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Hydrogen-Deuterium Exchange Effects on ß-Endorphin Release from AtT20 Murine Pituitary Tumor Cells

Masayuki Ikeda ^* , Shigeru Suzuki , Masahiro Kishio , Moritoshi Hirono ^*, Takashi Sugiyama , Junko Matsuura , Teppei Suzuki , Takayuki Sota , Charles N. Allen ^¶, Shiro Konishi ^|| and Tohru Yoshioka ^* 

^* Advanced Research Institute for Science and Engineering, Waseda University, 3-4-1 Okubo Shinjuku-ku, Tokyo 169-8555, Japan; Department of Molecular Behavioral Biology, Osaka Bioscience Institute, 6-2-4 Furuedai, Suita, Osaka 565-0874, Japan; Department of Molecular Neurobiology, School of Human Sciences, Waseda University, 2-579-15 Mikajima, Tokorozawa, Saitama 359-1192, Japan; School of Science and Engineering, Waseda University, 3-4-1 Okubo Shinjuku-ku, Tokyo 169-8555, Japan; ^¶ Center for Research on Occupational and Environmental Toxicology, Oregon Health and Science University, Portland, Oregon 97201-3098, USA; and ^|| Mitsubishi Kagaku Institute of Life Sciences, and Core Research for Evolutional Science and Technology (CREST), Japan Science and Technology Agency (JST), 11 Minamiooya, Machida-shi, Tokyo 194-8511, Japan

Correspondence: Address reprint requests to Tohru Yoshioka, PhD, Tel.: +81-3-5286-2761; Fax: +81-3-3205-6419; E-mail: yoshioka{at}human.waseda.ac.jp.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Abundant evidences demonstrate that deuterium oxide (D₂O) modulates various secretory activities, but specific mechanisms remain unclear. Using AtT20 cells, we examined effects of D₂O on physiological processes underlying ß-endorphin release. Immunofluorescent confocal microscopy demonstrated that 90% D₂O buffer increased the amount of actin filament in cell somas and decreased it in cell processes, whereas ß-tubulin was not affected. Ca²⁺ imaging demonstrated that high-K⁺-induced Ca²⁺ influx was not affected during D₂O treatment, but was completely inhibited upon D₂O washout. The H₂O/D₂O replacement in internal solutions of patch electrodes reduced Ca²⁺ currents evoked by depolarizing voltage steps, whereas additional extracellular H₂O/D₂O replacement recovered the currents, suggesting that D₂O gradient across plasma membrane is critical for Ca²⁺ channel kinetics. Radioimmunoassay of high-K⁺-induced ß-endorphin release demonstrated an increase during D₂O treatment and a decrease upon D₂O washout. These results demonstrate that the H₂O-to-D₂O-induced increase in ß-endorphin release corresponded with the redistribution of actin, and the D₂O-to-H₂O-induced decrease in ß-endorphin release corresponded with the inhibition of voltage-sensitive Ca²⁺ channels. The computer modeling suggests that the differences in the zero-point vibrational energy between protonated and deuterated amino acids produce an asymmetric distribution of these amino acids upon D₂O washout and this causes the dysfunction of Ca²⁺ channels.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Deuterium, a heavy and stable isotope of hydrogen, binds to oxygen to form deuterium oxide (D₂O), commonly known as heavy water. D₂O exists in the environment at 1/6700 of H₂O, and is expected to have some biological effects, deleterious and otherwise, on whole animals, animal cells, microorganisms, and macromolecules (Kushner et al., 1999), but many of these effects remain uncharacterized.

The effects of D₂O on cellular secretion have been extensively studied in various cell types without any consensus on the underlying mechanisms. For example, D₂O significantly reduces insulin release from pancreatic islets (Malaisse-Lagae et al., 1971; Freinkel et al., 1976; Montague et al., 1976; Hill and Rhoten, 1983), amylase release from pancreatic fragments (Calderon et al., 1980), and gastrin release from antral mucosal fragments (Harty et al., 1983). D₂O has been shown to modulate the amount and arrangement of microtubules (Malaisse-Lagae et al., 1971; Houston et al., 1974; Itoh and Sato, 1984) and secretory granules are transported from the basal to the apical cytoplasm via microtubules (Lacy et al., 1968; Malaisse et al., 1975; Sasaki and Tashiro, 1976), thus the D₂O-induced decrease in secretion has been ascribed to the stabilization of microtubules (Malaisse-Lagae et al., 1971; Montague et al., 1976; Hill and Rhoten, 1983). The mechanism underlying the effect of D₂O may not be this simplistic, however, because it has been shown that D₂O increases serotonin release from blood platelets (Bennett et al., 1979) and spleen cells (Sulowska and Wyczolkowska, 1991) and histamine release from mast cells (Fewtrell and Metzger, 1980; Maeyama et al., 1986) and basophilic leukemia cells (Urata et al., 1989).

Theoretically, D₂O has at least two independent physiological effects: i), an isotope exchange (hydrogen to deuterium) effect on functional proteins, and ii), a solvent isotope effect, meaning that the mobility of monovalent cations is reduced by 20%, whereas the dielectric constant is not changed in D₂O buffer (Bass and Moore, 1973). The computer simulation studies on the isotope exchange effect using the density functional theory demonstrated that with H₂O to D₂O exchange, the state of functional proteins changes quickly, but with D₂O to H₂O exchange, the state of proteins recovers very slowly, due to the stability of deuterated amino acids (Table 1). On the other hand, reported electrophysiological properties (membrane potential and channel conductances), which are under the influence of a solvent isotope effect, change reversibly and rapidly with the change from H₂O to D₂O and back again (Schauf and Bullock, 1979; Schauf and Chuman, 1986; Andjus et al., 1994). Therefore, isotope exchange effects and solvent isotope effects are possibly distinguishable as temporal pattern of D₂O actions. However, D₂O actions on physiological processes such as exocytosis have not been analyzed from this biophysical point of view.

View this table: [in this window] [in a new window]   TABLE 1  Electronic (self-consistent field) energy (Hartree), ZPVE (kcal/mol), and ZPVE (kcal/mol) of hydrophilic amino acids

With special cautions for the temporal coordinates of D₂O actions, we found biphasic and bidirectional effects of D₂O on high-K⁺-induced ß-endorphin secretions in AtT20 cells. The initial increase of secretion corresponds to the immediate redistribution of actin filaments, whereas the delayed inhibition of secretion corresponds to dysfunction of VSCCs upon D₂O washout.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Cell cultures
AtT20 cells (clone D16V) were cultured at 37°C in Dulbecco's modified eagle medium supplemented with 10% fetal bovine serum in a humidified atmosphere of 95% air and 5% CO₂. For Ca²⁺ imaging and electrophysiological experiments, cells were plated onto 35-mm glass-bottom petri dishes or 12-mm glass cover slips at a density of 5 x 10⁴ cells. The culture medium was exchanged every other day and on the day before the experiments.

Immunofluorescent confocal imaging
To observe the effects of D₂O on the cytoskeleton, AtT20 cells were incubated in H₂O-based buffered salt solution (BSS) consisting of (mM) 128 NaCl, 5 KCl, 2.7 CaCl₂, 1.2 MgCl₂, 1 Na₂HPO₄, 10 glucose, and 10 HEPES/NaOH (pH 7.3) for 20 min, and the H₂O BSS was replaced with 90% D₂O BSS for 5 or 15 min. AtT20 cells were then fixed in 4% phosphate-buffered paraformaldehyde for 15 min and washed three times with phosphate-buffered saline (PBS) consisting of (mM) 137 NaCl, 2.7 KCl, 1.5 KH₂PO₄, and 8.1 Na₂HPO₄ (pH 7.4). The fixed sample was then dipped in acetone for 3 min and dried to facilitate staining of F-actin with rhodamine phalloidin (Molecular Probes, Eugene, OR). The dried sample was incubated for 2 h at room temperature (24–26°C) with monoclonal antibodies against ß-tubulin (1:100, ICN Biomedicals, Aurora, OH) and neurofilament-M (1:100, Chemicon International, Temecula, CA) dissolved in PBS supplemented with 1% bovine serum albumin (BSA), 0.1% Triton-X, and 0.05% sodium azide. The sample was rinsed three times with PBS and incubated for 1 h at room temperature with fluorescein isothiocyanate-conjugated goat anti-mouse IgG (ICN Biomedicals, 1:200) and rhodamine phalloidin (1:200) dissolved in the bovine serum albumin-supplemented PBS. After washing out the secondary antibody with PBS, the sample was mounted with glycerol. A confocal imaging system (µ-Radiance, Bio-Rad, Hercules, CA) attached to a fluorescence microscope (AxioPlan2, Carl Zeiss, Oberkochen, Germany) was used with an argon/krypton laser and a 40x objective lens (Plan-Neofluar 40x/0.75, Carl Zeiss). The digital image was processed using Photoshop (Adobe) software. All staining experiments were repeated at least three times.

Ca²⁺ imaging
The culture medium was gently rinsed from dishes using BSS. The cells were then incubated in BSS supplemented with 5 µM fura-2 AM (Molecular Probes) for 30 min in a CO₂ incubator. Before photometry, the cells were rinsed again with BSS, incubated at 37°C for 15 min, and then continuously perfused with BSS at 30°C at a flow rate of 3 ml/min. Fluorescence images were produced by alternating 340- and 380-nm ultraviolet pulses (143-ms duration) and the resulting pairs of images were projected onto a cooled CCD camera (C4880, Hamamatsu Photonics, Hamamatsu, Japan) at 3–6-s intervals. A cocktail of 10 µM nifedipine (Sigma, St. Louis, MO) and 100 nM -conotoxin MVIIC (Alomone Labs, Jerusalem, Israel) was used to block VSCCs, because these VSCC blockers at these concentrations inhibit most VSCC subtypes in AtT20 cells (Loechner et al., 1996). Tetrodotoxin (TTX; Alomone Labs) was applied at a concentration of 1 µM to estimate possible involvement of Na⁺ channels in the Ca²⁺ oscillations. The frequency of the Ca²⁺ oscillations was estimated by the number of peaks within 180-s blocks recorded during the control and TTX treatment periods. All drugs were bath-applied with the perfusate. Further details of the Ca²⁺ imaging technique have been described previously (Ikeda et al., 2001; Nelson et al., 2001).

Electrophysiology
Recordings were performed at 3–4 days after plating by placing a cover slip fragment into a recording chamber located on the stage of an upright microscope equipped with phase-contrast optics. The chamber was perfused at 1 ml/min with BSS. Microelectrodes, pulled in two stages (Narishige puller, Narishige, Tokyo, Japan) from thin-walled borosilicate glass (World Precision Instruments, Sarasota, FL), had resistances of 5–7 M when filled with a (intracellular) solution, at pH 7.3 and 310 mOsm, consisting of (mM) Cs methanesulfonate 87, CsCl 20, CsOH 25, CaCl₂ 1, HEPES 10, EGTA 10, ATP 3, and GTP 0.3. For the recording of resting membrane potential of AtT20 cells, the current clamp mode of the whole-cell patch clamp method was used with EPC-8 patch amplifier (HEKA, Lambrecht/Pfalz, Germany). Membrane potential responses were filtered at 3 kHz and acquired using pCLAMP7 software (Axon Instruments, Foster City, CA). For the recording of K⁺ currents, the cells were held at -60 mV using the whole-cell configuration of the patch-clamp technique and the membrane voltage stepped to +50 mV once every 30 s. To activate voltage-gated Na⁺ currents the cells were voltage clamped at -83 mV and depolarizing steps (10-mV steps, 30 ms in duration) applied once each 4 s. For the recordings of Ba²⁺ currents, Ca²⁺-free BSS containing 10 mM BaCl₂ and 0.5 µM TTX was used as the external solution and 3 µM QX-314 was added to the internal solution. For the recordings of Ca²⁺ currents, internal solutions consisting of (mM) CsCl 120, MgCl₂ 3, HEPES 10, EGTA 10, ATP 3, GTP 0.4 and creatine phosphate 14, and external solutions consisting of (mM) TEA-Cl 142, CaCl₂ 2.5, MgCl2 1, HEPES 15, and glucose 9.9 were used. To observe intracellular hydrogen/deuterium exchange effects, Ca²⁺ current recordings were initiated with the internal solution of 90% D₂O, and thereafter the external solution was replaced to 90% D₂O-BSS by the bath perfusion. Voltage-gated Ca²⁺ currents were activated from a holding potential of -80 mV with depolarizing pulses (10-mV steps, 50 ms) applied once each 8 s. These currents were recorded using an Axopatch 1C (Axon Instruments), filtered at 2 kHz (-3 dB) with an eight-pole Bessel filter and digitized at 10 kHz on-line using an ITC16 A/D interface (Instrutech, Port Washington, NY) and Pulse software (HEKA). The cell capacitance and series resistance (15–30 M, 80%) were compensated. All records were corrected for a liquid junction potential of -13 mV. Further details of the recording methods have been described previously (Suzuki et al., 1997a,b; Hirono et al., 1999).

Radioimmunoassay for ß-endorphin release
AtT20 cells were seeded onto a 100-mm dish (1 x 10⁶ cells/dish) and cultured at 37°C for 24 h in the growth medium. After washing out the growth medium with BSS, the cells were incubated in 4 ml H₂O BSS, D₂O BSS, 30 mM K⁺ H₂O BSS, and/or 30 mM K⁺ D₂O BSS at room temperature. ß-Endorphin concentrations in H₂O BSS (3 min) or in 90% D₂O BSS (3 min) were assayed as undepolarized controls. Three groups were depolarized with high-K⁺ concentration and treated with D₂O. The first group (H₂O) was stimulated with 30 mM K⁺ H₂O BSS (3 min) without any D₂O treatment. The second group (D₂O) was stimulated with 30 mM K⁺/90% D₂O BSS for 3 min. The third group (D₂O wash out) was treated with 90% D₂O BSS for 3 min and then depolarized with 30 mM K⁺ H₂O BSS for 3 min during D₂O wash out.

Radioimmunoassays of ß-endorphin in these samples were then performed by researchers fully blind to the sample conditions. The antiserum was prepared in a rabbit by several biweekly injections of ß-endorphin emulsified in complete Freund's adjuvant. The ß-endorphin was labeled using the chloramines-T method and assayed as described previously (Furui et al., 1984). Briefly, the incubation buffer was mixed with 100 mg/ml silicic acid and vortexed vigorously for 5 min. After centrifugation at 3000 rpm at 4°C for 5 min, the pellet was rinsed twice with an equal volume of ice-cold distilled water. The pellet was then extracted with 80% acetone in 0.1 N HCl via vigorous mixing for 5 min and centrifuged at 3000 rpm for 5 min. The supernatant was decanted and evaporated under a flow of nitrogen. The extract was dissolved in 0.2 ml of assay standard buffer (0.1 M phosphate buffer, pH 7.5, containing 0.5% bovine serum albumin and 500 U/ml aprotinin). Standard and samples were each brought to a total volume of 0.25 ml with the addition of standard buffer to achieve the appropriate dilution of antiserum. After incubation for 72 h at 4°C, ¹²⁵I-labeled ß-endorphin was added and the mixture was incubated for 24 h at 4°C. The ¹²⁵I-ß-endorphin was precipitated by the addition of 0.1 ml 0.1% rabbit gamma globulin, 0.5 ml 25% polyethylene glycol, and 0.1 ml standard buffer, and then centrifuged at 3000 rpm for 5 min. The radioactivity of the pellets was used as a quantitative estimation of the net ß-endorphin concentrations.

The concentration of ß-endorphin measured from each sample (4 ml) after 14–16 h was used to normalize experimental variations among flasks. One measurement from the H₂O group (n = 6) and one from the D₂O-pretreated group (n = 6) were rejected according to Smirnoff's normality test (5% confidence). Duncan's multiple range test after a one way analysis of variance (ANOVA) was used for the comparison among the groups. A Student's t-test was used for statistical comparison unless otherwise noted.

Quantum chemical method
Ab initio calculations were carried out based on the Kohn-Sham density function theory (Kohn and Sham, 1965) using the Becke3-Lee-Yang-Parr (B3LYP) function (Becke, 1993; Lee et al., 1988) and the 6-31G(d,p) basis set. Among available functions, the B3LYP function has been widely used in density-function-theory-based calculations and proven to yield very good descriptions of the vibration frequencies of small organic molecules. Harmonic vibration frequencies were calculated via analytic second derivatives of the energy with respect to the nuclear coordinates. We calculated the zero-point vibration energy of specific amino acids and water molecules, and evaluated the changes in the zero-point vibration energy that accompany the substitution of hydrogen atoms with deuterium atoms. No scaling factor for the zero-point vibration energy was used. All calculations were carried out using Gaussian 98 software (Frisch et al., 1998) on an AlphaSationXP1000 (Compaq Computer K.K., Tokyo, Japan) workstation.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Effects of D₂O on the cytoskeleton
To investigate the effects of D₂O on the cytoskeleton, we examined rhodamine phalloidin labeling of actin filaments and immunostaining of ß-tubulin (Fig. 1). Actin filaments were observed as filamentous structures that were found to be more highly concentrated in the cell processes than in cell somas (Fig. 1, A and B, a). Treatment with 90% D₂O BSS for 5 min immediately increased the amount of actin filaments in the cell somas and eliminated its filamentous structure (Fig. 1 A). The relative amount of actin filaments in the cell processes was decreased with D₂O treatment (Fig. 1 B, b). Longer treatment with 90% D₂O BSS (15 min) resulted in an increase in the actin filaments in the cell processes again, however, the filamentous structure was still lacking (Fig. 1 A). After D₂O washout, the actin filaments were equally distributed in the cell processes and cell somas (Fig. 1 B, c). After D₂O washout, the filamentous structure of the actin filaments recovered, but the shape of the filaments in the cell processes was similar to that of varicose filaments (Fig. 1, A and B, c). The original shape of the actin filaments did not recover by 30 min after D₂O washout. Despite the marked changes in the distribution and structure of actin filaments, the amount, distribution, and structure of ß-tubulin (Fig. 1 A) or neurofilament-M (data not shown) were not changed by D₂O treatment.

View larger version (58K): [in this window] [in a new window]   FIGURE 1  (A) Effects of 90% D2O treatment on actin filaments (rhodamine phalloidin labeling, red) and ß-tubulin (FITC-labeling, green) in AtT20 pituitary cells. Short (5 min) treatment with D2O increased the amount of actin filaments in the cell somas and decreased it in the cell processes. After a 15-min treatment with D2O, however, the amount of actin filaments in the cell processes increased. After D2O washout, actin filaments were equally distributed in the cell soma and processes. Short and long treatment with D2O also eliminated the filamentous structure of actin filaments, which returned immediately (<5 min) after D2O wash out, but resembled varicose filaments. D2O treatment had only a small effect on the amount, distribution, or structure of ß-tubulin. The typical changes in the distribution of actin filaments (marked as red frame a–c) are enlarged in B. Bar = 50 µm in A, 20 µm in B.

View larger version (16K): [in this window] [in a new window]   FIGURE 2  Effects of D2O on spontaneous action potential firing and voltage-gated Na+ channels. (A) Spontaneous action potentials were inhibited in 90% D2O BSS. This effect was observed in four separate cells. (B) Example of voltage-gated Na+ currents activated before, during, and after application of 90% D2O. The cell was held at -83 mV and the membrane voltage stepped to -13 mV. Note that the peak of the Na+ current was delayed compared to the control currents. (C) A current-voltage curve (average of three cells) showing the effects of the D2O solution on the Na+ current amplitude over a range of depolarizing voltage steps.

Effects of D₂O on spontaneous Ca²⁺ oscillations
Spontaneous Ca²⁺ oscillations were observed in approximately half the cells examined (389 of 743 cells, number of experiments = 16). The oscillation frequency varied widely (0.1–0.01 Hz). Cells were treated with 90% D₂O BSS for 90 s, and this resulted in a gradual inhibition of spontaneous Ca²⁺ oscillations for 4–6 min (Fig. 3 A). Longer treatment with 50% D₂O BSS (3 min; Fig. 3 A) or with lower concentrations of D₂O had little effect on the Ca²⁺ oscillations. Treatment with TTX (1 µM, for 10 min) had little effect on Ca²⁺ oscillations (n = 3), indicating that the Ca²⁺ oscillations were not directly driven by the action potentials.

View larger version (38K): [in this window] [in a new window]   FIGURE 3  Effects of D2O on spontaneous Ca2+ oscillations. (A) Treatment with 90% D2O inhibited spontaneous Ca2+ oscillations in AtT20 cells. The time course of intracellular Ca2+ concentration in three cells (Cells 1–3 as black, red, and green lines) that are marked in the transmitted light (TL) image on the top. Pseudocolor images on the top showing the changes in the intracellular Ca2+ concentration. Increasingly warmer colors indicate higher Ca2+ concentrations. Approximately half of the cells in this culture plate exhibited spontaneous Ca2+ oscillations, which were inhibited in 90% D2O BSS. The TL images before and after D2O treatment revealed that there was little morphological change. Treatment with 50% D2O BSS partially inhibited spontaneous Ca2+ oscillations. (B–D) The Ca2+ oscillations were repeatedly activated by 120-s exposure to caffeine (2 mM; white bars). (B) A Ca2+-ATPase inhibitor, thapsigargin (2 µM), inhibited neither spontaneous Ca2+ oscillations nor caffeine-induced Ca2+ oscillations. (C) Dantrolene (30 µM), an inhibitor for the ryanodine receptor, inhibited both spontaneous and caffeine-induced Ca2+ oscillations. It was not possible to wash out dantrolene during the recording period. (D) The 90% D2O solution reversibly inhibited spontaneous and caffeine-induced Ca2+ oscillations. All experiments were repeated in at least four separate trials.

Effects of D₂O on high-K⁺-induced membrane depolarization and Ca²⁺ influx
AtT20 cells that exhibited no spontaneous action potentials or infrequent Ca²⁺ oscillations were used in the following experiments. Exposure to 30 mM K⁺ depolarized the membrane potential from -69.0 ± 3.2 mV to -32.0 ± 3.6 mV (n = 4, Fig. 4, A and B). The resting membrane potential slightly depolarized (+4.4 ± 0.6 mV, n = 4) but high-K⁺-induced membrane depolarization was not significantly changed with treatment with 90% D₂O BSS (n = 3, Fig. 4 A). In addition, it was observed that VSCCs were not a primary determinant of the resting membrane potential, because blocking VSCCs with 10 µM nifedipine and 100 nM -conotoxin MVIIC failed to reduce the membrane potential (n = 3, Fig. 4 B). The K⁺ currents recorded at holding membrane potential of +50 mV (mean amplitude = 703 ± 42 pA) was reduced immediately by 90% D₂O BSS perfusion (-37.5 ± 6.7%, p < 0.01 by a paired Student's t-test, n = 4) and recovered to the control levels 8–9 min after D₂O washout (Fig. 4 C).

View larger version (18K): [in this window] [in a new window]   FIGURE 4  Effects of 90% D2O BSS on high-K+-induced membrane depolarization and K+ channel conductance. (A) Perfusion of 30 mM K+ for 45 s (open bars) evoked a rapid membrane depolarization. Broken line denotes the peak membrane potential before D2O treatment. This response was not significantly altered by treatment with 90% D2O (black bar). (B) A mixture of voltage-sensitive Ca2+ channel blockers (VSCCsX, gray bar) had no effect on the high-K+-induced membrane depolarization. Similar responses were observed in four individual cells. (C) The K+ current activated by depolarizing voltage steps was partially reduced by application of 90% D2O BSS (black bar). The cells were held at -60 mV and the membrane voltage stepped to +50 mV to observe K+ currents. The mean current amplitude ± SE was calculated from four cells. Broken line denotes the mean current amplitude before D2O treatment.

View larger version (34K): [in this window] [in a new window]   FIGURE 5  The effects of D2O on the high-K+-induced Ca2+ influx in AtT20 cells. (A) 30 mM K+ (open bars) induced a rapid, reproducible (7-min intervals) increase in intracellular Ca2+ concentration. (B) High-K+-induced Ca2+ influx was significantly inhibited by a mixture of voltage-sensitive Ca2+ channel blockers (VSCCsX). (C) Treatment with 50% D2O BSS had no effect on the high-K+-induced Ca2+ influx. (D) 90% D2O BSS inhibited the high-K+-induced Ca2+ influx only during D2O washout. (E) Only 1 min treatment with 90% D2O BSS was sufficient to inhibit several subsequent high-K+-induced Ca2+ responses triggered during D2O washout. (F) Longer treatment with 90% D2O BSS (up to 17 min) did not significantly inhibit the high-K+-induced Ca2+ influx triggered during treatment, but did result in a prolonged washout period during which high-K+-induced Ca2+ influx was inhibited. The time course of changes in intracellular Ca2+ concentrations from four individual cells was plotted in all frames.

View larger version (24K): [in this window] [in a new window]   FIGURE 6  The effects of D2O on voltage-sensitive Ca2+ channels were studied by voltage-clamp recordings of Ba2+ and Ca2+ currents. (A) The current-voltage curves for Ba2+ currents (average of seven cells) showing the little effects of 90% D2O on the Ba2+ current amplitude over a range of depolarizing voltage steps. Example of Ba2+ currents activated by depolarizing voltage steps applied before and during bath perfusion with 90% D2O is shown on the top. These Ba2+ currents were recorded by the whole-cell patch clamp method with internal solution of H2O. (B) The current-voltage curves for Ca2+ currents (average of 3–8 cells) showing the significant modulation of current-voltage relationship when Ca2+ currents were recorded with internal solution of 90% D2O. Note that inward currents at positive holding membrane potentials were significantly reduced with this recording condition. The current-voltage relationship of Ca2+ currents was recovered by the additional bath perfusion of 90% D2O-BSS. Example of Ca2+ currents is shown on the top as above in A.

Effects of D₂O on ß-endorphin release
Treatment with 90% D₂O BSS increased the release of ß-endorphin slightly (126.6 ± 20.6% of H₂O control), but not to a statistically significant degree (p = 0.11; Fig. 7). Exposure to 30 mM K⁺ resulted in a significantly greater release of ß-endorphin compared with baseline levels both in H₂O BSS (138.0 ± 8.6% of H₂O control, n = 5, p < 0.05) and 90% D₂O BSS (163.0 ± 24.3% of D₂O control, n = 6, p < 0.05). The high-K⁺-induced ß-endorphin release was inhibited during washout of 90% D₂O BSS (115.6 ± 9.2% of H₂O control, n = 5, p > 0.05).

View larger version (16K): [in this window] [in a new window]   FIGURE 7  Effects of D2O on ß-endorphin release from AtT20 cells. ß-Endorphin release was increased in 90% D2O BSS compared to H2O BSS. Exposure to 30 mM K+ increased ß-endorphin release in both H2O and 90% D2O BSS. Treatment with 90% D2O BBS for 3 min inhibited the high-K+-induced increase in ß-endorphin release, not during the treatment itself but during the D2O washout. The concentration of ß-endorphin released overnight (16 h) under control conditions in each flask was used to normalize the variations among flasks. *p < 0.05 by Student's t-test in comparison with corresponding control groups. Among three high-K+-treated groups, concentration of ß-endorphin was significantly higher in D2O than other two groups (F2,13 = 4.86, #p < 0.05 by Duncan's multiple range test after one way ANOVA).

Here we assume simple exchange reactions:

(A·A)–OH + D₂O (A·A)–OD + DOH (for Ser, Thr, and Tyr)

(A·A)–NH + D₂O (A·A)–ND + DOH (for Asn, Gln, Lys, Arg, and His).

These data show that a single deuterium substitution lowered the zero-point vibrational energy by 2.10 ± 0.02 kcal/mol and a dual deuterium substitution doubled the ZPVE for several amino acids examined. Compared to a bond to hydrogen, a deuterium-containing bond has lower zero point energy. The activation energy is greater and the rate of the reaction is correspondingly less for the deuterated compounds. Within the Born-Oppenheimer approximation, deuterated and nondeuterated molecules are electronically identical, thus isotope effects arise primarily from the ZPVE. These data indicate that for hydrophilic amino acids examined, the isotopic substitution of hydrogen by deuterium may be preferred over that of deuterium by hydrogen, which may result in slower deuterium to hydrogen exchange in these amino acids.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
This study examined the effects of D₂O treatment on the physiological processes underlying ß-endorphin release from AtT20 cells, and demonstrated several clear changes due to exposure to D₂O. First, changes that were observed during the actual treatment with D₂O were i), dynamic redistribution of actin filaments, ii), defilamentation of actin filaments, iii), partial inhibition of voltage-activated Na⁺ and K⁺ currents, iv), inhibition of spontaneous action potentials and Ca²⁺ oscillations, and v), increased high-K⁺-induced ß-endorphin release. During D₂O treatment, no changes were observed in i), the distribution of ß-tubulin, ii), high-K⁺-induced Ca²⁺ flux, and iii), high-K⁺-induced membrane depolarization. Therefore, the most probable mechanism to enhance ß-endorphin release within parameters analyzed here is a dynamic change in actin filaments. Second, more specific effects of D₂O were observed soon after D₂O treatment, during the washout of D₂O BSS, and the return to H₂O BSS. These changes were i), complete inhibition of high-K⁺-induced Ca²⁺ flux, and ii), inhibition of high-K⁺-induced ß-endorphin release. This delayed effect of D₂O cannot be explained by solvent isotope effects, and thus it is reasonable to consider the purely isotope exchange effects in VSCCs and/or their regulatory proteins.

Cellular responses under D₂O exposure
Although classical interpretations of D₂O-induced inhibition of cellular secretion have been based on D₂O-mediated stabilization of microtubules (Malaisse-Lagae et al., 1971; Montague et al., 1976; Hill and Rhoten, 1983), the effects of D₂O on the cytoskeleton appear to be even more prevalent and varied. Zimmermann et al. (1988) reported that D₂O induces the redistribution of filamentous actin and a change in morphology of human neutrophil granulocytes. More recently, Omori et al. (1997) examined the effects of D₂O on actin filaments using fluorescent staining of BALB 3T3 cells and observed a transient increase in actin filaments. In addition, they examined the effects of D₂O on actin polymerization in vitro and demonstrated that D₂O accelerates purified globular actin polymerization immediately upon exposure to D₂O, whereas the total amount of polymerized actin later during treatment with D₂O is the same as that in H₂O. Therefore, one possible mechanism underlying D₂O-induced actin filament redistribution may involve the hydrogen to deuterium exchange in globular proteins (Hermans and Scheraga, 1959; Scheraga, 1960), which results in more stable protein structures (Sing and Wood, 1969; Karasz and Gajnos, 1976), and thus is caused by a typical isotope exchange effect.

The results of this study indicate that among several cytoskeletal components, D₂O affected actin filaments specifically. The effect of D₂O on the amount of actin filaments was transient (<15 min), which is consistent with Omori et al. (1997). Although we cannot exclude the possible action of D₂O on microfilaments with a longer application of D₂O, the results of our study indicate that it is possible to affect actin filaments selectively with our experimental protocol in AtT20 cells. In addition, we found a decrease in actin filaments in the cell processes immediately upon exposure to D₂O in AtT20 cells. Secretory vesicles are transferred to the cell processes for exocytosis in AtT20 cells (Tooze and Burke, 1987). The disassembly of the actin barrier at the plasma membrane is thought to be essential for the fusion of secretory vesicles (Pfeiffer et al., 1985; Koffer et al., 1990; Lejen et al., 2001; Wilson et al., 2001); therefore, D₂O-induced disassembly of the actin barrier may be the one possible mechanism to facilitate vesicle fusion, and ß-endorphin release. Therefore, D₂O-induced increases in secretion observed in this study and in various other cell types (Bennett et al., 1979; Sulowska and Wyczolkowska, 1991; Fewtrell and Metzger, 1980; Maeyama et al., 1986; Urata et al., 1989) may be explained in part by actin-mediated mechanisms.

The D₂O-induced reduction in spontaneous action potentials and slight hyperpolarization of the resting potential both rapidly returned to control values upon D₂O washout. The direct action of D₂O on Na⁺ channel gating has been demonstrated previously in Myxicola giant axons (Schauf and Bullock, 1979; Schauf and Chuman, 1986). Consistently, our study also demonstrated slow activation and inactivation of Na⁺ channel kinetics during D₂O exposure in AtT20 cells. Since Na⁺ channel activity is directly modulated by actin polymerization (Mironov and Richter, 1999; Negulyaev et al., 2000), it is also possible that D₂O indirectly effects on Na⁺ channels via actin filaments. In addition, a reduction in the mobility of Na⁺ in D₂O buffer is known as the solvent isotope effects (Bass and Moore, 1973). Therefore, both isotope exchange effects and solvent isotope effects may mediate the D₂O effects on spontaneous action potentials.

During D₂O exposure, we also found partial and reversible inhibition of spontaneous Ca²⁺ oscillations in AtT20 cells. Application of TTX did not inhibit spontaneous Ca²⁺ oscillations, indicating that these Ca²⁺ oscillations were not caused by voltage-dependent mechanisms. Since spontaneous Ca²⁺ oscillations were activated by caffeine and inhibited by dantrolene, the Ca²⁺ oscillations appears to be dependent on Ca²⁺ release from ryanodine-sensitive internal Ca²⁺ stores in AtT20 cells. D₂O inhibited both spontaneous and caffeine-activated Ca²⁺ oscillations, whereas neither high-K⁺-induced Ca²⁺ influx nor voltage-dependent Ca²⁺ currents were affected during D₂O exposure. Therefore, D₂O may affect proteins regulating Ca²⁺ release from internal stores, although the specific sequence for the inhibition of spontaneous Ca²⁺ oscillations still needs to be analyzed.

In the presence of high-K⁺, exposure to D₂O increased ß-endorphin release, whereas the D₂O-induced enhancement of ß-endorphin release was not significant without high-K⁺ stimulation. Considering observed D₂O actions on physiological processes underlying ß-endorphin secretion, two interpretations are possible: i), the modulation of actin filaments alone is not sufficient to increase ß-endorphin release, or ii), the inhibition of spontaneous action potentials and Ca²⁺ oscillations during D₂O treatment masked a concomitant D₂O-induced, actin-mediated increase in ß-endorphin release. However, the identification of primary D₂O action for secretory activities during D₂O exposure is difficult in living cells, since various solvent isotope effects and isotope exchange effects are simultaneously produced during D₂O exposure and many of these effects are interrelated.

Cellular responses during D₂O washout
During D₂O washout, the high-K⁺-induced Ca²⁺ influx was completely inhibited for several minutes and gradually recovered (Fig. 5). In contrast, the high-K⁺-induced membrane depolarization of the membrane potential was not significantly affected during or after D₂O exposure and agreed well with that predicted by the Nernst equation for K⁺ (Fig. 4 A). Although K⁺ channel conductance estimated by the voltage-clamp recordings was partially reduced upon D₂O exposure, no further reductions in K⁺ currents were observed upon D₂O washout (Fig. 4 C). Therefore, complete inhibition of the high-K⁺-induced Ca²⁺ influx upon D₂O washout appears to involve VSCCs themselves and not via the effects on K⁺ channels. Because secretion from AtT20 cells depends on Ca²⁺ dynamics that are regulated largely by VSCCs in AtT20 cells (Richardson and Schonbrunn, 1981; Richardson, 1983), it is reasonable to assume that the inhibition of high-K⁺-induced ß-endorphin release may be associated with the reduction of the high-K⁺-induced Ca²⁺ influx.

To further clarify the effects of D₂O on VSCCs, we examined Ba²⁺ and Ca²⁺ current recordings by the voltage clamp electrophysiology. Interestingly, with the conventional whole-cell patch-clamp recordings with an electrode pipette filled with H₂O internal solution, we could not observe any significant reductions in the current amplitudes during or after D₂O exposure (Fig. 6 A). However, when the D₂O was dialyzed into the cells by the patch electrode, the current-voltage relationship of Ca²⁺ currents was modulated with a significant decrease in the inward currents (Fig. 6 B). The results suggest that asymmetric distribution of D₂O (intracellular) and H₂O (extracellular) across the plasma membrane is critical for the inhibition of VSCCs. Since intracellular D₂O concentration in intact cells is higher than the extracellular D₂O concentration during D₂O washout, the inhibition of high-K⁺-induced Ca²⁺ influx may be explained by the asymmetric distribution of D₂O that was produced exclusively during D₂O washout.

Andjus et al. (1994) reported that asymmetric distribution of D₂O (extracellular) and H₂O (intracellular) across the plasma membrane of freshwater alga activated Ca²⁺ channels in these cells. This effect of D₂O may involve direct activation of Ca²⁺ channels, because the open channel probability of Ca²⁺ channels reconstituted in lipid bilayers is also increased in D₂O. The authors suggested that transient osmotic-like stress produced by the more rapid transmembrane diffusion of D₂O may mediate the activation of Ca²⁺ channels (Brooks, 1937). However, we could not find any cell shape changes during and after the D₂O exposure (Fig. 3), and thus the involvement of stress-sensitive mechanisms in the high-K⁺-induced Ca²⁺ influx can be ruled out. In addition, solvent isotope effects cannot explain the delayed D₂O effects on high-K⁺-induced Ca²⁺ influx. Therefore, replacement of hydrogen with deuterium in constitutive proteins of VSCCs or in their regulatory proteins asymmetrically across plasma membrane may be the most considerable mechanism for the modulation of VSCCs.

Despite the significant inhibition of high-K⁺-induced Ca²⁺ influx during D₂O washout, the same was not observed when the cells were treated in the presence of 90% D₂O BSS, even with long incubation periods (up to 17 min), suggesting that the inhibition of Ca²⁺ flux was not associated with the onset of exposure to D₂O or the duration of D₂O treatment but with the return to H₂O. Consistently, Ca²⁺ currents recorded with equable D₂O substitutions in intracellular and extracellular solutions displayed normal current-voltage relationship (Fig. 6 B). These results suggest that the Ca²⁺ conductance of VSCCs is greatly affected in the intermediate state of isotope exchange effects when the constitutive proteins of VSCCs or their regulatory proteins contain asymmetrically deuterated amino acids during D₂O washout, but not in the steady state when amino acids are completely deuterated during D₂O treatment in the constitutive proteins of VSCCs or in their regulatory proteins (models in Fig. 8).

View larger version (25K): [in this window] [in a new window]   FIGURE 8  (A) Schematic illustration of changes in the zero-point vibrational energy when hydrogen is exchanged for deuterium in hydrophilic amino acid residues of proteins. Solid curve denotes the electronic energy of the system. Single deuterium substitution for hydrogen reduces the binding energy 2.1 kcal/mol (ZPVE) in hydrophilic amino acids. The total reduction of vibrational energy reduction in functional proteins depends on the number of deuterated amino acids replaced. (B) For the energy differences between protonated and deuterated amino acids, the replacement of H2O buffer with D2O buffer immediately converts protonated amino acids to deuterated amino acids, whereas deuterated amino acids slowly convert to protonated amino acids after replacement of D2O buffer with H2O buffer. The difference of exchange rates produces long-lasting transient states with asymmetrically deuterated amino acids. The increase in the high-K+-induced ß-endorphin release is in phase with the deuterated amino acids, and the most probable cause is due to the loss of actin filament barriers in the cell processes, although this effect coincides with many other effects including solvent isotope effects. The decrease in the high-K+-induced ß-endorphin release is in phase with asymmetrically deuterated amino acids, and the most probable cause is the inhibition of voltage-sensitive Ca2+ channels.

There are many known sites and mechanisms of action for D₂O on cellular secretion. Both D₂O-induced inhibition and activation of secretion have been reported in many different cell types. The results of this study demonstrated biphasic and bidirectional effects of D₂O on ß-endorphin release that may be linked to the effects of D₂O on actin filaments and VSCCs. Since the D₂O concentration is only 1/6700 of H₂O in our environment, hydrogen/deuterium exchange effects on natural living systems may be negligible. However, the results answered the complex D₂O actions on cellular secretions in experimental models that were controversial for decades.

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
This work was supported in part by a grant for Research for the Future Program (96L00310 to T.Y.) from Japan Society for the Promotion of Science, a grants-in-aid for scientific research (B14380372) by the Ministry of Education, Culture, Sports, Science and Technology, Japan to M.I., and a grant from the National Institutes of Health (NS036607) to C.N.A.

	FOOTNOTES

 
Tohru Yoshioka's present address is Dept. of Molecular Neurobiology, School of Science and Engineering, Shillman Hall, Room 601, Waseda University, 169-0072 Japan.

Submitted on December 5, 2002; accepted for publication September 10, 2003.

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