Characterization of a Na+-Dependent Betaine Transporter With Clminus Channel Properties in Squid Motor Neurons

Christopher N. Petty and Mary T. Lucero

Department of Physiology, University of Utah School of Medicine, Salt Lake City, Utah 84108


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Petty, Christopher N. and Mary T. Lucero. Characterization of a Na+-dependent betaine transporter with Cl- channel properties in squid motor neurons. Most marine invertebrates, including squids, use transporters to accumulate organic osmolytes such as betaine, to prevent water loss when exposed to elevated salinity. Although a limited number of flux studies have shown the Na+ dependence of betaine transport, nothing is known about the electrogenic properties of osmolyte transporters. We used whole cell and perforated-patch voltage-clamp techniques to characterize the electrical properties of the betaine transporter in giant fiber lobe motor neurons of the squid Lolliguncula brevis. Betaine activated a large, Cl--selective current that was reversibly blocked by 100 µM niflumic acid (97 ± 2% block after 40 s, SD; n = 7) and partially inhibited by 500 µM SITS (29 ± 11%; n = 5). The Cl- current was Na+ dependent and was virtually eliminated by isotonic replacement of Na+ with Li+, NMDG+, or Tris+. Concentration-response data revealed an EC50 in a physiologically relevant range for these animals of 5.1 ± 0.9 mM (n = 11). In vertebrates, the betaine transporter is structurally related to the GABA transporter, and although GABA did not directly activate the betaine-induced current, it reversibly reduced betaine responses by 34 ± 14% (n = 8). Short-term changes in osmolality alone did not activate the Cl- current, but when combined with betaine, Cl- currents increased in hypertonic solutions and decreased in hypotonic solutions. Activation of the betaine transporter and Cl- current in hypertonic conditions may affect both volume regulation and excitability in L. brevis motor neurons. This study is the first report of a novel betaine transporter in neurons that can act as a Cl- channel.


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Transporters are integral membrane proteins that facilitate the movement of neurotransmitters, osmolytes, sugars, amino acids, and ions across the plasma membrane. The model of transporter function has changed over the years from that of a shuttle for coupled movement of substrate to something similar to an ion channel (Lester et al. 1994). The line between ion channel and transporter has become even less distinct, with recent findings that transporters can go into an uncoupled mode where transmitter-activated ionic currents occur, exceeding those predicted by stoichiometric models (DeFelice and Blakely 1996). Examples of the latter include transporters for serotonin (Galli et al. 1997; Mager et al. 1994), glutamate (Fairman et al. 1995; Larsson et al. 1996; Picaud et al. 1995; Wadiche 1995), GABA (Cammack et al. 1994, 1996; Mager et al. 1993, 1996), dopamine (Sonders et al. 1997), and norepinephrine (Galli et al. 1995). In this paper we report on a Na+-dependent, betaine transporter that is coupled to or contains a Cl- channel.

Glycine-betaine (betaine) is a trimethylamine produced by the oxidation of choline. Betaine is used as a nonperturbing osmolyte by plants, bacteria, invertebrates, and vertebrates to compensate for hypertonic stress (Burg 1995; Yancey et al. 1982). The vertebrate betaine transporter is part of the Na+/Cl--dependent carrier superfamily, with the closest homology to the GABA transporter (Yamauchi et al. 1992). The betaine transporter has been cloned from a number of vertebrate cell types, including canine Madin-Darby kidney cells (Yamauchi et al. 1992), rabbit renal papilla cells (Ferraris et al. 1996), and human brain (Borden et al. 1995). Northern blot studies show that it is distributed in kidney, liver, lung, spleen, brain (Takenaka et al. 1994), and macrophages (Warskulat et al. 1995; Zhang et al. 1996). An invertebrate betaine transporter has not been cloned.

In vertebrates, betaine transport requires Na+ and Cl- and is modulated by changes in osmolarity (Nakanishi et al. 1990). The invertebrate betaine transporter found in the gills of marine mussels (Mytilus californianus) is also dependent on Na+, and transport activity decreases with decreased osmolarity (Wright et al. 1992).

We studied the electrical properties of the Na+-dependent betaine transporter in giant fiber lobe (GFL) motor neurons of the squid Lolliguncula brevis, a euryhaline cephalopod. These osmoconforming squids rapidly adjust their blood osmolality to <= 2% of the ambient sea water, allowing them to cross osmotic gradients that range from 525 to 1,089 mOsm without harm (Hendrix et al. 1981). Betaine concentrations in cephalopod axoplasm can be very high (70 mM) compared with blood (4 mM) (Deffner 1961), implicating betaine as an important osmolyte for squids. We found that the betaine transporter in L. brevis is capable of moving into an uncoupled mode with large ion channel-like fluxes. Interestingly, for the betaine transporter that we studied, the fluxes are carried by Cl-, similar to the Cl- fluxes in glutamate transporters (Fairman et al. 1995) rather than the cationic fluxes of GABA transporters (Mager et al. 1996). In addition, activity of the Cl- current is dependent on changes in osmolarity, and this may be one mechanism used by euryhaline invertebrates to rapidly equilibrate to changing salinities of their natural environment.


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Cell preparation and culture conditions

The well-known squid giant motor axons are formed by the fusion of hundreds of normal sized axons that emanate from cell bodies located in the posterior tip of the GFL of the stellate ganglia (Young 1939). The animals used in this study, L. brevis, were obtained from the National Resource Center for Cephalopods (Galveston, TX). Isolation and culture of GFL cell bodies was similar to that described by Gilly et al. (1990). Briefly, the animals were decapitated, and the GFLs of the squid L. brevis were excised under a dissecting microscope and treated with nonspecific protease (10 mg/ml Sigma type XIV) in sterile filtered artificial seawater (ASW) for 40 min. After a 3- to 5-min rinse in ASW, cell bodies (40-60 µm diam) were teased from the GFL with glass micropipettes and plated onto a drop of culture media on concanavalin A-coated (10 mg/ml; Sigma type IV) glass coverslips and allowed to settle for 15 min before adding 2 ml media and placing in the incubator at 22°C. In the majority of cells, the axon was either cutoff at the level of the cell body or reabsorbed, and all recordings were made from round cells. Culture media was changed daily, and cells were used for experiments between 1 and 3 days in culture.

Solutions

The external bath solutions and internal pipette solutions are listed in Tables 1 and 2, respectively. The osmolality of 780 mOsm/kg H2O was chosen for our standard recording conditions because it is in the midrange of osmolalities of the seawater that this osmoconforming species of squid inhabits. The culture media consisted of Leibovitz's L-15 (Gibco, Grand Island, NY) supplemented with salts to bring the osmolality to 780 mOsm/kg H2O, 2 mM HEPES (pH 7.6), 4 mM 1-glutamine, 200 units/ml penicillin G, and 200 mg/ml streptomycin sulfate (Irvine Scientific, Santa Ana, CA). The media was set to pH 7.6 with NaOH and immediately sterile filtered. All chemicals were obtained from Sigma Chemical (St. Louis, MO) unless stated otherwise. A 20-mM stock solution of the chloride channel blocker niflumic acid was made by sonication in DMSO and then diluted to 100 µM in ASW. Control application of DMSO had no effect.


                              
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Table 1. Composition of external bath solutions


                              
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Table 2. Composition of internal pipette solutions

Gramicidin and whole cell recordings

Gramicidin perforated-patch recordings were used to preserve [Cl-]i (Myers and Hayden 1972) and internal second messengers, and were made according to the technique of Abe et al. (1994). The gramicidin stock solution was made fresh daily and consisted of 1 mg gramicidin/100 µl methanol; 10 µl of this stock solution was added to 1 ml internal solution (Table 2) and was kept in the dark and on ice.

Three to 6 MOmega resistance electrodes were pulled from thick-walled borosilicate filament glass (Sutter Instrument, San Rafael, CA) to make both whole cell (Hamill et al. 1981) and gramicidin perforated-patch recordings. Bath solutions (21-23°C, the temperature at which these animals live) were perfused through the chamber at a rate of 1-2 ml/min. Test solutions were delivered with a Warner SF-77 rapid solution changer (Warner Instrument, Hamden, CT). Measuring the time course of current responses during solution changes to ASW in which the NaCl was replaced by KCl revealed that it took 200-300 ms to completely change the solution on the GFL cells. A 3-M KCl-agar bridge was used to ground the bath solutions. The 1- to 13-mV liquid junction potential between bath and pipette solutions was calculated with Axoscope (Axon Instruments, Foster City, CA), and all reversal potentials reported in the text and in Fig. 1E were appropriately corrected. Calculated reversal potentials were based on ionic concentrations.



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Fig. 1. Betaine responses are selective for Cl-. A: raw current traces of a whole cell voltage-clamped GFL neuron in response to 4,000-ms voltage steps from -90 up to -30 mV in 20-mV increments (initial holding voltage was -70 mV). The command voltage at the end of each voltage step became the holding voltage during the 20-s interval between voltage steps. The time that 5 mM betaine superfused the cell (1,000 ms) is indicated by the solid bar in this and all subsequent figures. B: same current traces as in A after baseline subtraction and deletion of the first 500 ms. Solutions: TMA-Glut internal//580 mOsm artificial seawater (ASW) ext. C: current traces obtained from a different cell. Voltage steps were applied for 4,000 ms from -90 to +30 mV in 20-mV increments. Solutions: TMA-Cl int.//ASW ext. D: I-V relationship plotted for the peak currents in B (black-square) and C (). E: plot of observed Erev vs. calculated ECl shows that the betaine-induced currents followed the Nernst potential for Cl-. Error bars indicate ±SD. Numbers indicate number of cells. Solutions: TMA-Cl int.//ASW ext., observed Erev = -9 ± 6 mV, n = 14, open circle ; TMA-Cl int.//50% Li-ASW ext., observed Erev = -10 ± 2 mV, n = 4, ; Gluc. 21 int.//580 mosm ASW ext., observed Erev = -76 ± 5 mV, n = 3, black-square; TMA Gluc int.//ASW ext., observed Erev = -52 ± 4 mV, n = 4, ; TMA Glut int.//580 mosm ASW ext. observed Erev = -53 ± 1 mV, n = 3. F: betaine-induced current responses are shown from a perforated-patched GFL neuron with the same voltage protocol as in A. Solutions: Gramicidin TMA-Cl int.//ASW ext.

Data acquisition

Voltage-clamp recordings were made as described by Danaceau and Lucero (1998). Cells were voltage clamped to a given voltage for 4,000 ms. To allow time for voltage-gated currents to stabilize, betaine and other stimulants were applied 1,500 ms after the initiation of the voltage step. An example of a raw current trace is shown in Fig. 1A. In all of the following figures, we eliminated the first 500 ms containing transient voltage-gated currents. We baseline subtracted the steady-state, voltage-dependent and linear-leak currents by setting the current amplitudes recorded for 300 ms before a response to 0 pA so that only the betaine-sensitive currents are shown (Fig. 1B).


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Betaine activates a Cl- selective current

We applied 5 mM betaine to GFL neurons isolated from the squid L. brevis that were whole cell voltage clamped to -90 mV and internally perfused via the patch pipette with a high Cl- (366 mM) internal solution. Under these conditions we found that betaine activated inward currents as large as 15 nA, with average current values of 6.7 ± 4.1 nA (SD, n = 14). When normalized to cell capacitances that averaged 119 ± 33 pF (n = 14), the current densities averaged 63.7 ± 50.5 pA/pF (n = 14).

To determine the ionic selectivity of betaine responses, we used patch pipettes filled with various concentrations of Cl- (see Table 2) and measured the reversal potential (Erev) of betaine responses. By using ion substitution, the Erev of Na+, K+, and Ca2+ were set to very positive values (more than +60 mV) to measure the Cl- current reversal in isolation. Betaine-induced currents reversed near the predicted Cl- reversal potential (ECl) of -49 mV during whole cell recordings with a K+-free 51-mM Cl- pipette solution (TMA-Glut) and a 330-mM Cl- bath solution (Fig. 1, B and D, black-square, and E, black-triangle). Changing the Cl- concentrations shifted the reversal potential of betaine current responses. Figure 1C shows that betaine-induced currents reversed at -10 mV in a different cell, with a K+-free 366 mM Cl- pipette solution (TMA-Cl) and ASW bath solution (calculated ECl -5 mV). The current-voltage (I-V) relationships for the peak currents in Fig. 1, B and C, are shown in Fig. 1D. The plot of the average Erev of betaine responses obtained by using different [Cl-]i versus the ECl, calculated with the Nernst equation and the Cl- concentrations of the pipette and bath solutions, is shown in Fig. 1E. The data closely follow the solid line, which indicates the values for a perfectly selective Cl- channel. Changes in Ca2+concentration (data not shown) or Na+ and K+ concentrations had no effect on the Erev of betaine-induced currents (Fig. 2B).



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Fig. 2. Betaine-activated currents are dependent on external Na+. A: averaged current responses to 5 mM betaine in ASW with equimolar substitution of LiCl were normalized to peak responses in 470 mM Na ASW (error bars indicate ±SE; n = 6). B: current-voltage relationships of betaine responses are shown for a cell exposed to the [Na+]o indicated over the voltage range of -90 mV to +10 mV.

In comparing the I-V relationships in which Erev was negative to -40 mV (n = 10) with those with Erev close to 0 mV (n = 14), we observed a consistent nonlinearity (inward rectification) when the ECl was set to less negative potentials (Fig. 1D, ). This nonlinearity was not observed when ECl was more negative (Fig. 1, D, black-square, and F) and is likely due to the decrease in membrane resistance at depolarized potentials rather than a voltage dependence of the betaine-induced current.

Once we determined that the betaine responses were Cl- selective, we used the gramicidin perforated-patch technique, which allows whole cell measurement of current responses without perturbing internal Cl- concentration to determine the cell's natural [Cl-]i. When 5 mM betaine was applied to a cell in gramicidin perforated-patch mode, currents were inward at voltages negative to -60 mV but outward at voltages positive to -60 mV (Fig. 1F). On average, betaine responses recorded in gramicidin perforated-patch mode reversed at -46 ± 6 mV (n = 19), giving a calculated [Cl-]i of ~71 mM. The calculated [Cl-]i of L. brevis GFL neurons is similar to the [Cl-]i of the axoplasm from the squid Loligo (151 mM) (Deffner 1961), considering that our measurements were made in reduced salinity ASW (780 mOsm).

Betaine-Cl- responses are Na+ dependent

To investigate the dependence of betaine-Cl- responses on external Na+, we made equimolar substitutions of Li+ for Na+ in 470 mM Na ASW (the Na+ concentration of normal 1,000-mOsm seawater). We found that external Na+ is required for betaine responses. Figure 2A shows a plot of normalized and averaged betaine-induced current responses recorded at -70 mV in whole cell mode with different [Na+]o. A Hill equation fit to the data gave an EC50 of 224 mM ± 37 mM (n = 6). The I-V relationships plotted in Fig. 2B show the effects of reducing external Na+ over the voltage range from -90 to +10 mV. Although the size of the betaine responses decreased as external sodium was eliminated, Erev was unaffected, indicating that although Na+ is required it is not a major permeant ion. Identical results were obtained by substituting two additional and structurally different monovalent cations, Tris+ (n = 13) or NMDG+ (n = 28), suggesting that the elimination of Na+ rather than some sort of nonspecific blocking effect by Li+ reduced the betaine-Cl- responses. These results indicate that the betaine-induced Cl- current in GFL neurons is Na+ dependent.

Betaine-induced Cl- currents are dose dependent

To determine whether betaine acts in a physiologically relevant range, we measured the EC50 for the betaine response. Figure 3A shows current responses to 1-s applications of 1, 5, and 50 mM betaine. The amplitude and kinetics of betaine-induced Cl- currents increased with increasing betaine concentration. Figure 3B shows the concentration-response curve for betaine, obtained by normalizing the betaine responses for a given cell to that of the response of 100 mM of betaine in the same cell and plotting against the log of the betaine concentration. The data were fit with the Hill equation yielding an EC50 for the betaine response of 5.1 ± 1.0 mM (n = 11) and a Hill coefficient of 1.5 ± 0.3 (n = 11). This EC50 is in a physiologically relevant range assuming that Lolliguncula brevis have similar (mM) concentrations of betaine in their blood as Loligo (4 mM) (Deffner 1961).



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Fig. 3. Betaine-induced currents are dose dependent with an EC50 of 5.1 mM and a Hill coefficient of 1.5. A: superimposed current traces of betaine-induced currents after 1-s application of 1, 5, and 50 mM betaine at -70 mV. B: concentration-response curve was obtained by plotting normalized peak betaine-induced currents at -70 mV against the log of betaine concentration. A Hill equation was fit to the data (smooth line) to determine the EC50 and the Hill coefficient. Error bars indicate ±SE. Numbers indicate number of cells.

Pharmacology of betaine-induced Cl- currents

Our selectivity studies indicated that betaine activated a Cl--selective current. We tested whether the Cl- channel blocker niflumic acid could block betaine current responses. In Fig. 4A we show that application of 100 µM niflumic acid completely and reversibly blocked the betaine-elicited Cl- current. Figure 4B shows the averaged percent current remaining from seven to nine cells, with current responses normalized to the peak of the control betaine application. The amplitude of the control response (100%) was lowered to 6 ± 6% (n = 9) with the 20-s application of niflumic acid and down to 3 ± 2% (n = 7) with 40-s niflumic acid application. After 60 s of washing out niflumic acid, the average current response recovered to 92 ± 10% (n = 9) of the original response. Bath application of 500 µM SITS reversibly reduced 5 mM betaine current responses by 29 ± 11% (n = 5). Collectively, our results show that betaine is activating a Na+-dependent, Cl-- selective current. To rule out the possibility that the Cl- current was associated with betaine activation of a verapamil-sensitive P-glycoprotein (Tominaga et al. 1995), we applied 25 µM verapamil to four cells and saw no effect on the betaine-elicited current. In addition, we found that elimination of external Ca2+ by applying betaine in a 0-Ca2+ bath solution with 10 mM EGTA had no effect on the betaine-induced currents (n = 7). Finally, external application of 1 mM ATP did not reduce the betaine-induced Cl- current (n = 8).



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Fig. 4. The Cl- channel blocker niflumic acid reversibly blocks betaine-induced currents. A: current traces showing application of betaine in the presence and absence of 100 µM niflumic acid at -80 mV. a: control betaine response before niflumic acid application. b: betaine response after 20 s of continuous application of niflumic acid. c: betaine response after 40 s of continuous niflumic acid application. d: betaine response after 20 s of washout. e: betaine response after 40 s of washout. f: betaine responses after 60 s of washout. B: betaine responses were normalized to the control peak current and averaged across cells; a-f, same as previously. Error bars indicate ±SD. Numbers indicate number of cells.

Betaine responses are reversibly reduced by GABA

Because of the structural similarity of GABA and betaine, we tested whether GABA affected betaine responses. Figure 5 shows data from a cell in which application of 5 mM GABA alone did not activate a current; however, coapplication of 5 mM GABA and 5 mM betaine reversibly reduced the amplitude of the betaine response by 42%. On average, 5 mM GABA reduced 5 mM betaine responses by 34 ± 14% (n = 8). Application of 5 mM choline activated a small current with kinetics similar to the betaine-elicited current (n = 4), whereas 5 mM glycine, proline, alanine, or taurine did not elicit any current (n = 4). A subset of GFL neurons contained ionotropic GABA receptors and responded to 5 mM GABA with rapid, transient Cl- currents; however, those cells were not included in the present analyses.



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Fig. 5. Betaine responses are reduced by GABA. Current traces showing responses to 5-mM application of GABA alone, betaine alone, 5 mM GABA + 5 mM betaine, and betaine after GABA + betaine. Stimulus application is indicated by the solid bar.

Betaine-induced Cl- currents are sensitive to changes in osmolality

On the basis of our hypothesis that betaine activates a osmoregulatory Na+-dependent transporter containing a Cl- channel, we predicted that the Cl- current should be sensitive to the same osmotic changes that would affect betaine transport. To test this prediction, we applied betaine under three osmotic conditions that span the range of salinities that these animals can tolerate (580, 780, and 980 mOsm ASW) (Hendrix et al. 1981). Figure 6A shows the effect of increasing and decreasing osmolality on 5-mM betaine responses with whole cell voltage-clamp recordings. All of the recordings were made at -70 mV, and D-mannitol was used to increase osmolality so that Cl- and Na+ concentrations of the three solutions were the same. Our normal recording bath and pipette solutions were set at 780 mOsm. When the external solution bathing a GFL cell was changed from hypertonic 980 mOsm ASW (Fig. 6, ) to either 780 () or 580 mOsm ASW (triangle ), the betaine-induced Cl- currents were eliminated. In contrast, when the bath solution was changed from a hypotonic 580 mOsm ASW to either 780 or 980 mOsm ASW, the betaine-induced Cl- currents returned within the 20-s application interval. Similar results were obtained in 21 cells. These data indicate that the betaine-induced current acted exactly as an osmotically sensitive betaine transporter would act; in the presence of betaine, it turned on in hypertonic conditions (betaine uptake to prevent shrinkage) and it shut off in hypotonic conditions.



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Fig. 6. Betaine-induced currents are sensitive to changes in osmolality. A: peak current is plotted vs. time for 5-mM betaine application obtained while changing external osmolality (all solutions have 580 mOsm/kg H2O salts and use 200 or 400 mM mannitol to reach 780 and 980 mOsm, respectively). Current size decreased when exposed to hypotonic medium. B: comparison of normalized betaine-induced currents vs. time when switching from hypertonic to hypotonic medium. The hypertonic medium osmolarity was increased from 580 to 980 mOsm with mannitol (impermeant), urea (very permeant), and glucose (intermediate permeability). Data shown are from 3 different cells.

To further investigate osmotic effects on the betaine-activated Cl- current, we tested whether substitution of glucose or urea for mannitol affected the betaine-current responses. Because glucose is transported and therefore more permeant than mannitol, we predicted that in glucose solutions, the osmotic effects may be compensated for as glucose is transported into or out of the cell. Similarly, urea, which is freely permeant, should show even faster compensation for osmotic changes. In Fig. 6B we normalized and superimposed peak current responses to betaine application for a single transition from 980 to 580 mOsm ASW. When mannitol was used to increase osmolality to 980 mOsm, the betaine-induced Cl- current was eliminated in the 580-mOsm ASW. When glucose was used, the majority of the betaine-induce Cl- current recovered within 60 s of hypotonic solution exposure. When urea was used, the cells responded as if the osmolality of the solution was unchanged, as would be expected for a freely permeant substance. The effects of glucose and urea were repeatable and observed in all five cells tested. These data strongly indicate that the betaine-elicited Cl- current shows the same dependency on osmotic changes as expected for a betaine transporter. Furthermore, short-term changes in tonicity in the absence of betaine (2-3 min) had no effect on the cell, indicating that the Cl- current activated by betaine is not directly shrink nor swell activated. Prolonged incubations in hypotonic seawater (>5 min) appeared to activate volume-sensitive ion channels because the cells became very leaky. These slow, betaine-independent conductances were not investigated further.


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The work in this study showed that betaine, an organic osmolyte, activates a Cl- current in GFL motor neurons of the squid Lolliguncula brevis. Our data showed that the betaine-induced current is highly selective for Cl- and that it can be completely blocked by the Cl--channel blocker niflumic acid and partially blocked by SITS. The strict dependence on betaine, Na+, and hypertonicity suggests that the Cl- current is carried through the betaine transporter. However, the large amplitude of the Cl- current rules out the possibility that it is a stoichiometrically coupled transporter current. In contrast to a coupled transport process, we believe that as in many of the neurotransmitter transporters, the betaine transporter is capable of acting like an ion channel and generating a nonstoichiometric Cl- flux.

We designed a model similar to that proposed by Lester et al. (1994) for neurotransmitter transporters, and Wadiche et al. (1995) and Larsson et al. (1996) for glutamate transporters, which incorporates the findings of Moeckel et al. (1997) for betaine transporters. In our model, the transporter and the ion channel are inactive in the absence of sodium and betaine (Fig. 7A). When present, sodium and betaine act as coligands, binding to the outside of the transporter (Fig. 7B). The Hill coefficient of 1.5 may reflect the requirement for the binding of two Na+ ions as the rate-limiting step in betaine binding, as was described for the GABA transporter (Mager et al. 1996). Once Na+ and betaine bind, a conformational change occurs that opens a Cl- ion channel, allowing Cl- flux in and out of the cell, depending on the membrane potential and ECl (Fig. 7C). As Na+ and betaine are transported across the membrane and leave their binding sites, the Cl- channel closes (Fig. 7D). Our model is supported by our recent betaine flux studies on whole GFL tissue from L. brevis that show Na+-dependent betaine uptake. As in the current work, the [3H]betaine uptake is blocked by 100 µM niflumic acid (n = 5) and is dependent on external Cl- (J. Poulsen, D. Steel, and M. Lucero, unpublished observations). An alternative model that we cannot rule out is that the betaine transporter and the Cl- channel are two separate proteins that are blocked by niflumic acid, and the gating of the Cl- channel is controlled by betaine, Na+, and salinity in an identical manner as the betaine transporter.



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Fig. 7. Model of the squid neuronal betaine transporter showing the binding of substrates (2 betaine and >1 Na+) and conformational change, allowing for uncoupled Cl- flux. A: transporter is empty and closed. B: Na+ and betaine bind to activate the transporter. C: transporter has undergone a conformational change and opens into a Cl--selective channel. Cl- will move into or out of the cell, depending on the membrane potential. D: once Na+ and betaine are transported to the inside of the cell, the transporter empties, closes, and returns to the configuration shown in A.

The mammalian betaine transporter is part of the Na+/Cl--dependent carrier superfamily, with close homology to the GABA transporter (Lester et al. 1994; Yamauchi et al. 1992). Physiologically, our transporter is similar to that of GABA transporters in that it requires Na+ for activation but differs in that the ionic component of the GABA transporter can be activated in the absence of GABA and appears to be a cationic channel (Cammack et al. 1994; Mager et al. 1996). The serotonin transporter requires Na+, Cl-, and K+ (Nelson and Rudnick 1979) and transports Na+ (Galli et al. 1997; Mager et al. 1994). The dopamine transporter is also Na+/Cl- dependent and conducts a proton/alkali cationic leak that is Na+/Cl- independent (Sonders et al. 1997). Of the various transporter-ion channels described, the betaine-induced Cl- current is most similar to the glutamate transporter Cl- current (Picaud et al. 1995), which is in a different gene family from the monoamine transporters (Kanner 1993). The betaine-induced Cl- current in squid motor neurons differs from the glutamate transporter in that niflumic acid does not block the Cl- current associated with the glutamate transporter (Wadiche et al. 1995) and internal K+ is necessary for glutamate-induced Cl- currents (Picaud et al. 1995), whereas the presence and absence of K+ do not affect betaine-induced Cl- currents.

Our study represents the first time that the electrical properties of the betaine transporter were studied in native cells. Our results show that betaine-Cl- currents are Na+ dependent and increase when placed in hypertonic media, thereby giving us an ionic marker for betaine transport. These findings corroborate previous studies showing hypertonicity induced increases in betaine uptake (Ferraris et al. 1996; Moeckel et al. 1997; Takenaka et al. 1994; Wright et al. 1992) and indicate that betaine may have a prominent role in rapid adjustments of cellular osmolarity in squid motor neurons in response to osmotic stress. What role might Cl- currents play in the rapid osmotic adjustments mediated by betaine? Activation of the betaine-induced Cl- conductance could modulate neuronal excitability by holding the resting potential close to the Cl- reversal potential (-46 mV). In hypertonic conditions, a negative membrane potential would support betaine uptake. If the cell were to depolarize or hyperpolarize, Cl- current through the transporter would tend to restore the potential and help maintain a stable, negative resting potential in the face of changing ionic conditions.

Recent work was done on volume-sensitive Cl- channels (ICl.swell) (see reviews by Strange et al. 1996, 1998) and on volume-sensitive organic osmolyte/anion channels (VSOACs) that mediate efflux of organic osmolytes (Kirk and Strange 1998). These ion channels are swell activated, i.e., activated by hypotonic mediums, whereas hypertonic media have no effect and in many cell types are blocked by extracellular ATP. VSOACs show little selectivity among osmolytes, suggesting a common anion channel pathway. In our experiments, short-term changes (2-3 min) in the tonicity alone had no effect on ionic currents recorded from GFL neurons; 1 mM ATP did not reduce the betaine-induced Cl- current. Taurine, proline, and alanine did not activate the Cl- current, and hypotonic media rapidly eliminated the betaine response. Our results indicate that the betaine-induced conductance is not associated with swell-activated ion channels.

Our findings indirectly show that the betaine transporter in L. brevis plays a role in adapting to hypertonic conditions that may include modulation of neuronal excitability and support the evolving view of transporters acting as ion channels. It will be interesting to perform similar studies on GFL neurons from squid species that do not tolerate osmotic changes.


    ACKNOWLEDGMENTS

We thank Drs. L. DeFelice, W. Michel, D. Steel, and B. Olivera for helpful suggestions, and J. Lucero, D. Piper, J. Danaceau, J. Poulsen, and J. Lew for technical support.

This work was supported by National Institutes of Health Grant DC-02587-03 to M. T. Lucero and by a short-term research training grant, 5T35HL-07744, to C. N. Petty.


    FOOTNOTES

Address for reprint requests: M. T. Lucero, Dept. of Physiology, 410 Chipeta Way, Salt Lake City, UT 84108.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 21 August 1998; accepted in final form 4 January 1999.


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ABSTRACT
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