Relationship between the membrane potential of the contractile vacuole complex and its osmoregulatory activity in Paramecium multimicronucleatum
Pacific Biomedical Research Center, Snyder Hall 306, University of
Hawaii, 2538 The Mall, Honolulu, HI 96822, USA
* Present address: Department of Biology, University of Oslo, PO Box 1051,
Blindern, N-0316 Oslo, Norway
Present address: Physiological Institute, University of Würzburg,
Röntgenring 9, D-97070 Würzburg, Germany
Author for correspondence (e-mail:
naitoh{at}pbrc.hawaii.edu)
Accepted 18 July 2002
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Summary |
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Key words: contractile vacuole complex, membrane potential, V-ATPase, osmoregulation, fluid segregation activity, exocytotic cycle, Paramecium multimicronucleatum
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Introduction |
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Recently we described the periodic changes in the electrical potential that
exist across the CV membrane. These changes accompany the periodic exocytotic
activity of the CV (Tominaga et al.,
1998). A CV potential of approximately 80 mV, positive with
reference to the cytosol, was seen only when the CV membrane was attached to
the radial arms during the CV's fluid-filling phase. We therefore proposed
that the CV potential originates from the electrogenic activity of V-ATPases
in the membranes of the decorated spongiome that are concentrated along the
radial arms.
The primary objective of the present paper was to clarify the relationship
between the CV's electrical potential, V-ATPase activity and the rate of fluid
segregation in the CVC, RCVC. We examined the CV potential
under various external osmotic conditions and in the presence of inhibitors.
We found that the CV potential was directly correlated with the number of
functional V-ATPase complexes present in the CVC. RCVC,
however, was not directly correlated with this number. We propose that the
V-ATPase of the decorated spongiome provides energy for the translocation of
ions into the CVC lumen, thus setting up and maintaining the osmotic gradient
that, according to Stock et al.
(2002), allows cytosolic water
to enter the CVC lumen by osmosis.
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Materials and methods |
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Experimental solutions
Each experimental solution contained (mmoll-1 final
concentration) 2.0 KCl, 0.25 CaCl2 and 1.0 MopsKOH, pH 7.0.
The osmolarity of the experimental solution was adjusted by adding different
amounts of sorbitol to the solution while its ionic components remained
unchanged. Osmolarity was measured using a freezing-point depression osmometer
(Micro-Osmometer, Model MO plus, Advanced Instruments, Inc., Norwood, MA,
USA). The osmolarity of the solution without sorbitol was approximately 4
mosmoll-1. The osmolarities of the other experimental solutions
employed were 24, 64 and 124 mosmoll-1.
Inhibitor-containing solutions were prepared by diluting dimethylsulfoxide
(DMSO)-dissolved inhibitors (stock solutions) with the 4 mosmoll-1
experimental solution. The inhibitor solutions were (i) 30 nmoll-1
(final concentration) concanamycin B, an inhibitor of V-ATPase
(Woo et al., 1992) (a gift
from Dr K. Miwa, Central Research Laboratories, Ajinomoto Co., Japan;
[DMSO]<0.01% v/v), (ii) 40 µmoll-1 ethoxyzolamide (EZA), an
inhibitor of carbonic anhydrase (Deitmer
and Schlue, 1989
) (Sigma Chemical Co., St Louis, MO, USA;
[DMSO]
0.1% v/v) and (iii) 1 mmoll-1 furosemide, an inhibitor of
ion transport in the renal system and in erythrocyte membranes
(Brater, 1998
;
Lauf, 1984
) (Sigma Chemical
Co., St Louis, MO, USA; [DMSO]
0.1% v/v). Each inhibitor solution was
prepared immediately before experimentation.
Capturing a cell
An adapted cell surrounded by a minute amount of an experimental solution
was introduced into a droplet of mineral oil on a coverslip. Some of the
experimental solution surrounding the cell was removed through a micropipette
to prevent cell movement. The cell was then further arrested by inserting a
glass microneedle into it.
Recordings of data
Electrical signals
The tip of a glass capillary microelectrode filled with 3 moll-1
KCl (resistance approximately 70 M) was inserted into the cytosol. A
minute amount of the same experimental solution was then reintroduced into the
solution surrounding the cell through a micropipette to keep the cell from
becoming unduly compressed.
A fine-tipped glass capillary microelectrode filled with 3 mol
l-1 KCl (resistance approximately 120 M) was inserted into
the CV to measure the electrical potential difference between the CV fluid and
the cytosol. An approximately 20 ms electrical oscillation of the head
amplifier that was connected to the fine-tipped microelectrode was necessary
for successful insertion of the electrode into the CV. This electrode also
served to inject square-wave electric current pulses (0.2 nA, 100 ms duration)
into the CV to determine its input resistance. The input resistance was
determined by dividing the potential shift due to injection of a current pulse
by the current intensity. A glass microcapillary with a tip diameter of
approximately 30 µm was filled with a 2% agar-based experimental solution
(resistance approximately 1 M
) and placed into the surrounding solution
and grounded.
Two electrical signals, one from the electrode inserted into the cytosol (corresponding to the plasma membrane potential) and the other from the electrode inserted into the CV (corresponding to the sum of two potentials, one across the CV membrane and the other across the plasma membrane) were fed into a computer (Power Macintosh 7600/132, Apple Computer Inc. Cupertino, CA, USA) through an A/D-D/A converter (ITC-16; Instrutech Corp., Great Neck, NY, USA). These electrical signals were used to obtain the potential difference between the CV fluid and the cytosol (the CV potential).
Images of the CV
Images of the CV obtained using Nomarski optics (x40 objective on a
model DMIRB inverted microscope; Leica Inc. Deerfield, IL, USA) were
continuously recorded on VHS videotape using a video cassette recorder (AG
6300, Panasonic Industrial Co., Secaucus, NJ, USA) through a CCD camera
(CCD-72, Dage MTI Inc., Michigan City, IN, USA).
Software
Software for feeding the electrical signals into the computer, for
processing the signals and for generating the current pulses required for
experimentation was developed on the basis of IgorPro (WaveMetrix, Inc., Lake
Oswego, OR, USA) and PulseControl XOP software packages
(Herrington et al., 1995).
Adobe Photoshop 5 was used to analyse images of the CVC.
Measurement of RCVC
The experimental chamber, with an approximate volume of 20 µl, was first
filled with a solution of 0.02% (v/v) poly-L-lysine (Sigma, St Louis, MO,
USA). Cells suspended in an experimental solution were introduced into the
chamber at one end, while the poly-L-lysine solution was removed from the
chamber at the other end by absorption with filter paper. Cells that adhered
to the bottom surface of the chamber were subjected to further solution
exchange and video-recorded for determining RCVC
(Stock et al., 2001).
On the replayed images of the CV, we measured the time between two successive fluid discharges (T) and the maximum diameter of the spherical CV immediately before the start of fluid discharge (Dmax). RCVC was calculated by dividing the maximum volume of the CV estimated from Dmax by T and is given in fl s-1
Fluorescent microscopy of the decorated spongiome in the CVC
For examination of the CVC by fluorescence microscopy, formaldehyde-fixed
(3% in 50 mmoll-1 phosphate buffer, pH 7.4) and cold (-20°C)
acetone-permeabilized cells were treated using a monoclonal antibody raised
against the decorated spongiome (DS-1)
(Allen et al., 1990). This was
followed by treatment with fluorescein-isothiocyanate (FITC)-conjugated rabbit
anti-mouse IgG. Unbound antibody was washed away using excess buffer solution.
The cells were observed using a Zeiss microscope equipped with epifluorescence
illumination and a filter appropriate for FITC (B-2E Nikon). Photographs were
obtained using Kodak Tri-X film. Values in the text are presented as means
± S.E.M. (N).
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Results |
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The input resistance of the CV (Fig.
1A; filled circles) was approximately 160 M during the
rounding phase and decreased to approximately 60 M
during the
fluid-filling phase. It was approximately 30 M
during the
fluid-discharging phase.
When a cell was mechanically compressed against the coverslip by removing excess experimental solution and thereby lowering the surface boundary of the mineral oil, the CVC's membrane dynamics during exocytotic cycles were somewhat distorted. Fig. 1B shows a representative trace for the CV membrane potential during six exocytotic cycles (numbered 1-6) in a mechanically compressed cell. In such compressed cells, the electrode tip tended to escape from the CV and to lodge in the cytosol during the fluid-discharging phase. The tip then re-entered the CV as the CV swelled after the start of the fluid-filling phase. Recording of the CV potential was, therefore, interrupted during this phase. Interruptions to the potential recordings are shown by double slashes in Fig. 1B.
The CV potential in a compressed cell increased in a stepwise manner to a maximum value (usually 80-90 mV) soon after the start of the fluid-filling phase (Fig. 1B; cycles 2 and 4). The CV potential decreased, also in a stepwise manner, after the start of the rounding phase (Fig. 1B; cycles 1, 2 and 5).
Reattachment of the radial arms to the CV in mechanically compressed
cells
In mechanically compressed cells, reattachment of the radial arms to the CV
after the start of the fluid-filling phase occurred asynchronously. A
representative example of asynchronous attachment of the radial arms to the CV
during the fluid-filling phase is shown in
Fig. 2. In the frame at the
upper left labeled 0, which corresponds to the start of a series of 16 frames,
the CV was surrounded by four swollen radial arms or ampullae (the proximal
ends of the radial arms next to the CV; RA1-RA4). An ampulla is known to swell
before it attaches to the CV (Hausmann and
Allen, 1977) and then to narrow after it attaches to the CV. RA1
at 0.6 s, RA2 at 1.0 s, RA3 at 1.8 s and RA4 at 2.6 s (numbers are time in
seconds after the start of recording of this series) were narrower than those
in the respective previous frames. All radial arms reattached to the CV within
2.8 s. Reattachment of the radial arms in noncompressed cells occurs
more-or-less synchronously (data not shown).
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Effects of the osmolarity of the adaptation solution on the CV
potential, Rcvc and the immunological labeling of the decorated spongiome
As shown in Fig. 3A, the CV
potential during the fluid-filling phase (filled squares) was the same in
cells that had been adapted to different osmolarities for 18 h:
83.3±2.1 (N=4), 83.9±4.0 (N=8),
87.3±2.0 (N=5) and 83.4±4.9 mV (N=6) in cells
adapted to 4, 24, 64 and 124 mosmoll-1, respectively. There were no
significant differences between these four values (t-test,
P>0.1). By contrast, RCVC differed in cells
adapted to different osmolarities: 98.2±50.3 (N=5),
69.9±14.7 (N=10), 19.9±6.2 (P<0.01,
N=5) and 20.0±11.8 fl s-1 (P<0.01,
N=6) in cells adapted to 4, 24, 64 and 124 mosmoll-1,
respectively.
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Representative fluorescence images of the decorated spongiome of cells adapted to 4, 64 and 124 mosmoll-1 are shown in Fig. 3B. No differences in the brightness or appearance of the fluorescence images of the decorated spongiome were observed between cells adapted to different osmolarities.
Recovery of the CV potential, RCVC and immunological
labeling of the decorated spongiome after treatment with a hypertonic solution
had ended
Cells adapted to 4 mosmoll-1 for 18 h were exposed to a 124
mosmoll-1 solution for 30 min (hypertonic stimulation). The cells
were then re-exposed to 4 mosmoll-1 adaptation solution. The CV
potential during the fluid-filling phase and the RCVC were
measured 20, 40, 60, 120 and 240 min after reexposure to 4
mosmoll-1. Fluorescence images of the decorated spongiome were
taken at times corresponding to the times when the CV potential and
RCVC were measured. An image of a cell exposed to 124
mosmoll-1 for 30 min was taken to show the appearance of its
decorated spongiome at time 0 of re-exposure of the cell to 4
mosmoll-1. An image of a cell adapted to 4 mosmoll-1 was
also taken to serve as a control of the appearance of the decorated spongiome.
The CV was invisible at time 0, the time when the CV potential and
RCVC are both assumed to be zero.
As shown in Fig. 4A, the CV potential (filled squares) increased with time after the cells had been re-exposed to 4 mosmoll-1. The potential was 44.0±3.1 (N=5), 50.3±8.4 (N=5), 63.6±2.7 (N=5), 68.8±5.1 (N=6) and 70.6±3.4 mV (N=9) 20, 40, 60, 120 and 240 min after re-exposure to the 4 mosmoll-1 solution, respectively. RCVC (open circles) also increased with time after cell re-exposure to 4 mosmoll-1 in parallel with the increase in the CV potential. RCVC was 57.7±7.5 (N=5), 80.5±7.1 (N=5), 114.7±8.8 (N=6), 97.0±11.5 (N=7) and 95.3±36.8 fl s-1 (N=7) 20, 40, 60, 120 and 240 min after re-exposure to 4 mosmoll-1, respectively.
|
A representative series of fluorescence images of the decorated spongiome
is shown in Fig. 4B. The
decorated spongiome, or at least the V-ATPases
(Fok et al., 1995), was
disrupted and the DS-1 labeling was dispersed into the cytosol during
hypertonic stimulation of cells (image labeled 0). A labeled decorated
spongiome began to reappear along portions of the radial arms proximal to the
CV 20 min after the cells had been re-exposed to 4 mosmoll-1 (image
labeled 20). The decorated spongiome extended along the radial arms over time,
and its fluorescence image after 120 min of re-exposure to 4
mosmoll-1 was very similar to that in cells adapted to 4
mosmoll-1 for 18 h (compare image labeled 120 with that labeled
control).
Effects of hypotonic stimulation on the CV potential and
RCVC
Cells adapted to an osmolarity of 124 mosmoll-1 for 18 h were
exposed to a 4 mosmoll-1 solution for 30 min (hypotonic
stimulation). The CV potential during the fluid-filling phase and
RCVC were then measured. As shown in
Table 1, no significant
(P>0.1) change in the CV potential occurred in response to
exposure of cells adapted to 124 mosmoll-1 to 4
mosmoll-1. By contrast, RCVC was significantly
(P<0.05) increased from approximately 20 to 103 fl s-1
by hypotonic stimulation.
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Effects of inhibitors on the CV potential and
RCVC
Cells adapted to 4 mosmoll-1 were exposed to inhibitors in 4
mosmoll-1 solution for 10-30 min. The CV potential during the
fluid-filling phase and RCVC were then measured. The
inhibitors employed were 30 nmoll-1 concanamycin B, 1.0
mmoll-1 furosemide and 40 µmoll-1 EZA.
As shown in Table 2, exposure of cells to concanamycin B for 30 min brought about a decrease in the CV potential from approximately 82 to 40 mV (a 51% decrease; P<0.05). By contrast, RCVC decreased from approximately 108 to 62 fl s-1 (a 43% decrease; P<0.05). The CV potential decreased from approximately 82 to 52 mV (a 37% decrease; P<0.05) after a 10 min exposure to furosemide, while RCVC decreased from approximately 108 to 62 fl s-1 (a 43% decrease; P<0.05). Exposure of cells to EZA for 10 min caused a decrease in the CV potential from approximately 82 to 58 mV (a 27% decrease; P<0.05), while RCVC decreased from 108 to 62 fl s-1 (a 43% decrease; P<0.05). Exposure of cells to 4 mosmoll-1 solution containing 0.1% DMSO, the concentration of this solvent required to dissolve the inhibitors, had no effect on either the CV potential or RCVC (compare the control values for the CV potential and for RCVC in Table 2 with values obtained in the absence of DMSO shown in Fig. 3).
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Discussion |
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As shown in Fig. 1B, the CV potential increased in a stepwise manner to its highest value (70-90 mV) during the early fluid-filling phase in a mechanically compressed cell (see exocytotic cycles 2 and 4). In contrast, as shown in Fig. 1A, in a non-compressed cell, the CV potential generally reached its maximum value of approximately 80 mV without showing discrete stepwise changes during the early fluid-filling phase (see exocytotic cycles 1, 2 and 4; there was a single step in cycle 3).
In a compressed cell, the CV potential decreased in a stepwise manner to its minimum value (10-20 mV) during the early rounding phase (Fig. 1B; exocytotic cycles 1, 2 and 5). In contrast, as shown in Fig. 1A, in a non-compressed cell, the CV potential reached its minimum value (approximately 10 mV) without stepwise changes during the early rounding phase. Detachment of the radial arms from the CV during the early rounding phase also occurred asynchronously in such compressed cells (video data not shown). We conclude that stepwise changes in the CV potential, as seen in mechanically compressed cells, corresponds to the attachment or detachment of individual radial arms from the CV.
These findings support our previous hypothesis that the CV potential
observed during the fluid-filling phase is dependent on the electrogenic
activity of the radial arms (Tominaga et
al., 1998). An equivalent electrical circuit explaining these
stepwise changes in the CV potential is presented in
Fig. 5B and is discussed
below.
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Correlating the CV potential during the fluid-filling phase with
the presence of the V-ATPase-bearing decorated spongiome in the radial arms
and with RCVC,
As shown in Fig. 4A, we have
demonstrated that the CV potential (filled squares) gradually increases in
parallel with an increase in RCVC (open circles) in cells
re-exposed to a hypotonic (4 mosmoll-1) adaptation solution after
an exposure of 30 min to a hypertonic (124 mosmoll-1) solution.
This increase in the CV potential to the control value of approximately 80 mV
was accompanied by the reappearance of a monoclonal antibody
(DS-1)-immunolabeled decorated spongiome, as illustrated in
Fig. 4B. DS-1 labels the
V-ATPase on the decorated spongiome (Allen
et al., 1990; Fok et al.,
1995
). These findings support the idea that the CV potential in
the fluid-filling phase is generated in the decorated spongiome membrane,
where the V-ATPase complexes are situated
(Tominaga et al., 1998
).
Utilizing DS-1, Ishida et al.
(1993) demonstrated that an
important site in the fluid segregation activity of the CVC is situated in the
decorated spongiome. Ishida et al.
(1996
) then found that
hypertonic stimulation of Paramecium cells led to the separation of
the V-ATPase-bearing decorated spongiome from the CVC. The
RCVC dropped to near zero at the time that the
fluorescence of the decorated spongiome was disrupted. Re-exposure of cells to
the previous hypotonic solution resulted in the reappearance of the
immunologically labeled decorated spongiome around the radial arms as well as
to the resumption of the control rate of fluid segregation.
According to Stock et al.
(2001), the osmolarity of the
cytosol increases rather slowly (taking over 12 h) after cells have been
exposed to a hypertonic solution. It is therefore assumed that the cytosolic
osmolarity of cells adapted to 4 mosmol l-1 does not change
significantly during a 30 min exposure to a hypertonic (124 mosmol
l-1) solution. It is also assumed that the water permeability of
the plasma membrane does not change significantly during this brief exposure
to a hypertonic solution (Stock et al.,
2001
). The rate of osmotic water influx across the plasma membrane
after reexposure to a 4 mosmol l-1 solution is, therefore, assumed
to be similar to that before the 30 min exposure to 124 mosmol l-1.
The RCVC must equal the rate of osmotic water influx prior
to the cells' exposure to hypertonic solution, i.e. at a time when the
decorated spongiome is still intact, or the cell would swell or shrink. The
lower RCVC during the early phase of exposure to
hypertonic solution, compared with the later stages, can be attributed to the
decreased number of functionally active V-ATPase complexes associated with the
CVC. This decrease can be attributed to the disruption of the decorated
spongiome by hypertonic stimulation. As the number of functional V-ATPase
complexes increases over time (as shown by the gradual thickening and
lengthening of fluorescence images of the decorated spongiome;
Fig. 4B), the fluid segregation
also increases, but presumably only to a rate determined by the rate of water
entry across the plasma membrane. Thus, the numbers of V-ATPase complexes
present on the decorated spongiome and RCVC are not
necessarily proportional.
Based on these findings, we conclude that the electrical potential across
the spongiome membrane is proportional to the number of functional V-ATPase
complexes present in the decorated spongiome. Our finding that concanamycin B,
a potent inhibitor of V-ATPase activity
(Woo et al., 1992), caused a
significant decrease in both the maximum CV potential during the fluid-filling
phase and RCVC (Table
2) implies that electrogenic V-ATPase activity is involved in the
fluid expulsion activity of the decorated spongiome. The stoichiometric
relationship between the CV potential and the number of functional ATPase
complexes present should be examined further.
Based on the K+ and Cl- activities in the CVs in
vivo, Stock et al. (2002)
proposed that K+ and Cl- must be transported in
significant amounts across the CVC membrane for water to be transported
osmotically from the cytosol to the CVC lumen. These ions would be needed to
maintain RCVC at an appropriate level. To understand the
mechanism that promotes water transport across the CVC membrane, it is
important to know how the V-ATPase-mediated electrical potential across the
spongiome membrane is associated with this membrane's hypothetical
K+ and Cl- transport activity or of other ion transport
activities.
The CV potential during the fluid-filling phase remains at the same
maximum level in cells adapted to different osmolarities even though their
RCVC varies
As is clearly shown in Fig.
3A, the CV potential during the fluid-filling phase (filled
squares) was maximal and approximately the same (80-90 mV) for cells that had
been adapted to different osmolarities (4, 24, 64 and 124 mosmol
l-1). By contrast, RCVC changed significantly
(P<0.01) as the adaptation osmolarity changed, i.e. it was
approximately 98, 70, 20 and 20 fl s-1 for cells adapted to 4, 24,
64 and 124 mosmol l-1, respectively. The decorated spongiome
appeared to be normal and unaltered in all cells, even when adapted to
different osmolarities (Fig.
3B). Moreover, the CV potential during the fluid-filling phase
remained unchanged at its maximum level of approximately 80 mV, even though
RCVC increased from approximately 20 to 103 fl
s-1 when cells adapted to 124 mosmol l-1 were exposed to
a 4 mosmol l-1 solution (hypotonic stimulation,
Table 1).
These findings imply that (i) the number of functional V-ATPase complexes
in the decorated spongiome is nearly constant and at its maximum level in all
cells adapted to different osmolarities and (ii) the number of complexes is
unaffected by hypotonic stimulation even though RCVC
increases dramatically. Maximal V-ATPase activity might somehow energize the
hypothetical K+ and Cl- transport system in the CVC
membrane (Stock et al., 2002),
which can provide the osmotic gradient across the CVC membrane and thereby
promote osmotic flow of excess cytosolic water into the CVC lumen.
Furosemide and EZA reduce both the CV potential and
RCVC
Furosemide is known to inhibit ion transport in the loop of Henle
(Brater, 1998) and erythrocytes
(Lauf, 1984
;
Canessa et al., 1986
;
Garay et al., 1988
). Stock et
al. (2002
) found that
furosemide inhibited transport of K+ and Cl- across the
plasma membrane of Paramecium and caused a decrease in
RCVC. We show here that an external application of
furosemide (1.0 mmol l-1) reduces the CV potential to approximately
63% of its maximum control value (82 mV;
Table 2).
RCVC is reduced to approximately 57% of its control value
(108 fl s-1; Table
2). Stock et al.
(2002
) reported that
furosemide decreased cytosolic K+ and Cl- activities and
suggested that these decreases were partially responsible for the decrease in
RCVC. It is also possible that furosemide directly
inhibits V-ATPase. The effect of cytosolic K+ activity on V-ATPase
activity will need to be examined.
External application of EZA, which is known to inhibit carbonic anhydrase
(Deitmer and Schlue, 1989),
reduced the CV potential to approximately 71% of its control value (82 mV;
Table 2) and
RCVC to approximately 57% of its control value (108 fl
s-1; Table 2). It is
possible that carbonic anhydrase is involved in supplying H+ for
proton translocation by the V-ATPase of the decorated spongiome. In this case,
a reduced supply of protons to the V-ATPases could account for the decrease in
the CV potential.
The results of these inhibitor experiments also strongly support our suggestion that the V-ATPase-mediated electric potential of the CVC membrane is closely involved in establishing the osmotic gradient across the CVC membrane that is essential for osmotic flow of excess cytosolic water into the CVC lumen.
Electrical equivalent circuit for the CVC, simulation of the stepwise
changes in CV potential as the radial arms attach to the CV and estimation of
electric current generated by the V-ATPase activity in a single radial
arm
V-ATPase activity can be regarded as a source of constant electric current.
A schematic representation of the CVC showing the electric currents due to
V-ATPase activity is shown in Fig.
5A, and its corresponding equivalent electrical circuit is shown
in Fig. 5B. Each radial arm
(RA) generates a constant electric current,
iH+, which produces an electrical potential
difference across the radial arm membrane, eRA, that can
be written as:
![]() | (1) |
![]() | (2) |
![]() | (3) |
The input resistance of approximately 60 M of the CV during the
fluid-filling phase (Fig. 1A)
corresponds to the electrical resistance for the overall membrane of the CVC,
rCVC. The input resistance of approximately 160 M
of the CV
during the rounding phase (Fig.
1A) corresponds to the input resistance of the CV only,
rCV. rRA can be written as:
![]() | (4) |
As previously mentioned, the CV potential is approximately 80 mV in the
fluid-filling phase (Fig. 1A).
An electric current needed to maintain 80 mV of potential across the CVC
membrane of 60 M resistance would be 1.3x10-9 A
according to Ohm's law. If we assume the mean number of radial arms to be 10
for each CVC, the current generated by a single arm would be
1.3x10-10 A. This current would correspond to the transport
of 8.3x108H+s-1 across the membrane of
a single radial arm by its V-ATPases. This value is obtained by dividing
1.3x10-10 Cs-1 by e, the elementary
electric charge (1.6x10-19 C). A current of
1.3x10-10 A across a membrane with a potential difference 80
mV is equivalent to 1.1x10-11 W. These values could be useful
for an eventual understanding of the molecular mechanism of ion transport
across the CVC membrane.
CV membrane potential during the rounding and fluid-discharging
phases of the CVC cycle
The small potential (approximately 10 mV) seen during the rounding phase (R
in Fig. 1A), corresponds to an
unexplained potential across the CV membrane itself. We recently found that
the K+ and Cl- activities in the CV fluid are always
approximately 2.5 times those in the cytosol
(Stock et al., 2002). It is
therefore possible that this 10 mV CV membrane potential corresponds to a
potential value between two equilibrium potentials, one for K+
(approximately -23 mV) and the other for Cl- (approximately 23 mV).
Studies on the putative ion channels in the CV membrane are needed to explain
how an electrical potential is developed during the rounding phase. However,
as pointed out by Tominaga et al.
(1998
), the potential measured
during the fluid-discharging phase, 26 mV, corresponds to the plasma membrane
potential since the CV pore is open during this phase and, therefore, the CV
fluid is electrically connected to the external fluid.
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Acknowledgments |
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References |
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