Specificity of TRH receptor coupling to G-proteins for regulation of ERG K⁺ channels in GH₃ rat anterior pituitary cells

Abstract

The identity of the G-protein coupling thyrotropin-releasing hormone (TRH) receptors to rat ether-à-go-go related gene (r-ERG) K⁺ channel modulation was studied in situ using perforated-patch clamped adenohypophysial GH₃ cells and dominant-negative variants (Gα-QL/DN) of G-protein α subunits. Expression of dominant-negative Gα_q/11 that minimizes the TRH-induced Ca²⁺ signal had no effect on r-ERG current inhibition elicited by the hormone. In contrast, the introduction of dominant-negative variants of Gα₁₃ and the small G-protein Rho caused a significant loss of the inhibitory effect of TRH on r-ERG. A strong reduction of this TRH effect was also obtained in cells expressing either dominant-negative Gα_s or transducin α subunits, an agent known to sequester free G-protein βγ dimers. As a further indication of specificity of the dominant-negative effects, only the dominant-negative variants of Gα₁₃ and Rho (but not Gα_s-QL/DN or Gα_t) were able to reduce the TRH-induced shifts of human ERG (HERG) activation voltage dependence in HEK293 cells permanently expressing HERG channels and TRH receptors. Our results demonstrate that whereas the TRH receptor uses a G_q/11 protein for transducing the Ca²⁺ signal during the initial response to TRH, this G-protein is not involved in the TRH-induced inhibition of endogenous r-ERG currents in pituitary cells. They also identify G_s (or a G_s-like protein) and G₁₃ as important contributors to the hormonal effect in these cells and suggest that βγ dimers released from these proteins may participate in modulation of ERG currents triggered by TRH.

Regulation of ether-à-go-go related gene (ERG) K⁺ channel activity by the hypothalamic neuropeptide thyrotropin-releasing hormone (TRH) constitutes an essential point of hormonal control of electrical activity, and hence of intracellular Ca²⁺ levels ([Ca²⁺]_i) and the secretory response in anterior pituitary cells (Barros et al. 1994, 1997; Weinsberg et al. 1997; Bauer, 1998; Bauer et al. 1998, 1999). The human ERG (HERG) channel has been also recognized as an important determinant of action potential characteristics in heart muscle and its inhibition by inherited mutations or a panoply of cardiac- and noncardiac-related prescribed drugs has been associated with an increased risk of cardiac arrhythmia and sudden death (Chiang & Roden, 2000; Keating & Sanguinetti, 2001; Redfern et al. 2003; Finlayson et al. 2004). Interestingly, the observation that arrhythmogenic syncopes are usually associated with physical, emotional or auditory stress suggests a link between hormonal (e.g. adrenergic) stimulation and cardiac ion channel function including HERG (Thomas et al. 2004). Furthermore, ERG channels also seem to play a key role setting the electrical behaviour of other cell types including neurones and glial, chromaffin, pancreatic β and tumour cells (Zhou et al. 1998a; Emmi et al. 2000; Rosati et al. 2000; Gullo et al. 2003; Sacco et al. 2003; Lastraioli et al. 2004). Nevertheless, unlike the relatively well known molecular and kinetic characteristics of ERG channels and in spite of their physiological and pathological relevance, many of their molecular mechanisms of regulation by different physiological agents remain unclear.

In native lactotrophs and clonal GH adenohypophysial cells, endogenous ERG currents are inhibited by activation of the G protein-coupled TRH receptor (TRH-R; Bauer et al. 1990, 1994; Barros et al. 1992, 1993; Schäfer et al. 1999; Schledermann et al. 2001). The TRH-R is coupled to a G-protein of the G_q/11 family (reviewed in Gershengorn & Osman, 1996) resulting in phospholipase C (PLC) activation and generation of myo-inositol 1,4,5-trisphosphate (IP₃) and diacylglycerol (DAG) from phosphatidylinositol 4,5-bisphosphate (PIP₂). It is also known that TRH is able to activate several PKC isozymes in GH₃ cells (Kiley et al. 1991; Akita et al. 1994). However, TRH-induced inhibition of ERG current in these cells does not depend on PKC or PKA activation (Bauer et al. 1990, 1994; Barros et al. 1992, 1993; Schäfer et al. 1999; Schledermann et al. 2001). Whether G_q/11 protein transduction, typically linked in many cells (including adenohypophysial cells) to Ca²⁺ signalling, plays a role in ERG channel regulation remains controversial. Thus, a pathway for ERG regulation by TRH involving a G₁₃- and Rho-mediating signalling cascade has been described in whole-cell voltage-clamped GH₄C₁ cells (Storey et al. 2002), but the relative importance of this transduction pathway as compared with the ‘classical’ G_q/11-mediated TRH-induced signalling was not assessed. Furthermore, G_q/11 but not G_i/o or G₁₃ has been recently shown to mediate muscarinic inhibition of ERG currents in tsA-201 cells coexpressing rat ERG1 channels and M1 muscarinic receptors (Hirdes et al. 2004). In adenohypophysial cells, the G_q/11-mediated coupling of TRH-R to PIP₂ hydrolysis leads to an initial elevation of [Ca²⁺]_i via IP₃ that mediates a peak of secretion associated with a transient hyperpolarization of the cell membrane due to activation of Ca²⁺-dependent K⁺ channels (Gómez-Varela et al. 2003b). However, normal inhibition of ERG channel activity in response to TRH has been observed in individual cells in which the initial Ca²⁺ response is totally absent (Barros et al. 1991, 1992, 1994). Although the PKC branch of the PLC signalling cascade is necessary for reversal of the TRH-induced ERG inhibition in GH₃ cells (Gómez-Varela et al. 2003b) and PIP₂ depletion could lead to HERG current reduction in HEK293 cells (Bian et al. 2001), the TRH-induced inhibition of the GH₃ cell r-ERG current also takes place after blockade of PIP₂ consumption with a PLC inhibitor (Gómez-Varela et al. 2003b). These results and the reported modification of the TRH effects on r-ERG in cholera toxin-treated GH₃ cells (Barros et al. 1994; Bauer et al. 1994) open the possibility that, at least in adenohypophysial cells, a transduction cascade(s) involving either a G_s-like protein or a G₁₃- and Rho-based pathway, couples the TRH-R to endogenous ERG channel inhibition.

In this report we use double mutants (Gα-QL/DN) of G-protein α subunits able to act as dominant-negative inhibitors against specific G-proteins (Yu et al. 2000) to explore the specificity of TRH-R coupling to G-proteins for ERG K⁺ channel inhibition in GH₃ rat anterior pituitary cells. Our results demonstrate that whereas the TRH-R certainly uses a G_q/11 protein for transducing the Ca²⁺ signal during the initial response to the hormone, this G-protein is not involved in the TRH-induced inhibition of ERG currents. Dominant-negative variants of Gα₁₃ and Rho, but not of Gα_q/11, are able to significantly reduce the inhibitory effect of TRH on ERG. Furthermore, a prominent reduction is observed upon introduction of dominant-negative Gα_s. Interestingly, a strong reduction of the TRH-induced inhibition is also observed in cells overexpressing transducin α subunits (Gα_t), an agent known to sequester free G-protein βγ dimers (Crespo et al. 1994; Faure et al. 1994; Palomero et al. 1998). The specificity of the dominant-negative and Gα_t effects is demonstrated by their failure to modify the Ca²⁺ response in the same cells. Furthermore, dominant-negative Gα_q/11 (but not Gα_t or dominant-negative Gα_s, Gα₁₃ and Rho) was able to reduce TRH-induced release of Ca²⁺ from intracellular stores in HEK293 cells permanently expressing HERG channels and TRH-Rs. In these cells, however, only the dominant-negative forms of Gα₁₃ and Rho (but not Gα_t or dominant-negative Gα_s) were able to antagonize the modifications in activation voltage dependence induced by TRH on HERG currents. On the other hand, only Gα_t and dominant-negative Gα_q/11 expression reduced the TRH-induced HERG current inibition at positive voltages. Apart from emphasizing the specificity of the different dominant-negative constructs, this also suggests that the cellular background and/or the channel isoform may influence the transduction mechanism(s) involved in hormonal regulation of ERG.

Methods

Plasmids and chemicals

The original plasmid containing the cDNA for the HERG channel was a generous gift of Dr E. Wanke (University of Milan, Italy). pEGFP-N3 plasmid was obtained from Clontech. pcDNA3.1 plasmids containing dominant-negative forms of Gα_q (Gα_q-Q209L/D277N), Gα₁₃ (Gα₁₃-Q266L/D294N), Gα_s (Gα_s-Q227L/D295N) and RhoA (RhoA-T19N 3xHA-tagged-NH2) were obtained from Guthrie (Guthrie cDNA Resource Center; currently transferred to University Missouri-Rolla cDNA Resource Center, Rolla, MO, USA). Gα_t was cloned in pcDNA3 as an EcoRI/XhoI fragment transferred from pcDNAI (provided by Dr J. S. Gutkind, N. I. of Dental Research, N.I.H., Bethesda, MD, USA). TRH and nystatin were purchased from Sigma. E-4031 and anti-HERG polyclonal antibodies were from Alomone Laboratories; Fura-2 and Fura-2/AM were from Molecular Probes.

GH₃ cell culture and transfection

GH₃ rat anterior pituitary cells (ATCC-CCL 82.1) were plated in 35-mm diameter tissue culture plastic dishes containing sterile glass coverslips coated with poly l-lysine and grown at 37°C in a humidified atmosphere of 95% air and 5% CO₂. The culture medium consisted of a 1: 1 mixture of Dulbecco's modified Eagle's medium and Ham's F-12 nutrient mixture (Sigma) supplemented with 100 U ml⁻¹ penicillin, 1.1 mg ml⁻¹ streptomycin and a serum mixture of 15% horse serum and 2.5% fetal bovine serum. The coverslips constituted the bottom of a small recording chamber (0.2–0.3 ml) that was continuously perfused with saline at a rate of about 1 ml min⁻¹. Cells trypsinized 24 h prior to transfection lying in poly-l-lysine-coated coverslips were transiently transfected using Lipofectamine 2000 according to the manufacturer's instructions (Invitrogen, Carlsbad, CA, USA). Unless otherwise indicated, 5.0 μg of plasmids containing the different constructs and pEGFP-N3 codifying green fluorescent protein (eGFP) as a marker for transfection in a 10: 1 ratio were used. The mixture of 5.5 μg of total DNA and Lipofectamine was incubated in serum-free medium for 20 min and added to the plates containing the cells in serum-containing medium without antibiotics. Recordings were performed 24–48 h after transfection.

Generation and isolation of permanently transfected HEK 293 cell clones

HERG channel cDNA was subcloned into HindIII/BamHI sites of the pcDNA3 vector (Invitrogen). Monolayer cultures (∼50% confluent) of human embryonic kidney cells (HEK293; ATCC CRL-1573) were transfected with this construct using Lipofectamine (Gibco). Three days after transfection, the cells were trypsinized and diluted in a medium containing 1 mg ml⁻¹ geneticin. Subsequently they were cultured until cell colonies were visible. Individual colonies were picked with cloning cylinders and tested for HERG currents. A clone named H36 was selected for further transfection. Cells of clone H36 were cotransfected with plasmid pcDNA3.1/Hygro(+) (Invitrogen) containing the cDNA for the TRH-R (de la Peña et al. 1992) inserted between the HindIII/XbaI sites of the vector. Hygromycin B (150 μg ml⁻¹) was used to select H36 clones coexpressing HERG channels and TRH-Rs. We chose for further work a clone named HEK-H36/T1 showing: (a) robust HERG currents under voltage-clamp, (b) reproducible calcium responses when perfused with TRH after loading the cells with the fluorescent Ca²⁺ indicator Fura-2, and (c) a predominant level of a 155 kDa band of HERG protein, corresponding to the mature and more glycosylated HERG likely to be located in the plasma membrane (Zhou et al. 1998b; Petrecca et al. 1999), immunodetected in cell extracts with HERG-specific antibodies. Cells were grown at 37°C in a humidified atmosphere of 95% air and 5% CO₂. HEK 293 cells were cultured in the same medium as GH₃ cells supplemented with 100 U ml⁻¹ penicillin, 0.1 mg ml⁻¹ streptomycin and 10% fetal bovine serum. HEK-H36/T1 cells were maintained in the presence of 1 mg ml⁻¹ geneticin sulphate and 150 μg ml⁻¹ hygromycin B (Gibco) and plated on the poly-l-lysine-coated coverslips for recording. Transient transfection of HEK-H36/T1 cells was performed following the procedures indicated above for the GH₃ cells.

Electrophysiological recordings, solutions and data analysis

Current recordings were performed at room temperature with the perforated-patch variant of the patch-clamp technique as previously described (Barros et al. 1991, 1992, 1994, 1997; Gómez-Varela et al. 2003b). Electrodes were fabricated from borosilicate or kimax disposable micropipettes (Boralex, Rochester Scientific, Rochester, NY; Fisherbrand, Fisher Scientific, Pittsburg, PA or Kimble glass Inc., Vineland, NJ, USA). Electrode resistance amounted 2–5 MΩ when filled with the pipette solution containing (mm): 65 KCl, 30 K₂SO₄, 10 NaCl, 1 MgCl₂, 50 sucrose and 10 Hepes (pH 7.4 with KOH). The tip of the pipette was initially filled with nystatin-free solution and the remainder of the pipette was back-filled with the same solution also containing 250 μg ml⁻¹ nystatin, added from a stock of 50 mg ml⁻¹ nystatin freshly dissolved in dimethylsulphoxide. These solutions were sonicated just before use. The course of perforation was followed by monitoring the progress of capacitive transients under voltage-clamp mode, setting the pipette voltage at a value of −70 mV. Access resistance, as estimated from the capacitive compensation circuitry on the amplifier, reached 10–30 MΩ within 5–20 min after the seal was made. Solution junction potentials were nulled before seal formation. Once patch permeabilization reached the indicated levels, the extracellular solution was changed as indicated and the cell was voltage-clamped at the desired holding potential. An EPC-7 patch-clamp amplifier (HEKA Elektronic, Lambrecht, Germany) was used to record membrane currents. Stimulation, data acquisition and analysis were carried out using Pulse and PulseFit software (HEKA Elektronic) running on Macintosh computers. Current records were sampled every 1 ms and digitally filtered at 500 Hz. r-ERG current data are shown without correction for leakage and capacitative transients. A P/n method was used for leak and capacitive current subtraction of the HERG recordings in HEK-H36/T1 cells. Further data processing was performed with PulseFit and Igor Pro (WaveMetrics, Lake Oswego, OR, USA).

The standard extracellular saline used for perforation and monitoring [Ca²⁺]_i contained (mm): 137 NaCl, 4 KCl, 1.8 CaCl₂, 1 MgCl₂, 10 glucose, and 10 Hepes (pH 7.4 with NaOH). Recordings of r-ERG currents in GH₃ cells were performed after changing the extracellular medium to high-K⁺, Ca²⁺-free solution once permeabilization of the patches had been completed. This solution contained (mm): 140 KCl, 4 MgCl₂, 10 EGTA and 10 Hepes titrated to pH 7.4 with KOH. Inward currents were studied during hyperpolarization pulses to −100 mV from a holding potential of −10 mV. The hyperpolarization pulses were preceded by a 100 ms ramp from 0 to −50 mV that can yield an estimation of the membrane conductance within this voltage range and would tend to potentiate the otherwise voltage-dependent effect of TRH (Bauer et al. 1990; Barros et al. 1992, 1994, 1997). To prevent variations due to differences in deactivation rates from cell to cell, the magnitude of the inward currents was estimated with the PulseFit software as the total inward charge computed between cursors located at 0.5 and 100% duration of the hyperpolarization pulses. HERG currents were recorded in HEK-H36/T1 cells in standard extracellular saline following the pulse protocols indicated on the figures. Kinetic parameters of activation and deactivation were obtained as previously described (Barros et al. 1998; Viloria et al. 2000). The voltage dependence of current activation was assessed using standard tail current analysis. Tail current magnitudes normalized to maximum were fitted with a Boltzmann function:

where V is the test potential, V_½ is the half-activation voltage, and k is the slope factor. Steady-state voltage dependencies of activation were obtained as previously described (Viloria et al. 2000) applying depolarization pulses of variable magnitude and up to 10 s duration from two holding potentials: +40 mV to hold the channels fully open and −80/−100 mV to hold them fully closed. The position of the Boltzmann curves under true steady-state conditions was estimated as an extrapolated mean from the curves obtained at both holding potentials to ensure that they were a function exclusively of depolarization pulse characteristics, regardless of the previous (open or closed) state of the channels. The time course of activation was monitored using an indirect envelope of tail current protocol (Viloria et al. 2000), varying the duration of the depolarization pulse and following the variation in the magnitude of the tail currents recorded after going back to a negative voltage. The rates of deactivation were determined from negative-amplitude biexponential fits to the decaying phase of tail currents. The first cursor of the fitting window was advanced to the end of the initial hook due to the recovery of inactivation.

Intracellular calcium measurements

Measurements of intracellular Ca²⁺ concentrations ([Ca²⁺]_i) were performed in cells platted in poly-l-lysine-coated coverslips as indicated above. In this case the coverslips were transferred to wells containing standard extracellular saline plus 5 μm Fura-2/AM (Molecular Probes) and loaded with the dye for about 60 min at room temperature. After loading with Fura-2, cells were washed with saline to remove non-hydrolysed Fura-2/AM and left for another 30 min before recording to facilitate AM hydrolysis by cellular esterases. Fluorescence measurements were performed in a Axiovert 100 microscope equipped with a Plan-NeoFluar 40×/0.75 objective and epifluorescence accessories (Carl Zeiss), attached to a fluorescence imaging system (TILL-Photonics GmbH, Martinsried, Germany). Control of the monochromator (Polychrome IV) and the 12-bit cooled CCD camera (IMAGO) was performed using TILLvisION imaging software. Cells were excited through a dichroic mirror reflecting less than 395 nm light. Fluorescence signals were filtered through a 410 nm long-pass filter. Cycles of sample excitation were repeated every 500 ms consisting in 10/20 ms periods of irradiation with 340, 360 and 380 nm light. The ratio of the emission intensities (340 nm/380 nm) was used as a measure for changes in intracellular Ca²⁺. When eGFP-transfected cells were used, a correction for eGFP fluorescence due to residual eGFP excitation at 340–380 nm was performed. Using eGFP-containing cells without Fura-2 we estimated previously that 13% of the fluorescence recovered above 510 nm (using a GFP filter set with a 500 nm dichroic and a 510 nm long-pass filter) following eGFP excitation at 488 nm is also present upon eGFP excitation at 380 nm using the conventional Fura-2 set-up. Less than 1% was recovered when eGFP was excited at 340 nm. Subsequently, the eGFP-derived fluorescence at 380 nm was estimated in every individual cell loaded with Fura-2 from the (13%) fluorescence intensity at 488 nm (a wavelength at which Fura-2 is not excited). This amount was subtracted from the total fluorescence at 380 nm to isolate the Fura-2-specific signal. Ca²⁺ concentrations were estimated from the 340 nm/380 nm fluorescence ratio by comparison with Fura-2 standards (Barros et al. 1994).

Statistics

Data values given in the text and in figures with error bars represent the mean ± s.e.m. for the number of indicated cells. Comparison between data groups was at first performed by parametric Student's unpaired t test (2-tailed) or ANOVA. Due to dispersion of the data in individual cells after some treatments (e.g. Gα₁₃-QL/DN and RhoA T/N transfections in Fig. 4), non-homogeneous variances (as evidenced after a Bartlett's test) were sometimes obtained. Therefore, alternate Welch's test assuming Gaussian populations with unequal s.d.s and a non-parametric Wilcoxon or Mann-Whitney test that does not make any assumption about the scatter of the data were also used to evaluate significance of mean differences between cell populations. For a posteriori comparison of two specific samples a Bonferroni or a Dunn test for multiple comparisons was also used following one-way ANOVA or Kruskal-Wallis non-parametric ANOVA tests, respectively. In all cases, P-values < 0.05 were considered as indicative of statistical significance.

Figure 4. Percentage inhibition of r-ERG currents induced by TRH in GH₃ cells expressing dominant-negative Gα subunits and Gα_t.

Current inhibitions were estimated from total inward charge as described in Fig. 3. Data from individual cells averaged on the bars are shown as open circles. Values from cells transfected with dominant negative variants of Gα_q, Gα₁₃, RhoA and Gα_s, or transducin α subunits (Gα_t) are shown. Data from cells transfected with vector pcDNA3B (Control 3B) and from untransfected cells (NO transf.) are also shown for comparison. Significant variations among population medians as evidenced by Mann-Whitney test or ANOVA and post hoc multiple comparison test are indicated.

Results

Validation of the dominant-negative strategy using Gα_q-QL/DN and the TRH-induced Ca²⁺ response of the GH₃ cells

To examine the transduction pathway leading to ERG inhibition upon TRH stimulation, we expressed xanthine nucleotide binding mutants of different G-protein α subunits. These mutants carrying a double mutation (leucine and asparagine substituting for a glutamine and an aspartate, respectively) possess a lowered affinity by guanine nucleotides and an enhanced affinity by xanthine nucleotides that make them form stable and specific complexes with cognate receptors and compete with endogenous wild-type G-proteins (Yu et al. 2000). We first probed the effectivity of this approach using dominant-negative Gα_q (Gα_q-Q209L/D277N) and exploring the initial (Phase 1) TRH-dependent response of the GH₃ cells. This phase corresponds to a transient release of stored Ca²⁺ into the cytosol due to production of IP₃ by PLC-catalysed PIP₂ hydrolysis, leading to an initial and transient hyperpolarization of the cell membrane by activation of Ca²⁺-dependent K⁺ channels (Gómez-Varela et al. 2003b). We reasoned that blockade of this transduction pathway with Gα_q-QL/DN should minimize all the cellular responses that characterize this phase such as (i) the transient membrane hyperpolarization, (ii) the transient increase in potassium currents that determine this hyperpolarization, and (iii) the transient [Ca²⁺]_i increase that activates the Ca²⁺-dependent K⁺ currents. As shown in Fig. 1A the transient hyperpolarization of the cell membrane induced by TRH in perforated-patch current-clamped GH₃ cells is nearly abolished in cells expressing Gα_q-QL/DN (see averaged voltage traces in the insets). Interestingly, as shown in the individual cell recordings illustrated in Fig. 1A, the increase in the rate of production of action potentials (Phase 2 of hormone action) that follows the initial hyperpolarization was maintained in the presence of the dominant-negative. As a second validation of the dominant-negative effect, we studied the appearance of membrane currents immediately after TRH addition in patch-perforated voltage-clamped GH₃ cells bathed in high-K⁺ Ca²⁺-free solution. Figure 1B shows that the current increases induced by TRH were absent in cells expressing Gα_q-QL/DN. Thus, these increases represent Gα_q-mediated PLC activation and activation of K⁺ currents by Ca²⁺ released from intracellular stores via IP₃, because they were determined without extracellular Ca²⁺. Furthermore, the effect of Gα_q-QL/DN was specific since the TRH-induced currents remained unaltered in cells expressing the dominant-negative α subunit of G₁₃, a G-protein predictably unrelated to the PLC–IP₃ Ca²⁺ signalling pathway.

Figure 1. Blockade by dominant-negative Gα_q-QL/DN of the initial phase 1 of response induced by TRH in GH₃ cells.

A, effect of dominant-negative Gα_q on TRH-induced Phase 1 of hyperpolarization. Representative recordings of membrane potential are shown in two cells either expressing the dominant-negative form of Gα_q (lower trace) or not (upper trace). Application of 1 μm TRH is marked with a horizontal line on top of the traces. Note the maintenance of Phase 2 of increased electrical activity in the Gα_q-QL/DN-transfected cell. Traces averaged point by point from several cells are shown in the insets. Continuous current traces averaged point by point and their corresponding s.e.m. are shown. In this cases, traces were synchronized to the time of TRH addition as indicated with an arrow. B, effect of dominant-negative Gα_q and Gα₁₃ on Ca²⁺-dependent K⁺ currents elicited in GH₃ cells by TRH during the initial Phase 1 of response. Continuous current traces averaged point by point and their corresponding s.e.m. are shown for cells transfected with vector pcDNA3B (Control 3B), or with plasmids codifying the dominant-negative mutants of Gα_q (Gα_q-QL/DN) and Gα₁₃ (Gα₁₃-QL/DN). Traces were synchronized to the time of 1 μm TRH addition as indicated with an arrow. High-K⁺, low-Ca²⁺ extracellular solution and a potential of −30 mV were used for better detection of inward K⁺ currents activated by Ca²⁺ released from intracellular stores via IP₃.

For an additional demonstration of the effectiveness and specificity of the dominant-negative approach we studied the effect of the Gα-QL/DN proteins on hormone-induced [Ca²⁺]_i increases. For this purpose, the fluorescence of Fura-2 was monitored in the transfected cells as indicated in Methods. As an internal control, TRH-induced variations in fluorescence of the cells present in the same microscope field but not expressing the transfection marker (eGFP) were monitored. We determined first that transfection or expression of the eGFP biosensor itself does not modify the [Ca²⁺]_i increases induced by the hormone, using GH₃ cells transfected with eGFP and pcDNA3B plasmid lacking any G-protein codifying insert. Data from a representative experiment indicate that whereas addition of 1 μm TRH raised [Ca²⁺]_i from a basal averaged value of 105 ± 32 nm (n = 35) to an initial maximum of 236 ± 32 nm in the 35 cells of a microscope field lacking eGFP fluorescence, the [Ca²⁺]_i level was increased from 80 ± 31 to 240 ± 29 nm in the 13 cells from the same field showing a clear eGFP expression. On the other hand, in both cases most of the cells showed a significant increase in peak [Ca²⁺]_i regardless of the presence or the absence of eGFP expression.

The aforementioned results strongly differ from those obtained with cells transfected with eGFP and a plasmid coding for Gα_q-QL/DN (Fig. 2A). In this case, TRH raised the basal [Ca²⁺]_i level of 73 ± 9 nm to 141 ± 14 nm in the 26 cells showing no detectable eGFP expression. In contrast, only a peak value of 61 ± 5 nm[Ca²⁺]_i from a basal level of 49 ± 6 nm was obtained upon TRH addition in the 11 cells of the same field showing a clear eGFP fluorescence. It is also important to note that in this case only 3 of the 11 cells expressing eGFP showed any significant increase in [Ca²⁺]_i as compared with the 18 in which [Ca²⁺]_i was clearly increased from 26 cells not expressing the transfection marker. Furthermore, whereas only a modest [Ca²⁺]_i increase from 61 ± 6 to 94 ± 4 nm was observed in those three cells expressing eGFP, a prominent initial peak of Ca²⁺ that raised [Ca²⁺]_i from 73 ± 9 to 172 ± 7 nm was obtained in the 18 cells in which eGFP (and hence Gα_q-QL/DN) expression was not detectable. Analogous results were obtained in two additional experiments. This demonstrates that (i) a much lower percentage of cells expressing the dominant-negative variant of Gα_q respond to TRH, and (ii) even in the few cells expressing Gα_q-QL/DN that show a detectable response to the hormone, the magnitude of the [Ca²⁺]_i increase is clearly smaller than that of cells in which the transfection marker (and hence the dominant-negative Gα_q) has not been expressed.

Figure 2. Effect of dominant-negative Gα subunits or Gα_t expression on Ca²⁺ liberation from GH₃ cell intracellular stores in response to TRH.

The time course of variations in [Ca²⁺]_i levels is shown for Fura-2 loaded cells from a microscope field transfected with Gα_q-QL/DN (A), Gα_s-QL/DN (B), Gα₁₃-QL/DN (C), or Gα_t (D). Addition of 1 μm TRH is indicated by horizontal lines on top of the traces. Continuous traces averaged point by point and their corresponding s.e.m. are shown. Averaged basal levels are signalled by dashed lines. Data correspond to cells showing fluorescence of the transfection marker (eGFP⁺, left) or not (eGFP⁻, right). Variations of [Ca²⁺]_i levels in the cell subpopulations showing a detectable peak Ca²⁺ increase after visual inspection are illustrated. In all cases the number of averaged cells with respect to the total number of cells present in the field is boxed. Data from the whole cell population present in the microscope field are shown in the insets of panel A. Averaged values from all data points before TRH addition and that of the initial maximum are indicated. Analogous results were obtained in two additional experiments.

Specificity of dominant-negative Gα subunits on TRH-induced Ca²⁺ response

The results presented above indicate that expression of the dominant-negative form of Gα_q constitutes an efficient way to suppress the Gq-dependent initial Ca²⁺ response to TRH of the GH₃ cells. Whereas coupling of endogenous and heterologously expressed TRH-Rs to Gq for activation of PLC-mediated PIP₂ hydrolysis has been widely documented, it has been also reported that in GH cells the TRH-R can interact with a number of different G-proteins that may include G₁₃, G_i and G_s or a G_s-like protein (Storey et al. 2002; reviewed by Gershengorn & Osman, 1996). To check the possible specificity of the dominant-negatives we also studied the Ca²⁺ response in cells expressing the QL/DN variants of Gα_s and Gα₁₃. The elevations of [Ca²⁺]_i in response to TRH remained the same in cells expressing either Gα_s-QL/DN (Fig. 2B) or Gα₁₃-QL/DN (Fig. 2C) as compared with cells from the same microscope field lacking eGFP expression. Similar results were obtained in cells expressing transducin α subunits (Fig. 2D; see also below). Furthermore, as in control cells transfected with pcDNA3B (see above) or in untransfected cells (not shown), most of the cells expressing dominant-negatives of Gα_s or Gα₁₃ showed significant increases in peak [Ca²⁺]_i. This indicates that whereas coupling of TRH-R to G_q/11 is indispensable for the TRH-evoked Ca²⁺ response, coupling to a G-protein of the G_s or G₁₃ type is not required for this effect.

Effect of different dominant-negative Gα subunits on TRH-induced inhibition of endogenous r-ERG currents in GH₃ cells

To investigate the transduction cascade linking the TRH-R to r-ERG channel inhibition we always used perforated-patch conditions to minimize cell dialysis and to preserve intact the intracellular components necessary for the hormonal response. To isolate the r-ERG current present in GH₃ cells and to quantify its inhibition by TRH, high-K⁺ low-Ca²⁺ extracellular solutions and established voltage protocols were used. Thus, due to the fast inactivation of ERG channels at depolarized potentials and the presence of several outwardly rectifying voltage- and calcium-dependent K⁺ currents in GH₃ cells, currents were studied during hyperpolarization pulses to −100 mV from a holding potential of −10 mV (see Methods). High-K⁺ low-Ca²⁺ extracellular solutions were also used to increase the amplitude of the inwardly rectifying r-ERG currents and to reduce Ca²⁺ currents and activation of Ca²⁺-dependent K⁺ currents (Bauer et al. 1990, 1999; Barros et al. 1992, 1997; Weinsberg et al. 1997). Furthermore, since under these conditions the TRH-induced current inhibition is almost exclusively exerted on ERG currents, 5 μm of the ERG-specific blocker E-4031 (a concentration that totally blocks the r-ERG current) was added at the end of the experiments to subtract the E-4031-insensitive currents from the initial ones and compare the difference between the TRH- and E-4031-blocked currents in every individual cell (see Gómez-Varela et al. 2003b). Representative examples of TRH effects on r-ERG currents in cells transfected with different G-protein α subunit variants are shown in Fig. 3. Only successfully expressing cells identified by their eGFP fluorescence were used for recording. As shown in Figs 3A and 4, the inhibitory effect of TRH on the E-4031-sensitive current was not modified by the transfection procedure. Thus, a value of 76.8 ± 2.7% (n = 20) was obtained in cells transfected with eGFP and pcDNA3B plasmid lacking any G-protein coding insert, as compared with the 78.6 ± 5.4% (n = 8) inhibition obtained in cells showing no detectable eGFP expression. Similar inhibitions have been previously reported under identical conditions using untransfected cells (Gómez-Varela et al. 2003b). Most importantly, the TRH-induced inhibition was the same in cells expressing dominant-negative Gα_q-QL/DN (74.8 ± 6.3%, n = 6). It is important to note that failure to significantly modify the TRH-induced inhibition of r-ERG was not due to lack of efficient expression of Gα_q-QL/DN, since the early increases in Ca²⁺-dependent K⁺ currents induced by TRH (a Gq-dependent and PLC/IP₃/Ca²⁺-related effect, see above) were absent in the same cells (Fig. 3B). Apart from adding further support to specificity of the Gα_q-QL/DN for the Ca²⁺ response, this indicates that whereas the TRH receptor certainly uses a G_q/11 protein for transduction of the Ca²⁺ signal during the initial response to the hormone, this G-protein is not involved in the TRH-induced inhibition of endogenous r-ERG currents in GH₃ cells.

Figure 3. Effect of dominant-negative Gα subunits and Gα_t on TRH-induced inhibition of endogenous r-ERG currents in GH₃ cells.

The time course of relative r-ERG current reduction by TRH and E-4031 is shown for six different cells transfected with vector pcDNA3B (A), or with plasmids encoding the dominant-negative mutants of Gα_q (B), Gα₁₃ (C), RhoA (D) and Gα_s (E), or the α subunit of transducin (F). Currents were recorded during 1 s hyperpolarizing pulses to −100 mV from a holding potential of −10 mV. The hyperpolarization step was preceded by a 100 ms ramp from 0 to −50 mV (Gómez-Varela et al. 2003b). Current estimations were performed from total inward charge during the hyperpolarization steps at −100 mV as described in Methods. Averaged charge values before any addition and those corresponding to the minimum following addition of 5 μm E-4031 were considered as 0 and 100%, respectively. Filled circles correspond to the current traces shown above. The first one or two data points following addition of TRH, when total inward currents become transiently enhanced by activation of Ca²⁺-dependent K⁺ channels due to massive liberation of Ca²⁺ from intracellular stores, have been deleted for clarity. Perfusion of 1 μm TRH and addition of E-4031 to the recording chamber are indicated. Values for the data in the absence and presence of TRH are indicated by horizontal dashed lines in the time courses. A continuous membrane current recording at a holding potential of −30 mV around the time of hormone addition is shown in B and marked with a thick arrow. Note the total absence of transient Ca²⁺-dependent K⁺ currents following TRH addition in the same cell showing a clear reduction of r-ERG currents. Similar results were obtained in the five additional cells expressing Gα_q-QL/DN.

Recent experiments in GH₄C₁ cells using a constitutively active mutant of Gα₁₃ and whole-cell recording showed a partial inhibition of r-ERG currents equivalent to that induced by TRH under similar conditions (Storey et al. 2002). Using our perforated-patch conditions, the inhibition of r-ERG by TRH was attenuated when the G₁₃ pathway was antagonized with dominant-negative Gα₁₃. Thus, an inhibition of 56.4 ± 6.9% (n = 12, P < 0.01 versus control pcDNA3B-transfected cells, Welch and Mann-Whitney tests) in the E-4031-sensitive r-ERG currents was induced by TRH in cells transfected with 1.6 μg of plasmid encoding Gα₁₃-QL/DN (4: 1 ratio versus EGFP plasmid; not shown). This value was slightly decreased to 43.8 ± 6.4 (n = 15) when 5.0 μg of the same plasmid were used (Figs 3C and 4). This effect was specific, as demonstrated by the total absence of Gα₁₃-QL/DN influence on TRH-induced Ca²⁺ responses (see above). Interestingly, the magnitude of the TRH-induced inhibition in individual cells showed a huge dispersion at both DNA concentrations, which made it appear that the 27 cell sample was composed of two subpopulations. Whereas nearly half of the cells showed TRH-induced r-ERG inhibitions above 50% (equivalent to those of controls and untransfected cells), a clear reduction of the hormonal effects leaving the TRH-induced inhibition below 50% took place in the other half of the cell population (see Fig. 4).

It has been shown that the activation of G₁₃- (and G_q-) dependent pathways is able to signal to different effectors through the small G-protein RhoA (Seasholtz et al. 1999; Sah et al. 2000; Dutt et al. 2002; Vogt et al. 2003). Expression of the dominant-negative RhoA-T19N also significantly attenuated r-ERG modulation by TRH, leaving a mean inhibition of 53.4 ± 6.3% (n = 11, P < 0.01 versus control, Welch and Mann-Whitney tests; Figs 3D and 4). Again, a considerable dispersion of the data was obtained in this case. Nevertheless, these data confirm previous results in GH₄C₁ cells and suggest that a transduction cascade involving G₁₃ (but not G_q) and Rho may participate in the inhibition of ERG channels by TRH.

Previous results in GH₃ cells showed a modification of the TRH-induced effects on r-ERG in cells treated with the G_s-modifying agent cholera toxin (Barros et al. 1994; Bauer et al. 1994). Although a coupling of TRH-R to G_s with subsequent activation of adenylyl cyclase has been reported in GH₃ cells, evidence against (i) stimulation of the enzyme by TRH and (ii) specific down-regulation of Gα_s following long-term expositions of GH₃ cells to TRH has been obtained also (reviewed in Gershengorn & Osman, 1996). Furthermore, a cholera toxin-dependent degradation of Gα_s has been demonstrated in GH₃ cells (Chang & Bourne, 1989), but it is also known that cholera toxin treatment can cause a significant reduction in the number of TRH-Rs (Yajima et al. 1988). This prompted us to check whether antagonization of the Gs pathway with dominant-negative Gα_s could cause any effect on the TRH-induced r-ERG current reductions. As shown in Figs 3E and 4, a prominent reduction of the inhibitory effect of TRH on r-ERG was observed in the presence of dominant-negative Gα_s-QL/DN. In this case, mean inhibition amounted only to 33 ± 3.9% (n = 10, P < 0.0001 versus control, Student's t and Mann-Whitney tests). This value is also significantly smaller than that obtained in the presence of RhoA-T19N (P < 0.02, Mann-Whitney test). It is important to note that in the presence of dominant-negative Gα_s not only is the reduction of the TRH-induced inhibition the biggest observed, but also the dispersion of the data is notably reduced (Fig. 4). This supports the conclusion that G_s plays a crucial role on the inhibitory effects of TRH in GH₃ cells.

Effect of transducin α subunit expression on the TRH-induced inhibition of r-ERG in GH₃ cells

It has been recognized that receptors that activate G₁₃ also couple to G_q and G₁₁. It is also well known that Gα₁₃ and Gα_q/11 signals induce Rho activation and subsequent cellular responses by inducing GDP–GTP exchange in Rho guanine nucleotide exchange factors in response to extracellular stimuli (Seasholtz et al. 1999; Sah et al. 2000; Dutt et al. 2002; Vogt et al. 2003). Thus it seemed surprising that dominant-negative G_s (but not G_q), G₁₃ and Rho share antagonism on the TRH-induced effects on r-ERG. To check the possibility that reduction of TRH-induced inhibition involves a component common to G_s and G₁₃ but not available in heterotrimeric G_q, we studied the consequences of expressing transducin α subunits (Gα_t), a scavenger of βγ dimers after they are released from G-proteins by receptor stimulation (Crespo et al. 1994; Faure et al. 1994; Palomero et al. 1998). The inhibition of r-ERG by TRH was strongly attenuated by Gα_t (33.5 ± 4.4%; n = 14, P < 0.0001 versus control, Student's t and Mann-Whitney tests; Figs 3F and 4) up to levels equivalent to those observed with dominant-negative Gα_s. This effect of Gα_t was totally specific, since both the percentage of cells showing a detectable increase in peak [Ca²⁺]_i and the magnitude of the TRH-induced Ca²⁺ response remained the same regardless of the presence or the absence of Gα_t (Fig. 2D). As discussed below, this suggests that free βγ subunits released from G_s (and perhaps shared by G₁₃) heterotrimers may be responsible for the TRH-induced inhibition of endogenous r-ERG currents in GH₃ cells.

Generation of a HEK293 cell clone permanently expressing HERG channels and TRH receptors and characterization of the TRH response

Unlike the strong and quite reproducible current inhibitions induced by TRH on the endogenous ERG current, only modest effects of the hormone have been reported on other kinetic characteristics of ERG either endogenous or heterologously expressed in GH₃ cells (Barros et al. 1992; Bauer et al. 1998; Schledermann et al. 2001). This fact and the sometimes variable effect of dominant negatives in the individual cells prevented us from reaching any consistent conclusion about the way in which different G proteins could be affecting other current parameters in these cells.

HEK293 cells permanently expressing HERG constitute an interesting and widely used model system to study easily the biochemical and electrophysiological properties of the channel (Zhou et al. 1998b). For this reason, we initially isolated several cell clones expressing HERG. Subsequently, they were cotransfected with a plasmid containing the TRH-R cDNA (de la Peña et al. 1992). Single colonies were selected and tested for both HERG current under voltage-clamp and TRH-induced Ca²⁺ responses with the fluorescent Ca²⁺ indicator Fura-2 (see Methods). A cell line named HEK-H36/T1 showing high HERG current density and also systematic Ca²⁺ increases when exposed to TRH was used for the experiments reported here. As shown in Fig. 7, the HEK-H36/T1 cells showed voltage-activated K⁺ currents typical of HERG under the perforated-patch conditions chosen to maintain as intact as possible the hormonal responses. Apart from HERG currents, we frequently detected also other outward K⁺ currents during the depolarization steps. The magnitude and the degree of inactivation of these currents during the depolarization pulses were highly variable. Nevertheless, their contribution was negligible along the tail current time course, as evidenced by their absence after treating the cells with the specific HERG inhibitor E-4031 (Fig. 5A). Thus only HERG current characteristics would be referred to by limiting the analysis to tail currents. Addition of TRH to HEK-H36/T1 cells loaded with Fura-2 triggered a transient increase in cytoplasmic Ca²⁺ that slowly declined to basal values upon hormone washout. On the average, the amplitude of this Ca²⁺ response was equivalent to that elicited in the same cells by carbachol (data not shown) probably acting through the endogenous muscarinic receptors of HEK293 cells. The TRH-induced Ca²⁺ responses were absent in the cell clones containing HERG channels but not coexpressing TRH-Rs.

Figure 7. Effect of dominant-negative Gα subunits and Gα_t expression on Ca²⁺ liberation from HEK-H36/T1 cell intracellular stores in response to TRH.

Protocols and data evaluation were performed as described in Fig. 2. Only data from the cell subpopulations showing a visually detectable peak Ca²⁺ increase are plotted. In all cases the number of averaged cells respect to the total number of cells present in the field is boxed. Analogous results were obtained in two additional experiments.

Figure 5. Effect of TRH on HERG current activation in HEK-H36/T1 cells.

A, effect of TRH on voltage dependence of HERG current availability. The voltage dependence of activation was studied in the absence (Control) or the presence of 100 nm TRH (TRH) by varying the amplitude of a 1 s prepulse from a holding potential of −80 mV, and measuring the magnitude of the peak tail current at a constant repolarizing voltage of −50 mV. Membrane currents recorded in the presence of TRH plus 1 μm E-4031 (E-4031) are also shown for comparison. Superimposed current traces for depolarizing steps between −50 and +40 mV (Control) or −40 and +60 mV (TRH and E-4031) are shown on the left. Control and TRH traces obtained after subtracting the E-4031-resistant currents are also shown on the right. Pulses were applied once every 20 s. Current traces have been subtracted for leak. Recording of currents in the presence of TRH started 2 min after introducing the neuropeptide in the recording chamber. Averaged fractional activation curves obtained before and after addition of TRH are shown at the bottom. Normalized outward tail current magnitudes at the peak are plotted versus test pulse potential (E_m). Data normalized to maximum of tail current magnitude without TRH are presented. The continuous lines correspond to Boltzmann curves h(V) =I_max(1/(1 + exp(V−V_½)/k)), that best fitted the data. The relative position of the V_½ values and their numeric magnitude are indicated in the graph. B, effect of TRH on HERG current activation rate. The time course of voltage-dependent activation was studied by varying the duration of a depolarizing prepulse and measuring the magnitude of the peak tail current at a constant repolarizing voltage of −50 mV. Families of currents at a depolarizing potential of +40 mV are shown for a cell before (Control) and after adding 100 nm TRH (TRH). Current traces corresponding to depolarization steps of 0, 20, 40, 80, 160, 320, 640, 1280, 2560 and 5120 ms are shown superimposed. The dependence of activation rates on depolarization membrane potential is shown at the bottom. The magnitude of the peak tail current upon repolarization was determined from recordings as shown at the top. The time necessary to attain half-maximum current magnitude at different depolarization potentials is plotted in the absence or the presence of TRH.

The simultaneous presence of TRH-R and HERG channels in HEK-H36/T1 cells prompted us to check the effect of the hormone on HERG currents in this putatively simple and defined cellular background. TRH reduced tail HERG currents within 1–3 min after introducing the hormone in the recording chamber (Fig. 5A). The current level was lowered by 36 ± 2% (n = 38) when measured at the peak of the tails that followed pulses of 1 s duration at +40 mV. Furthermore, a strong shift in current availability voltage dependence to more positive voltages was also caused by TRH. Thus the V_½ value of the Boltzmann functions describing the I–V curves was shifted from −1.5 ± 1 to +25.5 ± 2 mV (n = 14) under these conditions (Fig. 5A). It is important to note that the diminished tail currents remaining after treatment with TRH were entirely due to the operation of HERG channels, since they were abolished by E-4031.

Due to the slow activation and deactivation kinetics of HERG channels, no steady-state conditions would be expected within 1 s depolarizations. A possible cause of the shift in activation voltage dependence could be a displacement of the V_½ values to more depolarized potentials due to a slower activation rate leading to a less steady-state condition (Schönherr et al. 1999; Viloria et al. 2000). In fact, it is known that the activation rate of HERG channels in Xenopus oocytes is slowed by activation of coexpressed TRH-Rs (Barros et al. 1998). For this reason, we also tested the effects of TRH on HERG activation rates in HEK-H36/T1 cells using an indirect envelope of tail currents protocol (Barros et al. 1998; Viloria et al. 2000). As shown in Fig. 5B, the time required to attain a half-maximum current magnitude was increased by TRH near an order of magnitude between +40 and +60 mV. Thus the difference in the inflection potentials of the I–V curves could be due, at least in part, to the different activation rates around the V_½ values. Nevertheless, an 18 mV shift from −22.5 ± 2.2 to −4.1 ± 2.8 mV (n = 6) was also detected in response to TRH when the I–V curves were generated from tail currents following long depolarization steps of 10 s duration (Fig. 6A). Finally, the position of the curves under real steady-state was extrapolated as the mean of those obtained from hyperpolarized (−80 mV) and depolarized (+40 mV) holding potentials (Schönherr et al. 1999; Viloria et al. 2000). In this case a 15 mV shift in V_½ (from −25.0 to −10.0 mV, n = 2) was obtained (Fig. 6B). Altogether, this suggests that two distinct effects on activation contribute to the shifts when studied with 1 s short depolarization steps: a genuine shift of the voltage dependence of activation and a marked slowing of activation rates moving the channels further away from steady-state conditions.

Figure 6. Effect of TRH on HERG activation voltage dependence under steady-state conditions.

A, effect of TRH on activation voltage dependence studied using long depolarization steps of 10 s. Tail currents upon repolarization to −50 mV were obtained following 10 s depolarizations to different voltages as indicated at the top. Only the end of the depolarizing steps and the tail currents at −50 mV are shown for clarity. Traces correspond to a cell before (Control) and 3 min after introduction of 100 nm TRH (TRH). Averaged I–V curves before and after the hormonal treatments are shown at the bottom. The magnitude of the peak tail current upon repolarization was determined from recordings as shown at the top and normalized to maximum of tail current magnitude without TRH. B, effect of TRH on HERG steady-state voltage dependence of activation. Steady-state voltage dependence of activation was studied following the protocols shown in the two upper panels, with a prepulse of varying magnitude and a duration ranging from 1 to 10 s, followed by a pulse test to −50 mV. Holding potentials of +40 mV to keep the channels fully open and −80 mV to hold them fully closed were used as indicated. Families of currents during the test pulse for Control and TRH-treated cells following 10 s prepulses to different potentials are shown. Fractional activation curves before and after TRH treatment are shown at the bottom. Open and filled symbols correspond to data obtained from −80 and +40 mV holding voltages, respectively. Data obtained at prepulse duration of 1 s (circles), 5 s (squares), and 10 s (triangles) are plotted. In every curve current values were normalized to tail current magnitude in response to the more depolarized potential of the prepulse series. The dashed lines represent the deduced position of the activation curves under steady-state conditions obtained as a mean of those corresponding to both holding voltages and prepulses of 10 s duration. The V_½ values for the steady-state curves are indicated by arrows in the graphs.

The deactivation properties of HERG can be obtained from the time course of current decay during voltage steps to negative potentials, following depolarizing pulses at a fixed time and voltage designed to activate (and inactivate) the channels. In this case, the closing time constants can be estimated by fitting a bi-exponential function to the decaying tail current phase that follows the initial peak once channel inactivation has been removed by the repolarization. Both the fast and the slow exponential components of current deactivation were significantly accelerated by TRH at all voltages between −40 and −120 mV. As an example, the values of deactivation time constants at −50 mV for the fast and the slow components of closing were reduced by the hormone from 213 ± 22 to 119 ± 11 ms (n = 5, P < 0.01) and from 965 ± 135 to 579 ± 77 ms (P < 0.05, Student's t test), respectively.

The clear effects of TRH on HERG opening and closing contrast with the almost complete absence of hormone-induced modifications on inactivation properties. Thus only a slight but non-significant acceleration of inactivation rates was observed upon addition of TRH. Furthermore, both control and TRH-treated cells showed identical rates of inactivation recovery when the time course of the initial current increase in response to hyperpolarizing pulses was compared (not shown).

Effect of dominant-negative Gα subunits and Gα_t on TRH-induced Ca²⁺ responses and modifications of HERG I–V curves in HEK-H36/T1 cells

The influence of the dominant-negatives and Gα_t expression on the hormonal effects in HEK-H36/T1 cells was studied following procedures analogous to those described for endogenous channels in adenohypophysial GH₃ cells. Again, dominant-negative Gα_q, but not Gα_t or dominant-negative Gα_s, Gα₁₃ and RhoA, reduced TRH-induced increases of cytoplasmic Ca²⁺ in the HEK-H36/T1 cells (Fig. 7). The modulation of HERG channels by TRH in HEK-H36/T1 cells was not changed by the transfection procedure used for dominant-negative expression (Control in Fig. 8). Surprisingly, the TRH-induced HERG current inhibition at positive voltages was not significantly altered in the presence of dominant-negative Gα₁₃, RhoA or Gα_s, but was strongly reduced by Gα_q-QL/DN or Gα_t expression (Fig. 8A and C). On the other hand, the positive shifts in activation voltage dependence were reduced by RhoA-TN and nearly abolished by Gα₁₃-QL/DN (Fig. 8A and B). Although the shift in the I–V curves was almost absent in Gα_q-QL/DN-transfected cells, the basal position of the curves before adding hormone was clearly displaced to positive voltages in the cells expressing the dominant-negative form of Gα_q (Fig. 8A). This complicates the interpretation of the dominant-negative Gα_q influence on the hormonal effect in the I–V relation. Finally, expression of the βγ dimer scavenger Gα_t left unaltered the TRH-induced voltage dependence shift in 10 of 14 tested cells, but blocked completely the reductions in maximal tail current magnitude triggered by the hormone. These results not only indicate a variation of the dominant-negative effects in different cellular and/or channel background, but also that these effects are differently manifested as a function of the channel parameter being considered.

Figure 8. Effect of dominant-negative Gα subunits and Gα_t on TRH-induced modifications of HERG I–V curves in HEK-H36/T1 cells.

A, effect of Gα subunit expression on HERG activation voltage dependence. Voltage dependence of activation was studied in the absence (open symbols) or the presence (filled symbols) of 1 μm TRH by varying the magnitude of a 1 s depolarizing pulse as detailed in the legend of Fig. 5A. Data normalized to maximum of tail current magnitude without TRH are shown. V_½ values obtained from the Boltzmann curves that best fitted the averaged data (continuous lines) and the number of recorded cells are indicated in the graphs. B, comparison of TRH-induced shifts on activation voltage dependence in cells expressing different Gα subunits. Data from the same cells used to generate the averaged I–V curves shown in panel A were used. Values of TRH-induced shifts from the different individual cells were averaged and are shown in the histogram. C, effect of Gα subunit expression on TRH-induced inhibition of HERG currents at positive voltages. Data from the same cells used to generate the averaged I–V curves shown in panel A were used. Values of TRH-induced reductions in maximal currents from the different individual cells were averaged and are shown in the histogram. Statistically significant variations versus controls in Student's t or Mann-Whitney tests are indicated in B and C.

Discussion

In this report we used a dominant-negative strategy to explore the specificity of TRH-R coupling to G-proteins for inhibition of the native r-ERG K⁺ channels present in GH₃ rat anterior pituitary cells. Based in the reported insensibility of the TRH responses to pertussis toxin treatment (Bauer et al. 1994) we focused our efforts in the G_q, G₁₃ and G_s families of G-proteins. Using the same approach we also compared the results with those obtained in an heterologous expression system, namely a HEK293 cell line (HEK-H36/T1) permanently expressing HERG channels and TRH-Rs. In both cases, xanthine nucleotide-binding (GαX) double mutants (Gα-QL/DN) of G-protein α subunits able to act as dominant-negative inhibitors against specific G-proteins (Yu et al. 2000) were used. Our results demonstrate that transduction of the Ca²⁺ signal during the GH₃ cell initial response to TRH is potently antagonized by dominant-negative Gα_q/11 and that this Ca²⁺ response remains unaltered in the presence of the dominant-negative variants of Gα₁₃, RhoA and Gα_s. On the other hand, dominant-negative variants of Gα₁₃ and Rho, but not of Gα_q/11, are able to significantly reduce the inhibitory effect of TRH on ERG. A more prominent reduction of the TRH-induced ERG inhibition is observed upon introduction of dominant-negative Gα_s. These results indicate that the TRH receptor certainly couples to a G_q/11 protein for transduction of the Ca²⁺ signal, but that this G-protein is not involved in the TRH-induced inhibition of ERG currents. This is coherent with previous results indicating that appearance of Phase 2 of increased electrical activity in GH₃ cells takes place in the presence or the absence of a detectable initial Ca²⁺ response (Phase 1; see Barros et al. 1994; Bauer et al. 1994). Our data also indicate that (i) failure to detect any influence of the Gα₁₃, Rho and Gα_s dominant-negatives on the Ca²⁺ signal is not due to lack of their functional expression, (ii) coupling of the TRH-R to one (or more) heterotrimeric G protein(s) carrying Gα_s and/or Gα₁₃ subunits takes place in the GH₃ cells, (iii) Gα_s and perhaps a Gα₁₃- and Rho-dependent pathway can participate in the transduction cascade linking TRH-R activation to r-ERG modulation in GH₃ cells, and (iv) there is a clear specificity in the ability of dominant-negatives to antagonize different physiological responses triggered by TRH in these cells.

In principle, the observed specificity of the dominant-negatives seems rather surprising according to the proposed mechanism for negative dominance in the Gα-QL/DN mutants. Introduction of the QL/DN mutations shifts nucleotide binding specificity from guanine nucleotides to xanthine nucleotides. Because the low concentrations of xanthine nucleotides in vivo, essentially nucleotide-free Gα-QL/DN proteins would exist in cells, and these would form stable complexes with cognate receptors and inhibit them by competing with endogenous wild-type G proteins (Yu et al. 2000). Accordingly, it could be expected that all signalling pathways lying on a given receptor became inhibited by any Gα-QL/DN variant interacting with it. Our results demonstrate that this is not the case for the TRH-R. The presence of a heterogeneous population of binding sites for TRH in GH₃ cells has been previously reported (Gautvik & Lystad, 1981) and two TRH-R isoforms derived from the same gene by differential splicing mechanisms have been shown to be expressed in GH₃ cells (de la Peña et al. 1992). However, no significant differences in binding or signalling properties of these receptors seem to exist (de la Peña et al. 1992; Lee et al. 1995; Gershengorn & Osman, 1996). Furthermore, the presence of two different molecular species of TRH-R could not explain the dominant-negative specificity found in HEK-H36/T1 cells in which only an isoform of the receptor is expressed. Clearly, further work will be necessary to understand the reason(s) for the observed specificity of the dominant-negative effects.

Regardless of the exact mechanism by which the negative dominance takes place, the demonstration of the functionality and specificity of the Gα-QL/DN variants allowed us to explore the TRH-R coupling to defined G proteins for transducing the inhibitory signal to ERG channels. Our results suggest that Gα_s plays a crucial role transducing the TRH signal to native r-ERG channels in GH₃ cells, since dominant-negative Gα_s causes the greatest reduction of the hormonal effect as well as the smallest data dispersion in individual cells. These data are consistent with previous results showing that the TRH-induced r-ERG inhibition is enhanced by short-term treatment with cholera toxin (an agent able to specifically modify Gα_s functionality) and nearly abolished when the treatment with the toxin is prolonged (Barros et al. 1993, 1994; Bauer et al. 1994). They will be also coherent with the reported inhibition of TRH-induced PLC stimulation in Xenopus oocytes using nucleotides antisense to Gα_s (de la Peña et al. 1995; but see Gershengorn & Osman, 1996). The reasons why coupling of TRH-R to G_s in GH₃ cells only modestly stimulates adenylyl cyclase (Paulssen et al. 1992; Gershengorn & Osman, 1996) and why prolonged stimulation with TRH down-regulates G_q/11 but has no effect on the Gα_s protein levels (Kim et al. 1994) remain to be established. As suggested previously (Barros et al. 1993, 1994; Bauer et al. 1994), it is possible that a G_s-like protein that is not G_s itself, but sensitive to cholera toxin and blocked by Gα_s-QL/DN expression, is involved in GH₃ cell r-ERG inhibition by TRH.

Using the dominant-negative approach we also studied the possible implication of Gα₁₃ and RhoA, two entities recently proposed as mediators of r-ERG modulation by TRH (Storey et al. 2002). In this case, expression of dominant-negative Gα₁₃-QL/DN significantly reduced the TRH-induced inhibition, although a slightly smaller effect was observed than that with Gα_s-QL/DN and only in around 50% of the recorded cells was the hormonal effect clearly antagonized. It is unlikely that this situation is caused by an unspecific inhibition of G_s by dominant-negative Gα₁₃ because smaller TRH-induced inhibitions and wider dispersion of the data were also observed with dominant-negative RhoA, a component located downstream of G₁₃ in many cellular systems and (supposedly) not directly related to G_s. On the other hand, it seems difficult to conceive a transduction mechanism involving Gα_s, Gα₁₃ and RhoA (but not Gα_q) either simultaneously or indistinctly. Our data following the overexpression of Gα_t, an agent known to sequester free G-protein βγ dimers (Crespo et al. 1994; Faure et al. 1994; Palomero et al. 1998), offer a possible explanation of this apparent paradox. Thus, prominent reductions of the TRH-induced r-ERG inhibition equivalent to those obtained with dominant-negative Gα_s are observed in Gα_t-transfected GH₃ cells. The specificity of the Gα_t effect was demonstrated by its failure to modify the Ca²⁺ response in the same cells. It is tempting to speculate that a specific set of free βγ subunits released from G_s heterotrimers (or a similar combination present in G₁₃ but not in G_q) may be responsible for the TRH-induced inhibition of endogenous r-ERG currents in GH₃ cells. Interestingly, both G_α and G_βγ subunits have been implicated in activation of Rho (Niu et al. 2003). Furthermore, GH₃ cells constitute perhaps the best characterized example in which specific hormone receptors have been shown to use G protein heterotrimers of different αβγ subunit composition to modulate an ionic channel (reviewed in Robishaw & Berlot, 2004).

Comparison of TRH and dominant-negative effects in native GH₃ cells and a heterologous expression system such as the HEK-H36/T1 cells reveals some quantitative and qualitative differences. These include (i) considerably smaller TRH-induced current inhibitions in HEK-H36/T1 cells, (ii) a clear inability of dominant-negative Gα_s to alter TRH-induced modifications of both activation voltage dependence and HERG current magnitude in HEK-H36/T1 cells, (iii) a significant reduction of the I–V curve shifts induced by TRH in HEK-H36/T1 cells upon expression of Gα₁₃-QL/DN and RhoA-T/N without any concomitant alteration of current inhibition, and (iv) the maintenance of a TRH-induced positive shift of the I–V curves in Gα_t-transfected HEK-H36/T1 cells in which the presence of the βγ scavenger abolishes the HERG current inhibition induced by the hormone. Use of perforated-patch conditions excludes uncontrolled alterations of the intracellular environment as a cause for these differences. Instead, our results point to differences in the cellular background and/or the molecular identity of the channel protein as determinant of them. The parallel effects of TRH on ERG channels either endogenous or overexpressed in GH₃/B₆ cells (Schledermann et al. 2001) suggest that the cellular background may influence the transduction mechanism(s) involved in hormonal regulation of the channels. This is further supported by recent results showing that G_q/11, but not G_i/o or G₁₃, mediates hormonal inhibition of ERG currents in tsA-201 cells coexpressing rat ERG1 channels and another G_q/11-coupled receptor, the M1 muscarinic receptor (Hirdes et al. 2004). Some differences between the signal cascade mediating the TRH-induced modulation of HERG in Xenopus oocytes and the physiological pathway modulating the native ERG channels in GH₃ cells have been also reported (Barros et al. 1998). It is important to highlight that the dominant-negative effects reported here may vary also as a function of the kinetic parameter being considered. Thus whereas only an attenuation of the TRH-induced I–V shifts is observed in HEK-H36/T1 cells in which the G₁₃ pathway is blocked with dominant-negative G₁₃ or RhoA, only the hormone-induced current inhibition but not the I–V shift is antagonized in Gα_t-transfected cells. The possibility that more than one hormone-activated mechanism exists for regulation of different channel properties is reinforced by recent data showing that separate protein segments in the amino terminus and/or different structural rearrangements of the channel molecule are necessary for the TRH-induced modifications of HERG activation and deactivation gating in Xenopus oocytes (Gómez-Varela et al. 2003a).

In summary, the results presented in this report lead us to propose that free βγ subunits released from G_s (and perhaps shared by G₁₃) heterotrimers are responsible for TRH-induced inhibition of the endogenous r-ERG current in GH₃ cells, in which reductions of current magnitude constitute the major component of the hormonal response. Differences in the subunit composition of the βγ dimers associated with G_q could explain the inability of G_q heterotrimers to serve a similar function in these cells. Variations in the cellular background upon heterologous expression of the channels may influence the transduction mechanism(s) involved in hormonal regulation of ERG, but also the kinetic characteristics modified in response to an agonist. Thus, as suggested by the similarities in the effects of Gα_q-QL/DN and Gα_t, βγ dimers released from a G protein different from G_s (probably G_q) are probably involved in the relatively small current inhibition detected in HEK-H36/T1 cells. However, it is a Gα₁₃- and RhoA-dependent pathway what seems to determinate the prominent alterations in HERG activation voltage dependence triggered by TRH in these cells heterologously expressing HERG channels and TRH-Rs.