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Noradrenaline acting on α1-adrenoceptor mediates REM sleep deprivation-induced increased membrane potential in rat brain synaptosomes
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Neurochemistry International 52 (2008) 734–740 www.elsevier.com/locate/neuint
Noradrenaline acting on a1-adrenoceptor mediates REM sleep deprivation-induced increased membrane potential in rat brain synaptosomes Gitanjali Das, Birendra Nath Mallick * School of Life Sciences, Jawaharlal Nehru University, New Delhi 110067, India Received 22 June 2007; received in revised form 1 September 2007; accepted 5 September 2007 Available online 11 September 2007
Abstract We hypothesized that one of the functions of REM sleep is to maintain brain excitability and therefore, REM sleep deprivation is likely to modulate neuronal transmembrane potential; however, so far there was no direct evidence to support the claim. In this study a cationic dye, 3,30 diethylthiacarbocyanine iodide was used to estimate the potential in synaptosomal samples prepared from control and REM sleep deprived rat brains. The activity of Na–K–ATPase that maintains the transmembrane potential was also estimated in the same sample. Further, the roles of noradrenaline and a1-adrenoceptor in mediating the responses were studied both in vivo as well as in vitro. Rats were REM sleep deprived for 4 days by the classical flower-pot method; large platform and recovery controls were carried out in addition to free-moving control. The fluorescence intensity increased in samples prepared from REM sleep deprived rat brain as compared to control, which reflected synaptosomal depolarization after deprivation. The Na–K–ATPase activity also increased in the same deprived sample. Furthermore, both the effects were mediated by noradrenaline acting on a1-adrenoceptors in the brain. This is the first direct evidence showing that REM sleep deprivation indeed increased neuronal depolarization, which is the likely cause for increased brain excitability, thus supporting our hypothesis and the effect was mediated by noradrenaline acting through the a1-adrenoceptor. # 2007 Elsevier Ltd. All rights reserved. Keywords: DiSC2; Excitability; Membrane depolarization; Na–K–ATPase; Noradrenaline; REM sleep deprivation; Synaptosomes
Rapid eye movement (REM) sleep plays a crucial role in the development and sustenance of the central nervous system as evidenced by its presence across evolution at least from birds to mammals (Amlaner and Ball, 1994; Zepelin, 1994; Siegel et al., 1999) and its decrease with age (Gaudreau et al., 2005). REM sleep is reported to decrease in several diseases including Alzheimer’s, Parkinson’s (Montplaisir et al., 1995; Gagnon et al., 2002) and in several psychiatric disorders (Vogel, 1999). Its loss has been associated with several signs and symptoms like increased anxiety, aggression, irritability, confusion, loss of concentration (for review see Gulyani et al., 2000) and increased sensitiveness to tactile stimuli where subjects tend to flinch, jump and squeal (Kushida et al., 1989). REM sleep loss has been reported to affect mood; behavior and threshold for electroconvulsive shock in both animal and human subjects
* Corresponding author. Tel.: +91 11 26704522; fax: +91 11 26717558. E-mail address: [email protected] (B.N. Mallick). 0197-0186/$ – see front matter # 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.neuint.2007.09.002
(Kushida et al., 1989; Gulyani et al., 2000; Clark, 2005). Recent studies have shown that longer duration of REM sleep deprivation (REMSD) results in morphological changes in neurons (Majumdar and Mallick, 2005) and neuronal death (Biswas et al., 2006; Cordova et al., 2006). Based on these altered behavioral and physiological changes we hypothesized that REMSD alters brain excitability and as a corollary we proposed that one of the functions of REM sleep is to maintain the threshold of neuronal and brain excitability as well as responsiveness (Mallick et al., 1994); however, its mechanism of action was unknown. A reciprocal relationship exists between the level of excitability and membrane potential in neurons, where the latter is the cause and an estimate of the former. The higher the positivity of the intracellular potential, greater is the excitability level and lower is the threshold of responsiveness of the neuron. As a possible mechanism of action for such changes in excitability we showed that REMSD increases (Gulyani and Mallick, 1993) the activity of the neuronal membrane bound
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enzyme Na–K–ATPase, a key factor that maintains transmembrane potential. On the other hand, REMSD increases the level of noradrenaline (NA) in the brain (for review see Pal et al., 2005). However, no direct study was available to show the effect of REMSD on membrane potential of neurons, which might be responsible for the REMSD-induced altered excitability and its relationship with the increased level of NA. Technical limitation of recording transmembrane potential of the same neuron in vivo in freely behaving animals, before and after a reasonable period of REMSD, is a major hurdle for such study. Hence, in this in vivo and in vitro studies we estimated the membrane potential using a cationic dye and correlated it with Na–K–ATPase activity in the same synaptosomal fraction from the control and REMSD rat brains. Furthermore, we correlated the effect of NA in the presence and absence of its antagonist on those parameters. 1. Methodology 1.1. Materials Noradrenaline (NA); a1-adrenoceptor antagonist, prazosin (PRZ); b-adrenoceptor antagonist, propranolol (PRN); a2adrenoceptor agonist, clonidine (CLN); Na–K–ATPase inhibitor, ouabain; mono-cationic fluorescent probe, 3,3diethylthiacarbocyanine iodide (DiSC2) and dimethyl sulfoxide (DMSO) were procured from Sigma–Aldrich, USA. All other chemicals were of analytical grade. 2. Animals All the animal experimental protocols were approved by the Institutional Animal Ethics Committee. Male inbred Wistar rats (250–280 g), supplied with food and water ad libitum, maintained at 12/12 h light/dark cycle were used in this study. Standard flower-pot method was used for 4 days REMSD as reported earlier (Gulyani and Mallick, 1993). In brief, the rats were maintained on small (6.5 cm) platform raised over surrounding water. To rule out non-specific effects control rats were maintained under identical conditions for the same period in the same room on raised larger (13 cm) platform (LPC) surrounded by water. Free-moving home cage rats were used as normal control (FMC) for the baseline value. Another control set included rats deprived of REM sleep for 4 days that were then allowed to recover from REM sleep loss for 3 days in their normal home cages, the recovery control (REC). Thus, in each set there was one rat each of FMC, LPC, REMSD and REC and five such sets of studies were carried out. Additionally, in separate series of experiments intraperitoneal injection of PRZ and CLN were done in five different sets, each having one FMC and one REMSD rats. For in vitro experiments synaptosomes prepared from the FMC rat brain were treated with NA alone or in the presence of either PRZ or PRN. Since PRN did not prevent the NA-induced effects in in vitro studies, it was not used for the in vivo studies. Thus, every effort was made to minimize the use of number of rats and finally, data from 45 rats are presented in the study.
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2.1. Synaptosome preparation The deprived as well as the control rats were decapitated after cervical dislocation. The brains were quickly removed and homogenized in 10 volumes of ice-cold buffer containing 0.32 M sucrose and 12 mM Tris at pH 7.4 for synaptosome preparation from the whole brain (Mallick and Adya, 1999). In brief, the brain homogenate was centrifuged for 5 min at 3000 g and the supernatant was further centrifuged at 11,000 g for 20 min. The pellet obtained was suspended in 1 ml of the homogenizing buffer, loaded onto a preformed sucrose density gradient of 1.2 M and 0.8 M and ultracentrifuged in a swing-out rotor at 100,000 g for 2 h. The band obtained at the interface of 1.2 M and 0.8 M sucrose was taken as synaptosome, which was divided into two parts. One part was re-suspended in the homogenizing buffer while the other part was re-suspended in the HEPES buffer containing 140 mM NaCl, 5 mM KCl, 5 mM NaHCO3, 1 mM MgCl26H2O, 1.2 mM Na2HPO4, 10 mM glucose, 20 mM HEPES at pH 7.4. The former was used for assaying Na–K–ATPase activity while the latter for synaptosomal membrane potential study. The protein concentration in the synaptosomes was estimated by the Lowry’s method (1951) using bovine serum albumin as standard. 2.2. Modulation of membrane potential and Na–K–ATPase activity by NA The studies were conducted both in vivo and in vitro conditions. Both membrane potential as well as Na–K–ATPase activities were estimated from the same synaptosomal fraction of FMC, LPC, REMSD and REC rat brains. For the in vivo experiments either PRZ (4 mg/kg) or CLN (0.1 mg/kg) was injected (i.p.) into both the FMC and REM sleep deprived rats 8 h before sacrifice. For the in vitro studies synaptosomes prepared from the FMC rat whole brain were incubated with NA (100 mM) alone or in the presence of either PRZ (50 mM) or PRN (50 mM) and membrane potential as well as Na–K– ATPase activities estimated. 2.3. Estimation of membrane potential DiSC2 is a membrane permeable mono-cationic fluorescent dye, which accumulates within the synaptosomes that resembles the inside of a neuron and therefore is relatively negative (compared to the extracellular compartment). It has been convincingly shown that DiSC2 movement inside the cells is directly proportional to the intracellular negativity (Wang et al., 2001). The anions inside the synaptosomes bind to DiSC2 and consequently quench its fluorescence. Hence, greater the intracellular negativity more was the DiSC2 quenching, resulting in reduced net fluorescence intensity. Thus, intensity of fluorescence was inversely proportional to the intracellular anion concentration and as a corollary increased DiSC2fluorescence intensity meant membrane depolarization (Waggoner, 1976; Hare and Atchison, 1992). Therefore synaptosomal (reflection of intracellular) membrane potential was estimated
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by measuring the intensity of the cationic fluorescent probe DiSC2 as described previously by Perkinton and Sihra (1998) and was expressed as fluorescence units/50 mg synaptosomes. In brief, the DiSC2 stock solution was made in DMSO and working solution was prepared fresh every time by diluting the stock solution in the HEPES buffer such that the final concentration of the dye loaded in the synaptosomes was of 4 mM. The control and REMSD synaptosomes (50 mg) were separately but simultaneously incubated with DiSC2 (4 mM) for 10 min at 37 8C, following which the reaction was terminated by addition of ice-cold HEPES buffer and the volume was made up to 2 ml. The DiSC2-loaded synaptosomes were stored on ice till the fluorescence intensity was measured using a Cary Eclipse Spectrofluorimeter (Varian, Palo Alto, USA) with the filters set at 646 nm and 674 nm for excitation and emission, respectively. 2.4. Estimation of Na–K–ATPase activity The Na–K–ATPase activity was estimated as reported earlier (Gulyani and Mallick, 1993). Synaptosomes (40 mg) were incubated with the reaction buffer containing 100 mM NaCl, 20 mM KCl, 5 mM MgCl2, 3 mM ATP and 50 mM Tris at pH 7.4 in the presence and absence of 1 mM ouabain (blocker of Na–K–ATPase) at 37 8C for 15 min. After 15 min, the reaction was stopped by addition of 1 ml of 10% ice-cold trichloroacetic acid. ATP was used as the substrate and the concentration of the liberated inorganic phosphate was estimated by the Fiske and SubbaRow (1925) method. Thus, the ouabain sensitive ATPase activity was estimated and expressed as mM Pi released/ mg protein h. 3. Statistics Data has been presented as mean S.E.M. Statistical analysis was carried out using SigmaStat 3.5 software (Jandel Scientific, San Rafael, CA, USA). Differences between the mean values from each experimental and treated group compared with FMC and Ctl groups were evaluated using analysis of variance (ANOVA-one way) coupled with Student– Newman–Keuls test. The p-value less than 0.05 were considered significant. 4. Results 4.1. Effect of REMSD on synaptosomal potential The DiSC2-fluorescence intensity in the synaptosomal samples prepared from FMC, LPC, REMSD and REC rats were 0.38 0.05; 0.37 0.05; 0.69 0.05 and 0.49 0.06 fluorescence units/50 mg protein, respectively. The increased fluorescence value in the REMSD sample reflected relative (compared to control) depolarized state of the synaptosomes. The REMSD value was significantly high compared to both FMC (F = 16.30, d.f. = 4, p < 0.01) and LPC (F = 18.72, d.f. = 4, p < 0.01) synaptosmes, which however returned to the baseline level after recovery of REM sleep. The DiSC2-
Fig. 1. Percent changes in DiSC2-fluorescence intensity and Na–K–ATPase activity in synaptosomes prepared from LPC, REMSD and REC rat brain as compared to FMC (taken as 100%) are shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text.
fluorescence intensities of LPC (F = 0.01, d.f. = 4, p = 0.90) and REC (F = 1.65, d.f. = 4, p = 0.23) were comparable to FMC (Fig. 1). 4.2. Effect of REMSD on synaptosomal Na–K–ATPase activity The Na–K–ATPase activity in synaptosomes of FMC, LPC, REMSD and REC were 12.60 0.92; 13.16 1.63; 18.86 0.86 and 11.38 1.65 mM Pi released/mg protein h, respectively. REMSD significantly increased the synaptosomal Na–K–ATPase activity (F = 23.28, d.f. = 4, p < 0.01) as compared to the FMC. The enzyme activities in LPC (F = 0.08, d.f. = 4, p = 0.78) and REC (F = 0.40, d.f. = 4, p = 0.57) samples were also comparable to that of the FMC (Fig. 1). 4.3. Effect of NA and its antagonist in vivo 4.3.1. Fluorescence intensity As mentioned above, REMSD increased the DiSC2fluorescence intensity; however, the increased intensity was prevented by intraperitoneal injection of PRZ (0.46 0.08 fluorescence units/50 mg protein; F = 0.79, d.f. = 4, p = 0.39, compared to FMC) and CLN (0.24 0.02 fluorescence units/ 50 mg protein; F = 4.19, d.f. = 4, p = 0.07, compared to FMC) into the REM sleep deprived rats. Such injections into the FMC control rats (0.34 0.08 fluorescence units/50 mg protein; F = 0.09, d.f. = 4, p = 0.77 for PRZ and 0.33 0.05 fluorescence units/50 mg protein; F = 0.38, d.f. = 4, p = 0.55 for CLN) were ineffective in modulating the fluorescence (Fig. 2a). 4.3.2. Na–K–ATPase activity Intraperitoneal injection of PRZ (14.22 0.55 mM Pi released/mg protein h; F = 2.68, d.f. = 4, p = 0.14, compared to FMC) and CLN (14.22 0.36 mM Pi released/mg protein h; F = 2.10, d.f. = 4, p = 0.19, compared to FMC) prevented the
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Fig. 2. (a) DiSC2-fluorescence intensity in synaptosomal samples prepared from FMC and REMSD rats with and without i.p. injection of PRZ and CLN was estimated. The percent changes in the intensities as compared to FMC (without any injection) value taken as 100%, are shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text. (b) Na–K– ATPase activity in synaptosomes prepared from FMC and REMSD rat brain with and without i.p. injection of PRZ and CLN was estimated. The percent changes in activities as compared to FMC (without any injection) value taken as 100%, have been shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text.
REMSD-induced increase in Na–K–ATPase activity (see above). However, PRZ (11.71 0.99 mM Pi released/mg protein h; F = 0.43, d.f. = 4, p = 0.53) or CLN (10.31 1.64 mM Pi released/mg protein h; F = 2.06, d.f. = 4, p = 0.19) alone was ineffective in altering the enzyme activities in the FMC rats (Fig. 2b). 4.4. Effect of NA and its antagonist in vitro 4.4.1. Fluorescence intensity Incubation of the FMC synaptosomes with NA (100 mM) increased DiSC2-fluorescence intensity to 0.73 0.07 fluorescence units/50 mg protein (F = 15.04, d.f. = 4, p < 0.01) compared to the untreated control (Ctl) samples (0.38 0.05 fluorescence units/50 mg protein). Further, such NA-induced
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Fig. 3. (a) DiSC2-fluorescence intensity was estimated in samples containing rat brain synaptosomes (Ctl) and after in vitro incubation of samples with NA alone as well as in the presence of either PRZ or PRN. The percent changes in intensity compared to Ctl value taken as 100%, are shown (N = 5). **p < 0.01, significant as compared to Ctl. Abbreviations: as in the text. (b) Percent changes in rat brain synaptosomal Na–K–ATPase activities after in vitro incubation of samples with NA alone as well as in the presence of either PRZ or PRN are shown (N = 5). **p < 0.01, significant as compared to Ctl. Abbreviations: as in the text.
increase in DiSC2-fluorescence intensity was prevented by preincubating the synaptosomes with PRZ (0.43 0.03 fluorescence units/50 mg protein; F = 0.59, d.f. = 4, p = 0.46, compared to Ctl) but not with PRN (0.66 0.04 fluorescence units/50 mg protein; F = 14.10, d.f. = 4, p < 0.01, compared to Ctl) (Fig. 3a). 4.4.2. Na–K–ATPase activity Incubation of the synaptosomes with NA (100 mM) significantly increased the Na–K–ATPase activity (20.46 1.42 mM Pi released/mg protein h; F = 21.28, d.f. = 4, p < 0.01) compared to its untreated Ctl samples (12.60 0.92 mM Pi released/mg protein h). This NA-induced increase in the enzyme activity was prevented by pre-incubating the synaptosomes with PRZ (13.33 1.31 mM Pi released/ mg protein h; F = 0.21, d.f. = 4, p = 0.66, compared to Ctl) but not by PRN (19.87 1.34 mM Pi released/mg protein h; F = 19.76, d.f. = 4, p < 0.01, compared to Ctl) (Fig. 3b).
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5. Discussion Transmembrane potential provides an overview of the state of affairs of a neuron, which reflects its physiological condition. It may modulate directly or indirectly a variety of neuronal functions including signaling, neurotransmitter release and neurogenesis (Choi, 1988; Hochner et al., 1989; Deisseroth et al., 2004). The depolarization of neurons brought about either by an increased influx of positive ions or efflux of the negative ions, is associated with neuronal excitability (Deisseroth et al., 2004). It is known that abnormal alteration in the ionic flux may release apoptotic factors like cytochrome c thereby activating the caspase cascade which ultimately may lead to its death (Moon et al., 2005). The Na–K–ATPase plays a key role in maintaining the transmembrane potential by an efflux of three Na+ in exchange for two K+ influx (Trachtenberg et al., 1981; Horisberger et al., 1991). Various neurotransmitters also modulate the neuronal transmembrane potential by regulating the cellular ion homeostasis either directly by activating ion channels or indirectly through various signaling molecules. NA is one such neurotransmitter that has been shown to modulate membrane depolarization in neurons (Pan et al., 1994). During REMSD, the level of NA increases in the brain because (i) the NA-ergic REM-OFF neurons, which normally cease activity during REM sleep, continue being active (Mallick et al., 1989); (ii) there is increased synthesis (Sinha et al., 1973; Majumdar and Mallick, 2003) and decreased breakdown (Thakkar and Mallick, 1993) of the NA synthesizing and hydrolyzing enzymes, respectively during REMSD; (iii) REM sleep is reduced if those neurons are continuously activated (Singh and Mallick, 1996) or not allowed to cease activity (Kaur et al., 2004); (iv) NA levels increase after REMSD (Porkka-Heiskanen et al., 1995); and (v) we have confirmed that the REMSD associated increased Na–K– ATPase activity was indeed due to the elevated levels of NA in the brain (Gulyani and Mallick, 1995; Mallick et al., 2000, 2002). Increased levels of NA bring about various cellular changes including alterations in biochemical, physiological and molecular processes, resulting in disturbed homeostasis that has far reaching effects on the behavior of a subject (Kopp et al., 1982; Heaney et al., 1999). The results of this study showed that upon REMSD there was increased DiSC2-fluorescence intensity compared to that of the FMC. This suggested that the neurons tended to remain in a relative depolarized state, thereby reducing its threshold for excitation. Intraperitoneal injection of PRZ and CLN, which blocks the NA action and release, respectively, prevented the REMSD-induced depolarization in vivo. Further, incubation of the synaptosomes with NA in vitro increased the DiSC2fluorescence, which however, was prevented by PRZ but not by PRN. Based on these observations we proposed that the REMSD-induced increased depolarization was mediated by NA acting through the a1-adrenoceptor. Simultaneously, in the same synaptosomal sample we observed that REMSD increased Na–K–ATPase activity and the effect was also mediated by NA acting on a1-adrenoceptor as reported by us earlier (Gulyani and Mallick, 1995; Mallick et al., 2000).
Thus, the fact that both Na–K–ATPase activity and synaptosomal potential were increased after REMSD in the same sample and both were mediated by NA acting on a1adrenoceptor, suggest that the two processes are linked phenomena. Independent studies have shown that NA depolarizes a membrane due to a net efflux of the chloride anions (Lamb and Barna, 1998), thereby activating the voltagedependent sodium channels (Takahashi et al., 1999). Thus, REMSD-induced elevated NA increases intracellular positivity possibly by an efflux of negative ions resulting in membrane depolarization. We have shown under similar conditions that REMSD stimulated the chloride-sensitive Mg-ATPase pump (Mallick and Gulyani, 1993), which is known to extrude the negative chloride ions (Shiroya et al., 1989), support the findings. The REMSD-induced NA mediated membrane depolarization then activates the voltage-dependent sodium channels (Catterall, 1992) to cause a net influx of sodium ions, which also adds to the increase in excitable state of the neuron. As the intracellular level of Na+ increases, the neuron tries to compensate and balance the Na+ overload inside the cell (Takahashi et al., 1999) by activating the Na–K–ATPase, which extrudes three Na+ in lieu of two K+ influx (Trachtenberg et al., 1981; Horisberger et al., 1991). Notwithstanding, both in vivo and in vitro experiments have shown that the REMSD-induced increased NA increases the Na–K–ATPase activity (Gulyani and Mallick, 1995; Mallick et al., 2000). Thus, the NA-induced stimulation of Na–K–ATPase may be a direct effect or secondary to efflux of anions. Although it is difficult to confirm if one mechanism follows the other, we suspect both the mechanisms exist and they may be activated under different conditions including various diseases. Therefore, the etiology and progression of various diseases may be different, though the symptoms expressed may be similar. Notwithstanding, the increased Na–K–ATPase activity alters the release of various neurotransmitters (Vizi et al., 1982) which maybe the cause of symptoms expressed during REMSD and associated disorders. The rats were REM sleep deprived by the flower-pot method, the most preferred method of choice globally for such studies. Notwithstanding, to rule out the effects of non-specific factors including stress-induced effects, we carried out other standard LPC and REC control experiments. Earlier we have shown under similar conditions that the increased Na–K–ATPase activity was neither due to increased muscular activity nor due to movement restriction on the small platform (Gulyani and Mallick, 1993). Although there may be some loss of non-REM sleep on the small platform, it is comparable to the LPC rats, at least after more than 48 h deprivation (Mendelson et al., 1974). Electrophysiological recordings have shown that this method indeed induces selective REM sleep loss and the effects were unlikely due to stress (Porkka-Heiskanen et al., 1995). Besides, the arguments in favor of the observed effects being specific to REMSD, as discussed earlier, hold true for this study as well (Vogel, 1975; Gulyani et al., 2000). We showed earlier that REMSD increases Na–K–ATPase activity (Gulyani and Mallick, 1993, 1995) as well as its expression (Majumdar et al., 2003) on neurons in different regions of the brain as well as in the whole brain; hence, we
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carried out this study in the whole brain synaptosomes. However, there are various types of neurons in the brain, e.g., excitatory and inhibitory, hence it is possible that the responses of some neurons may differ and also the effects would depend on the duration of deprivation. This view may be supported by the finding in slice preparation that although the dentate gyrus granule neurons were not affected, the hippocampal CA1 neurons showed reduced firing after REMSD (McDermott et al., 2003). In related in vivo study it has been shown that the firing of REM-OFF neurons decreased, while that of REM-ON neurons increased after REMSD (Mallick et al., 1989), while, other neurons responded to lower intensity of auditory stimulation (Mallick et al., 1991). Therefore, it is worth studying the threshold and excitability of neurons in different brain areas and specifically on neurochemically identified neurons. Though this method has been used widely for synaptosomal potential studies, it has a few limitations. The DiSC2 dye might accumulate in different organelles or it might remain bound to the synaptosomal membrane, thereby affecting the final fluorescence intensity. However, since we have studied relative changes in intensity, these are likely to be comparable in the control samples, unless there is any specific reason to believe that REMSD alters the synaptosomal membrane property affecting its interactions with DiSC2. In conclusion, we had proposed that REMSD alters brain excitability but direct evidence was lacking. Although it was known that REMSD increases NA that stimulates Na–K– ATPase activity, we did not know how the enzyme gets activated. It was also known that intracellular positivity stimulates the enzyme. In this study, we observed that indeed the intracellular positivity is increased after REMSD and this is mediated by NA acting through a1-adrenoceptor. Thus, the results of this study confirm and provide direct evidence in support of our hypothesis that REMSD increases brain excitability and that is mediated by at increased level of NA. Acknowledgements Research was supported by funding from Indian Council of Medical Research and University Grants Commission, India. GD received research fellowship from Council of Scientific and Industrial Research, India. References Amlaner, C.J.J., Ball, N.J., 1994. Avian sleep. In: Kryger, M.H., Roth, T., Dement, W.C. (Eds.), Principles and Practice of Sleep Medicine. WB Saunders Company, Philadelphia, pp. 81–94. Biswas, S., Mishra, P., Mallick, B.N., 2006. Increased apoptosis in rat brain after rapid eye movement sleep loss. Neuroscience 142, 315–331. Catterall, W.A., 1992. Cellular and molecular biology of voltage-gated sodium channels. Physiol. Rev. 72, 15–48. Choi, D.W., 1988. Calcium-mediated neurotoxicity: relationship to specific channel types and role in ischemic damage. Trends Neurosci. 11, 465– 469. Clark, C.P., 2005. Mood changes. In: Kushida, C.A. (Ed.), Sleep Deprivation: Basic Science, Physiology and Behavior, vol. 192. Marcel Dekker, New York, pp. 415–420.
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Mallick, B.N., Thakkar, M., Gulyani, S., 1994. Rapid eye movement sleep deprivation induced alteration in neuronal excitability: possible role of norepinephrine. In: Mallick, B.N., Singh, R. (Eds.), Environment and Physiology. Narosa Publishing House, New Delhi, India, pp. 196–203. Mallick, B.N., Adya, H.V., Faisal, M., 2000. Norepinephrine-stimulated increase in Na+, K+-ATPase activity in the rat brain is mediated through alpha1A-adrenoceptor possibly by dephosphorylation of the enzyme. J. Neurochem. 74, 1574–1578. Mallick, B.N., Majumdar, S., Faisal, M., Yadav, V., Madan, V., Pal, D., 2002. Role of norepinephrine in the regulation of rapid eye movement sleep. J. Biosci. 27, 539–551. McDermott, C.M., LaHoste, G.J., Chen, C., Musto, A., Bazan, N.G., Magee, J.C., 2003. Sleep deprivation causes behavioral, synaptic, and membrane excitability alterations in hippocampal neurons. J. Neurosci. 23, 9687– 9695. Mendelson, W.B., Guthrie, R.D., Frederick, G., Wyatt, R.J., 1974. The flower pot technique of rapid eye movement (REM) sleep deprivation. Pharmacol. Biochem. Behav. 2, 553–556. Montplaisir, J., Petit, D., Lorrain, D., Gauthier, S., Nielsen, T., 1995. Sleep in Alzheimer’s disease: further considerations on the role of brainstem and forebrain cholinergic populations in sleep–wake mechanisms. Sleep 18, 145–148. Moon, Y., Lee, K.H., Park, J.H., Geum, D., Kim, K., 2005. Mitochondrial membrane depolarization and the selective death of dopaminergic neurons by rotenone: protective effect of coenzyme Q10. J. Neurochem. 93, 1199– 1208. Pal, D., Madan, V., Mallick, B.N., 2005. Neural mechanism of rapid eye movement sleep generation: cessation of locus coeruleus neurons is a necessity. Acta Physiol. Sin. 57, 401–413. Pan, Z.Z., Grudt, T.J., Williams, J.T., 1994. Alpha 1-adrenoceptors in rat dorsal raphe neurons: regulation of two potassium conductances. J. Physiol. 478, 437–447. Perkinton, M.S., Sihra, T.S., 1998. Presynaptic GABA(B) receptor modulation of glutamate exocytosis from rat cerebrocortical nerve terminals: receptor decoupling by protein kinase C. J. Neurochem. 70, 1513–1522. Porkka-Heiskanen, T., Smith, S.E., Taira, T., Urban, J.H., Levine, J.E., Turek, F.W., Stenberg, D., 1995. Noradrenergic activity in rat brain during rapid
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Neurochemistry International 52 (2008) 734–740 www.elsevier.com/locate/neuint
Noradrenaline acting on a1-adrenoceptor mediates REM sleep deprivation-induced increased membrane potential in rat brain synaptosomes Gitanjali Das, Birendra Nath Mallick * School of Life Sciences, Jawaharlal Nehru University, New Delhi 110067, India Received 22 June 2007; received in revised form 1 September 2007; accepted 5 September 2007 Available online 11 September 2007
Abstract We hypothesized that one of the functions of REM sleep is to maintain brain excitability and therefore, REM sleep deprivation is likely to modulate neuronal transmembrane potential; however, so far there was no direct evidence to support the claim. In this study a cationic dye, 3,30 diethylthiacarbocyanine iodide was used to estimate the potential in synaptosomal samples prepared from control and REM sleep deprived rat brains. The activity of Na–K–ATPase that maintains the transmembrane potential was also estimated in the same sample. Further, the roles of noradrenaline and a1-adrenoceptor in mediating the responses were studied both in vivo as well as in vitro. Rats were REM sleep deprived for 4 days by the classical flower-pot method; large platform and recovery controls were carried out in addition to free-moving control. The fluorescence intensity increased in samples prepared from REM sleep deprived rat brain as compared to control, which reflected synaptosomal depolarization after deprivation. The Na–K–ATPase activity also increased in the same deprived sample. Furthermore, both the effects were mediated by noradrenaline acting on a1-adrenoceptors in the brain. This is the first direct evidence showing that REM sleep deprivation indeed increased neuronal depolarization, which is the likely cause for increased brain excitability, thus supporting our hypothesis and the effect was mediated by noradrenaline acting through the a1-adrenoceptor. # 2007 Elsevier Ltd. All rights reserved. Keywords: DiSC2; Excitability; Membrane depolarization; Na–K–ATPase; Noradrenaline; REM sleep deprivation; Synaptosomes
Rapid eye movement (REM) sleep plays a crucial role in the development and sustenance of the central nervous system as evidenced by its presence across evolution at least from birds to mammals (Amlaner and Ball, 1994; Zepelin, 1994; Siegel et al., 1999) and its decrease with age (Gaudreau et al., 2005). REM sleep is reported to decrease in several diseases including Alzheimer’s, Parkinson’s (Montplaisir et al., 1995; Gagnon et al., 2002) and in several psychiatric disorders (Vogel, 1999). Its loss has been associated with several signs and symptoms like increased anxiety, aggression, irritability, confusion, loss of concentration (for review see Gulyani et al., 2000) and increased sensitiveness to tactile stimuli where subjects tend to flinch, jump and squeal (Kushida et al., 1989). REM sleep loss has been reported to affect mood; behavior and threshold for electroconvulsive shock in both animal and human subjects
* Corresponding author. Tel.: +91 11 26704522; fax: +91 11 26717558. E-mail address: [email protected] (B.N. Mallick). 0197-0186/$ – see front matter # 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.neuint.2007.09.002
(Kushida et al., 1989; Gulyani et al., 2000; Clark, 2005). Recent studies have shown that longer duration of REM sleep deprivation (REMSD) results in morphological changes in neurons (Majumdar and Mallick, 2005) and neuronal death (Biswas et al., 2006; Cordova et al., 2006). Based on these altered behavioral and physiological changes we hypothesized that REMSD alters brain excitability and as a corollary we proposed that one of the functions of REM sleep is to maintain the threshold of neuronal and brain excitability as well as responsiveness (Mallick et al., 1994); however, its mechanism of action was unknown. A reciprocal relationship exists between the level of excitability and membrane potential in neurons, where the latter is the cause and an estimate of the former. The higher the positivity of the intracellular potential, greater is the excitability level and lower is the threshold of responsiveness of the neuron. As a possible mechanism of action for such changes in excitability we showed that REMSD increases (Gulyani and Mallick, 1993) the activity of the neuronal membrane bound
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enzyme Na–K–ATPase, a key factor that maintains transmembrane potential. On the other hand, REMSD increases the level of noradrenaline (NA) in the brain (for review see Pal et al., 2005). However, no direct study was available to show the effect of REMSD on membrane potential of neurons, which might be responsible for the REMSD-induced altered excitability and its relationship with the increased level of NA. Technical limitation of recording transmembrane potential of the same neuron in vivo in freely behaving animals, before and after a reasonable period of REMSD, is a major hurdle for such study. Hence, in this in vivo and in vitro studies we estimated the membrane potential using a cationic dye and correlated it with Na–K–ATPase activity in the same synaptosomal fraction from the control and REMSD rat brains. Furthermore, we correlated the effect of NA in the presence and absence of its antagonist on those parameters. 1. Methodology 1.1. Materials Noradrenaline (NA); a1-adrenoceptor antagonist, prazosin (PRZ); b-adrenoceptor antagonist, propranolol (PRN); a2adrenoceptor agonist, clonidine (CLN); Na–K–ATPase inhibitor, ouabain; mono-cationic fluorescent probe, 3,3diethylthiacarbocyanine iodide (DiSC2) and dimethyl sulfoxide (DMSO) were procured from Sigma–Aldrich, USA. All other chemicals were of analytical grade. 2. Animals All the animal experimental protocols were approved by the Institutional Animal Ethics Committee. Male inbred Wistar rats (250–280 g), supplied with food and water ad libitum, maintained at 12/12 h light/dark cycle were used in this study. Standard flower-pot method was used for 4 days REMSD as reported earlier (Gulyani and Mallick, 1993). In brief, the rats were maintained on small (6.5 cm) platform raised over surrounding water. To rule out non-specific effects control rats were maintained under identical conditions for the same period in the same room on raised larger (13 cm) platform (LPC) surrounded by water. Free-moving home cage rats were used as normal control (FMC) for the baseline value. Another control set included rats deprived of REM sleep for 4 days that were then allowed to recover from REM sleep loss for 3 days in their normal home cages, the recovery control (REC). Thus, in each set there was one rat each of FMC, LPC, REMSD and REC and five such sets of studies were carried out. Additionally, in separate series of experiments intraperitoneal injection of PRZ and CLN were done in five different sets, each having one FMC and one REMSD rats. For in vitro experiments synaptosomes prepared from the FMC rat brain were treated with NA alone or in the presence of either PRZ or PRN. Since PRN did not prevent the NA-induced effects in in vitro studies, it was not used for the in vivo studies. Thus, every effort was made to minimize the use of number of rats and finally, data from 45 rats are presented in the study.
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2.1. Synaptosome preparation The deprived as well as the control rats were decapitated after cervical dislocation. The brains were quickly removed and homogenized in 10 volumes of ice-cold buffer containing 0.32 M sucrose and 12 mM Tris at pH 7.4 for synaptosome preparation from the whole brain (Mallick and Adya, 1999). In brief, the brain homogenate was centrifuged for 5 min at 3000 g and the supernatant was further centrifuged at 11,000 g for 20 min. The pellet obtained was suspended in 1 ml of the homogenizing buffer, loaded onto a preformed sucrose density gradient of 1.2 M and 0.8 M and ultracentrifuged in a swing-out rotor at 100,000 g for 2 h. The band obtained at the interface of 1.2 M and 0.8 M sucrose was taken as synaptosome, which was divided into two parts. One part was re-suspended in the homogenizing buffer while the other part was re-suspended in the HEPES buffer containing 140 mM NaCl, 5 mM KCl, 5 mM NaHCO3, 1 mM MgCl26H2O, 1.2 mM Na2HPO4, 10 mM glucose, 20 mM HEPES at pH 7.4. The former was used for assaying Na–K–ATPase activity while the latter for synaptosomal membrane potential study. The protein concentration in the synaptosomes was estimated by the Lowry’s method (1951) using bovine serum albumin as standard. 2.2. Modulation of membrane potential and Na–K–ATPase activity by NA The studies were conducted both in vivo and in vitro conditions. Both membrane potential as well as Na–K–ATPase activities were estimated from the same synaptosomal fraction of FMC, LPC, REMSD and REC rat brains. For the in vivo experiments either PRZ (4 mg/kg) or CLN (0.1 mg/kg) was injected (i.p.) into both the FMC and REM sleep deprived rats 8 h before sacrifice. For the in vitro studies synaptosomes prepared from the FMC rat whole brain were incubated with NA (100 mM) alone or in the presence of either PRZ (50 mM) or PRN (50 mM) and membrane potential as well as Na–K– ATPase activities estimated. 2.3. Estimation of membrane potential DiSC2 is a membrane permeable mono-cationic fluorescent dye, which accumulates within the synaptosomes that resembles the inside of a neuron and therefore is relatively negative (compared to the extracellular compartment). It has been convincingly shown that DiSC2 movement inside the cells is directly proportional to the intracellular negativity (Wang et al., 2001). The anions inside the synaptosomes bind to DiSC2 and consequently quench its fluorescence. Hence, greater the intracellular negativity more was the DiSC2 quenching, resulting in reduced net fluorescence intensity. Thus, intensity of fluorescence was inversely proportional to the intracellular anion concentration and as a corollary increased DiSC2fluorescence intensity meant membrane depolarization (Waggoner, 1976; Hare and Atchison, 1992). Therefore synaptosomal (reflection of intracellular) membrane potential was estimated
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by measuring the intensity of the cationic fluorescent probe DiSC2 as described previously by Perkinton and Sihra (1998) and was expressed as fluorescence units/50 mg synaptosomes. In brief, the DiSC2 stock solution was made in DMSO and working solution was prepared fresh every time by diluting the stock solution in the HEPES buffer such that the final concentration of the dye loaded in the synaptosomes was of 4 mM. The control and REMSD synaptosomes (50 mg) were separately but simultaneously incubated with DiSC2 (4 mM) for 10 min at 37 8C, following which the reaction was terminated by addition of ice-cold HEPES buffer and the volume was made up to 2 ml. The DiSC2-loaded synaptosomes were stored on ice till the fluorescence intensity was measured using a Cary Eclipse Spectrofluorimeter (Varian, Palo Alto, USA) with the filters set at 646 nm and 674 nm for excitation and emission, respectively. 2.4. Estimation of Na–K–ATPase activity The Na–K–ATPase activity was estimated as reported earlier (Gulyani and Mallick, 1993). Synaptosomes (40 mg) were incubated with the reaction buffer containing 100 mM NaCl, 20 mM KCl, 5 mM MgCl2, 3 mM ATP and 50 mM Tris at pH 7.4 in the presence and absence of 1 mM ouabain (blocker of Na–K–ATPase) at 37 8C for 15 min. After 15 min, the reaction was stopped by addition of 1 ml of 10% ice-cold trichloroacetic acid. ATP was used as the substrate and the concentration of the liberated inorganic phosphate was estimated by the Fiske and SubbaRow (1925) method. Thus, the ouabain sensitive ATPase activity was estimated and expressed as mM Pi released/ mg protein h. 3. Statistics Data has been presented as mean S.E.M. Statistical analysis was carried out using SigmaStat 3.5 software (Jandel Scientific, San Rafael, CA, USA). Differences between the mean values from each experimental and treated group compared with FMC and Ctl groups were evaluated using analysis of variance (ANOVA-one way) coupled with Student– Newman–Keuls test. The p-value less than 0.05 were considered significant. 4. Results 4.1. Effect of REMSD on synaptosomal potential The DiSC2-fluorescence intensity in the synaptosomal samples prepared from FMC, LPC, REMSD and REC rats were 0.38 0.05; 0.37 0.05; 0.69 0.05 and 0.49 0.06 fluorescence units/50 mg protein, respectively. The increased fluorescence value in the REMSD sample reflected relative (compared to control) depolarized state of the synaptosomes. The REMSD value was significantly high compared to both FMC (F = 16.30, d.f. = 4, p < 0.01) and LPC (F = 18.72, d.f. = 4, p < 0.01) synaptosmes, which however returned to the baseline level after recovery of REM sleep. The DiSC2-
Fig. 1. Percent changes in DiSC2-fluorescence intensity and Na–K–ATPase activity in synaptosomes prepared from LPC, REMSD and REC rat brain as compared to FMC (taken as 100%) are shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text.
fluorescence intensities of LPC (F = 0.01, d.f. = 4, p = 0.90) and REC (F = 1.65, d.f. = 4, p = 0.23) were comparable to FMC (Fig. 1). 4.2. Effect of REMSD on synaptosomal Na–K–ATPase activity The Na–K–ATPase activity in synaptosomes of FMC, LPC, REMSD and REC were 12.60 0.92; 13.16 1.63; 18.86 0.86 and 11.38 1.65 mM Pi released/mg protein h, respectively. REMSD significantly increased the synaptosomal Na–K–ATPase activity (F = 23.28, d.f. = 4, p < 0.01) as compared to the FMC. The enzyme activities in LPC (F = 0.08, d.f. = 4, p = 0.78) and REC (F = 0.40, d.f. = 4, p = 0.57) samples were also comparable to that of the FMC (Fig. 1). 4.3. Effect of NA and its antagonist in vivo 4.3.1. Fluorescence intensity As mentioned above, REMSD increased the DiSC2fluorescence intensity; however, the increased intensity was prevented by intraperitoneal injection of PRZ (0.46 0.08 fluorescence units/50 mg protein; F = 0.79, d.f. = 4, p = 0.39, compared to FMC) and CLN (0.24 0.02 fluorescence units/ 50 mg protein; F = 4.19, d.f. = 4, p = 0.07, compared to FMC) into the REM sleep deprived rats. Such injections into the FMC control rats (0.34 0.08 fluorescence units/50 mg protein; F = 0.09, d.f. = 4, p = 0.77 for PRZ and 0.33 0.05 fluorescence units/50 mg protein; F = 0.38, d.f. = 4, p = 0.55 for CLN) were ineffective in modulating the fluorescence (Fig. 2a). 4.3.2. Na–K–ATPase activity Intraperitoneal injection of PRZ (14.22 0.55 mM Pi released/mg protein h; F = 2.68, d.f. = 4, p = 0.14, compared to FMC) and CLN (14.22 0.36 mM Pi released/mg protein h; F = 2.10, d.f. = 4, p = 0.19, compared to FMC) prevented the
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Fig. 2. (a) DiSC2-fluorescence intensity in synaptosomal samples prepared from FMC and REMSD rats with and without i.p. injection of PRZ and CLN was estimated. The percent changes in the intensities as compared to FMC (without any injection) value taken as 100%, are shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text. (b) Na–K– ATPase activity in synaptosomes prepared from FMC and REMSD rat brain with and without i.p. injection of PRZ and CLN was estimated. The percent changes in activities as compared to FMC (without any injection) value taken as 100%, have been shown (N = 5). **p < 0.01, significant as compared to FMC. Abbreviations: as in the text.
REMSD-induced increase in Na–K–ATPase activity (see above). However, PRZ (11.71 0.99 mM Pi released/mg protein h; F = 0.43, d.f. = 4, p = 0.53) or CLN (10.31 1.64 mM Pi released/mg protein h; F = 2.06, d.f. = 4, p = 0.19) alone was ineffective in altering the enzyme activities in the FMC rats (Fig. 2b). 4.4. Effect of NA and its antagonist in vitro 4.4.1. Fluorescence intensity Incubation of the FMC synaptosomes with NA (100 mM) increased DiSC2-fluorescence intensity to 0.73 0.07 fluorescence units/50 mg protein (F = 15.04, d.f. = 4, p < 0.01) compared to the untreated control (Ctl) samples (0.38 0.05 fluorescence units/50 mg protein). Further, such NA-induced
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Fig. 3. (a) DiSC2-fluorescence intensity was estimated in samples containing rat brain synaptosomes (Ctl) and after in vitro incubation of samples with NA alone as well as in the presence of either PRZ or PRN. The percent changes in intensity compared to Ctl value taken as 100%, are shown (N = 5). **p < 0.01, significant as compared to Ctl. Abbreviations: as in the text. (b) Percent changes in rat brain synaptosomal Na–K–ATPase activities after in vitro incubation of samples with NA alone as well as in the presence of either PRZ or PRN are shown (N = 5). **p < 0.01, significant as compared to Ctl. Abbreviations: as in the text.
increase in DiSC2-fluorescence intensity was prevented by preincubating the synaptosomes with PRZ (0.43 0.03 fluorescence units/50 mg protein; F = 0.59, d.f. = 4, p = 0.46, compared to Ctl) but not with PRN (0.66 0.04 fluorescence units/50 mg protein; F = 14.10, d.f. = 4, p < 0.01, compared to Ctl) (Fig. 3a). 4.4.2. Na–K–ATPase activity Incubation of the synaptosomes with NA (100 mM) significantly increased the Na–K–ATPase activity (20.46 1.42 mM Pi released/mg protein h; F = 21.28, d.f. = 4, p < 0.01) compared to its untreated Ctl samples (12.60 0.92 mM Pi released/mg protein h). This NA-induced increase in the enzyme activity was prevented by pre-incubating the synaptosomes with PRZ (13.33 1.31 mM Pi released/ mg protein h; F = 0.21, d.f. = 4, p = 0.66, compared to Ctl) but not by PRN (19.87 1.34 mM Pi released/mg protein h; F = 19.76, d.f. = 4, p < 0.01, compared to Ctl) (Fig. 3b).
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5. Discussion Transmembrane potential provides an overview of the state of affairs of a neuron, which reflects its physiological condition. It may modulate directly or indirectly a variety of neuronal functions including signaling, neurotransmitter release and neurogenesis (Choi, 1988; Hochner et al., 1989; Deisseroth et al., 2004). The depolarization of neurons brought about either by an increased influx of positive ions or efflux of the negative ions, is associated with neuronal excitability (Deisseroth et al., 2004). It is known that abnormal alteration in the ionic flux may release apoptotic factors like cytochrome c thereby activating the caspase cascade which ultimately may lead to its death (Moon et al., 2005). The Na–K–ATPase plays a key role in maintaining the transmembrane potential by an efflux of three Na+ in exchange for two K+ influx (Trachtenberg et al., 1981; Horisberger et al., 1991). Various neurotransmitters also modulate the neuronal transmembrane potential by regulating the cellular ion homeostasis either directly by activating ion channels or indirectly through various signaling molecules. NA is one such neurotransmitter that has been shown to modulate membrane depolarization in neurons (Pan et al., 1994). During REMSD, the level of NA increases in the brain because (i) the NA-ergic REM-OFF neurons, which normally cease activity during REM sleep, continue being active (Mallick et al., 1989); (ii) there is increased synthesis (Sinha et al., 1973; Majumdar and Mallick, 2003) and decreased breakdown (Thakkar and Mallick, 1993) of the NA synthesizing and hydrolyzing enzymes, respectively during REMSD; (iii) REM sleep is reduced if those neurons are continuously activated (Singh and Mallick, 1996) or not allowed to cease activity (Kaur et al., 2004); (iv) NA levels increase after REMSD (Porkka-Heiskanen et al., 1995); and (v) we have confirmed that the REMSD associated increased Na–K– ATPase activity was indeed due to the elevated levels of NA in the brain (Gulyani and Mallick, 1995; Mallick et al., 2000, 2002). Increased levels of NA bring about various cellular changes including alterations in biochemical, physiological and molecular processes, resulting in disturbed homeostasis that has far reaching effects on the behavior of a subject (Kopp et al., 1982; Heaney et al., 1999). The results of this study showed that upon REMSD there was increased DiSC2-fluorescence intensity compared to that of the FMC. This suggested that the neurons tended to remain in a relative depolarized state, thereby reducing its threshold for excitation. Intraperitoneal injection of PRZ and CLN, which blocks the NA action and release, respectively, prevented the REMSD-induced depolarization in vivo. Further, incubation of the synaptosomes with NA in vitro increased the DiSC2fluorescence, which however, was prevented by PRZ but not by PRN. Based on these observations we proposed that the REMSD-induced increased depolarization was mediated by NA acting through the a1-adrenoceptor. Simultaneously, in the same synaptosomal sample we observed that REMSD increased Na–K–ATPase activity and the effect was also mediated by NA acting on a1-adrenoceptor as reported by us earlier (Gulyani and Mallick, 1995; Mallick et al., 2000).
Thus, the fact that both Na–K–ATPase activity and synaptosomal potential were increased after REMSD in the same sample and both were mediated by NA acting on a1adrenoceptor, suggest that the two processes are linked phenomena. Independent studies have shown that NA depolarizes a membrane due to a net efflux of the chloride anions (Lamb and Barna, 1998), thereby activating the voltagedependent sodium channels (Takahashi et al., 1999). Thus, REMSD-induced elevated NA increases intracellular positivity possibly by an efflux of negative ions resulting in membrane depolarization. We have shown under similar conditions that REMSD stimulated the chloride-sensitive Mg-ATPase pump (Mallick and Gulyani, 1993), which is known to extrude the negative chloride ions (Shiroya et al., 1989), support the findings. The REMSD-induced NA mediated membrane depolarization then activates the voltage-dependent sodium channels (Catterall, 1992) to cause a net influx of sodium ions, which also adds to the increase in excitable state of the neuron. As the intracellular level of Na+ increases, the neuron tries to compensate and balance the Na+ overload inside the cell (Takahashi et al., 1999) by activating the Na–K–ATPase, which extrudes three Na+ in lieu of two K+ influx (Trachtenberg et al., 1981; Horisberger et al., 1991). Notwithstanding, both in vivo and in vitro experiments have shown that the REMSD-induced increased NA increases the Na–K–ATPase activity (Gulyani and Mallick, 1995; Mallick et al., 2000). Thus, the NA-induced stimulation of Na–K–ATPase may be a direct effect or secondary to efflux of anions. Although it is difficult to confirm if one mechanism follows the other, we suspect both the mechanisms exist and they may be activated under different conditions including various diseases. Therefore, the etiology and progression of various diseases may be different, though the symptoms expressed may be similar. Notwithstanding, the increased Na–K–ATPase activity alters the release of various neurotransmitters (Vizi et al., 1982) which maybe the cause of symptoms expressed during REMSD and associated disorders. The rats were REM sleep deprived by the flower-pot method, the most preferred method of choice globally for such studies. Notwithstanding, to rule out the effects of non-specific factors including stress-induced effects, we carried out other standard LPC and REC control experiments. Earlier we have shown under similar conditions that the increased Na–K–ATPase activity was neither due to increased muscular activity nor due to movement restriction on the small platform (Gulyani and Mallick, 1993). Although there may be some loss of non-REM sleep on the small platform, it is comparable to the LPC rats, at least after more than 48 h deprivation (Mendelson et al., 1974). Electrophysiological recordings have shown that this method indeed induces selective REM sleep loss and the effects were unlikely due to stress (Porkka-Heiskanen et al., 1995). Besides, the arguments in favor of the observed effects being specific to REMSD, as discussed earlier, hold true for this study as well (Vogel, 1975; Gulyani et al., 2000). We showed earlier that REMSD increases Na–K–ATPase activity (Gulyani and Mallick, 1993, 1995) as well as its expression (Majumdar et al., 2003) on neurons in different regions of the brain as well as in the whole brain; hence, we
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carried out this study in the whole brain synaptosomes. However, there are various types of neurons in the brain, e.g., excitatory and inhibitory, hence it is possible that the responses of some neurons may differ and also the effects would depend on the duration of deprivation. This view may be supported by the finding in slice preparation that although the dentate gyrus granule neurons were not affected, the hippocampal CA1 neurons showed reduced firing after REMSD (McDermott et al., 2003). In related in vivo study it has been shown that the firing of REM-OFF neurons decreased, while that of REM-ON neurons increased after REMSD (Mallick et al., 1989), while, other neurons responded to lower intensity of auditory stimulation (Mallick et al., 1991). Therefore, it is worth studying the threshold and excitability of neurons in different brain areas and specifically on neurochemically identified neurons. Though this method has been used widely for synaptosomal potential studies, it has a few limitations. The DiSC2 dye might accumulate in different organelles or it might remain bound to the synaptosomal membrane, thereby affecting the final fluorescence intensity. However, since we have studied relative changes in intensity, these are likely to be comparable in the control samples, unless there is any specific reason to believe that REMSD alters the synaptosomal membrane property affecting its interactions with DiSC2. In conclusion, we had proposed that REMSD alters brain excitability but direct evidence was lacking. Although it was known that REMSD increases NA that stimulates Na–K– ATPase activity, we did not know how the enzyme gets activated. It was also known that intracellular positivity stimulates the enzyme. In this study, we observed that indeed the intracellular positivity is increased after REMSD and this is mediated by NA acting through a1-adrenoceptor. Thus, the results of this study confirm and provide direct evidence in support of our hypothesis that REMSD increases brain excitability and that is mediated by at increased level of NA. Acknowledgements Research was supported by funding from Indian Council of Medical Research and University Grants Commission, India. GD received research fellowship from Council of Scientific and Industrial Research, India. References Amlaner, C.J.J., Ball, N.J., 1994. Avian sleep. In: Kryger, M.H., Roth, T., Dement, W.C. (Eds.), Principles and Practice of Sleep Medicine. WB Saunders Company, Philadelphia, pp. 81–94. Biswas, S., Mishra, P., Mallick, B.N., 2006. Increased apoptosis in rat brain after rapid eye movement sleep loss. Neuroscience 142, 315–331. Catterall, W.A., 1992. Cellular and molecular biology of voltage-gated sodium channels. Physiol. Rev. 72, 15–48. Choi, D.W., 1988. Calcium-mediated neurotoxicity: relationship to specific channel types and role in ischemic damage. Trends Neurosci. 11, 465– 469. Clark, C.P., 2005. Mood changes. In: Kushida, C.A. (Ed.), Sleep Deprivation: Basic Science, Physiology and Behavior, vol. 192. Marcel Dekker, New York, pp. 415–420.
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