Life at acidic pH imposes an increased energetic cost for a eukaryotic acidophile
1 The Josephine Bay Paul Center for Comparative Molecular Biology and
Evolution, Marine Biological Laboratory, Woods Hole, MA 02543, USA
2 BioCurrents Research Center, Program in Molecular Physiology, Marine
Biological Laboratory, Woods Hole, MA 02543, USA
3 Sea Education Association, PO Box 6, Woods Hole, MA 02543, USA
4 Centro de Biología Molecular, Universidad Autónoma de
Madrid, Cantoblanco, Madrid 28049, Spain
* Author for correspondence (e-mail: mmesserli{at}mbl.edu)
Accepted 25 April 2005
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Summary |
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Key words: acidophile, cytosolic pH, membrane potential, energetic cost, Chlamydomonas sp
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Introduction |
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In order to identify the adaptations used to survive in acid, we need to
consider first which components of acidophiles are in contact with low pH. For
example, acidophilic bacteria have cell surface enzymes that have acid pH
optima. A surface iron oxidase from bacterial strain TI-1 has a pH optimum of
3.0 (Takai et al., 2001) while
a surface thiosulfate dehydrogenase from Acidithiobacillus
thiooxidans has a pH optimum of 3.5
(Nakamura et al., 2001
).
Protistan extremophiles must also cope with acid conditions on the surface of
the plasma membrane. Ion channels and transporters are in contact with the low
pH of the extracellular medium and this would minimally require molecular
modifications as compared to similar proteins from closely related species
growing at neutral pH. In considering the transmembrane H+
gradient, the internal conditions are as important. There are conflicting
reports of the cytosolic pH from acidophiles. Cyanidium caldarium
(Beardall and Entwisle, 1984
;
Enami et al., 1986
) and
Dunaliella acidophilum (Gimmler
et al., 1989
), grown at extracellular pH of 2.1 and 0-1.0,
respectively, maintain cytosolic pH of 6.6 and 7.0, respectively. In contrast,
Picrophilus oshimae (van de
Vossenburg et al., 1998
), Bacillus acidocaldarius
(Thomas et al., 1976
),
Sarcina ventriculi (Goodwin and
Zeikus, 1987
) and Euglena mutabilis
(Lane and Burris, 1981
) grown
at pH 0.8-4.0, 3.0, 3.0 and 2.8, respectively, are reported to maintain acidic
cytosolic pH of 4.6, 5.5, 4.25 and 5.0-6.4, respectively. The methods used to
determine the cytosolic pH of these later examples were a measure of total
cellular pH, where the presence of acid-containing organelles could lead to
more acidic estimates of cytosolic pH. Identification of cytosolic enzymes
from these organisms that are optimally functional at acidic pH would support
these measurements.
Maintenance of a neutral pH cytosol in an extracellular environment at pH 2
indicates that a 105-fold [H+] gradient must exist
across the plasma membrane. Apart from acidophiles, there are only a few other
reported examples of such large H+ gradients across cellular
membranes, such as in the mammalian stomach (104- to
106-fold) and the acidic vacuole in plant cells
(105-fold). These H+ gradients are achieved by a
combination of active transport and low permeability to H+
(Boron et al., 1994;
Muller et al., 1996
).
Maintenance of large, transmembrane H+ gradients would be an
energetically costly endeavor if the membrane were relatively leaky to
H+. Some measurements have shown that lipid bilayers are orders of
magnitude less permeable to H+ at acidic pH than neutral pH
(Gutknecht, 1984
). Other
measurements indicate that the difference at acidic pH is not so large. For
example, the permeability of thermoacidophilic archeal membranes to
H+ show less than an order of magnitude difference between pH 6 and
pH 2.5 and are only an order of magnitude less permeable than egg
phosphatidylcholine membranes at neutral pH
(Komatsu and Chong, 1998
).
These measurements have been made on purified lipid and are not representative
of cellular membranes that contain about 50% protein, including water-filled
ion channels, H+-cotransporters and other weakly acidic or weakly
basic proteins that may increase permeability to H+. Based on this
information, we hypothesize that eukaryotic acidophiles have an increased
energetic cost to survive at acidic pH due to the transmembrane H+
gradient, which may be orders of magnitude larger in the same unicellular
organism growing at pH 2 vs pH 7.
In this study we characterize the electrochemical H+ gradient that exists across the plasma membrane of a eukaryotic acidophile at pH 2 and pH 7. Cytosolic pH was measured using the ratiometric fluorescent indicator 2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF), and the plasma membrane potential difference was measured using intracellular electrodes. We also monitored the relative metabolic activity of these organisms growing in environments at pH 2 and pH 7 by measuring O2 and ATP consumption in the dark to determine the energetic cost of living in acid.
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Materials and methods |
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Relative growth rate measurements
Instantaneous growth rates were acquired for both Chlamydomonas
sp. and Chlamydomonas reinhardtii at different pH values in order to
determine their pH tolerance. Instantaneous growth rate (r) is
calculated as the difference between natural logarithms of chlorophyll
fluorescence (f) at different points in time (t) during
exponential growth according to the following equation:
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Cytosolic pH measurements
The cell-permeant form of the ratiometric H+-indicator,
2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF)
(Molecular Probes, Eugene, OR, USA), was used to measure cytosolic pH of
Chlamydomonas sp. Cells were loaded with BCECF by incubation at pH 2
with 10-12 µmol l-1 BCECF for 1 h, resulting in a final
cytosolic concentration that gave a similar light intensity at the pH
insensitive wavelength as 6 µmol l-1 dye held between two
coverslips 10 µm apart. Cells were rinsed 3x with culture medium,
concentrated by centrifugation and resuspended in growth medium in order to
remove the unloaded dye. A similar procedure was used to load the neutral
growing Chlamydomonas reinhardtii (catalog no. WW-15-2041, Carolina
Biological Supply, Burlington, NC, USA) as a positive control. C.
reinhardtii, however, was loaded at pH 5 for the same amount of time.
Imaging was performed on cells immobilized in 1% low temperature-gelling
agarose (Boehringer Mannheim, Germany) on the bottom of small cultures dishes
with a #0 coverglass bottom window. Images were acquired with a wide-field
Attofluor RatioVision System (Rockville, MD, USA) coupled to a Zeiss 135
Axiovert inverted microscope. Images were collected through a x100, 1.3
N.A. lens using 440±10 nm and 488±5 nm excitation with
525±12 nm emission. The ratio of 488/440 was masked to remove
background and out-of-focus light. At the BCECF excitation/emission
wavelengths the chlorophyll contributed no autofluorescence. Image acquisition
parameters were adjusted so that the small amount of autofluorescence
contributed by the cells was no longer detected. The cytosolic pH is presented
as the mean ± S.E.M.
Calibration of fluorescent H+ indicator
In vitro calibration of BCECF was performed by loading 6 µmol
l-1 BCECF in Fisher brand phosphate buffers pH 5-9 (Suwanee, GA,
USA). A small volume of buffer, 4.8 µl, was placed between one large #0
coverslip and one 22x22 mm coverslip so that the depth of buffer
including dye was 10 µm, to simulate the thickness of the cells. Images
were collected as described above and the average ratio of the central 90% of
the image was used to calibrate the settings.
In vivo calibrations were performed to confirm that the cytosolic pH indicator responded to changes in cytosolic pH in the same manner as the Fisher brand buffers. Standard calibration procedures using the ionophores valinomycin and nigericin did not collapse transmembrane K+ or H+ gradients in this organism at either neutral or acidic pH. As an alternative, we used weak acids and a weak base to change and clamp the cytosolic pH at different known pH values. The weak acids and base diffuse through the plasma membrane in their uncharged form, thus releasing (acid) or gaining (base) H+ within the cytosol. The neutral form of the weak acid or weak base will reach equilibrium inside and outside of the cell so that the cytosolic pH will reach equilibrium with extracellular pH. Carbonyl cyanide m-chlorophenylhydrazone (CCCP), a protonophore, was then used to eliminate small H+ gradients and charge build-up across the membrane. Acetic acid, pKa 4.76, was used at pH 5 and 6, p-nitrophenol, pKa 7.1, for pH 7 and ammonia pKa 9.2 for pH 8 and 9.
Membrane potential difference measurements
Plasma membrane potential differences were measured for cells growing in
MAM at pH 2 and pH 7 without trace elements or vitamins. The cell holding
chamber consisted of two 24x60 mm coverslips spaced 3-3.5 mm apart by
three standard sized microscope slides at each end, allowing culture medium to
be held between the two coverslips for several hours. The chamber allowed us
to position a holding pipette and recording electrode parallel to the
microscope stage. A large-tip holding pipette was fire polished to just under
10 µm outer diameter. Culture media and cells were drawn into the holding
pipette such that cells larger than 10 µm could not enter and were held at
the tip of the pipette. Plasma membrane potential differences were measured
using 1.5 mm outer diameter theta glass. These electrodes were pulled to
resistances of 60-90 M when 1 mol l-1 KCl was used both as
backfill and outside the pipette. Current-voltage (I-V) relationships
were generated with these electrodes to ensure that the cells had been
impaled. Intracellular recordings were acquired with Axoclamp 2B controlled
with Pclamp 9 (Axon Instruments, Union City, CA, USA). The theta glass was
backfilled with 50 mmol l-1 K2SO4 during
recordings. Only measurements from cells that showed flagellar and contractile
vacuole activity before and during impalement were used. To increase the
chance of a successful impalement, larger cells were selected for these
experiments by filtration through an 8 µm Nuclepore filter (Pleasanton, CA,
USA). Cells were resuspended from pH 2 medium into pH 7 medium, giving a final
pH of 6.7. After impalement at pH 6.7, the holding pipette was removed and
recordings were collected while the cell was impaled on the measuring
electrode. An ISMATEC IPC 8 channel peristaltic pump (Glattbrugg-Zürich,
Switzerland) was used to exchange pH 6.7 medium with pH 2 medium and then pH 7
medium. Membrane potential differences and I-V curves were collected
at both pH 2 and pH 7. All membrane potential differences are presented as the
mean ± S.E.M.
Measuring O2 consumption
O2 consumption was measured using a miniaturized Whalen-style
polarographic probe combined with spatial self-referencing, a signal-to-noise
enhancement method. The self-referencing polarographic O2
electrode, described by Jung et al.
(1999), consists of a recessed
platinum reactive surface in the tip of a 2-3 µm opening glass
microelectrode. Oxygen is reduced at the Pt surface polarized between -0.6 to
-0.8 V. In pH 2 medium the current used to maintain the -0.8 V was so great
that it saturated the amplifier. The current was brought into range by using a
Pt wire reference electrode rather than the Ag/AgCl reference described in the
original publication. The Pt wire reference was used at both acidic and
neutral pH. Electrodes were polarized using an Ionview amplifier with a
polarographic headstage running Ionview software (BRC, Woods Hole, MA, USA).
Oxygen consumption could not be measured reliably from single cells so cells
were immobilized in 1% low-temperature-gelling agarose (Boehringer Mannheim,
Germany) in order to measure consumption from a population of cells at a
density between 1010-1011 cells l-1. After
mixing an equal volume of cells in medium with 2% agarose the mixture was
taken up into a 1.5 mm diameter capillary glass and allowed to cool before
pushing the molded agarose cylinder back into the medium. Measurements were
taken within an hour. Movement of gels between solutions of different pH
resulted in pH gradients near the surface of the gels that disappeared within
30 min. This prevented us from measuring O2 consumption within 30
min after transfer to different pH medium but allowed the pH in the core of
the gel to reach the pH of the bath. The agarose did not appear to harm the
cells as they continued to divide, embedded within the agarose, for weeks.
Measurements of O2 consumption were made on cells in a simplified
culture medium consisting of MAM without trace elements or vitamins. Sulfate
was used as the pH buffer at pH 2 while Hepes was used as the pH buffer at pH
7, as described above. Measurements were performed at ambient temperature,
23°C and in the dark to eliminate O2 efflux due to
photosynthesis.
Monitoring ATP consumption
Cells normally cultured at pH 2 were transferred to pH 7 medium,
concentrated, and washed 3 times with pH 7 medium. Azide-treated cells were
incubated for 15 min in 159 µmol l-1 azide, at pH 7. This
resulted in 1 µmol l-1 of cell-permeant azide, hydrazoic acid,
pKa 4.7, based on the following equation
AH=[A]/[(Ka/[H+])+1], a rearrangement of the
Ka equilibrium expression, where AH is protonated weak
acid and [A] is the total concentration of weak acid. A tenfold lower
concentration of azide did not have an effect on ATP levels while a tenfold
higher concentration resulted in greater than 50% loss of ATP. After loading,
cells were rinsed 6 times to lower the extracellular hydrazoic acid
concentration to less than 2 nmol l-1 and then dispensed into pH 2
and pH 7 media so that the final concentration of cells matched the controls.
Control samples were dispensed into pH 2 and pH 7 media from the first cell
concentration step in pH 7 medium.
Total cellular ATP measurements were performed using the Promega ENLITEN kit (Madison, WI, USA) based on the ATP-dependent light output of the Luciferin/Luciferase reaction. Photons were counted with a Zylux FB12 Tube Luminometer (Zylux Corporation, Maryville, TN, USA) averaging counts for 10 s after a 2 s delay. We collected and averaged three 10 s readings from each sample. Luminometer dark counts averaged 15 relative light units (RLU) while counts due to the presence of the Luciferin/Luciferase alone gave a range of 50-400 RLU. Signals from samples ranged between 104-106 RLU. Cell samples were processed by combining 250 µl of cells in native medium with 250 µl of 10% trichloroacetic acid (TCA) to lyse cells and inactivate ATPases. After brief vortexing the sample was left at room temperature (23°C) for 10 min before vortexing again. The sample was neutralized by adding 10 µl of lysate to 40 µl Trisacetate buffer, pH 8.5. After brief mixing with the pipette tip, 10 µl of this sample was combined with 90 µl of enzyme and buffer provided in the kit, mixed by pipetting and immediately placed in the luminometer counting chamber. This method diluted the initial sample 100-fold, yielding measurements from 105-106 cells.
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Results |
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Acidophile membrane potential difference is close to zero
The transmembrane electrochemical H+ gradient normally comprises
both concentration and electrical differences across the plasma membrane. The
electrical component was characterized in the acidophile using intracellular
recordings. Theta-glass, double-barreled microelectrodes were used to measure
the membrane potential difference and also to clamp the membrane voltage to
generate I-V plots. When the backfilling solution was 1-3 mol
l-1 KCl, impalements often led to ejection of flagella and cell
swelling. The number of successful impalements increased when the backfill was
changed to 50 mmol K2SO4 as used in small fungi cells
(Blatt and Slayman, 1983), with
the pH set to 6.8. Larger cells growing at pH 2 were selected by filtration,
suspended in a larger volume of pH 7 medium (final pH 6.7) and impaled. Cells
showing flagellar activity were selected for impalement. At the higher
magnification used for impalement, contractile vacuole activity could be
identified. Only cells showing flagellar and contractile vacuole activity
before and during impalement are reported. After impalement a flow system was
used to change the pH of the medium to pH 2 and then back to pH 7 while
monitoring the membrane potential difference. The membrane potential
difference immediately after impalement at pH 6.7 was -3.3±2.6 mV, and
increased to +20.8±2.3 mV upon exchange to pH 2 medium, returning to
-3.6±1.8 mV (N=8) when cells were further exposed to pH 7
medium. The large change in pH affected the surface charge on the recording
electrode glass and contributed an artifactual error to the measured potential
difference. This error was measured by keeping the tip potential constant
through immersion in pH 7 medium in a holding pipette and then exchanging the
medium with a flow system over the immersed shaft of the recording electrode.
Exchange of the medium from pH 7 to pH 2 gave rise to an increase of
+22.2±0.6 mV (N=4 electrodes) in the measured potential
despite the fact that the tip of the electrode was kept in the same medium.
When this artifact is removed from the measurement of the membrane potential
differences recorded from cells at pH 2, the measured membrane potential
difference is -1.5±2.3 mV.
The I-V relationship of the cells was determined under these conditions to ensure correct placement of the microelectrode. At pH 7, cells showed smaller inward and outward currents compared to cells at pH 2. Fig. 5A,B shows representative current traces from a Chlamydomonas sp. at pH 7 and pH 2, respectively. Currents were recorded at voltages ±100 mV about the measured potential difference. Fig. 5C shows the mean currents for seven cells, normalized for cell size, including the corrected voltage offset at pH 2. Impaled cells were best modeled as ovoid with an average length of 12 µm and width of 10 µm. Surface area due to flagella was not included. Cell conductance changes linearly at negative holding potentials but rises more rapidly with increasing voltage at positive holding potentials.
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Acidic conditions increase ATP consumption
Measuring O2 consumption as a relative indicator of metabolic
cost was a coarse method, as cells can generate ATP by other means without
consuming O2, and other cellular enzymes consume O2
without producing ATP. A more sensitive method for determining the differences
in metabolic cost was to measure the rates of ATP consumption at pH 2 and pH
7. As the total cellular ATP is the difference between ATP production and ATP
consumption, we loaded cells with sodium azide, an electron transport
inhibitor, to arrest ATP production, and then followed the rate of ATP
consumption by cells at pH 2 and pH 7. According to our hypothesis, cells in
pH 2 medium should consume their cellular ATP pool more quickly than cells
growing in pH 7 medium. After incubating cells in azide for 15 min, then
rinsing away the azide, cells were moved to fresh medium buffered at pH 2 and
pH 7 and the total cellular ATP levels were measured over time. Both groups of
azide-treated cells showed an average 30% reduction in the total amount of ATP
compared to the levels in untreated controls perhaps due to cell death within
the population. The ATP standard curve gave an average 8.3±0.4-fold
increase in light emission per order of magnitude increase in ATP
concentration. We found that, on average, cells growing at pH 2 contain
5.2x109 molecules ATP/cell (N=10 populations of
cells). A summary of the time course of changes in cellular ATP levels is
displayed in Fig. 6. All data
are shown as the ratio between paired groups to eliminate variation between
experiments due to the loss of enzyme activity over time and differences in
cell density. The first time point is taken just less than 5 min after the
shift to pH 2 and pH 7 medium. The control curve (filled squares) in
Fig. 6A is the ratio of the
ATP-dependent light emission from cells in pH 2 medium to cells in pH 7
medium. Neither control group was treated with azide. There was no significant
difference in ATP levels between cells growing in pH 2 compared to pH 7 for
the first 65 min after moving to a different pH (P>0.2, 0.5, 0.4,
0.2 for 5, 25, 45 and 65 min, respectively, N=10). However, at 105
min and 165 min there was, relatively, 17% and 10% more ATP measured from
cells in pH 2 than pH 7. The raw data show that this is due to a decrease in
ATP levels in cells at pH 7. The second curve (open squares) in
Fig. 6A shows the ratio of the
ATP levels of cells at both pH treated with azide while the third curve
(circles) is the normalization of the ratio of azide-treated cells to their
control pH group for each trial.
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We now consider only the normalized ATP curve (circles). There is an 8% decrease in the total cellular ATP of azide-treated cells at pH 2 compared to azide-treated cells at pH 7 between 0 and 5 min (P<0.001; paired t-test, N=7). ATP levels decrease further in pH 2 at 25 min after transfer, where cells have 16% less ATP than cells growing at pH 7 (P<0.002). Recovery of ATP levels begins to occur at 45 min and is obvious by 65 min after treatment with azide, when ATP levels are only 14% and 7% less than in cells growing at pH 7(P<0.02 and 0.03, respectively). ATP levels have fully recovered in cells growing at pH 2 by 105 and 165 min to the extent that there is not a significant difference in ATP between cells in pH 2 and pH 7 (P>0.3 for both). Fig. 6B shows the rate of change of ATP levels, which is the slope of the normalized curve in Fig. 6A. We assume that there is no difference in the amounts of cellular ATP for azide-treated cells at t=0 such that the normalized curve is equal to 1 at t=0. In Fig. 6B the slopes of the individual segments of the normalized curve in Fig. 6A are plotted midway between the original time points. Negative values indicate a faster rate of ATP depletion while positive values indicate a faster rate of ATP production. The greatest rate of ATP depletion occurred as soon as the azide-treated cells were moved to pH 2. ATP decreased until about 30 min, after which time it slowly increased until the ATP levels in the pH 2 grown cells matched the ATP levels in the pH 7 cells. As azide is a weak acid it begins diffusing out of the cells as soon as it has been removed from the extracellular medium. The loss of azide from the cytosol makes it possible for the cells placed in pH 2 medium to recover their cellular ATP levels. This implies that the most accurate measurement of the difference in ATP consumption occurs at the beginning of the recording, when ATP production is most effectively inhibited.
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Discussion |
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There could be many reasons for the lack of growth of
Chlamydomonas sp. above pH 7.0, including restrictions due to the
working pH range of cell wall enzymes or even reversal of the electrochemical
H+ gradient. For example, the cell-wall-removing enzymes of C.
reinhardtii have relatively no activity outside of their working pH
ranges. The vegetative lytic enzyme has relatively no activity below pH 7
(reported by Jaenicke et al.,
1987) or pH 5 (reported by
Matsuda et al., 1995
), while
the gametic lysin does not work below pH 3 or above pH 12
(Jaenicke et al., 1987
). The
reduced growth of C. reinhardtii below pH 5.5, measured here, is
consistent with the more acidic working pH range of the autolysin enzyme
reported by Matsuda et al.
(1995
). With respect to the
H+ gradient, the electrochemical driving force reverses at
extracellular pH 6.6, due to the average cytosolic pH of 6.6 and near zero
membrane potential difference, indicating that transport processes coupling
H+ influx to nutrient uptake would no longer occur. One possible
explanation for this wide range of tolerance is that single cells must be able
to deal with the acidic pH of the river. However, cells growing at high
density, such as mats, may be able to neutralize the pH of the local
environment, as has been demonstrated with C. reinhardtii, the
nonacidophilic species used in these studies, which was able to neutralize the
culture medium originally set at pH 4 and pH 6
(Lustigman et al., 1995
).
Chlamydomonas sp. maintains a mean cytosolic pH of 6.6 in
extracellular medium of pH 2.0, so it is 3.2 times more acidic than C.
reinhardtii, which maintains a mean cytosolic pH of 7.1 in an
extracellular pH 7.0. The neutral cytosolic pH is similar to other acidophilic
algae like Cyanidium caldarium
(Beardall and Entwisle, 1984;
Enami et al., 1986
) and
Dunaliella acidophilum (Gimmler
et al., 1989
). The cytosolic pH of the acidophile does not change
dramatically between extracellular medium at pH 2 or pH 7. This indicates that
the cytosolic milieu of these acidophilic algae are not too different from
similar organisms growing under neutral conditions. It also indicates that the
cells are preferentially maintaining a slightly acidic cytosol even under more
neutral conditions, reversing the electrochemical driving force on
H+. We did measure a higher variation in cytosolic pH from cells
growing in medium at pH 2, i.e. a range of nearly 1 order of magnitude. This
may be a unique property of the acidophile. While lighting conditions change
cytosolic pH in C. reinhardtii
(Braun and Hegemann, 1999
), the
conditions used for imaging Chlamydomonas sp. and C.
reinhardtii in this paper were identical.
The electrochemical H+ gradient of the acidophile is lower than
predicted based on the negative resting potential difference of other
protists. A weakly negative membrane potential difference was measured using
theta-glass microelectrodes with 50 mmol K2SO4 backfill,
-1.5±2.3 mV at pH 2 and -3.6±1.8 mV at pH 7. Tip potentials
measured between pH 7 MAM and 100 mmol l-1 KCl were +5 mV (data not
shown) for the theta-glass electrodes backfilled with 50 mmol
K2SO4. It is possibile that the near zero membrane
potential difference could be due to excessive damage during impalement,
although we do not think that this was the case. Excessive damage during
impalement led to ejection of flagella, perhaps in a Ca2+- or
H+-dependent manner, similar to C. reinhardtii
(Quarmby and Hartzell, 1994).
If damage occurred when impaled at neutral pH, flagellar ejection would have
occurred when the pH was lowered to 2. We employed rigorous impalement
criteria to keep poor impalements out of the analyzed data. The I-V
plots also help confirm that the microelectrodes are in fact in the cell.
Interestingly the I-V plots acquired from the acidophile look similar
to I-V plots acquired from Neurospora under acid load
(Sanders et al., 1981
).
Specifically, membrane conductance is greater during Neurospora acid
load, a phenomenon observed in the current study when Chlamydomonas
sp. was in acidic conditions. Further study of ionic conductances in
Chlamydomonas sp. may lead to the first characterization of ion
channels in a eukaryotic acidophile.
Microelectrodes were used to measure the membrane potential difference of
another acidophile, Dunaliella acidophila
(Remis et al., 1992). Here 1
mol l-1 choline chloride was used as the backfilling solution to
measure an average membrane potential difference of +48.5 mV at pH 1.0.
However a -36.5 mV membrane potential difference was measured at pH 7. This
may reflect different mechanisms of maintaining cytosolic neutrality, for
example H+ efflux through a plasma-membrane-bound pump vs
H+ sequestration via an intracellular H+ pump
on cytoplasmic organelles such as the V-type H+ ATPase pump
associated with the contractile vacuole. The contents of the contractile
vacuole of Paramecium multimicronucleatum are more acidic (pH 6.4)
than the cytosol (pH 7.0) when grown in medium at pH 7.0
(Stock et al., 2002
). Of the
two compartments of the contractile vacuole, the decorated spongiome labels
with antibodies to the V-ATPase H+ transporter while the smooth
spongiome that fuses with the plasma membrane does not (for a review, see
Allen and Naitoh, 2002
). This
indicates that H+ can be taken up into an intracellular compartment
and released without the use of a plasma-membrane-bound H+
transporter. Dunaliella acidophila possesses a well-characterized
P-type H+ ATPase to maintain cytosolic neutrality
(Sekler et al., 1991
). The
H+-ATPase transcripts increase with decreasing extracellular pH
(Weiss and Pick, 1996
). D.
acidophila also contains two contractile vacuoles, but they may not be
very active due to the hypersaline environments in which the organism is
found. Chlamydomonas sp. also has at least two contractile vacuoles
that may help to neutralize cytosolic pH; it is grown in more dilute
conditions than D. acidophila so may rely more heavily on
H+ extrusion via the contractile vacuole rather than a
plasma-membrane-bound H+ transporter. Use of a H+
transporter that never comes into contact with the harsh extracellular
conditions may explain the ability of Chlamydomonas sp. to survive
such a wide pH range.
Transmembrane ionic gradients are maintained by low permeability to ions and ionic transporters that pump ions against their electrochemical gradients. The greater the permeability to H+ the more energy the cell must expend in order to maintain the gradient. This could occur either by direct ATP-dependent transport of H+ out of the cytosol, or via coupled export, where a symporter or antiporter uses the driving force of a different ion to remove H+ from the cytosol. In the latter case, energy would be needed to reset the gradient of the coupled ion. In order to test the hypothesized increase in energy demand we compared the rates of O2 consumption and ATP consumption in cells growing at pH 2 with those growing at pH 7. Initially we found that the rates of O2 consumption, measured using an O2 electrode, were not different in cells growing at acidic pH. Also, under normal conditions, the cellular ATP levels are similar at pH 2 and pH 7, at least for 1 h after transfer to medium of different pH. However, upon reduction of ATP production by an electron transport inhibitor we were able to measure a relatively high rate of ATP consumption by cells in medium at pH 2. The inhibitor is a weak acid, which diffuses out of the loaded cells over time, so that the relatively high rate of ATP consumption dropped to zero within 40 min and then reversed, yielding relatively high production after 40 min (Fig. 6B); cells therefore recovered near normal amounts of ATP after around 135 min. The most accurate measurement of the difference in the rate of ATP consumption is thus during the first few minutes, when the greatest inhibition of ATP production occurs. Cells at pH 2 consume, on average, nearly 0.03% more of the total cellular ATP pool per second than cells at pH 7. For comparison, we calculate that the cells at pH 2 consume O2 at a rate that could produce nearly 0.4% of the total cellular ATP pool s-1 if all of the consumed O2 was used to generate ATP. Considering these two values we find that the cells at pH 2 consume about 7% more of the ATP produced each second than cells at pH 7; i.e. cells at pH 2 are working 7% harder than cells at pH 7. This small difference in ATP consumption could account for our inability to detect differences in the rate of O2 consumption, as the standard error of the measurements was just over 6% of the basal O2 consumption.
If we assume that the increase in ATP consumption is primarily used to
maintain the transmembrane H+ gradient we can estimate the
permeability of the plasma membrane to H+ using the relationship
that net H+ flux across the plasma membrane
(JH+) is the difference between the passive
influx (PH+dC) and the active efflux
(relatively higher rate of ATP consumption/cell surface area), where
PH+ is the cellular permeability coefficient
for H+ and dC is the electrochemical H+
gradient:
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Our results show that Chlamydomonas sp. survives a wide range of extracellular pH, maintains a relatively neutral cytosolic pH, and burns ATP at a faster rate when living in acidic than in neutral conditions. The protection mechanism could be entirely due to active H+ extrusion into a cytosolic vacuole, a hypothesis based on the relatively higher rate of ATP consumption, near zero membrane potential difference and tolerance to a wide range of extracellular pH. The contractile vacuoles of Chlamydomonas sp. may be able to help maintain near neutral cytosolic pH without H+ transporters or exchangers being exposed to the harsh extracellular environment. This implies that the primary structure of H+ transporters in this acidophile may be no different from those found in neutral growing protists. The plasma membrane and cell wall are still in contact with the extracellular environment and certainly there is a higher conductance in the plasma membrane under acidic conditions. Two classes of enzymes are thus identified that may have evolved to function at acidic pH: plasma membrane channels and cell wall lysins.
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Allen, R. D. and Naitoh, Y. (2002). Osmoregulation and contractile vacuoles of protozoa. Int. Rev. Cytol. 215,351 -394.[Medline]
Amaral Zettler, L. A., Gómez, F., Zettler, E., Keenan, B. G., Amils, R. and Sogin, M. L. (2002). Eukaryotic diversity in Spain's River of Fire. Nature 417, 137.[CrossRef][Medline]
Beardall, J. and Entwisle, L. (1984). Internal pH of the obligate acidophile Cyanidium caldarium Geitler (Rhodophyta?). Phycologia 23,397 -399.
Blatt, M. R. and Slayman, C. L. (1983). KCl leakage from microelectrodes and its impact on the membrane parameters of a nonexcitable cell. J. Membr. Biol. 72,223 -234.[Medline]
Boron, W. F., Waisbren, S. J., Modlin, I. M. and Geibel, J.
P. (1994). Unique permeability barrier of the apical surface
of parietal and chief cells in isolated perfused gastric glands. J.
Exp. Biol. 196,347
-360.
Braun, F.-J. and Hegemann, P. (1999). Direct measurement of cytosolic calcium and pH in living Chlamydomonas reinhardtii cells. Eur. J. Cell Biol. 78,199 -208.[Medline]
Doemel, W. N. and Brock, T. D. (1971). The physiological ecology of Cyanidium caldarium. J. Gen. Microbiol. 67,17 -32.
Deamer, D. W. and Akeson, M. (1994). Role of water in proton conductance across model and biological membranes. In Advances in Chemistry, vol. 235 (ed. M. Blank and I. Vodyanoy), pp. 41-54. Washington, DC: American Chemical Society.
Durán, C., Marín, I. and Amils, R. (1999). Specific metal sequestering acidophilic fungi. In Biohydrometallurgy and the Environment: Towards the Mining of the 21st Century (ed. R. Amils and A. Ballester), pp.521 -530. Amsterdam: Elsevier.
Enami, I., Nagashima, H. and Fukuda, I. (1975). Mechanisms of the acido- and thermo-phily of Cyanidium caldarium Geitler II. Physiological role of the cell wall. Plant Cell Physiol. 16,221 -231.
Enami, I., Akutsu, H. and Kyogoku, Y. (1986). Intracellular pH regulation in an acidophilic unicellular alga, Cyanidium caldarium: 31P-NMR determination of intracellular pH. Plant Cell Physiol. 27,1351 -1359.
Fuggi, A., Pinto, G., Pollio, A. and Taddei, R. (1988). Effects of NaCl, Na2SO4, H2SO4 and glucose on growth, photosynthesis, and respiration in the acidophilic alga Dunaliella acidophila (Volvocales, Chlorophyta). Phycologia 27,334 -339.
Gimmler, H., Weis, U., Weiss, C., Kugel, H. and Treffny, B. (1989). Dunaliella acidophila (Kalina) Masyuk - an alga with a positive membrane potential. New Phytol. 113,175 -184.
Goodwin, S. and Zeikus, J. G. (1987). Physiological adaptations of anaerobic bacteria to low pH: metabolic control of proton motive force in Sarcina ventriculi. J. Bacteriol. 169,2150 -2157.[Medline]
Gutknecht, J. (1984). Proton/hydroxide conductance through lipid bilayer membranes. J. Membr. Biol. 82,105 -112.[Medline]
Jaenicke, L., Kuhne, W., Spessert, R., Wahle, U. and Waffenschmidt, S. (1987). Cell-wall lytic enzymes (autolysins) of Chlamydomonas reinhardtii are (hydroxy)proline-specific proteases. Eur. J. Biochem. 170,485 -491.[Abstract]
Jung, S. K., Gorski, W., Aspinwall, C. A., Kauri, L. M. and Kennedy, R.T. (1999). Oxygen microsensor and its application to single cells and mouse pancreatic islets. Anal. Chem. 71,3642 -3649.[CrossRef][Medline]
Komatsu, H. and Chong, P. L.-G. (1998). Low permeability of liposomal membranes composed of bipolar tetraether lipids from thermoacidophilic archaebacterium Sulfolobus acidocaldarius.Biochem. 37,107 -115.[CrossRef][Medline]
Lane, A. E. and Burris, J. E. (1981). Effects of environmental pH on the internal pH of Chlorella pyrenoidosa, Scenedesmus quadricauda, and Euglena mutabilis. Plant Physiol. 68,439 -442.
López-Archilla, A. I. and Amils, R. (1999). A comparative ecological study of two acidic rivers in southwestern Spain. Microb. Ecol. 38,146 -156.[CrossRef][Medline]
López-Archilla, A. I., Marin, I. and Amils, R. (2001). Microbial community composition and ecology of an acidic aquatic environment: The Tinto River, Spain. Microb. Ecol. 41,20 -35.[Medline]
Lustigman, B., Lee, L. H. and Weiss-Magasic, C. (1995). Effects of cobalt and pH on the growth of Chlamydomonas reinhardtii. Bull. Environ. Contam. Toxicol. 55,65 -72.[CrossRef][Medline]
Matsuda, Y., Koseki, M., Shimada, T. and Saito, T. (1995). Purification and characterization of a vegetative lytic enzyme responsible for liberation of daughter cells during the proliferation of Chlamydomonas reinhardtii. Plant Cell Physiol. 36,681 -689.[Medline]
Muller, M. L., Irkens-Kiesecker, U., Rubinstein, B. and Taiz,
L. (1996). On the mechanism of hyperacidification in lemon,
comparison of the vacuolar H+-ATPase of fruits and epicotyls.
J. Biol. Chem. 271,1916
-1924.
Nakamura, K., Nakamura, M., Yoshikawa, H. and Amano, Y. (2001). Purification and properties of thiosulfate dehydrogenase from Acidithiobacillus thiooxidans JCM7814. Biosci. Biotechnol. Biochem. 65,102 -108.[CrossRef][Medline]
Ohta, H., Shirakawa, H., Uchida, K., Yoshida, M., Matuo, Y. and Enami, I. (1997). Cloning and sequencing of the gene encoding the plasma membrane H+-ATPase from an acidophilic red alga, Cyanidium caldarium. Biochim. Biophys. Acta 1319, 9-13.[Medline]
Palmer, M. R., Pearson, P. N. and Cobb, S. J.
(1998). Reconstructing past ocean pH-depth profiles.
Science 282,1468
-1471.
Quarmby, L. M. and Hartzell, H. C. (1994). Two distinct, calcium-mediated, signal transduction pathways can trigger deflagellation in Chlamydomonas reinhardtii. J. Cell Biol. 124,807 -815.[Abstract]
Remis, D., Simonis, W. and Gimmler, H. (1992). Measurement of the transmembrane electrical potential of Dunaliella acidophila by microelectrodes. Arch. Microbiol. 158,350 -355.[CrossRef]
Sanders, D., Hansen, U. P. and Slayman, C. L.
(1981). Role of the plasma membrane proton pump in pH regulation
in non-animal cells. Proc. Natl. Acad. Sci. U.S.A.
78,5903
-5907.
Sekler, I., Gläser, H.-U. and Pick, U. (1991). Characterization of a plasma membrane H+-ATPase from the extremely acidophilic alga Dunaliella acidophila. J. Membr. Biol. 121, 51-57.[Medline]
Stock, C., Gronlien, H. K. and Allen, R. D. (2002). The ionic composition of the contractile vacuole fluid of Paramecium mirrors ion transport across the plasma membrane. Eur. J. Cell Biol. 81,505 -515.[Medline]
Sze, H., Li, X. and Palmgren, M. G. (1999).
Energization of plant cell membranes by H+-pumping ATPases:
regulation and biosynthesis. Plant Cell
11,677
-689.
Takai, M., Kamimura, K. and Sugio, T. (2001). A
new iron oxidase from a moderately thermophilic iron oxidizing bacterium
strain TI-1. Eur. J. Biochem.
268,1653
-1658.
Thomas, J. A., Cole, R. E. and Langworthy, T. A. (1976). Intracellular pH measurements with a spectroscopic probe generated in situ. Fed. Proc. 35, 1455.
van de Vossenburg, J. L. C. M., Driessen, A. J. M., Zillig, W. and Konings, W. N. (1998). Bioenergetics and cytoplasmic membrane stability of the extremely acidophilic, thermophilic archaeon Picrophilus oshimae. Extremophiles 2, 67-74.[CrossRef][Medline]
Weiss, M. and Pick, U. (1996). Primary
structure and effect of pH on the expression of the plasma membrane
H+-ATPase from Dunaliella acidophila and Dunaliella
salina. Plant Physiol. 112,1693
-1702.
Wetzel, R. G. (1975). Limnnology. Philadelphia: W. B. Saunders Company.
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