Functional expression of the alpha 7 and alpha 4 containing nicotinic acetylcholine receptors on the neonatal rat carotid body
10
0
0
Texto completo
(2) 116. R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. neuroactive ligands could also be involved in carotid chemosensory transmission (dopamine, adenosine and 5hydroxytryptamine among others; Goldman and Eyzaguirre, 1984; Oomori et al., 1994; Ureña et al., 1994; Iturriaga and Alcayaga, 2004). In addition to its postsynaptic effects, ACh can act directly on type-I cells by activating autoregulatory nicotinic and muscarinic ACh receptors (Wyatt and Peers, 1993; Dasso et al., 1997; Shirahata et al., 2007; Ortiz and Varas, 2010). At least in neonatal rat type-I cells, the activation of both types of cholinergic receptors modify the presynaptic element through different mechanisms: while activation of nAChRs evoke a fast (partially voltage dependent) entry of Ca2+ from the external media, the activation of muscarinic ACh receptors triggers the release of Ca2+ from internal stores (Dasso et al., 1997) and a Ca2+-independent modulation of the K+ background conductance (Ortiz and Varas, 2010), therefore possibly impacting in type-I cell excitability as well. Thus, ACh has not only a role as an excitatory transmitter, but it also acts as a modulator of type-I cell activity mainly by modifying intracellular Ca2+ levels ([Ca2+]i) and therefore transmitter release (Dinger et al., 1981; Conde and Monteiro, 2006). In particular, the activation of nAChRs upon presynaptic (type-I) cells would evoke rapid changes in membrane excitability and [Ca2+]i, potentially affecting the chemotransduction of physiological stimuli such as hypoxia and acidosis (which are also mediated via changes in type-I cell [Ca2+]i; Buckler and Vaughan-Jones, 1994a,b; Ureña et al., 1994) and the release of neurotransmitters (Conde and Monteiro, 2006). The nAChRs in the carotid body have not been fully characterized. Early studies with radiolabeled a-bungarotoxin suggest the presence of the a7 nAChR subunit within the cat CB (Dinger et al., 1981, 1985, Obeso et al., 1997). More recently, immunohistochemical and RT-PCR evidences have shown the presence of a3, a4, b2 and b4 subunits and the absence of a7 subunits of nAChRs in cat CBs (Higashi et al., 2003; Shirahata et al., 2007). Additionally, several nAChR subunits have been reported in the C57BL/J6 mice CB: a3, a4, a5, a7, b2, and b4 (Cohen et al., 2002; Kahlin et al., 2010); and more recently a3, a7, and b2 nAChR subunits mRNA have been reported in human carotid bodies (Fagerlund et al., 2010). In the rat model, pharmacological evidences suggest a main role for the heteromeric a4b2 nAChRs in the regulation of CB adenosine release (Conde and Monteiro, 2006). In spite of this eventual specie-specific differences, electrophysiological studies of isolated type-I cells show resemblances between rat and cat nAChRs (Wyatt and Peers, 1993; Higashi et al., 2003), suggesting that the same nAChR subtypes are present in both species. It is, therefore, clear that functional nAChRs are present in type-I cells and that they could play a role as modulators of carotid body activity. However functional nAChRs are assembled from multiple subunits, thus detection of a receptor subunit (protein or mRNA) does not necessarily prove the presence of functional receptors. In the present study, we tested the hypothesis that nAChRs regulate catecholamine release from neonatal rat carotid body. By using real time chronoamperometric detection we shown that activation of a7 and/ or a4-containing nAChR subtypes promotes catecholamine release from intact CB. In addition, by means of electrophysiological and immunoreaction approaches we identified the presence of a7 and a4-containing nAChR subtypes in CB type-I cells. Interestingly, our results show that either a7 or a4-containing nAChR are sufficient by themselves to promote catecholamine release; suggesting a dual nicotinic mechanism of regulation of catecholamine signaling in the CB system: direct Ca2+ entry through the homomeric subtype; and an indirect, voltage-dependent Ca2+ entry that depends upon heteromeric receptor activation and the overall excitability of the type-I cell.. 2. Methods 2.1. Ethical standards The experimental procedures were approved by the Bioethical and Biosafety Committee of the Faculty of Biological Sciences of the Pontificia Universidad Católica de Chile and were conducted in accordance with the guidelines of the National Fund for Scientific and Technological Research (FONDECYT) and the National Institutes of Health guide for the care and use of Laboratory animals (NIH Publications No. 80–23, revised 1978). 2.2. Carotid body and type-I cells isolation A total number of 22 neonatal Sprague Dawley rats (10–12 days old) were used in this study. Rat pups were anesthetized with ketamine/xylazine (75/7.5 mg/kg, i.p.). Once deeply anesthetized, as tested by foot pinch withdrawal reflex, the carotid bodies were extracted and placed in ice cold modified Hank’s balanced salt solution (4 °C, pH = 7.43). These CBs were then used either intact for chronoamperometric recordings (N = 9) or disassociated for patch-clamp studies (N = 30). At the end of CBs extraction the animals were killed by beheading whilst still anaesthetized. For type-I cells isolation, the extracted CBs were enzymatically dissociated using collagenase type I (0.4 mg/ml, Sigma) and trypsin (0.2 mg/ml, Sigma) for about 20 min at 37 °C and then mechanically triturated with a sterile glass Pasteur pipette, as previously described (Buckler, 1997). The cell suspension was then centrifuged, resuspended in Ham’s F-12 medium or a 50/50 mixture of Ham’s F12 and Dulbecco’s modified Eagle’s medium (DMEM, supplemented with 10% heat-inactivated fetal calf serum, 100 i.u/ml, penicillin 100 lg/ml streptomycin and 84 U/l insulin) and plated out onto poly-D-lysine coated glass coverslip. Cells were kept at 37 °C with 5% CO2 in humidified air until use (2–8 h). 2.3. High-speed chronoamperometric recordings Isolated intact CBs were dissected from surrounding connective tissues and pre-incubated by 10 min in a Tyrode solution (described below) containing dopamine (2 mM). Pre-loading with dopamine increases and makes more homogenous the initial dopamine content of the CB, therefore reducing the variability in catecholamine release between different CBs (not shown). Then, the CB was gently fixed with a pin to the bottom of the recording chamber (total volume 500 lL) and superfused at a stable flow rate of 3 ml/min with a standard HEPES-buffered Tyrode solution (in mM: NaCl 140, KCl 4.5, HEPES 10, MgCl2 1.0, CaCl2 2.5, glucose 11; pH 7.4) at 37 °C and continuously bubbled with a gas mixture of 20% O2 and 80% nitrogen. The bath was grounded with an Ag–AgCl pellet (no active ground electrode was employed since it makes no difference in the signal-to-noise ratio when used, not shown). The electrochemical measurement of catecholamine (CA) outflow was done through a microcomputer-controlled high-speed chronoamperometric system (IVEC-10, Medical System Corp., Greenvale, NY, USA). Working electrodes (Rocky Mountain Centre for Sensor Technology, Denver, CO, USA), consisting of multiple carbon fibers (tip diameter, 30 lm) were gently inserted into the carotid body on the opposite side to the inflow of saline solution. A square-wave application of 0.7 V, with respect to the reference Ag–AgCl electrode, was applied for 100 ms at a rate of 5 Hz. The resulting oxidation current was digitally integrated during the last 80 ms of each pulse (after complete settle of the oxidation current), averaged for five cycles, displayed at a rate of 1 Hz and stored in the computer. The reduction current generated when the potential.
(3) R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. returned to 0 V was digitized, integrated and averaged in the same manner. The IVEC-10 system expresses the integrated currents in arbitrary units, which are linearly proportional to CA concentrations. These reduction/oxidation (red/ox) current ratios were used to identify the electroactive species under oxidation, however in the present study we did not claim to discriminate between different CA species releases from the carotid bodies (but see Section 4.2). The carbon fiber electrodes were coated 10–15 times with Nafion (Aldrich Chemical Co., Milwaukee, WI, USA) – a polysulphonated derivative of Teflon, highly permeable to dopamine, but not to dihydroxyphenylacetic acid (DOPAC) or ascorbic acid (Gerhardt et al., 1984; Doherty and Gratton, 1992) to reduce their sensitivity to DOPAC and ascorbic acid. Our electrodes had sensitivity to dopamine >500 times higher than that to ascorbic acid (not shown), which is oxidized but not reduced. The sensitivity and linearity of our Nafion-coated carbon fiber electrodes were determined in vitro by calibrating their response to dopamine hydrochloride (final concentration 2–12 lM in the HEPES-Tyrode solution at pH 7.4, supplemented with 0.1 M perchloric acid to prevent oxidation). Only electrodes showing highly linear responses (r > 0.99, P < 0.05) to increasing concentrations of dopamine were used. The threshold for dopamine determinations was typically 10–20 nM. It is worth to mention that the red/ox current ratio value for the biogenic amine 5-hydroxytryptamine in these Nafion-coated carbon electrodes was significantly lower when compared to the red/ox ratio for dopamine (0.07 ± 0.01 and 0.48 ± 0.18 for serotonin and dopamine respectively, P < 0.05 Kruskal–Wallis test). Therefore we assume that the electrochemical signal recorded is due to reduction and oxidation of catecholamines. The calibration of electrodes was made previous to each experiment. Effluxes of CAs (CAefflux) evoked by different stimuli, expressed as DCAefflux over baseline, were computed from changes of the oxidation currents recorded by the carbon electrodes. After positioning the recording electrode, a 30 min stabilization period was defined before starting with any experimental maneuver. All drugs used were freshly prepared in standard HEPES-buffered Tyrode solution and diluted to desired concentration (see Section 3). None of the drugs used (as the mentioned concentration) were oxidized/reduced by the applied electrode potential, as tested in the recording chamber before placing the tissue (not shown). Nicotine was applied by fast bolus injection (200 lL) into the bath solution. In order to have an adequate diffusion of the blockers used, these were applied by continuous superfusion just after the end of the response to nicotine and for at least 3 min before the second nicotine challenge. 2.4. Whole cell voltage-clamp recordings Seeded isolated carotid body cells were placed on a recording chamber (total volume approx. 400 lL) and positioned on the stage of an inverted microscope under constant superfusion (3–5 ml/ min) with the same standard HEPES-buffered Tyrode solution described for chronoamperometric recordings, at 32–35 °C and continuously bubbled with a gas mixture of 20% O2 and 80% nitrogen. The bath was grounded with an Ag–AgCl pellet. Voltage-clamp recordings were performed using an Axopatch 200B amplifier (Axon Instruments, CA, USA). The recorded signal was filtered at 2 kHz and acquired at 20 kHz using a Cambridge Electronic Design power 1401 A/D converter and pClamp 9.2 acquisition software (Axon Instruments, CA, USA). Voltage commands were generated using pClamp 9.2 software. Borosilicate glass pipettes (GC150F-10, Warner Instruments Corp.), were pulled in a Flaming/Brown P-87 Micropipette Puller (Sutter Instrument Co.) and polished by heat. Whole-cell current recordings were carried out using electrodes filled with an internal solution containing (in mM): 5 NaCl, 135 KCl, 1 CaCl2, 11 EGTA, 10 HEPES, 2 K2ATP,. 117. pH = 7.2. Electrode resistance ranged from 4–7 MX. Seal formation and membrane breakthrough were monitored by observing under voltage-clamp mode the response to a 5 mV voltage pulse lasting 50 ms. Liquid junction potential between microelectrodes and bath solution was corrected. Series resistance (usually 5–15 MX) and capacitive transients were electronically compensated and periodically monitored throughout recordings to ensure proper compensation. According to our previous results, pipette potential was held at 60 mV. Cellular integrity was tested regularly by monitoring access resistance (using the seal test pulse described above) and by evaluating the presence of voltage-gated currents (obtained by imposing membrane potentials ranging from 100 to +40 mV, in 10 mV steps magnitude, 100 ms duration, see insert in Fig. 3A). All drugs used were freshly prepared in HEPES-buffered Tyrode standard solution and diluted to desired concentration (see Section 3). nAChR agonists were applied for 15 s by gravity ejection from a nearby pipette whose tip was located at about 100 lm from the cell surface. nAChR blockers were diluted to required concentration in the same external HEPES-buffered Tyrode standard solution and applied continuously through the conventional superfusion system for at least 3 min before tested. Control experiments using no nicotinic agents were performed in order to test if the rapid exchange of solutions could affect the recorded currents. 2.5. Immunocytochemistry Seeded isolated CB cells were immersed in cold (4 °C) 4% paraformaldehyde fixative for 10 min, transferred to 10% sucrose for 12 h and then storage in sucrose 30%. The a4 or a7 subunits were recognized by rabbit polyclonal antibody against a4 or a7 nAChRs subunits (anti-a4 or anti-a7, Abcam, USA). Recognition of the typeI cells was done by using polyclonal antibody (anti-TH, Millipore, USA). Fixed cells were washed several times in phosphate buffered saline (PBS: 137 mM NaCl; 10 mM Na2HPO4; 2.7 mM KCl; 1.8 mM KH2PO4; pH = 7.4), incubated in bleaching solution (50 mM ammonium chloride in PBS) for 30 min, washed again 4 times for 5 min, and incubated in blocking solution (3% donkey serum, 0.2% Triton X-100, 0.01% thimerosal in PBS) for 1.5 h. Double staining (anti-TH plus anti-a4 or anti-a7 nAChRs subunits) was obtained by incubation with the primary antibodies anti-TH (diluted at 1:1000) and antibodies anti-a4 or anti-a7 (both diluted at 1:200) in blocking solution at room temperature overnight. Then, sections cells were rinsed 4 times for 5 min with PBS, incubated with a secondary antibody Alexafluor 555-donkey polyclonal anti-rabbit IgG (Invitrogen, USA), Alexa 488-donkey polyclonal anti-sheep IgG (Invitrogen, USA) and FITC-goat polyclonal anti-chicken IgG (Abcam, USA) diluted 1:1000 during 2 h at room temperature, in blocking solution supplemented with 1.5% donkey serum at room temperature for 2 h, and washed again with PBS. Observation of samples cells was done in a laser scanning system (LSM 510; Carl Zeiss, Germany) mounted on a motorized Axiovert 100-M microscope (Carl Zeiss, Germany) and equipped with a multi-line argon laser (458 nm, 488 nm) and a helium–neon laser (543 nm). Images were acquired using a 40 objective, multitrack mode (505–550 nm BP, 585 nm LP) at 2 lm interval, and then averaged through LSM 510 software. 2.6. Drugs Dopamine hydrochloride, hexamethonium chloride, mecamylamine hydrochloride, nicotine bitartrate and 5-hydroxytryptamine hydrochloride were purchased from Sigma–Aldrich (MO, USA). a-bungarotoxin was purchased from Tocris Bioscience (Bristol, UK)..
(4) 118. R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. Erysodine was obtained as described elsewhere (IturriagaVásquez et al., 2010). Briefly, erysodine was isolated from seeds of Erythrina falcata Benth. All structures were confirmed by using one- and two-dimensional 1H and 13C NMR analyses. Purity and structure of the compound was established by high-resolution one- and two-dimensional NMR experiments and was typically 98% to 100%. 2.7. Data analysis Values given are MEAN ± SEM. For the voltage-clamp recordings, the peak (I) currents elicited by several concentrations of nicotine, cytisine and ACh were standardized to the membrane capacitance. Dose-dependent activation of the currents by nAChR agonist was. I=Imax ¼ 1=½1 þ ðEC 50 =½Xn Þ fitted to the Hill equation where Imax = maximal current evoked by a given agonist, EC50 = agonist concentration that evoked the halfmaximal current, X = agonist concentration delivered by the stimulus pipette and n = Hill coefficient. Net charge carried by the current triggered by nAChR activation was estimated by the area under the curve of the current trace using the software Igor Pro 5.0 (Wave-Metrics, Lake Oswego, OR, USA) The dose–response curves were compared using two-way ANOVA. In order to compare more of two grouping variables, we used nonparametric ANOVA, Kruskal–Wallis test followed by Dunn’s Multiple Comparison post hoc test. The significance level was set at P = 0.05. All curve fitting and statistical calculations were performed with GraphPad Prism 5.0 (GraphPad Software, USA). 3. Results 3.1. Electrochemical detection of catecholamine efflux from intact in vitro carotid body Catecholamine efflux (CAefflux) from an intact in vitro carotid body was unmodified during the 30 min of stabilization period in the recording chamber (see Section 2). Since the CBs are chemoreceptor organs, we first assessed their viability in vitro by challenging them with fast application of an acidic stimulus (pH 6.5/ 200 ll). In all the 9 CBs tested, the acidic stimuli evoked a rapid electrochemical response, similar to the one obtained with exogenous dopamine in the absence of the CB (red/ox current ratios of 0.42 ± 0.12 and 0.48 ± 0.18 for the response to acidic stimulus and exogenous dopamine respectively; P > 0.05, Kruskal–Wallis test, see Fig. 1A). This observation suggests that we are indeed detecting a cathecolaminergic efflux from the CB in response to low pH. Thus, the acidic stimulus increases CA efflux when compared with the response to the control bolus application (DCAefflux = 1.0 ± 0.2 lM, N = 7 at pH 6.5; DCAefflux = 0.11 ± 0.02 lM, N = 7 at pH 7.4; P < 0.05, Dunn’s Multiple Comparison post hoc test, posteriori to Kruskal–Wallis test, see Fig. 1B and C). Additionally we tested the excitability of the preparation by fast applying boluses of high K+-containing Tyrode solution (KCl 150 mM), that consistently evoked a transient increase in CA efflux (not shown). 3.2. Activation of nicotinic acetylcholine receptors increases catecholamine efflux from intact in vitro carotid body In 7 out of 9 CBs that respond to acid stimuli, intrastream injections of nicotine boluses (1 mM/200 lL) transiently increases the CAefflux (DCAefflux = 1.8 ± 0.3 lM, N = 7, P < 0.05, Dunn’s Multiple Comparison post hoc test, compared with control; see Fig. 1).. Fig. 1. (A) Reduction/oxidation current ratios (red/ox) obtained from the carbon electrodes measurements. Note that red/ox ratios for catecholamine release from carotid body (CB) in response to acidification (pH 6.5) or nicotine (Nic 1 mM) are very similar to red/ox ratio for dopamine (DA) applied directly on the carbon fiber electrode during calibration (without carotid body, w/o CB). (B) Acidic (pH 6.5) or nicotinic stimulation triggers an increase in CA efflux from whole CBs. Control bolus application of external solution at pH 7.4 triggers no response. All traces shown are from the same preparation. (C) Summary of the increase in CA efflux evoked by pH 7.4 (N = 7), pH 6.5 (N = 7) and nicotine 1 mM (Nic 1 mM; N = 7). Asterisks indicate statistical significant differences, ⁄P < 0.05, ⁄⁄⁄P < 0.001. n.s. indicates no statistical difference. Kruskal–Wallis test, Dunn’s multiple comparisons post hoc test.. The nicotine-evoked DCAefflux was 90.5 ± 19% higher than that evoked by pH 6.5. Again, the red/ox ratio of the electrochemical response evoked by nicotine 1 mM was similar to the red/ox ratio obtained with exogenous dopamine (Fig. 1A), according to the catecholaminergic nature of the specie released from the CB. Lower doses of nicotine failed to increase significantly the CA efflux (not shown). We then tested the effects of three different nAChR antagonists upon the nicotine-evoked CA release (see Section 2). The dose– response curves for the effects of the generic nAChR blocker hexamethonium (Hexa, 0.1–100 lM), the heteromeric nAChR.
(5) R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. 119. Fig. 2. Effects of nAChR antagonists a-bungarotoxin (Bgtx), Erysodine (Eryso) and hexamethonium (Hexa) on the CA efflux evoked by nicotine 1 mM (Nic 1 mM). Note that all three antagonists reduced CA release with similar efficacies (70% of maximal inhibition). Inset: representative record for the same CB showing the increased CA efflux evoked by nicotine (1 mM) and its reversible blockage by Eryso 100 nM. See main text for further details.. Table 1 Efficacy of nicotinic antagonist in inhibiting the catecholamine release in CBs stimulated by nicotine. Antagonist. Emax (% inhibition of control). Concentration (lM). Hexamethomium a-Bungarotoxin Erysodine. 76 ± 9.1 78 ± 9.5 71.1 ± 5.7. 10 0.1 10. Emax = maximal % of inhibition (media ± s.e.m); Concentration = antagonist concentration that caused the maximal inhibition.. blocker erysodine (Eryso, 0.01–10 lM) and the homomeric nAChR blocker a-bungarotoxin (Bgtx, 0.0001–1 lM) on the release of CA evoked by nicotine 1 mM, were performed. All three nAChR blockers caused a concentration-dependent decrease in the nicotineevoked DCAefflux from the CBs (Fig. 2). Interestingly, none of them were able to fully block the nicotine effect on CAefflux, having a similar maximal inhibitory efficacy (Hexa, 76.0 ± 9.1%, N = 5; Eryso, 71.1 ± 5.7%, N = 4; Bgtx, 78.0 ± 9.5%, N = 4, see Fig. 2 and Table 1).. Fig. 3. Nicotine evoked a fast inward current (A) and membrane depolarization (B) in isolated CB type-I cells (V holding = 60 mV). The addition of mecamylamine (Mec) 1 lM to the bath solution, completely suppressed the response to nicotine. Insert in A: only cells that showed voltage-gated currents were included in this study (see main text for details). Mecamylamine has no effect on voltage-gated currents (not shown). (C) Example of the whole cell inward currents evoked by increasing concentrations of nicotine (1–1000 lM) in the same cell. Upper bars indicate the nicotine application. Successive doses were applied at 3 min intervals at a holding potential of 60 mV. (D) Dose–response curves for the nAChR agonist nicotine (Nic and filled circles), cytisine (Cyt and open circles) and acetylcholine (plus 1 lM atropine, ACh, filled triangles). Peak current amplitudes were normalized to that in the presence of 1 mM nicotine to the indicated concentration of agonist. The dose–response curves for nicotine and cytisine are not different between them (P > 0.05, Two-way ANOVA) but both differ with the dose–response curve for ACh (P < 0.05, Two-way ANOVA). ⁄P < 0.05, ⁄⁄P < 0.01, Bonferroni post hoc test..
(6) 120. R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. 3.3. Nicotinic agonists evoke fast inward currents in isolated carotid body type-I cells If activation of nAChRs increases catecholamine release from CBs, then we should be able to show that a nAChR agonist can evoke fast inward currents in isolated type-I (catecholaminergic) cells. Therefore, we performed conventional whole-cell recordings upon freshly isolated CB type-I cells. We firstly assessed the viability of the cells by evoking voltage-gated currents (obtained by imposing membrane potentials ranging from 100 to +40 mV from a holding potential of 60 mV, see Section 2 and insert in Fig. 3A). According to what have been described for neonatal rat, CB type-I cells show only voltage-gated outward currents with no sign of inward currents (Wyatt and Peers, 1995; Buckler, 1997). As expected from our electrochemical results on whole CBs, fast application of nicotine upon isolated CB type-I cells (voltageclamped at 60 mV) evoked fast-desensitizing, dose-dependent inward currents (in 49 out of 68 cells tested) that were fully but reversibly blocked by the non-competitive nAChR antagonist mecamylamine (Fig. 3A). Under current clamp conditions (without current injected), nicotine evoked a fast depolarization that again was fully blocked by mecamylamine (1 lM, Fig. 3B). The application of nicotine onto isolated CB cells induced dose-dependent (1–1000 lM) inward currents (that were best fitted to a single Hill equation; r = 0.997, P < 0.05, see Section 2) with an estimated EC50 and Hill coefficient of 46.8 lM and 1.4 respectively (Fig. 3C and D). Maximal responses (mean peak inward current amplitude of – 166.4 ± 12.2 pA/pF, N = 13) were achieved at 100 lM nicotine. In contrast to a previous report that suggests a great variability in the magnitude of the responses to nicotine (Wyatt and Peers, 1993), we observed homogenous responses that varied slightly from cell to cell. Responses induced by increasing concentrations of the nicotinic agonists cytisine and acetylcholine are shown in Fig. 3D and summarized in Table 2. Although all three agonist produced. Table 2 Efficacy of potency of nicotinic agonists in stimulating an inward current in isolated cells from CB. Agonist. Peak (pA/pF). EC50 (lM). nHill. Acetylcholine Cytisine Nicotine. 126.3 ± 2.02 158.5 ± 3.01 166.4 ± 12.2. 212.7 19.3 46.8. 0.9 1.6 1.4. Peak = maximal inward current amplitude obtained by agonists (media ± s.e.m); EC50 = agonist concentration that produced 50% of maximal effect; nHill = Hill coefficient.. dose-dependent inward currents of similar amplitude, the rank of functional potency would be cytisine nicotine > acetylcholine (EC50 19.3; 46.8 and 212.7 lM, respectively). According to what we found in our chronoamperometric measurements, in all 5 cells tested, a-bungarotoxin partially blocked the response evoked by nicotine. In fact, superfusion of 10 nM of a-bungarotoxin for 3 min produced a 46.1% inhibition of the net current (area under the curve) evoked by 100 lM nicotine (Fig. 4). On the other hand, in other set of experiments, we observed that erysodine 100 nM produced a 75.1% inhibition of the net current evoked by 100 lM nicotine (Fig. 4). It is note to worth that the current evoked by nicotine that persisted during erysodine superfusion showed a fast time course that resembles the kinetics of an a7 nAChR (fast desensitization). According to this interpretation, the co-application of both a-bungarotoxin and erysodine (3 min superfusion) produced a 98.4% inhibition of the current evoked by 100 lM nicotine (Fig. 4). Interestingly in all 5 cells tested, a small inward current (6.9 ± 1.6 pA) remains unaffected in the presence of both antagonists. Since the generic nAChR antagonist mecamylamine fully blocks the currents evoked by nicotine (Fig. 3A), it is quite likely that this small resistant inward current is due to activation of a a-bungarotoxin- and erysodine-insensitive nAChR subtype.. Fig. 4. nAChR antagonist sensitivity of CB type-I cells. (A) Representative traces from three different type-I cells showing partial reversible block of the nicotinic response with a-bungarotoxin (Bgtx, left panel) and erysodine (Eryso, middle panel). The co-application of both blockers almost completely suppressed the response (right panel). (B) Summary of the effects of the two antagonist used separately or in combination upon net current (area under the curve, see Section 2). Please note that a small but significant response to nicotine remains in the presence of both blockers. (), () Indicates significative differences of P < 0.05 and P < 0.01 respectively between control and antagonist. (#) Indicates the difference between Bgtx + Eryso and each one separately. No significant difference between Bgtx and Eryso was observed. Two-way ANOVA, Bonferroni post hoc test..
(7) R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. 121. Fig. 5. Rat carotid body type-I cells express a4 and a7 nAChRs subunits. (A–C) and (D–F) Correspond to pseudo-color images (objective 63, zoom 3) from dissociated carotid body type-I cells in primary cultures showing the expression of tyrosine hydroxylase (TH; green) and a4 (C; red) or a7 (F; red) nAChR subunits. (B and E) Corresponds to single channel images for TH; C, single channel images for a4, and F, single channel images for a7nAChR subunits. (A and D) Merge images of respective single channel images showing co-expression (yellow) in cultures of dissociated carotid body cells. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.). 3.4. Immunocytochemical localization of a4 and a7 nAChR protein subunits in rat carotid body type-I cells The above functional studies strongly suggested that a7 and a4-containing nAChR subtypes are present in type-I cells and that they participate in the regulation of catecholamine release. If so, a7 and a4 subunits should be expressed in CB type-I cells. We therefore applied immunoreactivity techniques to localize a4 and a7 nAChR subunits in isolated dissociated cells from the CB (see Section 2). Cells incubated with preabsorbed antibody showed no immunoreactivity (not shown). To confirm localization of a7 and a4 staining, we performed double staining using antibodies against a7 or a4 subunits and an antibody against tyrosine hydroxylase (TH), the rate-limiting enzyme for catecholamine biosynthesis and a known phenotypic marker for type-I cells (González et al., 1994; Nurse, 2005). Consistent with our electrophysiological data, the majority of the CB cell bodies were positive for both TH and either a7 or a4 (Fig. 5). We did not assess directly if a same cell was positive for both nAChR subunits, but since for each subunit we detected positive immunoreaction in almost all the type-I cells studied, it is reasonable to assume that a7 and a4 are co-expressed in the same type-I cell, as suggested by our functional data. 4. Discussion The present study demonstrates, for the first time, that a7 and a4-containing nAChR subtypes contribute to the regulation of catecholamine transmission in the neonatal rat carotid body. We show that activation of the nAChRs generates fast inward currents and type-I cells membrane depolarization that ultimately leads to increased catecholamine release from the carotid body. In addition we provided evidences of a third functional nAChR subtype (resistant to a-bungarotoxin and erysodine) that accounts for a small fraction of the whole cell current evoked by nAChR agonists. Our results are in accordance with previous studies that indicate that acetylcholine released from type-I cells and/or autonomic microganglion cells (Gauda et al., 2004) has a role as an autocrine/paracrine regulator of the carotid body transmission. A novel aspect of our study is to shown that each type-I cell functionally expressed all these receptor subtypes, suggesting a. unique nicotinic phenotype composed by the contribution of at least three different components. It remains to be established the functional consequences of these observation, however it is worth noting that while the a7 nAChR is highly permeable to Ca2+, the heteromeric subtypes have a higher permeability to Na+ (Shirahata, 2007; Albuquerque et al., 2009). Thus, since activation of each (a7 and a4-containing) receptor subtype is able to promote catecholamine release, it is possible that the regulation of catecholamine release by nAChRs, in the neonatal rat CB, is achieved by a dual mechanism that involves direct Ca2+ entry (and therefore neurosecretion) through a7 nAChRs and an indirect, voltagedependent Ca2+ entry that depends on activation of a4-containing nAChR and the overall excitability of the type-I cell. 4.1. Activation of nicotinic acetylcholine receptors excites carotid body type-I cells Functional studies have shown that activation of nAChR in isolated type-I cells of the neonatal rat CB generates both inward currents (Wyatt and Peers, 1993) and increased [Ca2+]i (Dasso et al., 1997). However little is known about the different nAChR subtypes functionally expressed in these cells. Previous works have only reported the presence of a3, a4, a7 and b2 proteins or mRNA in human, mouse, cat and rat CBs (Shirahata et al., 2007; Niane et al., 2009; Kahlin et al., 2010; Fagerlund et al., 2010). In the present study we have confirmed that activation of nAChRs generates fast inward current and membrane depolarization of the type-I cells. In addition, we have shown that near to 70% of the CB cells studied responded to nicotine (value that is in close agreement with previous report (Wyatt and Peers, 1993; Dasso et al., 1997). Moreover, this nicotine-evoked response were partially blocked by both abungarotoxin, a highly selective antagonist of a7 (when compared with heteromeric) nAChRs, and erysodine, an Erythrina alkaloid closely related to the best known heteromeric nAChRs antagonist dihydro-b-erythroidine (DHbE), but at doses here used with higher potency to a4b2⁄ nAChR and better selectivity to discriminate from [125I] a-bungarotoxin binding sites in the rat brain (Decker et al., 1995; Iturriaga-Vásquez et al., 2010). Therefore, the nicotine response of type-I cells is always mediated by at least the activation of two types of nAChRs, a7 and a4-containing nAChRs. In.
(8) 122. R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. addition, our the immunocytochemistry assay reactivity studies show the presence of the a7 and a4 subunits on the vast majority of the isolated type-I cells of the CB, as it previously reported for neonatal rat CB (Niane et al., 2009), further confirming the observation of a homogenous population of type-I cells that express both a7 and a4-containing nAChRs. It should be noted, however, that we cannot exclude that a very small fraction (5%) of the nicotinic response is due to activation of a third nAChR subtype, resistant to a-bungarotoxin and erysodine (Fig. 4A, right panel) but sensitive to 1 lM mecamylamine. The fact that mecamylamine was able to fully suppress the response to nicotine suggests that there is a third nAChR subtype in type-I cells. This third component of the nicotine-evoked response could include a3-contaning nAChR, which has been reported in mice and cats CB (Cohen et al., 2002; Shirahata et al., 2007; Kahlin et al., 2010), is sensitive to micromolar concentrations of erysodine and highly sensitive to mecamylamine (Jensen et al., 2005). 4.2. The role of nAChRs in the cholinergic modulation of catecholamine neurotransmission in the carotid body It is well known that presynaptic nAChRs play a role in the release of several different neurotransmitters (including catecholamines) in the peripheral as well as in the central nervous systems (Gotti and Clementi, 2004; Shirahata et al., 2007; Albuquerque et al., 2009). Although we cannot identify which catecholamine was the main specie detected by the electrode, previous reports have shown that dopamine is the main biogenic amine present in the CB of most animal species. In the adult rat CB, for example, mean dopamine levels are roughly five times higher than noradrenaline (Vicario et al., 2000). Dopamine is synthesized, stored and released from the CB type-I cells in response to natural stimuli (González et al., 1994; Nurse, 2005). Though once dopamine was proposed as an excitatory transmitter in the CB, several lines of evidence suggest that dopamine has an inhibitory role in the carotid chemosensory transmission by several mechanisms. The complex role of dopamine includes postsynaptic regulation of the cholinergic and purinergic drive (Iturriaga and Alcayaga, 2004) and inhibitory effects of presynaptic D2 autoreceptors (Bairam et al., 2000; Carroll et al., 2005; Conde et al., 2008). On the other hand, noradrenaline (NE) is also present in CBs, and according to its relative lower levels when compared to dopamine, the noradrenergic marker dopamine b-hydroxylase has been found only in some type-I cells of the rat CB (Chen et al., 1985). Interestingly, nicotine elicits a preferential release of NE over dopamine (Vicario et al., 2000), suggesting that activation of nAChRs could also increases NE release from type-I cells (as well as from the noradrenergic nerve terminals of the superior cervical ganglion that innervates the CB). It is worth noting that dopamine could be released from nerve endings of C-fibers within the CB (Almaraz and Fidone, 1986). Therefore, postsynaptic nAChRs located in these nerve endings could also be participating in the increase in CAs levels after cholinergic stimulation. Since CBs release ACh in response to natural stimuli, the proposal that ACh accounts at least of part of the afferent excitatory drive arises naturally (Eyzaguirre and Zapata, 1968). However, ACh neurotransmission can not explain the total CB response to hypoxia, a main natural stimuli for this organ (Reyes et al., 2007; Donnelly, 2009). In this regard, Donnelly (2009) showed that the application of hexamethonium or mecamylamine failed to reduce the hypoxic response. He concluded that – at least in non-adult rats (P18 and P26) – the cholinergic drive is not related with hypoxic response. These results, among others, support the idea that the release of ACh from CBs could have a presynaptic, more than a postsynaptic role (Dinger et al., 1981; Wyatt and Peers, 1993; Conde and Monteiro, 2006; Ortiz and Varas, 2010).. An important consideration is how this functional nAChR phenotype changes with development and aging. In fact, mRNA levels for several nAChR subunits changed during the ontogenetic development, where a3 and a7 subunits levels increases, while a4 diminished, without modifications in b2 mRNA levels at least in the cat CB (Bairam et al., 2007). Moreover, nAChR activity and its overall role in the hypoxic chemoreflex seems to be age dependent, as shown in whole-body plethysmographic studies performed in neonatal rats (Niane et al., 2009). Therefore, we cannot rule out that the functional nAChR phenotype of neonatal rat CB described here could be a transitory state that changes when reaching adulthood. Our results show that rat carotid body superfused in vitro responds to acid, high K+ or nicotine stimuli with a transient increase in catecholamine release that decays quickly after stimuli is turned off, which is consistent with a fast catecholamine clearance. It should be remembered that Nafion coated electrode are highly selective for dopamine, noradrenaline and adrenaline and much less sensitive to DOPAC, serotonin and ascorbic or perchloric acid (Gerhardt et al., 1984). Interestingly, previous reports suggest that the identification of electroactive species ultimately relies on the red/ox ratio, indicating that in the CB the main catecholamine specie detected was dopamine (Gerhardt et al., 1984; Iturriaga et al., 1996; Vicario et al., 2000). If CAs act as a transmitter/modulators between type-I cells and chemosensory nerve endings, one would expect that the amount of CA released should be enough to activate catecholaminergic receptors. Our results show a mean DCAefflux from the CB of 1.0 ± 0.2 and 1.8 ± 0.3 lM after transient stimulation with low pH or 1 mM nicotine, respectively. Since these values correspond to the overflow of catecholamine(s) from the CB (plus the eventual contribution of nerve endings close to CB, as suggested by Almaraz and Fidone; 1986) it should be expected within the carotid synaptic cleft a higher CA concentration. But several factors prevent us from an accurate estimation of the ‘‘true’’ catecholamine concentration in this preparation (i.e. diffusion distance vs. re-uptake rate, etc.). Nevertheless, our results are in close agreement with previous reports in other dopaminergic systems showing that single electrical pulse in the mesolimbic pathway can increase dopamine levels ranging from 0.54 ± 0.9 to 1.0 ± 0.6 lM in the nucleus accumbens of freely-moving mice (Budygin, et al., 2002). In addition, we have observed extensive TH-immunoreactivity in CB tissue that supports the notion that CB is essentially a dopaminergic organ, as it has been proposed previously (González et al., 1994; Vicario et al., 2000; Nurse, 2005). Based on their pharmacological profiles, in the present study we have functionally characterized two subtypes of nAChRs contributing to catecholamine release regulation. Similar results have been described in cat CBs suggesting a common mechanism among these different species (Dinger et al., 1985; Obeso et al., 1997). We have shown that the nicotine-evoked increase in catecholamine release from intact CB is partially blocked by a-bungarotoxin. This agrees with the fact that a-bungarotoxin reduced but not suppressed the nicotine-evoked currents in isolated type-I cells. In addition, our data show that erysodine reduces the nicotine-evoked DCAefflux by 71.1 ± 5.7%, suggesting that in addition to a7, heteromeric nAChRs also contribute to the regulation of catecholamine release in the rat CB. It is note to worth that 100 nM erysodine reduced by 31.6% the nicotine-evoked currents in isolated type-I cells. At such submicromolar concentrations, erysodine is highly selective for a4b2* nAChRs and only at micromolar concentrations erysodine binds to a3-containing nAChRs (Decker et al., 1995; Iturriaga-Vásquez et al., 2010). Thus, our evidences suggest that a7 and a4-containing (possible a4b2*) nAChR subtypes participate in the regulation of CB catecholamine neurotransmission, possibly with a major contribution.
(9) R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. of a4-containing nAChRs, according to the total charge transported in response to nicotine application. In agreement with the role for nAChR as regulator of carotid neurotransmission, Conde and Monteiro (2006) reported that activation of nAChRs increases adenosine release from adult rat CB. Moreover, by using both DHbE and a-bungarotoxin, these authors suggested a major role for a4b2* nAChRs and a vestigial – if any- role for homomeric receptors in the regulation of adenosine release. This raises the interesting possibility that different nAChR subtypes selectively regulate CB neurotransmission. In spite of possible age-related differences, it should be noted that while we used high speed-chronoamperometric detection of catecholamine release (therefore real-time measurements), Conde and Monteiro (2006) used 10 min incubation time for HPLC measurement of adenosine content. It is possible then, within this 10 min period, several phenomena could account for the differences between our results and these previous report, mainly the well known fast a7 nAChR desensitization (Albuquerque et al., 2009; Shen and Yakel, 2009) and/or more complex effects related to adenosine autoreceptor activation within the CB that could trigger and overall inhibitory effect (Xu et al., 2006). It is noticeable that neither the generic blocker hexamethonium nor the selective blockers (erysodine and a-bungarotoxin) were able to eliminate the nicotine-evoked catecholamine release. Further inhibition of the response can be achieved for each blocker by increasing pre-incubation time but then blockers’ effects are poorly reversible (particularly for a-bungarotoxin, data not shown). Therefore, a possible explanation for the inability of blockers for suppressing the nicotine-evoked CAefflux can be a reduced accessibility of these three antagonists to reach the core of the CB tissue. On the other hand, the application of nicotine (range from 1 to 10 lg boluses) does not trigger a dopamine release in cat CBs, but high doses of ACh (500–1000 lg) evoked a 2 lM increase in DA release (Iturriaga et al., 2000), suggesting a speciespecific mechanism of catecholamine release regulation. Our results suggest that in physiological conditions ACh action on CB via nAChRs regulates CAs release, however probably this is not the only way and muscarinic receptor (mAChR) activation may be involved. Particularly, in rabbit carotid body, carbachol (1–50 lM), through M1 and M3 mAChRs activation, reduced DA content and increased DA release from carotid body (Bairam et al., 2000). In this regard, we showed that in neonatal rat carotid body, activation of mAChR (probably odd mAChRs) inhibits a background potassium current and increases intracellular calcium of type-I cells; both events pointing in the same direction: an increase in transmission release evoked by ACh (Ortiz and Varas, 2010). Therefore, the effect of ACh on CB seems to be more complex, involving the contribution of both nAChRs and mAChRs, with significant actions as presynaptic regulator of transmitter’s release.. Acknowledgements This work was supported by the Associate Vice-rectorship of Research and Doctoral Program of the Pontificia Universidad Católica de Chile (VRAID); National Fund for Scientific and Technological Development of Chile (FONDECYT) Grants #11060502 (P.I-V.) and #1090375 (J.L.E.) and Millennium Scientific Initiative Grant P05-001-F.. References Albuquerque, E.X., Pereira, E.F., Alkondon, M., 2009. Mammalian nicotinic acetylcholine receptors: from structure to function. Physiol. Rev. 89, 73–120. Almaraz, L., Fidone, S., 1986. Carotid sinus nerve C-fibers release catecholamines from the cat carotid body. Neurosci. Lett. 67, 153–158. Bairam, A., Neji, H., Marchal, F., 2000. Cholinergic dopamine release from the in vitro rabbit carotid body. J. Appl. Physiol. 88, 1737–1742.. 123. Bairam, A., Joseph, V., Lajeunesse, Y., Kinkead, R., 2007. Developmental profile of cholinergic and purinergic traits and receptors in peripheral chemoreflex pathway in cats. Neuroscience 146, 1841–1853. Buckler, K.J., Vaughan-Jones, R.D., 1994a. Effects of hypoxia on membrane potential and intracellular calcium in rat neonatal carotid body type I cells. J. Physiol. 476, 423–428. Buckler, K.J., Vaughan-Jones, R.D., 1994b. Effects of hypercapnia on membrane potential and intracellular calcium in rat carotid body type I cells. J. Physiol. 478, 157–171. Buckler, K.J., 1997. A novel oxygen-sensitive potassium current in rat carotid body type I cells. J. Physiol. 498, 649–662. Budygin, E.A., John, C.E., Mateo, Y., Jones, S.R., 2002. Lack of cocaine effect on dopamine clearance in the core and shell of the nucleus accumbens of dopamine transporter knock-out mice. J. Neurosci. 22, RC222. Carroll, J.L., Boyle, K.M., Wasicko, M.J., Sterni, L.M., 2005. Dopamine D2 receptor modulation of carotid body type I cell intracellular calcium in developing rats. Am. J. Physiol. Lung Cell Mol. Physiol. 288, L910–L916. Chen, I.L., Hansen, J.T., Yates, J.T., 1985. Dopamine beta-hydroxylase-like immunoreactivity in the rat and cat carotid body: a light and electron microscopic study. J. Neurocytol. 14, 131–144. Cohen, G., Han, Z.Y., Grailhe, R., Gallego, J., Gaultier, C., Changeux, J.P., Lagercrantz, H., 2002. Beta 2 nicotinic acetylcholine receptor subunit modulates protective responses to stress: A receptor basis for sleepdisordered breathing after nicotine exposure. Proc. Natl. Acad. Sci. USA 99, 13272–13277. Conde, S.V., Monteiro, E.C., 2006. Activation of nicotinic ACh receptors with alpha4 subunits induces adenosine release at the rat carotid body. Br. J. Pharmacol. 147, 783–789. Conde, S.V., Gonzalez, C., Batuca, J.R., Monteiro, E.C., Obeso, A., 2008. An antagonistic interaction between A2B adenosine and D2 dopamine receptors modulates the function of rat carotid body chemoreceptor cells. J. Neurochem. 107, 1369– 1381. Dasso, L.L., Buckler, K.J., Vaughan-Jones, R.D., 1997. Muscarinic and nicotinic receptors raise intracellular Ca2+ levels in rat carotid body type I cells. J. Physiol. 498, 327–338. Decker, M.W., Anderson, D.J., Brioni, J.D., Donnelly-Roberts, D.L., Kang, C.H., O’Neill, A.B., Piattoni-Kaplan, M., Swanson, S., Sullivan, J.P., 1995. Erysodine, a competitive antagonist at neuronal nicotinic acetylcholine receptors. Eur. J. Pharmacol. 280, 79–89. Dinger, B., González, C., Yoshizaki, K., Fidone, S., 1981. Alpha-bungarotoxin binding in cat carotid body. Brain Res. 205, 187–193. Dinger, B., Gonzalez, C., Yoshizaki, K., Fidone, S., 1985. Localization and function of cat carotid body nicotinic receptors. Brain Res. 339, 295–304. Doherty, M.D., Gratton, A., 1992. High-speed chronoamperometric measurements of mesolimbic and nigrostriatal dopamine release associated with repeated daily stress. Brain Res. 586, 295–302. Donnelly, D.F., 2009. Nicotinic acetylcholine receptors do not mediate excitatory transmission in young rat carotid body. J. Appl. Physiol. 107, 1806–1816. Eyzaguirre, C., Zapata, P., 1968. The release of acetylcholine from carotid body tissues. Further study on the effects of acetylcholine and cholinergic blocking agents on the chemosensory discharge. J. Physiol. 195, 589–607. Fagerlund, M.J., Kåhlin, J., Ebberyd, A., Schulte, G., Mkrtchian, S., Eriksson, L.I., 2010. The human carotid body: expression of oxygen sensing and signaling genes of relevance for anesthesia. Anesthesiology 113, 1270–1279. Gauda, E.B., Cooper, R., Johnson, S.M., McLemore, G.L., Marshall, C., 2004. Autonomic microganglion cells: a source of acetylcholine in the rat carotid body. J. Appl. Physiol. 94, 384–391. Gerhardt, G.A., Oke, A.F., Nagy, G., Moghaddam, B., Adams, R.N., 1984. Nafion-coated electrodes with high selectivity for CNS electrochemistry. Brain Res. 290, 39039. Goldman, W.F., Eyzaguirre, C., 1984. The effect of dopamine on glomus cell membranes in the rabbit. Brain Res. 321, 337–340. González, C., Almaraz, L., Obeso, A., Rigual, R., 1994. Carotid body chemoreceptors: from natural stimuli to sensory discharges. Physiol. Rev. 74, 829–898. Gotti, C., Clementi, F., 2004. Neuronal nicotinic receptors: from structure to pathology. Prog. Neurobiol. 74, 363–396. Higashi, T., Yamaguchi, S., McIntosh, J.M., Shirahata, M., 2003. Nicotinic acetylcholine receptor channels in cat chemoreceptor cells. Adv. Exp. Med. Biol. 536, 285–290. Iturriaga, R., Alcayaga, J., Zapata, P., 1996. Dissociation of hypoxia-induced chemosensory responses and catecholamine efflux in the cat carotid body superfused in vitro. J. Physiol. 497, 551–564. Iturriaga, R., Alcayaga, J., Zapata, P., 2000. Lack of correlation between cholinergicinduced changes in chemosensory activity and dopamine release from the cat carotid body in vitro. Brain Res. 868, 380–385. Iturriaga, R., Alcayaga, J., 2004. Neurotransmission in the carotid body: transmitters and modulators between glomus cells and petrosal ganglion nerve terminals. Brain Res. Rev. 47, 46–53. Iturriaga-Vásquez, P., Carbone, A., García-Beltrán, O., Livingstone, P.D., Biggin, P.C., Cassels, B.K., Wonnacott, S., Zapata-Torres, G., Bermudez, I., 2010. Molecular determinants for competitive inhibition of alpha4beta2 nicotinic acetylcholine receptors. Mol. Pharmacol. 78, 366–375. Jensen, A.A., Frolund, B., Liljefors, T., 2005. Neuronal nicotinic acetylcholine receptors: structural revelations, target identifications, and therapeutic inspirations. J. ed. Chem. 48, 4705–4745..
(10) 124. R.C. Meza et al. / Neurochemistry International 60 (2012) 115–124. Kahlin, J., Eriksson, L.I., Ebberyd, A., Fagerlund, M.J., 2010. Presence of nicotinic, purinergic and dopaminergic receptors and the TASK-1 K+-channel in the mouse carotid body. Respir. Physiol. Neurobiol. 172, 122–128. Niane, L., Joseph, V., Bairam, A., 2009. Role of cholinergic-nicotinic receptors on hypoxic chemoreflex during postnatal development in rats. Respir. Physiol. Neurobiol. 169, 323–332. Nurse, C.A., 2005. Neurotransmission and neuromodulation in the chemosensory carotid body. Auton. Neurosci. Basic Clin. 120, 1–9. Obeso, A., Gómez-Niño, M., Almaraz, L., Dinger, B., Fidone, S., González, C., 1997. Evidence for two types of nicotinic receptors in the cat carotid body chemoreceptor cells. Brain Res. 754, 298–302. Oomori, Y., Nakaya, K., Tanaka, H., Iuchi, H., Ishikawa, K., Satoh, Y., Ono, K., 1994. Immunohistochemical and histochemical evidence for the presence of noradrenaline, serotonin and gamma-aminobutyric acid in chief cells of the mouse carotid body. Cell Tissue Res. 278, 249–254. Ortiz, F.C., Varas, R., 2010. Muscarinic regulation of a TASK-like channel in rat carotid body chemoreceptor cells. Brain Res. 1323, 74–78. Reyes, E.P., Fernández, R., Larraín, C., Zapata, P., 2007. Effects of combined cholinergic-purinergic block upon cat carotid body chemoreceptors in vitro. Respir. Physiol. Neurobiol. 156, 17–22. Schultz, H.D., Li, Y.L., Ding, Y., 2007. Arterial chemoreceptors and sympathetic nerve activity: implications for hypertension and heart failure. Hypertension 50, 6–13.. Shen, J.X., Yakel, J.L., 2009. Nicotinic acetylcholine receptor-mediated calcium signaling in the nervous system. Acta Pharmacol. Sin. 30, 673–680. Shirahata, M., Balbir, A., Otsubo, T., Fitzgerald, R.S., 2007. Role of acetylcholine in neurotransmission of the carotid body. Respir. Physiol. Neurobiol. 157, 93–105. Ureña, J., Fernández-Chacón, R., Benot, A.R., 1994. Hypoxia induces voltagedependent Ca2+ entry and quantal dopamine secretion in carotid body glomus cells. Proc. Natl. Acad. Sci. USA 91, 10208–10211. Varas, R., Alcayaga, J., Iturriaga, R., 2003. ACh and ATP mediate excitatory transmission in cat carotid identified chemoreceptor units in vitro. Brain Res. 988, 154–163. Vicario, I., Rigual, R., Obeso, A., González, C., 2000. Characterization of the synthesis and release of catecholamine in the rat carotid body in vitro. Am. J. Physiol. Cell Physiol. 278, C490–C499. Wyatt, C.N., Peers, C., 1993. Nicotinic acetylcholine receptors in isolated type I cells of the neonatal rat carotid body. Neuroscience 54, 275–281. Wyatt, C.N., Peers, C., 1995. Ca2+-activated K+ channels in isolated type cells of the neonatal rat carotid body. J. Physiol. 483, 559–565. Xu, F., Xu, J., Tse, F.W., Tse, A., 2006. Adenosine stimulates depolarization and rise in cytoplasmic [Ca2+] in type I cells of rat carotid bodies. Am. J. Physiol. Cell P. 290, C1592–C1598. Zhang, M., Zhong, H., Vollmer, C., Nurse, C.A., 2000. Co-release of ATP and ACh mediates hypoxic signalling at rat carotid body chemoreceptors. J. Physiol. 525, 143–158..
(11)
Figure
Documento similar
The main findings found in the current dissertation suggest that: (i) the adapted version of the environmental questionnaire ALPHA is a reliable and valid
We then show the findings of a recent approach to Faddeev equations using chiral unitary dynamics, where an explicit cancellation of the two body off shell amplitude with three
Sequencing of messenger RNA encoding the nicotinic acetylcholine receptor
In Chapter 7 we present two methodologies to derive three-outcome few-body symmetric Bell inequalities valid for many-body systems: (i) one based on projecting the set of
Because nicotine and tobacco-specific nitrosamines contribute to carcinogenesis by activating nicotinic acetylcholine receptors (nAChRs) on lung tumors and epithelial cells,
In the case of Loop , the successors attribute for its body has to be the condition expression, that is, the control flow successor of the loop’s last statement will be the
We have extended this analysis to the suboesophageal Capa cells (SECA cells), and show that expression of the same terminal identity gene Capa can be achieved from different
(Notice that A is of Kleinian type if and only if every simple quotient of A is of Kleinian type.) Then we use this characterization and results from [8] to characterize the
