Vasopressin Neurons Respond to Hyperosmotic Stimulation with Regulatory Volume Increase and Secretory Volume Decrease by Activating Ion Transporters and Ca2+ Channels
Kaori Sato-Numataa,b Tomohiro Numatab Yoichi Uetac Yasunobu Okadad,e,f
aJapan Society for the Promotion of Science, Tokyo, Japan, bDepartment of Physiology, School of Medicine, Fukuoka University, Fukuoka, Japan, cDepartment of Physiology, School of Medicine, University of Occupational and Environmental Health, Fukuoka, Japan, d National Institute for Physiological Sciences, Okazaki, Japan, eDepartment of Physiology, School of Medicine, Aichi Medical University, Nagakute, Japan, fDepartment of Physiology, Kyoto Prefectural University of Medicine, Kyoto, Japan
Key Words
Vasopressin neuron • Regulatory volume increase • Secretory volume decrease • Calcium channel • Hyperosmotic stimulation
Abstract
Background/Aims: Arginine vasopressin (AVP) neurons play an important role for sensing a change in the plasma osmolarity and thereby responding with regulated AVP secretion in order to maintain the body fluid homeostasis. The osmo-sensing processes in magnocellular neurosecretory cells (MNCs) including AVP and oxytocin (OXT) neurons of the hypothalamus were reported to be coupled to sustained osmotic shrinkage or swelling without exhibiting discernible cell volume regulation. Since increasing evidence has shown some important differences in properties between AVP and OXT neurons, osmotic volume responses are to be reexamined with distinguishing these cell types from each other. We previously reported that AVP neurons identified by transgenic expression of enhanced green fluorescence protein (eGFP) possess the ability of regulatory volume decrease (RVD) after hypoosmotic cell swelling. Thus, in the present study, we examined the ability of regulatory volume increase (RVI) after hyperosmotic cell shrinkage in AVP neurons. Methods: Here, we used eGFP-identified AVP neurons acutely dissociated from AVP-eGFP transgenic rats. We performed single-cell size measurements, cytosolic RT-PCR analysis, AVP secretion measurements, and patch-clamp studies. Results: The AVP neurons were found to respond to a hyperosmotic challenge with physiological cell shrinkage caused by massive secretion of AVP, called a secretory volume decrease (SVD), superimposed onto physical osmotic cell shrinkage, and also to exhibit the ability of RVI coping with osmotic and secretory cell shrinkage. Furthermore, our pharmacological and molecular examinations indicated that AVP secretion and its associated SVD event are triggered by activation of T-type Ca2+ channels, and the RVI event is attained by parallel operation of Na+/H+ exchanger and Cl−/HCO3− anion exchanger. Conclusion: Thus, it is concluded that AVP neurons respond to hyperosmotic stimulation with the regulatory volume increase and the secretory volume increase by activating ion transporters and Ca2+ channels, respectively.
Introduction
Arginine vasopressin (AVP) is an antidiuretic hormone which maintains osmotic pressure stability in the body fluid. AVP-producing neurons are localized in the supraoptic nucleus (SON) and paraventricular nucleus (PVN) of the hypothalamus. Synthesized AVP in these AVP neurons is secreted by an exocytotic mechanism into the systemic circulation from the axon terminal extending to the posterior pituitary gland. The amount of AVP secreted into the systemic circulation is controlled in the hypothalamus by osmo-sensing information derived not only from the axons of osmosensory neurons localized in the organum vasculosum laminae terminalis (OVLT) and subfornical organ (SFO) but also directly from AVP neurons themselves in the SON and PVN.
Cell volume regulation is, in general, a prerequisite function coping with osmotic perturbation in animal cells to survive under both physiological and pathological conditions [1-5]. In contrast, the osmo-sensing processes in magnocellular neurosecretory cells (MNCs), which consist of AVP neurons and oxytocin (OXT) neurons, were, in particular, reported to be coupled to sustained osmotic shrinkage or swelling without exhibiting discernible cell volume regulation [6]. In rat AVP neurons distinguished from OXT neurons by transgenic expression of enhanced green fluorescence protein (eGFP), on the other hand, we recently found that an acute hypoosmotic challenge induces rapid osmotic cell swelling followed by slow cell volume regulation, called the regulatory volume decrease (RVD) which is essentially mediated by activation of volume-sensitive outwardly rectifying anion channel (VSOR) [7]. However, it remains to be examined whether the eGFP-identified AVP neurons can respond to acute hyperosmotic stress with the regulatory volume increase (RVI) after osmotic shrinkage.
AVP neurons secrete AVP not only from the axon terminal to regulate the body fluid homeostasis but also from the soma and dendrites within the brain [8-11] to exert a variety of autocrine/paracrine actions, such as depressing effects on the EPSCs in MNCs [12], optimizing effects on the phasic firing pattern of the neurons [13], promoting effects on the reconstitution of local arterioles within the hypothalamic nuclei [14], and facilitating effects on their RVD process by augmenting VSOR activity [7]. Both axonal terminal secretion and somato-dendritic release of AVP and OXT from MNCs are known to be sensitive to tetanus toxin (TeTx) [8, 15] and dependent on Ca2+ [8, 11, 15-19], indicating an involvement of exocytotic secretory mechanism. Sustained cell shrinkage, called a secretory volume decrease (SVD) [20], was observed in a number of secretory cells under stimulation with secretagogues until the stimuli are withdrawn [5, 20-27]. Thus, there is a possibility that such an SVD event is coupled to somato-dendritic AVP release from dissociated AVP neurons in response to a hyperosmotic challenge.
In the present study, we examined whether AVP neurons exhibit, in addition to osmotic cell shrinkage, SVD and RVI in response to acute hyperosmotic stimulation, by using AVP neurons isolated from transgenic rats expressing eGFP under the control of the AVP promoter [28] in the single-cell preparation free of OXT neurons, glia cells, and connective tissue cells. Our data indicated that AVP neurons respond to a hyperosmotic challenge not only with physical osmotic shrinkage but also a physiological SVD event, which is sensitive to TeTx and blockers for T-type calcium channels, and also do possess the RVI ability, which is sensitive to blockers for Na+/H+ exchanger (NHE) and Cl−/HCO3− anion exchanger (AE).
Materials and Methods
Animals and Preparation of Acutely Dissociated AVP Neurons
All procedures
involving animals were approved in advance by the Ethics Review Committee for
Animal Experimentation of Fukuoka University and were in accordance with the
guidelines of the Physiological Society of Japan. Non-transgenic female Wistar
rats (Charles River Laboratories Japan,
Yokohama, Japan) and heterozygous transgenic female Wistar rats, which
express an AVP-enhanced green fluorescent protein (eGFP) fusion gene [28],
were bred and housed under standardized conditions (12-h:12-h light/dark cycle)
with food and water. For all the experiments, 4–5-week old AVP-eGFP transgenic
female rats were used.
Acutely dissociated AVP neurons were prepared, as described previously [7], and incubated in Ringer solution containing (in mM): 140 NaCl, 5 KCl, 1 MgCl2, 2 CaCl2, 10 HEPES and 10 glucose (adjusted to pH 7.25 with Tris, 300 mosmol/kg-H2O, bubbled with 100% O2) at room temperature (22–26°C). In all of the experiments, eGFP expression was confirmed each time under a fluorescence microscope to identify given SON neurons as AVP neurons.
Single Cell Size Measurements
Single-cell size was measured at room temperature in AVP neurons adhered to a cover glass that were exposed to hyperosmotic solution (430 mosmol/kg-H2O) after a 45-min preincubation with isoosmotic solution (315 mosmol/kg-H2O). The experimental solutions contained (in mM): 120 NaCl, 2.5 KCl, 2 MgCl2, 2 CaCl2, 25 NaHCO3, 10 glucose (315 or 430 mosmol/kg-H2O adjusted with mannitol). Solutions were always prepared immediately before experimental use to achieve reproducible HCO3 concentrations and bubbling with 5% CO2 throughout the experiments. The cells were visualized with a charge-coupled device camera (ORCA-ER-1394, Hamamatsu Photonics) and recorded with AquaCosmos software (version 2.0: Hamamatsu Photonics). The CSA of the soma was measured as an indicator of cell size by ImageJ software. To inhibit exocytosis, we preincubated cells with isoosmotic solution containing 15 nM tetanus toxin (TeTx: Sigma-Aldrich, Tokyo, Japan) for 70 min, and then we performed experiments by applying hyperosmotic solution in the presence of TeTx.
RNA Isolation and RT-PCR
Using the RNeasy Micro Kit (Qiagen, Tokyo, Japan), total cellular RNAs were extracted from the cytosol suctioned into patch pipettes from 10 AVP neurons and pooled. RNA samples were reverse-transcribed using the ReverTra Ace qPCR RT Master Mix (TOYOBO, Osaka, Japan) according to the manufacturer’s protocols. Gene-specific primers used for PCR were designed with Primer3 software (http://frodo.wi.mit.edu/primer3/) and NCBI BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi) to identify complementary sequences in the rat genome. The primers sequences used in the RT-PCR experiments were listed in Table 1 along with the product sizes and GenBank accession numbers. As a positive control, we amplified the glyceraldehyde-3-phosphate-dehydrogenase (GAPDH) sequence. As a negative control, we performed RT-PCR without reverse transcriptase. PCR was performed with 0.4 U of KOD -Plus- Neo Taq (TOYOBO). Amplification was carried out in a thermal cycler (Tadvanced 96 SG: Biometra, Göttingen, Germany) under the following conditions: initial heating at 94°C for 2.5 min, followed by 40 cycles of denaturation at 94°C for 15 s, annealing at 60°C for 30 s and extension at 68°C for 1 min, and then final extension at 68°C for 5 min. The products of RT-PCR were electrophoresed on a 2% agarose gel, and then cloned into the pGEM-T Easy vector (Promega, Tokyo, Japan) after purification with the Wizard SV Gel and PCR Clean-Up System (Promega). Plasmids were purified with the Wizard Plus Minipreps DNA Purification System (Promega) and used as templates for sequencing with ABI PRISM 310 genetic analyzer (Applied Biosystems, Foster City, CA) or with ABI 3730xl genetic analyzer (FASMAC, Kanagawa, Japan)
Measurements of AVP Secretion
The amounts of AVP secretion from dissociated AVP neurons exposed to isoosmotic and hyperosmotic solutions were measured at 37°C using the arg8-Vasopressin Enzyme Immunoassay Kit (Assay Designs, Ann Arbor, MI) according to the manufacturer’s protocol. The experimental solutions were the same as used in the single cell size measurements. After a 20-min preincubation of a batch of ~3800 isolated neurons in 200 mL of iso-osmotic solution, 800 mL of isoosmotic or hyperosmotic solution was added to the cell suspension and the cells were incubated for 90 min. These solutions were then centrifuged at 400g for 10 min at 4°C. Extraction of AVP was performed with the supernatants, which were applied onto C-18 SEP-COLUMNs (Phoenix Pharmaceuticals, Burlingame, CA) according to the manufacturer’s protocol using buffers A and B (Phoenix Pharmaceuticals). Extracted AVP was dried with a vacuum freeze-dryer (VD-800F: Taitec). AVP was assayed with a Microplate reader (MultiscanMS-UV, Lab Systems) at a wavelength of 414 nm. To inhibit exocytosis, we preincubated cells in isoosmotic solution containing 15 nM TeTx for 70 min, and then we performed experiments by applying hyperosmotic solution containing TeTx.
Electrophysiology
The patch electrodes had a resistance of around 1–3 MΩ. Currents or voltages were recorded using an Axopatch 200B amplifier (Axon Instruments, Union City, CA) coupled to DigiData 1440A A/D and D/A converters (Axon Instruments). Currents or voltage signals were filtered at 5 kHz and digitized at 20 kHz. pClamp software (version 9.0.2: Axon Instruments) was used for command pulse control, data acquisition and analysis. For the measurements of Ca2+ channel currents, whole-cell voltage-clamp recordings were performed at room temperature. The time course of current activation was monitored by repetitively applying (every 10 s) alternating pulses (0.3-s duration) of −10 mV from a holding potential of −80 mV. To observe voltage dependence of the current profile, step pulses were applied (every 10 s) from the holding potential to test potentials of -60 mV to +70 mV for 0.3 s in 10 mV increments. For the peak value, we measured the largest value among the inward currents observed immediately after stimulation of the test pulse. For the steady-state value, we measured the value at 280 ms after stimulation of the test pulse. Series resistance (<10 MΩ) was compensated (to 70–80%) to minimize voltage errors. The intracellular (pipette) solution contained (in mM): 160 NMDG-phosphate, 4 MgCl2, 40 HEPES, 10 EGTA, 12 phosphocreatine, 0.1 leupeptine, 2 ATP-Tris salt and 0.4 GTP-Tris salt (280 mosmol/kg-H2O, adjusted to pH 7.2 with phosphoric acid). The isosmotic or hyperosmotic extracellular (bath) solution (300 or 380 mosmol/kg-H2O) contained (in mM): 60 TEA-Cl, 10 HEPES, 50 BaCl2, 10 glucose and 0 or 80 mannitol (adjusted to pH 7.3 with TEA-OH). For the measurements of membrane potential and spontaneous firing, perforated whole-cell current-clamp recordings were performed at 32–35°C controlled by using an automatic temperature controller (TC-324; Warner Instruments, Hamden, CT) with the pipette solution containing (in mM): 99 K2SO4, 31 KCl, 5 MgCl2, 0.2 EGTA, 5 HEPES (280 mosmol/kg-H2O, adjusted to pH 7.4 with KOH). Pipette solution was placed in the tip of the pipette by capillary action (~5 s), and then pipettes were backfilled with nystatin-containing (200 μg/mL) pipette solution. The isosmotic or hyperosmotic bath solution (300 or 380 mosmol/kg-H2O) contained (in mM): 120 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 glucose, 10 HEPES and 20 or 100 mannitol (adjusted to pH 7.3 with NaOH).
Statistical Analysis
Data are given as the means ± SEM of observations (n). Statistical differences of the data were evaluated using one-way ANOVA followed by a Bonferroni-type for multiple comparisons and considered to be significant at p < 0.05.
Chemicals
All chemicals except nickel were prepared on the day of the experiments by diluting stock solutions of 1000-fold concentrations. Ni2+ (Wako Pure Chemical) was directly made in the solution for each experiment and used during that day. The stocks of flufenamic acid (FFA: Sigma-Aldrich), 4,4’-diisothiocyanatostilbene-2,2’-disulfonic acid disodium salt hydrate (DIDS; Sigma-Aldrich), benzamil (Sigma-Aldrich), and 3-aminobenzamide (3-AB: Sigma-Aldrich) were made in dimethyl sulfoxide (DMSO: Wako Pure Chemical, Osaka, Japan) and kept at −20°C. The stocks of TeTx, ω-conotoxin GVIA (ω-CgTx: PEPTIDE INSTITUTE, INC., Osaka, Japan), and amiloride (Sigma-Aldrich) were made in distilled water and kept at −20°C.
Results
Persistent Osmotic Shrinkage in Dissociated AVP Neurons under Hyperosmotic Stimulation and Its Aggravation by Blocking the NHE and AE Activities
When dissociated AVP neurons were exposed to hyperosmotic extracellular solution (136.5% osmolarity), the cross-sectional area (CSA) of AVP neurons became rapidly smaller, and such hyperosmotic cell shrinkage persisted over 90 min, as shown in Fig. 1 (a: open squares; b: Control), being consistent with a previous report for MNCs [6]. Since AVP neurons apparently lack the RVI ability, we next examined molecular expression of NHE and AE transporters as well as of the epithelial Na+ channel (ENaC) and the transient receptor potential cation channel, subfamily M, member 2 (TRPM2), that are both known to be responsible for the hypertonicity-induced cation channel (HICC) [29, 30], in AVP neurons, because the volume-regulatory NaCl influx involved in the RVI event is known to be attained by these Na+ and Cl- transporters or Na+-permeable cation channels in a variety of animal cells [29, 31-33]. As shown in Fig. 2a, cytosol RT-PCR analysis showed significant expression of mRNAs for NHE1, NHE4, NHE5, AE2, AE3, and TRPM2 in AVP neurons. The sequences of above six transporter/channel genes expressed in AVP neurons were found to completely match by sequence analysis to the sequences corresponding to their respective rat NHEs, AEs and TRPM2. On the other hand, any ENaC mRNAs were not be detected in the cytosol of AVP neurons (Fig. 2a), although α-, β-, and γ-ENaC were all sizably detected in the SON region (Fig. 2b). The sequences of above three ENaC genes expressed in the SON region completely matched to the sequences corresponding to their respective rat ENaC channels. Thus, the effects of blockers for these transporters and channels on hyperosmotic cell shrinkage were next examined by measuring the CSA of AVP neurons. When applying an NHE blocker, amiloride (100 mM), or an AE blocker, DIDS (100 mM), osmotic cell shrinkage became prominently exaggerated, as shown in Fig. 1a (filled symbols). In the presence of amiloride or DIDS, as summarized in Fig. 1b (+amiloride/+DIDS), the shrinking CSA change never recovered even after 90 min after a hyperosmotic challenge and moreover became much more shrunken by around 30%. In contrast, applications of an ENaC blocker, benzamil [34], and a blocker of poly (ADP-ribose) polymerase (PARP), 3-AB [35], which thereby inhibits PARP-dependent TRPM2 channel activity [36], failed to affect the persistent cell shrinkage even 90 min after a hyperosmotic challenge (Fig. 1b). These results suggest that AVP neurons do possess the RVI ability, in which NHE and AE are involved, though they apparently exhibit sustained shrinkage under hyperosmotic stimulation.
Prominent RVI Event Sensitive to Blockers of NHE and AE in Dissociated AVP Neurons under the Conditions where Hyperosmolarity-Induced AVP Secretion was Inhibited
When AVP neurons were exposed to hyperosmotic solution containing flufenamic acid (FFA, 100 mM), which was shown to markedly suppress T- and N-type voltage-gated Ca2+ channel (VGCC) currents and thereby abolish action potential firings in AVP neurons under isotonic conditions [37], the RVI event surprisingly became evident (Fig. 3a: filled circles) in a manner sensitive to amiloride and DIDS at 100 mM (Fig. 3a: filled triangles and reversed triangles). A similar prominent RVI event was also observed in the presence of a blocker of vesicular exocytotic secretion, TeTx (15 nM), or a known blocker of VGCC, Ni2+ (3 mM), which was found to predominantly suppress T-type VGCC currents and thereby abolish action potential firings in AVP neurons under normotonic conditions [37], in dissociated AVP neurons under hyperosmotic conditions (Fig. 3b). As summarized in Fig. 3c, TeTx, Ni2+ and FFA recovered the CSA by 50 to 60% in dissociated AVP neurons 90 min after a hyperosmotic challenge, and the FFA-induced CSA recovery was abolished by amiloride and DIDS. Since release of AVP and OXT from MNCs are known to be inhibited by TeTx [8, 15] and to be dependent on Ca2+ influx through VGCCs [38-41], it is likely that the FFA effects on the CSA changes in AVP neurons after a hyperosmotic challenge are caused by inhibiting hyperosmotic AVP secretion. In fact, hyperosmolarity-induced AVP secretion was remarkably inhibited not only by TeTx and Ni2+ but also by FFA, as shown in Fig. 3d. These results indicate that AVP neurons respond to hyperosmotic stimulation with the SVD event caused by massive vesicular exocytotic release of AVP, superimposed onto osmotic cell shrinkage. It is noted that they exhibited the NHE- and AE-dependent RVI event in opposition to such dual volume-decreasing events.
Sensitivity of Action Potential Firing Activity to FFA and Ni2+ in Dissociated AVP Neurons under Hyperosmotic Stimulation
It is known that AVP secretion depends on action potential firing activity which stimulates Ca2+ entry via VGCCs in MNCs [42-46]. Also, we recently found that FFA and Ni2+ abolish action potential firings in rat AVP neurons under normotonic conditions [37]. Thus, we examined the effects of FFA and a known VGCC blocker, Ni2+, on the firing activity in dissociated AVP neurons after hyperosmotic stimulation by nystatin-perforated whole-cell current-clamp recordings. When the neurons were exposed to a hyperosmotic solution (126.7% osmolarity), the firing activity was markedly enhanced with inducing a slight depolarizing shift of the resting membrane potential, as shown in Fig. 4. These results are in good agreement with previous observations [45, 47-49]. The firing activity observed after hyperosmotic stimulation was prominently suppressed by application of FFA (Fig. 4a) and Ni2+ (Fig. 4b). The firing frequency was increased around three-fold by hyperosmotic stimulation and was almost completely abolished by application of FFA and Ni2+ (Fig. 4c). In contrast, hypertonicity-enhanced action potential firing activity in AVP neurons were, as shown in Fig. 4c, not affected by application of ω-CgTx (0.5 mM), which is a specific blocker of N-type Ca2+ channels [50-52] and was found to predominantly suppress N-type VGCC currents in AVP neurons under isotonic conditions [37]. These results suggest that FFA inhibits hyperosmolarity-induced AVP secretion and thereby the SVD event by inhibiting the firing activity presumably through suppression of Ni2+-sensitive T-type VGCCs in AVP neurons under hyperosmotic conditions.
Sensitivity of the Ca2+ Channel Activities to FFA and Ni2+ in Dissociated AVP Neurons under Hyperosmotic Stimulation
To examine whether FFA and Ni2+ actually suppresses VGCC currents in dissociated AVP neurons under not only isotonic but also hyperosmotic conditions, we then made conventional whole-cell recordings. Under exposure to 50 mM Ba2+, AVP neurons showed Ba2+ currents through VGCCs exhibiting the peak inward currents at around −10 mV under isotonic conditions (Fig. 5a: Iso), as observed in our recent studies under normotonic conditions [37]. A hyperosmotic challenge (126.7% osmolarity) did not significantly affect depolarization-induced Ba2+ currents (Fig. 5a: Hyper), indicating that the VGCCs in AVP neurons per se do not exhibit osmosensitivity. As found in our preceding study [37], VGCC currents in AVP neurons under normotonic conditions were largely suppressed by Ni2+ (3 mM) with shifting the voltage showing the peak current from around −10 mV to around +10 mV (Fig. 5b), whereas they were suppressed by ω-CgTx (0.5 mM) without shifting the voltage showing the peak current (Fig. 5c). Ni2+-insensitive currents observed under isosmotic conditions, which were shown to represent mainly N-type VGCC currents [37], were little affected by a hyperosmotic challenge (Fig. 5b). The Ni2+-insensitive current observed under hyperosmotic conditions was markedly suppressed by ω-CgTx (Fig. 5b). Also, ω-CgTx-insensitive currents observed under isosmotic conditions, which was shown to represent mainly T-type VGCC currents [37], were also little affected by a hyperosmotic challenge (Fig. 5c). The ω-CgTx-insensitive current observed under hyperosmotic conditions was mostly abolished by Ni2+ (Fig. 5c). These results indicate that VGCC currents observed in AVP neurons under hyperosmotic conditions are composed mainly of T-type and N-type Ca2+ channels, as was the case of those observed under isosmotic conditions [37].
When FFA was applied to AVP neurons, their VGCC currents observed under hyperosmotic conditions were markedly suppressed by FFA (Fig. 6a, d), just as the case of those observed under isotonic conditions [37]. In the presence of w-CgTx, FFA suppressed the remaining (mainly T-type) VGCC currents (Fig. 6b, d); and also, in the presence of Ni2+, FFA largely diminished the remaining (mainly N-type) VGCC currents (Fig. 6c, d) in AVP neurons under hyperosmotic conditions. Taken together, it is concluded that FFA and Ni2+ block voltage-gated Ca2+ channel activities in dissociated rat AVP neurons under hyperosmotic conditions and thereby inhibit the SVD event coupled to hypertonicity-induced AVP secretion.
Discussion
Release of AVP and OXT from MNCs in the hypothalamus is evoked by Ca2+ influx through VGCCs [41, 53, 54]. Previous electrophysiological studies on VGCC currents showed that the soma of rat SON MNCs express T-, N-, L-, P/Q- and R-type Ca2+ channels [55-59]. Among these types of VGCCs, somato-dendritic expression of T-, N- and L-type Ca2+ channels in AVP neurons were indirectly suggested by observing sensitivity to VGCC blockers of increases in ([Ca2+]i) in response to AVP [46] and pituitary adenylate cyclase-activating polypeptide (PACAP) [60]. Although direct electrophysiological studies showed expression of L-, N- and P/Q-type of Ca2+ channels in the neurohypophysial nerve endings of rat AVP neurons under unstimulated normotonic conditions [39, 41], such studies have not been conducted in the soma/dendrites in AVP neurons. Under dehydration conditions, somato-dendritic L-type Ca2+ channel currents were found to be increased in both AVP and OXT neurons in rats after 16-24 h water deprivation [61], but such studies have not been done upon an acute hyperosmotic challenge. Also, T-type Ca2+ channel currents were recorded in guinea pig AVP-containing MNCs in the slice preparation of the SON under normotonic conditions [62]. Recently, we showed that in rat AVP neurons under normotonic conditions, T-type Cav3.1 and N-type Cav2.2 channels are molecularly and functionally expressed, but Cav3.1 is primary involved in their action potential generation [37]. In the present study, for the first time, expression of VGCCs was examined upon acute hyperosmotic stimulation in dissociated rat AVP neurons that are largely devoid of nerve terminals and were identified by transgenic eGFP expression, and w-CgTx-sensitive (mainly N-type) and Ni2+-sensitive (mainly T-type) VGCC channels were found to be predominantly functioning in the plasma membrane of AVP neurons under hyperosmotic stimulation by whole-cell recordings (Fig. 5). Also, both N- and T-type Ca2+ channels in AVP neurons were found to be sensitive to FFA under hyperosmotic conditions (Fig. 6). In addition, the spontaneous firing in AVP neurons under hyperosmotic stimulation was virtually abolished by FFA and Ni2+ (Fig. 4), suggesting that the firing activity under acute hyperosmotic conditions is caused mainly by T-type VGCC channels.
The ability of cell volume regulation is fundamental to the survival and function of animal cells under not only physiological conditions but also pathological situations [1-4, 31, 63, 64]. The volume regulation processes are classified into RVD and RVI, that take place after cell swelling and shrinkage, respectively, and they are essentially involved not only in the relief of osmotic volume perturbation but also in cell shape changes associated with cell proliferation or mitosis, differentiation and migration [2, 55, 65, 66]. Persistent cell swelling and shrinkage lead to induction of necrotic and apoptotic cell death, respectively [4, 67]. On the other hand, rat SON MNCs were reported to exhibit sustained cell swelling and shrinkage in response to hypoosmotic and hyperosmotic stimulation, respectively, without showing discernible cell volume regulation [6]. However, we demonstrated that AVP neurons identified by transgenic eGFP expression can respond to a hypoosmotic challenge with a slow RVD event which is attained by activation of volume-sensitive outwardly rectifying anion channels (VSORs) by osmotic cell swelling and AVP receptor-mediated autocrine signaling [7]. In the present study, eGFP-identified AVP neurons were found to respond with sustained cell shrinkage to a hyperosmotic challenge without apparently showing volume recovery (Fig. 1a). However, when FFA was added to a hyperosmotic solution, a typical RVI event became evident after osmotic cell shrinkage in the AVP neurons (Fig. 3a). The RVI event observed in the presence of FFA was completely eliminated by application of amiloride and DIDS (Fig. 3a, c) that are blockers of NHE and AE, respectively. In contrast, as shown in Fig. 1b, changes in the CSA values of AVP neurons after exposure to a hyperosmotic solution were affected neither by benzamil, which is known to block ENaC much more strongly than amiloride [34], nor by 3-AB, which is known to block PARP [35] and thereby suppress PARP-dependent TRPM2 channel activity [36]. Thus, NHE and AE are likely involved in the RVI process in the presence of FFA in AVP neurons. Taken together, it appears that rat AVP neurons do possess the RVI ability which is largely attained by parallel operation of NHE and AE, and that the RVI event is hidden behind some additional cell shrinkage process which is sensitive to FFA.
Essentially similar RVI-actualizing effects of FFA were found to be duplicated by application of a vesicular transport inhibitor, TeTx, and a known T-type VGCC blocker, Ni2+, in AVP neurons upon a hyperosmotic challenge (Fig. 3b, c). In addition, not only TeTx but also Ni2+ and FFA were found to prominently inhibit somatodendritic AVP secretion (Fig. 3d). Exocytotic AVP secretion from both axon terminals and somata/dendrites is known to be regulated by the frequency and pattern of action potential firings generated in the soma due to activation of both Na+ and Ca2+ channels [45, 68, 69]. In fact, firing activity in eGFP-identified AVP neurons was enhanced by hyperosmotic stimulation in a manner sensitive to Ni2+ and FFA (Fig. 4). Thus, an FFA-sensitive component of cell shrinkage represents an SVD event [20] which was observed in a number of secretory cells during stimulation with a variety of secretagogues [5, 20-27]. Taken together, it is concluded that the AVP neurons exhibit three different types of volume responses to a hyperosmotic challenge; that is, physical osmotic cell shrinkage as well as physiological SVD and RVI events.
Ni2+ is known to block a number of VGCCs [70], preferentially T-type Ca2+ channels [71, 72]. FFA was shown to inhibit L-type Ca2+ channels in rat smooth muscle cells [73]. Thus, a possibility arises that FFA blocks Ni2+-sensitive VGCCs in rat AVP neurons, with inhibiting hyperosmolarity-induced enhancement of firing activity and AVP secretion coupled to an SVD event. In the present study, actually, both Ni2+ and FFA were found to prominently suppress VGCC currents in AVP neurons under hyperosmotic stimulation (Fig. 5 and 6). Pharmacological experiments showed that hyperosmotic VGCC currents in AVP neurons are mainly composed of ω-CgTx-sensitive N-type and Ni2+-sensitive T-type Ca2+ channel currents (Fig. 5a, c) and that FFA could block both N- and T-type Ca2+ channel currents (Fig. 6b-d). Thus, it appears that rat AVP neurons respond to acute hyperosmotic stimulation not only with osmotic cell shrinkage but also with the SVD event due to massive somatodendritic AVP secretion that was triggered by activation of T-type Ca2+ channels, and that rat AVP neurons do exhibit the RVI event which is likely attained by parallel operation of NHE and AE coping with these dual volume decreasing events.
Accumulating evidence has shown that a number of important properties are different between AVP and OXT neurons [74]. Therefore, future studies are warranted to clarify whether OXT neurons distinguished from AVP neurons also respond to acute hyperosmotic stimulation not only with osmotic cell shrinkage but also with RVI and SVD events, and which types of VGCCs are activated upon a hyperosmotic challenge.
Conclusion
In summary, we revealed that AVP neurons retain the ability of RVI largely involving NHE and AE activities and also do exhibit the SVD response due to massive AVP exocytosis under hyperosmolar conditions. The volume recovery due to the RVI mechanism and the volume decrease due to the SVD event occurred in parallel in a manner approximately cancelling out the volume changes of each other, and the volume recovery was therefore not apparently observed after the cell volume reduction by hyperosmotic stimulation. It is suggested that the SVD event was triggered by activation of Ni2+- and FFA-sensitive voltage-gated Ca2+ channels in AVP neurons under hyperosmotic conditions.
Acknowledgements
Author Contributions
K.S.-N. conducted all experiments and data analysis. T.N. and Y.U. helped to design the work and commented on the draft. K.S.-N. and Y.O. conceived and designed the work and wrote the manuscript.
Funding Sources
This work was supported in part by Grants-in-Aid for Scientific Research (KAKENHI) from the Japan Society for the Promotion of Science (No. 18J40103) and by Grant of The Clinical Research Promotion Foundation (2020).
Statement of Ethics
All procedures performed in this study were in accordance with the regulations established at the Fukuoka University, with due consideration given to animal welfare and safety and health, and approved by the Ethics Committee of Animal Care and Experimentation, Fukuoka University, Japan (approval No.: 1906025).
Disclosure Statement
The authors have no conflicts of interest to declare.
References
|
1 Hoffmann EK, Lambert
IH, Pedersen SF: Physiology of cell volume regulation in vertebrates. Physiol
Rev 2009;89:193-277. |
|
|
|
|
|
2
Lang F, Busch GL, Ritter M, Volkl H, Waldegger S, Gulbins E, Haussinger D:
Functional significance of cell volume regulatory mechanisms. Physiol Rev
1998;78:247-306. |
|
|
|
|
|
3
Okada Y: Cell volume regulation: the molecular mechanism and volume sensing
machinery. Elsevier 1998:17-21. |
|
|
|
|
|
4
Okada Y, Maeno E, Shimizu T, Manabe K, Mori S, Nabekura T: Dual roles of
plasmalemmal chloride channels in induction of cell death. Pflugers Arch
2004;448:287-295. |
|
|
|
|
|
5
Pedersen S, Hoffmann E, Novak I: Cell volume regulation in epithelial
physiology and cancer. Front Physiol 2013;4:233. |
|
|
|
|
|
6
Zhang Z, Bourque CW: Osmometry in osmosensory neurons. Nat Neurosci
2003;6:1021. |
|
|
|
|
|
7
Sato K, Numata T, Saito T, Ueta Y, Okada Y: V2 receptor-mediated autocrine
role of somatodendritic release of AVP in rat vasopressin neurons under
hypo-osmotic conditions. Sci Signal 2011;4:ra5. |
|
|
|
|
|
8
Dayanithi G, Weller U, Ahnert-Hilger G, Link H, Nordmann JJ, Gratzl M: The
light chain of tetanus toxin inhibits calcium-dependent vasopressin release
from permeabilized nerve endings. Neuroscience 1992;46:489-493. |
|
|
|
|
|
9
Ludwig M, Callahan MF, Neumann I, Landgraf R, Morris M: Systemic osmotic
stimulation increases vasopressin and oxytocin release within the supraoptic
nucleus. J Neuroendocrinol 1994;6:369-373. |
|
|
|
|
|
10
Ludwig M, Horn T, Callahan MF, Grosche A, Morris M, Landgraf R: Osmotic
stimulation of the supraoptic nucleus: central and peripheral vasopressin
release and blood pressure. Am J Physiol Endocrinol Metab 1994;266:E351-E356. |
|
|
|
|
|
11
Tobin VA, Ludwig M: The actin filament and dendritic peptide release. Biochem
Soc Trans 2007;35:1243-1246. |
|
|
|
|
|
12
Kombian SB, Mouginot D, Pittman QJ: Dendritically released peptides act as
retrograde modulators of afferent excitation in the supraoptic nucleus in
vitro. Neuron 1997;19:903-912. |
|
|
|
|
|
13
Gouzènes L, Desarmenien MG, Hussy N, Richard P, Moos FC: Vasopressin
regularizes the phasic firing pattern of rat hypothalamic magnocellular
vasopressin neurons. J Neurosci 1998;18:1879-1885. |
|
|
|
|
|
14
Alonso G, Gallibert E, Lafont C, Guillon G: Intrahypothalamic angiogenesis
induced by osmotic stimuli correlates with local hypoxia: a potential role of
confined vasoconstriction induced by dendritic secretion of vasopressin.
Endocrinology 2008;149:4279-4288. |
|
|
|
|
|
15
de Kock CP, Wierda KD, Bosman LW, Min R, Koksma JJ, Mansvelder HD, Verhage M,
Brussaard AB: Somatodendritic secretion in oxytocin neurons is upregulated
during the female reproductive cycle. J Neurosci 2003;23:2726-2734. |
|
|
|
|
|
16
Cazalis M, Dayanithi G, Nordmann JJ: Hormone release from isolated nerve
endings of the rat neurohypophysis. J Physiol 1987;390:55-70. |
|
|
|
|
|
17
Di Scala-Guenot D, Strosser MT, Richard P: Electrical stimulations of
perifused magnocellular nuclei in vitro elicit Ca2+-dependent,
tetrodotoxin-insensitive release of oxytocin and vasopressin. Neurosci Lett
1987;76:209-214. |
|
|
|
|
|
18
Ludwig M, Sabatier N, Bull PM, Landgraf R, Dayanithi G, Leng G: Intracellular
calcium stores regulate activity-dependent neuropeptide release from
dendrites. Nature 2002;418:85-89. |
|
|
|
|
|
19
Tobin V, Gouty LA, Moos FC, Desarmenien MG: A store-operated current (SOC)
mediates oxytocin autocontrol in the developing rat hypothalamus. Eur J
Neurosci 2006;24:400-404. |
|
|
|
|
|
20
Manabe K, Shimizu T, Morishima S, Okada Y: Regulatory volume increase after
secretory volume decrease in colonic epithelial cells under muscarinic
stimulation. Pflugers Arch 2004;448:596-604. |
|
|
|
|
|
21
Bachmann O, Heinzmann A, Mack A, Manns MP, Seidler U: Mechanisms of
secretion-associated shrinkage and volume recovery in cultured rabbit
parietal cells. Am J Physiol Gastrointest Liver Physiol 2007;292:G711-G717. |
|
|
|
|
|
22
Dissing S, Hansen HJ, Undén M, Nauntofte B: Inhibitory effects of
amitriptyline on the stimulation-induced Ca2+ increase in parotid acini. Eur
J Pharmacol 1990;177:43-54. |
|
|
|
|
|
23
Foskett JK: [Ca2+]i modulation of Cl− content controls cell volume in
single salivary acinar cells during fluid secretion. Am J Physiol Cell
Physiol 1990;259:C998-C1004. |
|
|
|
|
|
24
Lee RJ, Foskett JK: Mechanisms of Ca2+-stimulated fluid secretion by porcine
bronchial submucosal gland serous acinar cells. Am J Physiol Lung Cell Mol
Physiol 2010;298:L210-L231. |
|
|
|
|
|
25
Nakahari T, Murakami M, Sasaki Y, Kataoka T, Imai Y, Shiba Y, Kanno Y: Dose
effects of acetylcholine on the cell volume of rat mandibular salivary acini.
Jpn J Physiol 1991;41:153-168. |
|
|
|
|
|
26
Nakahari T, Murakami M, Yoshida H, Miyamoto M, Sohma Y, Imai Y: Decrease in
rat submandibular acinar cell volume during ACh stimulation. Am J Physiol
Gastrointest Liver Physiol 1990;258:G878-G886. |
|
|
|
|
|
27
Numata T, Sato-Numata K, Okada Y, Inoue R: Cellular mechanism for herbal
medicine Junchoto to facilitate intestinal Cl− /water secretion that
involves cAMP-dependent activation of CFTR. J Nat Med 2018;72:694-705. |
|
|
|
|
|
28
Ueta Y, Fujihara H, Serino R, Dayanithi G, Ozawa H, Matsuda K, Kawata M,
Yamada J, Ueno S, Fukuda A, Murphy D: Transgenic expression of enhanced green
fluorescent protein enables direct visualization for physiological studies of
vasopressin neurons and isolated nerve terminals of the rat. Endocrinology
2005;146:406-413. |
|
|
|
|
|
29
Numata T, Sato K, Christmann J, Marx R, Mori Y, Okada Y, Wehner F: The deltaC
splice-variant of TRPM2 is the hypertonicity-induced cation channel in HeLa
cells, and the ecto-enzyme CD38 mediates its activation. J Physiol
2012;590:1121-1138. |
|
|
|
|
|
30
Plettenberg S, Weiss EC, Lemor R, Wehner F: Subunits alpha, beta and gamma of
the epithelial Na+ channel (ENaC) are functionally related to the
hypertonicity-induced cation channel (HICC) in rat hepatocytes. Pflugers Arch
2008;455:1089-1095. |
|
|
|
|
|
31
Hoffmann EK, Simonsen LO: Membrane mechanisms in volume and pH regulation in
vertebrate cells. Physiol Rev 1989;69:315-382. |
|
|
|
|
|
32
Okada Y: Ion channels and transporters involved in cell volume regulation and
sensor mechanisms. Cell Biochem Biophys 2004;41:233-258. |
|
|
|
|
|
33
Wehner F, Tinel H: Role of Na+ conductance, Na+-H+ exchange, and
Na+-K+-2Cl− symport in the regulatory volume increase of rat
hepatocytes. J Physiol 1998;506:127-142. |
|
|
|
|
|
34
Kleyman TR, Cragoe EJ, Jr.: Amiloride and its analogs as tools in the study
of ion transport. J Membr Biol 1988;105:1-21. |
|
|
|
|
|
35
Purnell MR, Whish WJ: Novel inhibitors of poly (ADP-ribose) synthetase.
Biochem J 1980;185:775-777. |
|
|
|
|
|
36
Roberge S, Roussel J, Andersson DC, Meli AC, Vidal B, Blandel F, Lanner JT,
Le Guennec JY, Katz A, Westerblad H, Lacampagne A, Fauconnier J:
TNF-α-mediated caspase-8 activation induces ROS production and TRPM2
activation in adult ventricular myocytes. Cardiovasc Res 2014;103:90-99. |
|
|
|
|
|
37
Sato-Numata K, Numata T, Ueta Y, Okada Y: Expression and functions of N-type
Cav2.2 and T-type Cav3.1 channels in rat vasopressin neurons under normotonic
conditions. J Physiol Sci 2020;70:49. |
|
|
|
|
|
38
Fisher TE, Bourque CW: Calcium-channel subtypes in the somata and axon
terminals of magnocellular neurosecretory cells. Trends Neurosci
1996;19:440-444. |
|
|
|
|
|
39
Ortiz-Miranda SI, Dayanithi G, Velazquez-Marrero C, Custer EE, Treistman SN,
Lemos JR: Differential modulation of N-type calcium channels by micro-opioid
receptors in oxytocinergic versus vasopressinergic neurohypophysial
terminals. J Cell Physiol 2010;225:276-288. |
|
|
|
|
|
40
Wang D, Fisher TE: Expression of CaV 2.2 and splice variants of CaV 2.1 in
oxytocin- and vasopressin-releasing supraoptic neurones. J Neuroendocrinol
2014;26:100-110. |
|
|
|
|
|
41
Wang G, Dayanithi G, Kim S, Hom D, Nadasdi L, Kristipati R, Ramachandran J,
Stuenkel EL, Nordmann JJ, Newcomb R, Lemos JR: Role of Q-type Ca2+ channels
in vasopressin secretion from neurohypophysial terminals of the rat. J
Physiol 1997;502:351-363. |
|
|
|
|
|
42 Bicknell RJ: Optimizing release from peptide hormone secretory nerve terminals. J Exp Biol 1988;139:51-65. |
|
|
|
|
|
43
Fisher TE, Bourque CW: The function of Ca2+ channel subtypes in exocytotic
secretion: new perspectives from synaptic and non-synaptic release. Prog
Biophys Mol Biol 2001;77:269-303. |
|
|
|
|
|
44
Ludwig M, Bull PM, Tobin VA, Sabatier N, Landgraf R, Dayanithi G, Leng G:
Regulation of activity-dependent dendritic vasopressin release from rat
supraoptic neurones. J Physiol 2005;564:515-522. |
|
|
|
|
|
45
Poulain DA, Wakerley JB: Electrophysiology of hypothalamic magnocellular neurones
secreting oxytocin and vasopressin. Neuroscience 1982;7:773-808. |
|
|
|
|
|
46
Sabatier N, Richard P, Dayanithi G: L-, N- and T- but neither P- nor Q-type
Ca2+ channels control vasopressin-induced Ca2+ influx in magnocellular
vasopressin neurones isolated from the rat supraoptic nucleus. J Physiol
1997;503:253-268. |
|
|
|
|
|
47
Hussy N, Deleuze C, Desarménien MG, Moos FC: Osmotic regulation of neuronal
activity: a new role for taurine and glial cells in a hypothalamic
neuroendocrine structure. Prog Neurobiol 2000;62:113-134. |
|
|
|
|
|
48
Moriya T, Shibasaki R, Kayano T, Takebuchi N, Ichimura M, Kitamura N, Asano
A, Hosaka YZ, Forostyak O, Verkhratsky A, Dayanithi G, Shibuya I: Full-length
transient receptor potential vanilloid 1 channels mediate calcium signals and
possibly contribute to osmoreception in vasopressin neurones in the rat
supraoptic nucleus. Cell Calcium 2015;57:25-37. |
|
|
|
|
|
49
Sharif Naeini R, Witty MF, Seguela P, Bourque CW: An N-terminal variant of
Trpv1 channel is required for osmosensory transduction. Nat Neurosci
2006;9:93-98. |
|
|
|
|
|
50
Kasai H, Aosaki T, Fukuda J: Presynaptic Ca-antagonist omega-conotoxin
irreversibly blocks N-type Ca-channels in chick sensory neurons. Neurosci Res
1987;4:228-235. |
|
|
|
|
|
51
Plummer MR, Logothetis DE, Hess P: Elementary properties and pharmacological
sensitivities of calcium channels in mammalian peripheral neurons. Neuron
1989;2:1453-1463. |
|
|
|
|
|
52
Williams ME, Brust PF, Feldman DH, Patthi S, Simerson S, Maroufi A, McCue AF,
Veliçelebi G, Ellis SB, Harpold MM: Structure and functional expression of an
omega-conotoxin-sensitive human N-type calcium channel. Science
1992;257:389-395. |
|
|
|
|
|
53
Tobin VA, Douglas AJ, Leng G, Ludwig M: The involvement of voltage-operated
calcium channels in somato-dendritic oxytocin release. PLoS One
2011;6:e25366. |
|
|
|
|
|
54
Wang G, Dayanithi G, Newcomb R, Lemos JR: An R-type Ca2+ current in
neurohypophysial terminals preferentially regulates oxytocin secretion. J
Neurosci 1999;19:9235-9241. |
|
|
|
|
|
55
Akita T, Okada Y: Characteristics and roles of the volume-sensitive outwardly
rectifying (VSOR) anion channel in the central nervous system. Neuroscience
2014;275:211-231. |
|
|
|
|
|
56
Fisher TE, Bourque CW: Voltage-gated calcium currents in the magnocellular
neurosecretory cells of the rat supraoptic nucleus. J Physiol
1995;486:571-580. |
|
|
|
|
|
57
Fisher TE, Bourque CW: Distinct omega-agatoxin-sensitive calcium currents in
somata and axon terminals of rat supraoptic neurones. J Physiol
1995;489:383-388. |
|
|
|
|
|
58
Foehring RC, Armstrong WE: Pharmacological dissection of
high-voltage-activated Ca2+ current types in acutely dissociated rat
supraoptic magnocellular neurons. J Neurophysiol 1996;76:977-983. |
|
|
|
|
|
59
Joux N, Chevaleyre V, Alonso G, Boissin-Agasse L, Moos FC, Desarménien MG,
Hussy N: High voltage-activated Ca2+ currents in rat supraoptic neurones:
biophysical properties and expression of the various channel alpha1 subunits.
J Neuroendocrinol 2001;13:638-649. |
|
|
|
|
|
60
Shibuya I, Noguchi J, Tanaka K, Harayama N, Inoue Y, Kabashima N, Ueta Y,
Hattori Y, Yamashita H: PACAP increases the cytosolic Ca2+ concentration and
stimulates somatodendritic vasopresson release in rat supraoptic neurons. J
Neuroendocrinol 1998;10:31-42. |
|
|
|
|
|
61
Zhang W, Star B, Rajapaksha WRAKJS, Fisher TE: Dehydration increases L-type
Ca2+ current in rat supraoptic neurons. J Physiol Sci 2007;580:181-193. |
|
|
|
|
|
62
Erickson KR, Ronnekleiv OK, Kelly MJ: Role of a T-type calcium current in
supporting a depolarizing potential, damped oscillations, and phasic firing
in vasopressinergic guinea pig supraoptic neurons. Neuroendocrinology
1993;57:789-800. |
|
|
|
|
|
63
Delpire E, Gagnon KB: Water homeostasis and cell volume maintenance and
regulation. Curr Top Membr 2018;81:3-52. |
|
|
|
|
|
64
Okada Y: Channelling frozen cells to survival after thawing: opening the door
to cryo-physiology. J Physiol 2016;594:1523-1524. |
|
|
|
|
|
65
Okada Y: Volume expansion-sensing outward-rectifier Cl− channel: fresh
start to the molecular identity and volume sensor. Am J Physiol Cell Physiol
1997;273:C755-C789. |
|
|
|
|
|
66
Wehner F, Olsen H, Tinel H, Kinne-Saffran E, Kinne RK: Cell volume
regulation: osmolytes, osmolyte transport, and signal transduction. Rev
Physiol Biochem Pharmacol 2003;148:1-80. |
|
|
|
|
|
67
Okada Y, Maeno E, Shimizu T, Dezaki K, Wang J, Morishima S: Receptor-mediated
control of regulatory volume decrease (RVD) and apoptotic volume decrease
(AVD). J Physiol 2001;532:3-16. |
|
|
|
|
|
68
Bourque CW, Renaud LP: Calcium-dependent action potentials in rat supraoptic
neurosecretory neurones recorded in vitro. J Physiol 1985;363:419-428. |
|
|
|
|
|
69
Mason WT, Leng G: Complex action potential waveform recorded from supraoptic
and paraventricular neurones of the rat: evidence for sodium and calcium
spike components at different membrane sites. Exp Brain Res 1984;56:135-143. |
|
|
|
|
|
70
Zamponi GW, Bourinet E, Snutch TP: Nickel block of a family of neuronal
calcium channels: subtype- and subunit-dependent action at multiple sites. J
Membr Biol 1996;151:77-90. |
|
|
|
|
|
71
Lee JH, Gomora JC, Cribbs LL, Perez-Reyes E: Nickel block of three cloned
T-type calcium channels: low concentrations selectively block alpha1H.
Biophys J 1999;77:3034-3042. |
|
|
|
|
|
72
Obejero-Paz CA, Gray IP, Jones SW: Ni2+ block of CaV3.1 (alpha1G) T-type
calcium channels. J Gen Physiol 2008;132:239-250. |
|
|
|
|
|
73
Shimamura K, Zhou M, Ito Y, Kimura S, Zou LB, Sekiguchi F, Kitramura K,
Sunano S: Effects of flufenamic acid on smooth muscle of the carotid artery
isolated from spontaneously hypertensive rats. J Smooth Muscle Res
2002;38:39-50. |
|
|
|
|
|
74
Armstrong WE, Foehring RC, Kirchner MK, Sladek CD: Electrophysiological
properties of identified oxytocin and vasopressin neurones. J Neuroendocrinol
2019;31:e12666. |
|