used to immunoprecipitate α7-Venus nAChRs and then probed on Western blots with either anti-phosphotyrosine antibody to determine phosphorylated α7-Venus nAChRs or probed on a parallel blot with anti-HA antibody to determine total α7-Venus nAChRs. The first three lanes are negative controls of Jurkat cell samples expressing α7-Venus nAChRs and immunoprecipitated using IgG antibody. The next three lanes are samples immunoprecipitated with anti-HA antibody for Jurkat cells expressing α7-Venus nAChRs that had undergone control treatment, ConA treatment, and coexpressing a constitutively active form of fyn kinase (FKA). B, Quantitative analysis of the density of the bands of control versus ConA treatment in which the phosphotyrosine signal is normalized to the anti-HA signal shows that ConA treatment (n = 4) significantly enhanced tyrosine phosphorylation of α7-Venus nAChRs compared with control treatment (n = 4) (p = 0.02, Wilcoxon rank sum test). C, Whole-cell α7 nicotinic current traces elicited with 1 mm ACh (1 s) and recorded from transfected Jurkat cells decreases in peak current amplitude after TCR activation with 30 min of ConA (75 μg/ml) incubation. D, Averaged data show a significant reduction in the mean current response between control (n = 10) and ConA (n = 9) (p = 0.0004, Wilcoxon rank sum test) treated Jurkat cells. E, In Jurkat cells transfected with the mutant α7(Y–A) nicotinic receptors, in which Y442 in the M3–M4 cytoplasmic loop, was mutated to alanine, no difference was observed in peak nicotinic current responses between control and ConA-treated Jurkat cells. F, Bar charts illustrating no significant difference (p = 0.7, Student's t test) in ACh-mediated mutant α7(Y442A) nAChRs responses between control (n = 6) and ConA (n = 6) stimulated Jurkat cells.
Using Prosite analysis of the amino acid sequence of the M3-M4 cytoplasmic loop of the α7 nicotinic receptor, we identified a putative tyrosine phosphorylation site at Tyr 442 of the receptor (EEVRYIANR). There are two other tyrosines (Y317 and Y386) in the α7 M3-M4 loop (IVLRYHHHD and GNLLYIGFR, respectively), which were not recognized as consensus sites for tyrosine kinase phosphorylation by Prosite. Furthermore, Y442 is conserved over all 11 mouse neuronal nicotinic receptor subunits, whereas neither Y317 nor Y386 is conserved in any other neuronal nicotinic receptor subunit. Thus, we focused our attention to investigate whether the effects of TCR activation and fyn tyrosine kinase were directed against the consensus tyrosine phosphorylation site at Y442 of the α7 nicotinic receptor.
To study the role of phosphorylation of α7 Tyr 442 on TCR regulation of α7 nicotinic receptor function, we mutated α7 Tyr 442 into alanine (Y442A). Whole-cell recordings were performed on WT Jurkat cells expressing mutant α7 nictonic receptors (Fig. 4E,F). TCR activation with ConA stimulation for 30 min (300 ± 54 pA, n = 6) resulted in no significant difference in the ACh-evoked α7 nicotinic current responses compared with control treatment (255 ± 56 pA, n = 6) (p = 0.7, t test) (Fig. 4E,F). However, TCR activation with ConA in the Jurkat cells expressing WT α7 nictonic receptors showed a significant decrease in peak response (107 ± 35 pA, n = 10) compared with that of control treatment (502 ± 119 pA, n = 9) (p = 0.0004, Wilcoxon rank sum test) (Fig. 4C,D). These data strongly suggest that fyn/lck kinases modulate α7 nicotinic receptor function by directly targeting Y442 of α7 nAChRs.
TCR activation decreases the number of surface α7 nicotinic receptors
There is conflicting evidence regarding whether tyrosine kinases affect trafficking of α7 nicotinic receptors (Charpantier et al., 2005; Cho et al., 2005). Because we found that TCRs signal through fyn and lck kinases to decrease α7 receptor function, we examined whether the effect of TCR activation in Jurkat cells could be attributed to a decrease in the number of surface receptors. To test this, we imaged Alexa Fluor 647-conjugated α-BTX (Fl-BTx) binding on the surface of Jurkat cells transfected with α7-Venus under nonpermeablized conditions and normalized the surface bound Fl-BTx to the total cellular fluorescence from α7-Venus. ConA incubation (75 μg/ml for 30 min) significantly decreased the number of surface Fl-BTx bound α7 nAChRs (0.29 ± 0.05, n = 58) compared with that of the control-treated cells (0.50 ± 0.07, n = 58) (p = 0.0003, Wilcoxon rank sum test) (Fig. 5). However, the total cellular α7-Venus intensity remained the same for both treated and untreated cells (data not shown), implicating no change in the total level of protein.
TCR activation decreases the number of α7 nAChRs expressed at the cell surface. A–F, Confocal images of Jurkat cells transfected with α7-Venus. Surface labeling of α7-Venus is performed with Alexa Fluor 647-conjugated α-BTX (Fl-BTx) under nonpermeabilizing conditions. ConA (75 μg/ml for 30 min) stimulation (n = 58) resulted in a significant decrease in the amount of surface α7-Venus receptors as determined with Fl-BTx labeling compared with control treatment (n = 58) (p = 0.0003, Wilcoxon rank sum test) (G). H–M, Confocal images of Jurkat cells transfected with a mutant α7-Venus in which the putative tyrosine phophorylated site (Y442) was mutated to alanine (α7(Y–A)Venus). N, Upon incubation with ConA (75 μg/ml for 30 min) (n = 63), there was no significant change in Fl-BTx bound surface α7-Venus nAChRs compared with control treatment (n = 58) (p = 0.09, Wilcoxon rank sum test). Both bar charts are displayed in logarithmic scale and represent the mean ± SE with each circle showing the measure from each individual cell. O, Control experiment demonstrating the specificity of Fl-BTx. Preincubation with 10 nm MLA (n = 26) is able to compete with and significantly lower Fl-BTx labeling compared with control treatment (n = 26) (p < 0.0001, Wilcoxon rank sum test).
In Jurkat cells transfected with mutant α7-Venus in which the putative tyrosine phosphorylation site, Tyr 442, is mutated to alanine (α7(Y-A)Venus), ConA activation of TCRs did not show any significant change in surface Fl-BTx bound α7 nAChRs (0.30 ± 0.03, n = 63) compared with control treatment (0.22 ± 0.02, n = 58) (p = 0.09, Wilcoxon signed rank test) (Fig. 5H–N). We demonstrated the specificity of Fl-BTx labeling of α7-Venus receptors because preincubation of WT α7-Venus transfected Jurkat cells with 10 nm MLA (0.26 ± 0.05, n = 26) was successful in competing for binding sites and significantly lowered (p < 0.0001, Wilcoxon rank sum test) Fl-BTx signal compared with cells without MLA (0.79 ± 0.11, n = 26) (Fig. 5O).
Furthermore, we analyzed the specificity of Fl-BTx labeling of α7-Venus receptors by performing quantitative analysis of colocalization via calculation of the Mander's coefficients, M1 and M2, the proportion of colocalized α7-Venus pixels to Fl-BTx pixels and vice versa, respectively. We compared these Mander's coefficients with those calculated when the Fl-BTx was rotated 90° counterclockwise relative to the α7-Venus image, which should have less colocalization. We found that the Mander's coefficient of Fl-BTx colocalized to α7-Venus for both control (0.44 ± 0.03) and ConA (0.44 ± 0.03) treatment was significantly greater (p = 0.002, and p = 0.0005, Wilcoxon rank sum tests, respectively) than colocalization in the 90° rotated images for control (0.30 ± 0.03) and ConA treatments (0.28 ± 0.02).
Thus, these measurements of Mander's colocalization coefficients validated the specificity of Fl-BTx labeling of α7 receptors and confirmed that TCR activation decreases the surface expression of α7 nAChRs without changing the amount of total cellular receptors through phosphorylation of Tyr 442.
TCR activation decreases single-channel conductance of α7 nicotinic receptors
A second potential mechanism for the inhibition of nicotinic current after TCR activation is a decrease in intrinsic channel function. Because Y442 is located in the amphipathic helix of α7, which lines the ion permeation pathway, the addition of a phosphate group here could hinder ion conductance because of added steric hindrance. Hence, we used ionic current fluctuation analysis of whole-cell recorded nicotinic currents in brain slices to calculate single-channel conductance. Ionic current fluctuation analysis is a reliable tool to estimate single-channel conductance of ion channels (Sigworth, 1980, 1981; Gill et al., 1995). Using this method, we aimed to determine whether a decrease in single-channel conductance of α7 nicotinic receptors contributed to the mechanism of TCR-mediated decrease in α7 whole-cell current responses. Examples of whole-cell recorded α7 nicotinic currents elicited with PHA543613 and the corresponding AC-filtered α7 nicotinic current (showing current fluctuation) are shown for each experimental condition (Fig. 6). Current fluctuations are maximal during the peak of the nicotinic current response and represent maximal number of channel openings. PHA543613 is applied repeatedly, and the variance of the current fluctuations of each trace is plotted against the mean current of the whole-cell current at each time point. The slope of the relationship gives the unitary current and when divided by the driving force (Vh − Erev) equals the single-channel conductance (γ).
TCR activation attenuates single-channel conductance of α7 nAChRs. A, B, Current variance-current relations for fluctuations of PHA543613 (100 μm, 1 s) elicited α7 nicotinic whole-cell currents recorded for both control (A) and ConA treatments (B) from layer 1 interneurons from WT mice. Single-channel conductances (γ) were calculated from the slope of current variance versus mean current response. Inset, Whole-cell α7 nicotinic current waveforms and their respective AC-filtered waveforms for control (A) and ConA treatments (B). C, The mean single-channel conductance of α7 responses for ConA-treated slices (n = 8) was significantly lower than that of control (n = 12) (p = 0.02, Wilcoxon rank sum test). D, E, Current variance versus mean current relations for α7 nicotinic whole-cell currents recorded for both control (D) and ConA treatments (E) from layer 1 interneurons from TCR β KO mice. F, There was no significant difference in the mean single-channel conductance for α7 receptors between control (n = 7) and ConA-treated slices (n = 5) of TCR β KO mice (p = 0.5, Wilcoxon rank sum test).
The α7 receptor-mediated inward currents recorded from layer 1 prefrontal cortical interneurons of control-treated brain slices from WT mice (118 ± 34 pS, n = 12) were accompanied by a significantly greater calculated single-channel conductance than those of ConA-treated WT brain slices (30 ± 12 pS, n = 8) (p = 0.02, Wilcoxon rank sum test) (Fig. 6A–C). Exemplary graphs of current variance versus current mean were plotted and shown for α7 nicotinic receptor single-channel conductances showing 61 pS and 22 pS, respectively, for control and ConA incubated brain slices (Fig. 6A,B). To verify that TCR activation with ConA is responsible for the change in single-channel conductance, we performed fluctuation analysis of whole-cell recorded α7 nicotinic currents from TCR β KO mice (Fig. 6D–F). We calculated the single-channel conductance and observed no significant difference between brain slices treated with ConA (130 ± 68 pS, n = 4) compared with brain slices with control treatment (118 ± 23 pS n = 8) from TCR β KO mice (p = 0.5, Wilcoxon rank sum test) (Fig. 6F). Plots of exemplary data of current fluctuation analyses are shown for control-treated and ConA-treated brain slices of TCR β KO mice exhibiting single-channel conductances of 101 pS and 67 pS, respectively (Fig. 6D,E).
We further investigated the effects of TCR activation on modulating α7 channel gating kinetics recorded from layer 1 interneurons from WT brain slices. To examine the activation rate, we fitted the rise to peak of the α7 current with a single exponential and showed that there was no significant difference between the activation time constant for control treatment (6.4 ± 0.8 ms, n = 11) and ConA treatment (5.6 ± 0.7 ms, n = 7) (p = 0.8, Wilcoxon rank sum test) (Fig. 7). The time course of the α7 decay kinetics during PHA543613 application was well fit by the sum of two exponentials. Similarly, ConA application did not result in any significant change in either the slow (control: 57.0 ± 7.4 ms, n = 11 vs ConA: 62.1 ± 16.4 ms, n = 7) or fast time constants (control: 15.0 ± 2.5 ms, n = 11 vs ConA: 14.3 ± 2.2 ms, n = 7) (p = 0.9, Wilcoxon rank sum test, for both slow and fast time constants).
TCR activation does not alter gating kinetics of α7 nAChRs. A, B, Whole-cell recorded PHA-543613 (100 μm, 1 s) activated α7 nicotinic current traces from control-treated (Aa) and ConA-treated (Ba) layer 1 interneurons of medial prefrontal cortical brain slices. Insets, Higher time resolution comparable rates of activation rise times of α7 nicotinic currents for both control (Ab) and ConA treatments (Bb). C, There was no significant difference in the activation time constant between control (n = 11) and ConA (n = 7) treatments (p = 0.8, Wilcoxon rank sum test). D, Furthermore, ConA treatment did not result in a significant change in either the fast (control: 15.0 ± 2.5 ms, n = 11 vs ConA: 14.3 ± 2.2 ms, n = 7) or the slow (control: 57.0 ± 7.4 ms, n = 11 vs ConA: 62.1 ± 16.4 ms, n = 7) decay time constants (p = 0.9, Wilcoxon rank sum test, for both fast and slow time constants).
These data demonstrate that TCR activation inhibits α7-mediated whole-cell currents by decreasing their single-channel conductance but does not alter their gating kinetics.
Single-channel recordings verify that TCR activation reduces α7 nicotinic receptor single-channel conductance
To verify the whole-cell fluctuation analysis data that TCR activation decreases single-channel conductances of α7 nicotinic receptors, we performed single-channel recordings in cell-attached configuration from WT Jurkat cells exposed to either control or ConA (75 μg/ml for 30 min) solutions. Single-channel α7 nicotinic receptor-mediated currents were activated by having ACh (100 μm) in the patch pipette electrode solution. The extracellular solution was identical to the patch pipette recording solution minus ACh. The patch of membrane in cell-attached mode was voltage-clamped at a pipette potential of 60 mV. This would equal a transmembrane potential of −60 mV plus the resting membrane potential of the cell, which was on average −48 mV. Thus, the estimated transmembrane potential was −108 mV. When plotting histograms of the current amplitudes of the fitted open single-channel events, there was a significant reduction (p < 0.0001, Wilcoxon rank sum test) in α7 single-channel amplitudes of ConA-treated cells (2.4 ± 0.0 pA, n = 13902 open channel events >4 cells) compared with control treatment (6.0 ± 0.1 pA, n = 1471 open channel events >4 cells) (Fig. 8A–D). This corresponds to single-channel conductances of 22.0 ± 0.1 pS for ConA-treated cells and 55.5 ± 0.6 pS for control-treated cells. We also verified the conductance change by measuring the slope conductance of single-channel currents measured over various holding pipette potentials. Using this paradigm, we examined a control-treated cell with cell-attached single-channel currents having a higher slope conductance (41.4 pS) than a ConA-treated cell (14.6 pS) (data not shown).
Single-channel recordings show that TCR activation decreases single-channel conductance of α7 nAChRs. A, Examples of Jurkat cell-attached single-channel traces for control and ConA treatment. The ConA-treated single-channel responses have smaller single-channel current amplitudes than control treatment. The patch electrode solution contained 100 μm ACh. B, Histograms of fitted amplitudes from single-channel events from four separate patches show that ConA treatment had consistently smaller single-channel amplitudes than control treatment. C, D, Histograms of single-channel amplitudes showing the combined data of all patches recorded for control (n = 4) and ConA (n = 4) treated Jurkat cells showing that ConA treatment caused a significant reduction of the amplitudes of open single-channel events (p < 0.0001, Wilcoxon rank sum test). Only the amplitudes of open single-channel events are shown for clarity. E, F, Histograms of open duration of single-channel events compiled from all patches show no difference in distributions between control and ConA treatment. G, H, The histograms of closed duration of single-channel events show similar kinetics between ConA and control-treated cells.
To determine the effects of TCR activation on the gating kinetics of α7 nicotinic receptors, we plotted histograms of durations of single-channel open events and closed events for both control and ConA treatments (Fig. 8E–H) and then fitted the histograms to multiple exponential functions. Each histogram represents the combined data from four separate cell-attached patch-clamp recordings. We report the fitted time constants and the SEs of fit. For control-treated cells, the open channel duration histogram was fitted to a sum of two exponentials with time constants: τ1 = 0.12 ± 0.32 ms and τ2 = 0.66 ± 0.49 ms. For ConA-treated cells, the open channel duration histogram was fitted with a sum of two exponentials with τ1 = 0.07 ± 0.10 ms and τ2 = 0.42 ± 0.21 ms. Therefore, for the duration of the single-channel open events, there was no significant difference in the time constants between control (n = 4 patches) and ConA treatments (n = 4 patches) (p > 0.25, Wilcoxon rank sum test). Similarly, the durations of the closed single-channel events did not significantly differ between control (n = 4 patches) and ConA treatments (n = 4 patches) (p > 0.15, Wilcoxon rank sum test). Closed single-channel duration events for control-treated cells were fit with the sum of four exponentials with τ1 = 0.20 ± 0.12 ms, τ2 = 1.95 ± 0.13 ms, τ3 = 33.7 ± 0.2 ms, and τ4 = 1311.8 ± 0.2 ms. For ConA-treated cells, the closed single-channel durations were well fit with the sum of five exponentials: τ1 = 0.10 ± 0.10 ms, τ2 = 0.47 ± 0.13 ms, τ3 = 3.42 ± 0.09 ms, τ4 = 29.1 ± 0.1 ms, and τ5 = 227.0 ± 0.1 ms.
Thus, our single-channel data corroborate the whole-cell recorded current fluctuation analysis that TCR activation attenuates α7 mediated single-channel conductance while having little effect on gating kinetics.
TCR activation decreases action potential firing frequency of layer 1 cortical neurons
We examined the physiological consequence of decreasing α7 nicotinic receptor currents after TCR activation by measuring action potential firing rate of layer 1 cortical interneurons. Because most cortical neurons in brain slices lack the intrinsic ability to spontaneously fire action potentials resulting from severed afferent inputs, we performed current-clamp recordings in whole-cell configuration mode from layer 1 interneurons and applied depolarizing constant current steps (0–200 pA, 20 pA steps for 500 ms). To ascertain the excitability of the recorded neuron, we plotted input–output curves of current injection versus action potential frequency.
To test whether α7 nicotinic receptors contribute to the firing rate of layer 1 prefrontal cortical interneurons, we bath-applied 10 nm MLA (α7 nAChR competitive antagonist) (n = 7) and noticed a significant decrease in the action potential firing frequency compared with baseline control (n = 7) (p < 0.0001, MLA factor, two-way ANOVA) (Fig. 9A–C). All the recorded neurons included in the analysis had stable resting membrane potentials that were more negative than −60 mV and overshooting action potentials. There was no significant difference in the mean resting membrane potential and Rinput between control and MLA-treated slices (data not shown). If an α7 antagonist decreased neuronal excitability, then a specific α7 agonist, PHA543613, should increase neuronal excitability. Indeed, when PHA543613 (100 μm for 575 ms) was applied 75 ms preceding and during a 200 pA (500 ms) depolarization, there was a significant increase (p = 0.03, n = 7, paired t test) in action potential firing frequency (30 ± 2 Hz) in the same neuron compared with when there was only 200 pA depolarization but no costimulation with PHA543613 (27 ± 2 Hz) (Fig. 9G,H).
TCR activation modulates neuronal excitability of layer 1 cortical interneurons. A, B, Firing frequency of layer 1 cortical neurons is modulated by α7 nicotinic receptors as evidenced by the decrease in action potential firing in the presence of a specific α7 antagonist, MLA (10 nm) in brain slices from WT mice. Current-clamp recordings are shown for current steps (120, 140, and 160 pA). C, The action potential firing rate of layer 1 interneurons in response to current steps (0–200 pA) was significantly lower with MLA treatment (n = 7) compared with control treatment from WT mouse brain slices (n = 7) (*** p < 0.0001, MLA factor, two-way ANOVA). E, F, A significant increase in the action potential firing rate was observed in layer 1 cortical neurons from control-treated TCR β KO mice (n = 10) compared with control-treated WT mice (n = 7) (***p < 0.0001, genotype factor, two-way ANOVA) over current steps ranging between 0 and 200 pA. A, D, F, WT brain slices with ConA treatment (n = 9) resulted in a significant decrease in the action potential firing rate of layer 1 cortical interneurons compared with control-treated WT mouse brain slices (n = 7) (***p < 0.0001, two-way ANOVA) (current steps ranged between 0 and 200 pA). G, H, There was a significant increase (p = 0.03, n = 7, paired t test) in action potential frequency upon PHA543613 (PHA, 100 μm) coapplication with 200 pA than with 200 pA depolarizing stimulation alone from brain slice recordings obtained from WT mice. I, Current-clamp recordings from layer 1 interneurons from α7 nAChR KO mice show that ConA treatment (n = 6) displays similar neuronal excitability compared with control treatment (n = 11) (p = 0.25, ConA vs control treatment factor, two-way ANOVA). Each connected pair of circles represents the frequency before and after PHA application for one cell. J, In brain slices of WT mice, there was a significantly greater attenuation in action potential firing frequency with ConA treatment (n = 9) compared with ConA treatment of α7 nAChR KO mice (n = 6) (p = 0.005, Wilcoxon rank sum test). J, Each circle represents the mean ConA-mediated decrease in firing frequency for one specific current stimulation.
Because TCR activation results in the inhibition of α7 nicotinic receptor currents, to examine the involvement of TCRs in modulating the firing rate of layer 1 interneurons, we compared whole-cell current-clamp recordings of control-treated brain slices from WT mice (n = 7) with ConA-treated brain slices from WT mice (n = 9). We found a significant decrease in the mean firing frequency of ConA-treated WT brain slices (18 ± 4 Hz) compared with control-treated WT brain slices (26 ± 3 Hz) (p < 0.0001, treatment factor, two-way ANOVA) (Fig. 9A,D,F).
If ConA-mediated activation of TCRs caused a decrease in neuronal firing rate due to decreased α7 nicotinic receptor currents, then we would predict an increase in firing rate in TCR β subunit KO mice because baseline α7 nicotinic currents in TCR β KO mice have elevated α7 nicotinic currents in brain slices (Fig. 2C). A comparison of the mean action potential firing frequency showed a significant increase in neurons recorded from control-treated brain slices of TCR β KO mice (40 ± 5 Hz, n = 10) compared with neurons from control-treated brain slices of WT mice (26 ± 3 Hz, n = 7) (p < 0.0001, genotype factor, two-way ANOVA) (Fig. 9A,E,F).
We performed current-clamp experiments in layer 1 interneurons from brain slices of α7 nAChR KO mice to determine whether TCR modulation of neuronal excitability was through α7 receptors. ConA incubation (n = 6) in α7 nAChR null slices showed similar levels of neuronal excitability as control treatment (n = 11) for much of the range of stimulation intensities (p = 0.25, ConA vs control treatment, two-way ANOVA) (Fig. 9I). Indeed, when averaging over all current stimulation intensities the ConA-mediated decrease in frequency in neuronal firing from α7 nAChR null mice (n = 6) was significantly less than the ConA-mediated decrease in neuronal firing frequency in WT mice (n = 9) (p = 0.005, Wilcoxon rank sum test) (Fig. 9J). This supports that TCR activation is attenuating neuronal excitability through modulation of α7 nAChRs.
This set of data shows that α7 nAChRs contribute to the excitability of layer 1 cortical neurons and that TCR-mediated inhibition of α7 receptor function contributed to reduced neuronal excitability. The source of ACh to activate α7 nAChRs in the cortex may be from cut cholinergic terminals from neurons originating from the nucleus basalis magnocellularis. There has even been a recent report showing local cholinergic interneurons in the cerebral cortex (von Engelhardt et al., 2007).
Discussion
This study shows, for the first time, that the immune protein, the TCR, can modulate nAChR function and neuronal activity in the brain. Activating TCRs decreases the function of α7 nAChRs in Jurkat cells and in layer 1 interneurons of the medial prefrontal cortex. The mechanism of TCR-mediated dampening of α7 nicotinic currents was the result of downstream activation of src family tyrosine kinases, namely, fyn and lck kinases. TCRs' effect of attenuating α7 nAChR responses is mediated through Y442 of α7 because mutating Y442 to alanine in the α7 nAChR blocked TCRs' negative modulation. TCR activation leads to a decrease in the number of surface α7 nAChRs. TCR activation also decreases α7 single-channel conductance. Furthermore, TCRs and α7 nAChRs influence neuronal excitability. Inhibition of α7 receptors decreases the frequency of action potentials. Activation of TCRs also inhibited the neuronal firing frequency, whereas neurons from TCR β KO mice exhibited enhanced firing rate of action potentials. Thus, the TCR-mediated regulatory mechanism of α7 nAChR function illustrates a novel role of TCRs in the CNS affecting ligand-gated ion channel function and neuronal excitability.
TCRs modulate neural function and α7 nAChR activity
To our knowledge, this is the first demonstration that TCRs in the CNS can modify neuronal activity and that one of the mechanisms is through modulation of α7 nAChR function. We have shown that TCRs can decrease α7 currents in prefrontal cortical interneurons (Fig. 2). Our data also show that there is endogenous activation of TCR-mediated decrease of α7 responses in the CNS because TCR β subunit KO mice had significantly larger α7 currents than WT mice (Fig. 2). We also observed that TCRs modulate neuronal excitability. TCR activation resulted in a significant decrease in current-evoked action potential firing, whereas in TCR β subunit KO mice neurons had a significantly enhanced action potential firing (Fig. 9). The TCR is an octameric complex that includes the CD3ζ subunit. CD3ζ plays an important role in dendritic structure and development in cultured hippocampal and cortical neurons (Baudouin et al., 2008).
TCRs decrease α7 nAChR function through phosphorylation of tyrosine 442
Our Western blot and electrophysiology data support that TCRs have effect through phosphorylation of α7 receptors at tyrosine 442 (Fig. 4), which is the only putative tyrosine phosphorylation site within the cytoplasmic loop of α7 recognized by ProSite analysis. We show that both fyn and lck kinases are involved in TCR-mediated attenuation of α7 nicotinic responses because FKD or deletion of lck both prevented ConA from decreasing α7 activity (Fig. 3). Overexpressing FKD successfully competed with endogenously expressed Fyn kinase. Because genistein, a broad-spectrum inhibitor of tyrosine kinase, and FKD both augmented α7 currents (Fig. 3), our results are consistent with the results of Charpantier et al. (2005) and Cho et al. (2005). Furthermore, we were able to abolish the effect of ConA-stimulated TCR-mediated decrease in α7 responses by mutating tyrosine 442 to alanine. This is consistent with the results of Charpantier et al. (2005) who also mutated tyrosine 386 and tyrosine 442 to alanines but did not distinguish the effect of each tyrosine. We propose that tyrosine 442 is the key tyrosine that is phosphorylated and induces altered α7 nAChR function after TCR activation. However, our results are at odds with that of Cho et al. (2005), who mutated tyrosine 317, 386, or 442 to phenylalanine without any effect on genistein-mediated potentiation of α7. One potential caveat of site-directed mutagenesis is that mutating one amino acid into another may have more than the intended consequence. Mutating tyrosine to another amino acid may alter the structure of the protein because the functional properties of amino acid side groups and their degree of hydrophobicity play a role in stabilizing protein conformation (Yutani et al., 1987). In our study, as in those of Charpantier et al. (2005), the mutation was made to alanine, whereas Cho et al. (2005) mutated the residue to phenylalanine, which may explain the discrepancy.
Mechanisms of TCR-mediated decrease of α7 currents
Our exploration of the mechanisms of TCR-mediated decrease in nAChR function indicate two contributing parallel steps: (1) a decrease in surface α7 receptors; and (2) a decrease in single-channel conductance. Our results showing a decrease in cell surface receptors upon TCR stimulation using Fl-BTx binding are consistent with the results of Cho et al. (2005), who showed that inhibition of tyrosine kinase with genistein enhanced surface α7 receptors, whereas Charpantier et al. (2005) showed no alterations in surface α7 receptors when inhibiting tyrosine kinase activity. This TCR-mediated decrease in surface receptors is mediated by phosphorylation of tyrosine 442 because mutation of this residue to alanine in α7 completely abolished TCRs' effect of decreasing surface α7 receptors (Fig. 5).
Interestingly, tyrosine 442 lies within the amphipathic helix. The amphipathic helix lines the cytoplasmic side portals of cys-loop receptors and forms part of the ion permeation pathway (Hales et al., 2006). We found that TCR activation decreases the single-channel conductance of α7 receptors (Figs. 6 and 8). This allows a unique property of reversible modification of single-channel function through post-translational modification, namely, tyrosine phosphorylation. This is consistent with the results of Charpantier et al. (2005), who proposed that tyrosine phosphorylation of α7, which decreased macroscopic currents, was the result of alterations in channel function. We further examined whether the gating kinetics of α7 were also affected by TCR activation. We analyzed the activation and desensitization kinetics and open and closed channel durations (Figs. 7 and 8) and determined that α7 gating kinetics were unaffected by TCR activation. Thus, only single-channel conductance is decreased by TCR activation.
Physiological role of TCRs in the CNS
Our study demonstrates that an immune protein receptor complex, the TCR, has a neuronal function in the CNS. TCRs play an important role in modulating postsynaptic cholinergic neurotransmission by dampening α7 nicotinic currents (Fig. 2). Although we used an exogenous compound ConA for stimulation of TCRs, we also have evidence that endogenous activation of TCRs already occurs in the CNS, which dampens α7 currents. This is shown by significantly augmented α7 nAChR currents from layer 1 interneurons of TCR β KO mice compared with WT (Fig. 2). Furthermore, both TCR signaling and α7-mediated neurotransmission affect neuronal excitability because inhibition of α7 with MLA decreases action potential firing similar to TCR activation with ConA, whereas neurons from TCR β KO mice had elevated firing frequency (Fig. 9).
TCR modulation of α7 receptors involves a unique form of cellular signaling in the CNS. Unlike the receptor tyrosine kinases, insulin receptor (Ahmadian et al., 2004; Cho et al., 2005) and TrkB receptor (Zhou et al., 2004; Fernandes et al., 2008) signaling, which involve the release and binding of insulin and brain-derived neurotrophic factors to their respective receptors, the TCR is an octameric receptor complex, which binds to an antigen presented by either MHCI or MHCII found on the surface of the antigen-presenting cell. MHCI molecules are widely distributed in the CNS (Huh et al., 2000) and reside on neurons and microglia (Tooyama et al., 1990; Neumann et al., 1997; Corriveau et al., 1998). We propose that TCR-mediated tyrosine kinase signaling is based on cell–cell contacts and therefore senses neighboring cells by binding to MHCI of adjacent neurons or microglia. Interestingly, microglia are involved in the pruning of synaptic spines, as evidenced by synaptic material engulfed by microglia (Paolicelli et al., 2011). A purely speculative function of TCR activation is the downregulation of α7 receptors during microglial pruning of neuronal dendritic spines. This may be a protective mechanism to decrease calcium influx through the highly calcium-permeable α7 receptors during the trauma of spine pruning.
TCR signaling through cell–cell contacts may also be necessary for the modulation of ion channel expression during formation of synapses. Cell–cell contact signaling may be formed when MHCI molecules from a presynaptic neuron bind to TCRs on the postsynaptic neuron. Evidence supporting the role of TCR-MHCI complex in modulating neurotransmission includes the fact that MHCI-deficient mice display enhanced long-term potentiation in the hippocampus (Huh et al., 2000). Furthermore, CD3ζ subunits were shown to modulate AMPA glutamatergic neurotransmission and dendritic development (Xu et al., 2010) in neurons of the retina.
nAChRs play an important role in affecting not only neuronal excitability but also synaptic plasticity in many CNS areas (Fujii et al., 2000; Ji et al., 2001; Couey et al., 2007). Our results show that α7 nAChRs contribute to neuronal excitability of cortical interneurons. Therefore, TCR inhibition of nAChR activity can dynamically and precisely tune the excitability of neurons. Because other ligand-gated ion channels, including GABAA (Wan et al., 1997), NMDA (Wang and Salter, 1994) and AMPA receptors (Ahmadian et al., 2004), can be functionally modulated by src tyrosine kinases, this opens the possibility that TCRs can modulate neuronal excitability by impacting the activity of many other ion channels in the CNS.
Footnotes
This work was supported by a Natural Sciences and Engineering Research Council of Canada Discovery grant, a Heart and Stroke Foundation of Canada grant, a National Alliance for Research on Schizophrenia and Depression Young Investigator Award to R.N., the Victoria Foundation-Myre and Winifred Sim Fund, a Canadian Foundation for Innovation grant, the British Columbia Knowledge Development Fund, and a Natural Sciences and Engineering Research Council of Canada Research Tools and Instrumentation grant. We thank Dr. Kerry Delaney, Dr. Perry Howard, and Anthony Renda for helpful discussions; Dr. Bruce N. Cohen for technical advice on single-channel recordings; Dr. Barbara J. Morley (Boys Town National Research Hospital, Omaha, NE) for technical advice for α7-null mice husbandry; Zhiwei Shi and Qi Huang for excellent technical assistance; and Ariel Sullivan, Christina Barnes, Kristin Vandeloo, Carl Jensen, and all other members of the mouse facility at the University of Victoria for providing excellent care of our mice.
The authors declare no competing financial interests.
- Correspondence should be addressed to Dr. Raad Nashmi, University of Victoria, Department of Biology, PO Box 3020, Station CSC, Victoria, British Columbia V8W 3N5, Canada. raad{at}uvic.ca
