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Journal of Experimental Botany, Vol. 47, No. 300, pp. 967-973, July 1996 Journal of Experimental Botany Control of cytoplasmic pH under anoxic conditions and its implication for plasma membrane proton transport in Medicago sativa root hairs Hubert H. Felle1 Botanisches Institut I, Justus-Liebig-Universita't, Senckenbergstr. 17, D-35390 Giessen, Germany Received 15 November 1995; Accepted 19 March 1996 Abstract In root hairs of Medicago sativa, pH-sensitive microelectrodes have been applied to study cytoplasmic pH-regulation. To inhibitors like oligomycin, antimycin A, cyanide and the exchange of O2 for N2, the root hairs respond with a distinct cytoplasmic acidification. Whereas the cytoplasmic pH under aerobic conditions rests at 7.28 + 0.11 SE (n = 168), under conditions of (chemical) anoxia the cytoplasmic pH is shifted to a stable, well-regulated 6.78 ±0.08 SE (n = 81). Once this pH is attained in the presence of one inhibitor, addition of another has no effect. 2-deoxyglucose and W-acetylglucosamine, both inhibitors of glycolysis at the hexokinase level, increase cytoplasmic pH by about 0.3 pH units, as do glucogenic amino acids. It is suggested that aerobic energy metabolism does not contribute to acidosis of these cells. Since pH-shift and pump deactivation can be separated by using poor respiratory inhibitors, it is concluded that the switch from 'aerobic' to 'anaerobic' pH is not correlated with proton pump activity. Inversely, since cytoplasmic pH neither responds to pump activation by FC with alkalinization, nor to pump deactivation by cyanide with acidification, it is also concluded that changes in pump activity do not affect cytoplasmic pH. Key words: Cytoplasmic pH regulation, Medicago, pH-sensitive microelectrodes, proton transport, root hairs. Introduction In recent years, the importance of cytoplasmic pH as a messenger in intracellular signalling (Felle, 1989; Kurkdjian and Guern, 1989) and its role in regulating 1 Fax; +49 641 702 84419. Q Oxford University Press 1996 membrane transport (Johannes and Felle, 1987; Blatt, 1992) is more and more appreciated, and calls for an improved understanding of intracellular pH regulation, yet despite the many splendid reports and reviews (Smith and Raven, 1979; Roberts et al, 1984a, b; Raven, 1986; Davies, 1986; Fox and Ratcliffe, 1990; Guern et al, 1991) our understanding remains limited. In part, this may arise from the fact that pH studies are carried out on a variety of different plants which, growing under different conditions, have different strategies of pH regulation. Thus, the choice of what organisms and cells are studied and with which techniques, not only engenders the problem of oversimplification, but also encourages a certain polarity amongst investigators favouring the biophysical or biochemical pH-stat to be the predominant factor in cytoplasmic pH regulation. Since all life is based on aqueous chemistry, protons may be produced and consumed in metabolism and as such fulfil the basic requirements for a messenger between cellular reactions which otherwise have no other common link. This makes pH regulation different from any other intracellular ionic regulation process. Although cytoplasmic pH seems tightly regulated, it would be incorrect to conclude that it does not vary with time. Besides being a signal and/or a messenger, a well-controlled cytoplasmic pH is also an indication, or even a prerequisite, of underlying metabolic processes. Although evolution has developed a network of chemical and energetical transitions, in which the sum of protons produced, consumed, transferred or transported approximates zero, there are a variety of situations in a cell's life which can disturb this dynamic equilibrium. Since shortage of oxygen is a very common situation for a root (Roberts et al, 1984a), the effect of chemical and true anoxia on cytoplasmic pH of Medicago root hairs is investigated here, using direct 968 Felle probing with pH-sensitive microelectrodes. It is demonstrated that the pH under anoxia is well regulated without obvious contribution of the plasma membrane proton pump. Materials and methods General conditions Seedlings of Medicago saliva were grown for 48 h at 23 °C in a moist chamber. Excised roots or whole plantlets were mounted in a Plexiglass cuvette which was constantly perfused with the test medium. Unless stated otherwise, this comprised 0.1-1 mM KC1, 0.1 mM NaCl, and 0.1 mM CaCl2. The respective pH was adjusted by mixing 5 mM 2-(jV-morpholino)ethanesulphonic acid and 5 mM TRIS. Unless indicated otherwise, the external pH was 7.3 and adjusted with MES, wherever NaCN was used. After approximately 1 h the root hairs were adjusted to this medium. Oligomycin and antimycin A (Sigma) were predissolved in ethanol and stirred into the aqueous solution to obtain the final concentrations; ETOH concentrations never exceeded 0.1%. Electrophysiology and ion-selective Results Cytoplasmic pH under 'anaerobic' conditions (a) Chemical anoxia: Cytoplasmic pH of Medicago root hairs rests between 7.1 and 7.5 (7.28 + 0.11, n=168). Oligomycin, antimycin A and cyanide rapidly acidify the cytoplasm in a concentration-dependent manner up to 0.6 pH units (Figs 1, 2). The dose-response relationship is steep, i.e. whereas 1 /xM oligomycin has almost no effect, already 10/^M cause the maximal acidification. The new cytoplasmic pH remains stable as long as the respective inhibitor is present, but returns to its original value after removal of the inhibitor. Whereas for cyanide this recovery is rapid (Fig. 2), for oligomycin and antimycin A it may take 30 min or longer, which may be due to their lipid solubilities (Fig. 1). Depolarization and pH change can be separated: the acidification clearly precedes membrane potential changes, when oligomycin or antimycin A are added. For cyanide, this is not so evident, but can also be demonstrated for lower concen- microelectrodes The electrical set-up for the impalement of the root hairs, and the fabrication and application of the pH-sensitive microelectrodes has been described before (Felle and Bertl, 1986a). The test chamber was open on both sides for the horizontal approach of two or more separate electrodes, which was necessary for the simultaneous measurements of cytoplasmic pH and membrane potential. Since ion-selective microelectrodes pick up a mixed electrical signal, which consists of both the membrane potential difference and the free ion concentration, a separate microelectrode which measures the membrane potential, had to be inserted into the same cell. In some cases these electrodes were combined in a double barrel (Felle, 1987). The electrodes were connected with a high-impedance differential amplifier (FD 223; WPI, Sarasota, Fla. USA), which simultaneously measured and then subtracted both signals to obtain the net pH signal given in the figures. Due to the different response times of the two electrodes (the ion selective electrode being slower), this procedure may produce artefactual shifts of the pH-signal in the case of rapid changes in membrane potential. Since the pH-selective electrodes after the first impalement may alter their physicochemical properties, data were selected from electrodes which displayed satisfactory calibration after the respective test. The slope of pH-electrodes was routinely tested during the measurements inside the cells using the established reaction to weak acids. Although the membrane potential is recorded together with the cytoplasmic pH at all times, traces are only given when of particular interest to the argument. 0 to n 10 40 all Fig. 1. Cytoplasmic pH (pH c ) and membrane potential of Medicago root hairs, measured in the presence of different concentrations of oligomycin and 1 mM sodium cyanide (CN~), as indicated (arrows denote moment of addition). W = removal of both agents. Numbers on pH traces denote initial levels and at points of special interest following the external addition of the agents. ' + ' means that the prior added agent remained present. Traces are representative examples of 10 equivalent experiments. Nitrogen flushing Once the two electrodes were in place to be inserted, the open sides of the chamber were closed leaving a small aperture which only allowed minor lateral movements of the electrodes. To ensure a sufficient drop in pH of the medium, the entire chamber was jacketed and gently flushed with N2 (grade 5.5) while the N2-medium was perfused. N 2 was removed by flushing with air. tin (ail) Fig. 2. Cytoplasmic pH and membrane potential of Medicago root hairs, measured in the presence of different concentraUons of sodium cyanide (CN~) and antimycin A (AA), as indicated (arrows). W = removal of both agents. Numbers on traces, see legend to Fig. 1. Traces are representative examples of 8 equivalent experiments. pH regulation in root hairs 969 trations (Fig. 2). When oligomycin or antimycin A are added first, cyanide (in the presence of the previously added inhibitor) has either no or only a minor effect on cytoplasmic pH; however, cyanide still depolarizes the cells in the usual manner. Likewise, if cyanide is added first, no effect on cytoplasmic pH or membrane potential is observed following the addition of the other inhibitors. (b) Nitrogen: When the cells are flushed by nitrogen (Fig. 3), cytoplasmic pH rapidly drops by 0.5 units followed by a minor recovery, while the membrane potential drops to the so-called diffusion potential. Upon exchange of nitrogen by air the cells quickly recover and after about 5 min both cytoplasmic pH and membrane potential are back to normal. As long as oxygen is removed, cyanide and oligomycin have no effect on either cytoplasmic pH or membrane potential. The effect of acetic acid Weak acids with low anion lipid solubility, like acetic acid, are routinely being used to acidify the cytoplasm and clamp it to a value according to their pA^, and pH gradient across the respective membrane (Sanders and Slayman, 1982; Guern et al., 1986; Frachisse et al, 1988). In doing so, these substances may interfere with membrane transport and also with pH regulation. As expected, acetic acid at an external pH of 6.1 acidifies the cytoplasm and at the same time hyperpolarizes the cells (Fig. 4). Oligomycin, when added subsequently in the presence of acetic acid, has only an effect on cytoplasmic pH, provided the acidification induced by acetic acid did not reach the anoxia value (Fig. 4A, B). In fact, when acetic acid is washed out (while oligomycin remains), the cytoplasmic pH returns to the anoxia value after a transient increase. With antimycin A (not shown) and cyanide, equivalent effects can be demonstrated; the cyanide-induced depolarization occurs as usual. Figure 4C shows that there is a difference depending upon which order acetic acid and the inhibitors are added: when, for instance, oligomycin is added prior to acetic acid, the cytoplasm is clearly I 3 8-175 Fig. 3. Cytoplasmic pH and membrane potential of Medicago root hairs, measured before and after flushing the chamber with nitrogen (N 2 ) or air, and in the presence of 10/xM oligomycin (01) or 1 mM NaCN, respectively. Numbers on pH traces, see legend to Fig. 1. Traces are representative of 5 equivalent experiments. Fig. 4. Cytoplasmic pH and membrane potential of Medicago root hairs, measured in the presence of acetic acid (Ac), oligomycin (01) and sodium cyanide (CN~). (A) 4mM acetic acid, followed by 10 ^M oligomycin ( + O l ) a n d removal of acetic acid ( - A c ) . (B) 10 mM acetic acid, followed by lO^M oligomycin and 1 mM NaCN, respectively. (C) lOftM oligomycin, followed by 10 mM acetic acid. External pH was 6.1. Numbers on pH traces, see legend to Fig. 1. ' + ' means that the prior added agent(s) remained present. Traces are representative examples of 6 equivalent experiments. acidified by both agents, however, there is a clear tendency towards recovery after the addition of acetic acid. Pump activation and cytoplasmic pH Fusicoccin is a well-known tool to hyperpolarize the plasma membrane of higher plants, which is usually interpreted with an activation of the H + ATPase (Marre, 1979), but also seems to affect K + channels (Clint and Blatt, 1989). Since it is an important question, how cytoplasmic pH is regulated under conditions of anoxia, and it is still a matter of debate whether an activation change of the plasma membrane proton pump will alter the cytoplasmic pH (Guern et al., 1986), fusicoccin has been added in the presence of oligomycin, and as a control, without it. As shown in Fig. 5A, 1 ^M FC hyperpolarizes the plasma membrane by some 30 mV both in the presence of and without oligomycin, but does not alkalize the cytoplasm in any of the experiments. Also, in cases where the pump is deactivated (cyanide), neither a change in membrane potential nor in cytoplasmic pH (Fig. 5B) is observed. Inhibition of glycolysis 2-deoxyglucose and N-acetylglucosamine have been shown to inhibit glycolysis at the hexokinase level without 970 Felle cMtnl •=-200 IS S~iI7 without 2-deoxyglucose (Fig. 1). To 10 mM yV-acetylglucosamine the cells react in a similar way, however, the increase in pH is somewhat faster. So-called glucogenic amino acids, although not inhibitors of individual glycolytic steps, may block glycolysis through accumulation within the cytoplasm. When amino acids are added to the external medium, due to the strong inwardly directed proton motive force, they are transported into the cytoplasm by H + /amino acid symport (Jung and LQttge, 1980; Felle, 1981) and accumulated there. It is interesting to observe that the addition of alanine or serine does not acidify the cytoplasm, but results in an increase of some 0.2 to 0.3 units (Fig. 6B). Discussion Fig. 5. Cytoplasmic pH and membrane potential of Medicago root hairs, measured in the presence of ohgomycin (Ol), sodium cyanide (CN~) and fusicoccin (FC). (A) 5/xM oligomycin, followed by 1 ^M fusicoccin. Control: no oligomycin added. (B) After the addition of 1 mM cyanide, I ^M fusicoccin was added. W = removal of both agents Numbers on pH traces, see legend to Fig. 1. Traces are representative examples of 5 equivalent experiments. affecting respiration (Hochster, 1963). As shown in Fig. 6A, after a minor initial acidification, 10 mM 2-deoxyglucose slowly increases cytoplasmic pH by about 0.3 pH units. 10 jxM oligomycin, added in the presence of 2-deoxyglucose, acidifies the cytoplasm by about 0.8 pH units, which is considerably more than observed DOG 5 rain 0 5 rain Fig. 6. Cytoplasmic pH of Medicago root hairs, measured in the presence of (A) 10 mM deoxyglucose (DOG). 10 mM Ar-acetylglucosamine (NAc) and lOjiM oligomycin (Ol); (B) lOmM L-alauine (Ala) and L-serine (Ser). Numbers on pH traces, see legend to Fig. 1. Traces are representative examples of 4 equivalent experiments. The apparent ubiquity of membrane transport for the extrusion of proton equivalents from cells at first sight seems to confirm the view that the principal challenge of pH regulation is to relieve the cell of excess protons arising from proton leakage through the plasma membrane or from metabolism. As to the proton leak, no matter how protons enter the cells, be it through cotransport or other, the surplus of H"""-equivalents will sooner or later be exported. Whereas this view is widely accepted, considerable confusion seems to exist with regard to the 'proton producing' role of metabolism, a problem which has been extensively discussed and reviewed by Raven (1986). With respect to carbohydrate catabolism, it is demonstrated here, that a defined and stable acidification of the cytoplasm occurs when glycolysis is 'uncoupled' from respiratory electron transport by the poor respiratory inhibitors oligomycin and antimycin A, or when ATP synthesis of oxidative phosphorylation is inhibited using cyanide or by true anoxia. Although the inhibitors used are effective in different places, with regard to cytoplasmic pH the result is the same. It indicates that the acidifying protons probably arise from non-processed substrates, e.g. glycolytic compounds, which in turn identifies respiration as proton consuming. The alkalinization observed in the presence of the glycolytic inhibitors 2-deoxyglucose (DOG) and iV-acetylglucosamine and the subsequent acidification by oligomycin seem to support this view. Alternatively, one might argue that, under the conditions inferred, proteins are degraded leading to an increase in ammonium, which could explain some alkalinization under DOG and thus contribute to pH-regulation. Gevers (1977), and Busa and Nuccitelli (1984) pointed out that the glycolytic acidification may be due to hydrolysis of MgATP rather than to the production of lactic acid; if this were the case, steady-state aerobic metabolism would not contribute to the acidification of the cytoplasm as long as ATP hydrolysis and respiration are well coupled, and the cell or organism may influence cytoplasmic pH by hydrolysing more or less ATP or by shifting equilibria pH regulation in root hairs and thus the poise of the individual reactions of glycolysis and oxidative phosphorylation, respectively. However, a cytoplasmic concentration of MgATP of 0.1-0.2 mM stands against a buffer cytoplasmic capacity of about 30-50 mM per pH unit, which calls for an analysis of the ATP-turnover. The cytoplasmic pH under (chemical) anoxia is well defined and controlled The data presented here clearly indicate that acidification in the presence of oligomycin, antimycin A and cyanide does not proceed with time and, within the error margins, point to a well-defined value of 6.78 + 0.08 SE (« = 81; all inhibitors), a pH which may be necessary to cope in situations of severe natural hypoxia or anoxia, e.g. flooding. In fact, this acidification never exceeded 0.6 pH units, regardless of the type or concentration of the respective inhibitors used. Similar values were reported for Neurospora (Sanders and Slayman, 1982) and for Riccia (Felle and Bertl, 1986ft) using cyanide, by Roberts et al. (1984a, b) and Fox et al. (1995) in maize roots replacing oxygen for nitrogen. Therefore, it appears that for a variety of organisms this pH shift is common, reflecting the difference in cytoplasmic pH from aerobic to anaerobic metabolism, although Menegus et al. (1991) find that, under anoxia, rice seedlings only acidify by 0.4 units, while wheat acidifies by 0.8 units. As the aerobic pH, this 'anaerobic' pH is very stable, an observation which was not necessarily to be expected, recalling that under these conditions glycolytic products are not processed and hence accumulate within the cytoplasm. This pH stability is confirmed while manipulating the cytoplasmic pH with acetic acid (Fig. 4). Whereas the cells reacted to acetic acid as expected with acidification and hyperpolarization, addition of the inhibitors (in the presence of acetic acid) did not acidify the cytoplasm further in situations where the pH value had already approximated the anoxia value (Fig. 4B), but did, when the acidification was less (Fig. 4A)! These observations also indicate that the equilibria between the reactions which usually determine the cytoplasmic pH must have already been shifted in a way that inhibition of oxygen consumption or oxidative phosphorylation could only contribute as much as was left to reach the anoxia level. Also, the transient pH increase after removal of acetic acid (Fig. 4A) and the partial pH recovery in the presence of oligomycin and acetic acid (Fig. 4C) points in the same direction. Although not all pH changes were found to be strictly numerically additive, within an error margin of + 0.1 pH unit they fit the idea that the pH shift reflecting the switch from aerobic to anaerobic metabolism is indeed intrinsically well-defined and controlled. Roberts et al. (1989) and Fox et al. (1995) provide evidence that ethanol production is correlated with these 971 changes in cytoplasmic pH and argue that acidification is the signal to trigger the switch from lactate to ethanol production under anoxia by simultaneously inhibiting lactate dehydrogenase and activating pyruvate decarboxylase. The term 'switch' seems warranted because full acidification occurs within a rather narrow inhibitor concentration range. Although levels of glycolytic products have not been performed in these root hairs, the data given in the literature (Roberts et al., 1989; Fox et al., 1995) warrant the suggestion that, besides this switch to ethanol production, other processes must take place simultaneously to stabilize cytoplasmic pH under anoxia. For instance, upon increasing concentrations of lactate, malate and some amino acids (Roberts et al, 1992), the tendency for gluconeogenesis increases, i.e. the glycolysis within the known restrictions runs backwards and thus consumes protons. This possibility, also discussed by Sanders and Slayman (1982) for Neurospora, is supported by the finding that uptake of glucogenic amino acids (alanine, serine) leads to cytoplasmic alkalinization (Fig. 6B), an observation which has also been made in rhizoids of Riccia fluitans (Johannes and Felle, 1987). Cytoplasmic acidosis is not the result of pump deactivation Whereas at first sight the cyanide and nitrogen experiments appear to implicate that pump deactivation and acidification are in a causal relationship, the data shown in Figs 1 and 2 indicate that for oligomycin and antimycin A and for low cyanide concentrations this is not the case. In fact, experiments carried out in the presence of oligomycin and fusicoccin clearly demonstrate that the proton pump is still active enough to hyperpolarize the plasma membrane to values more negative than the resting potential (Fig. 5). This means that in the presence of oligomycin or antimycin A (not shown) energy levels are still sufficiently high to drive primary active membrane transport under the given experimental conditions. It proves that the anaerobic acidification is not a consequence of pump deactivation, e.g. caused by 'proton leakage', but clearly is an early event in the inhibition of respiration. Inversely, the cytoplasmic pH obviously is not influenced by pump activity (Felle, 1991), a view which is supported by experiments with fusicoccin (Fig. 5), and with cyanide in the presence of oligomycin (Fig. 1) or antimycin A (Fig. 2). When is a pH change registered as an error signal? Whereas a cytoplasmic acidification caused by external factors will lead to an immediate boost in pump activity (Fig. 4), as demonstrated in the past by several laboratories (Guern et al., 1991), evidence has just been brought forward that a metabolically based pH change seemingly has no such obvious effect on the pump. It seems, therefore, that not all changes or perturbations of the 972 Felle cytoplasmic pH are registered as an error signal, although certain transporters will nota bene have to react according to their basic responsiveness to protons. This is physiologically meaningful, but difficult to explain: pH perturbations coming from outside (external pH jumps, weak acids/bases) or even from chloroplasts after light/dark changes (Felle and Bertl, 1986A; Thaler et al, 1992) are quickly reacted to by the various short-term regulating units and as such are part of the biophysical pH-stat (Smith and Raven, 1978; Felle, 1987, 1988). Metabolically based pH changes may precede or are essential preconditions for a change in cellular activity of some sort, e.g. for growth stimulation or for a reaction to a microbial attack. Such a pH change is of course not to be countered and reversed, i.e. an override signal or command must exist to prevent a reaction of the pump or any other regulatory units. Whereas one may speculate on the nature of such a signal, it should be clear that the pump per se can not distinguish pH changes coming from different sources, i.e. its sensitivity towards protons must be modulated. In principle, such a regulation could be accomplished by a regulator protein. Since in the presence of FC the pump appears deregulated, i.e. it works with full power, the so-called FC receptor (Aducci et al, 1995; DeBoer and Korthout, 1995) would be a likely candidate. Alternatively, recalling the stable cytoplasmic pH under hypoxia, it seems obvious that the pump activity always reflects the metabolic status of the cell, which through shifting chemical equilibria may set the cytoplasmic pH. In such a network the pump would be at the periphery of pH regulation and under metabolic control. Acknowledgements This work was supported gemeinschaft. by the Deutsche Forschungs- References Aducci P, Marra M, Fogliano V, Fullone MR. 1995. 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