Download Control of cytoplasmic pH under anoxic

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
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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
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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-
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