Continuous monitoring of the cytoplasmic pH in Methanobacterium thermoautotrophicum using the intracellular factor F420 as indicator

Peter von Felten1 and Reinhard Bachofen1

Institute of Plant Biology, University of Zürich, Zollikerstraße 107, CH-8008 Zürich, Switzerland1

Author for correspondence: Reinhard Bachofen. Tel: +41 1 634 82 80. Fax: +41 1 634 82 04. e-mail: bachofen{at}botinst.unizh.ch


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
The absorption spectrum of factor F420 changes depending on the pH and the redox state of the cytoplasm. Specific wavelengths were used to calibrate absorption changes to allow the measurement of changes in the cytoplasmic pH in Methanobacterium thermoautotrophicum. Upon a hydrogen pulse, a rapid efflux of protons was observed. Under these energized conditions, the {Delta}pH amounts to 0·2–0·4 pH units at pH 6·6, and 0·6–0·8 pH units at pH 6·0. It decays within 10–20 s. In parallel, a sodium gradient is formed which has a slightly longer lifetime. Both {Delta}pH and {Delta}{Psi} contribute to the proton-motive force present during methanogenesis. The energy-conversion rate, as indicated by the decay of the energized state of the cell, is fastest under growth conditions, i.e. at pH 6·9 and at a temperature of 58 °C.

Keywords: cytoplasm, pH gradient, intracellular pH, protonmotive force, methanogenic bacteria

Abbreviations: BESA, bromoethanesulfonic acid; CF, carboxyfluorescein; TCS, tetrachlorosalicylanilide


   INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
The pH of the cell cytoplasm is a critical parameter controlling a variety of cellular processes. In most organisms, the pH in the cytoplasm is maintained over a range of approximately two pH units around neutrality (Padan et al., 1981 ). However, this pH homeostasis is dependent on energy. The pH surrounding cells or cell aggregates is the main environmental factor that strongly determines growth and metabolism. It is crucial to the size of the pH gradient across the cell membrane, which forms an essential part of the proton-motive force driving biological energy conversion.

In suspension cultures of micro-organisms, the external pH is easily followed by using electrodes. In contrast, the determination of cytoplasmic pH in cells as small as bacteria is more difficult. Certain methods are based on the distribution of radiolabelled weak acids or bases whereby the cells (after incubation) are separated from the medium by rapid filtration or centrifugation. As sampling is periodic, the data from these methods are not continuous (Padan et al., 1981 ). Spectrometric methods, however, either absorption or fluorescence spectrometry, use specific pH indicators to monitor the cytoplasmic pH continuously.

The first descriptions of pH estimations using dyes (fluorogenic esters) were given by Thomas et al. (1979 , 1982 ) for tumour cells and bacteria. The prerequisites of the method are as follows: (1) the cells must be permeable to these colourless and non-fluorescent esters; (2) the indicators must be concentrated in the cells; (3) an intracellular esterase must cleave off an absorbing or fluorescing species; (4) the membrane must be impermeable to the negatively charged species formed by the hydrolysis (so that the indicator remains in the cells, at least for the duration of the experiment); (5) a calibration curve must be obtained; and (6) no other cellular compounds should interfere (by absorbance or fluorescence) with the marker compound. Generally, not all of these prerequisites are fulfilled. To prevent leakage of the pH-indicator, a dye forming covalent bonds with cytoplasmic compounds has been developed (Breeuwer et al., 1996 ). However, the question as to whether or not indicator dyes interact with cytoplasmic proteins and cause erroneous results remains open to debate (Yassine et al., 1997 ). To compensate for unequal uptake of the dye and varying esterase activity, fluorescence ratios from two wavelengths were analysed to follow the pH (Aono et al., 1997 ). Recently, fluorescence ratio microscopy imaging even made it possible to follow the pH of single bacterial cells in a mixed culture (Siegumfeldt et al., 1999 ). These studies demonstrate the versatility of spectroscopic techniques in the investigation of pH homeostasis and the dynamics of pH changes during energy transduction.

The metabolism (including energy transduction) of methanogenic bacteria has been studied intensively in recent decades, as reviewed, for example, by Deppenmeier et al. (1996) and Schäfer et al. (1999) . In the process of the stepwise reduction of CO2 to CH4 by H2, protons are extruded, giving rise to a pH gradient, which, along with a Na+ gradient and the membrane potential, is an important component of the driving force for ATP synthesis in a chemiosmotic mechanism. Changes in membrane potential upon the energization of whole cells of Methanobacterium thermoautotrophicum have been measured by Butsch & Bachofen (1984) . Using the dye carboxyfluorescein (CF), Bachofen & Butsch (1986) demonstrated, qualitatively, the formation of a pH gradient upon cell energization. The length of the signal correlated with the partial pressure of hydrogen gas introduced, whereas its magnitude was independent of the partial pressure over the range 20–80% (v/v) hydrogen in the gas mixture.

In the present work, the endogenous factor F420 was used as an intrinsic pH indicator. It fulfils most of the requirements cited above for a cytoplasmic pH indicator.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
All chemicals used were of highest analytical grade and were obtained from Fluka, Sigma or Merck. Gases were obtained from Carbagas.

Organism, growth medium and growth conditions.
M. thermoautotrophicum strain Hveragerdi (DSM 3590) was isolated earlier in our laboratory (Butsch & Bachofen, 1984 ). Stock cultures were kept at -80 °C. The medium was based on that of Schönheit et al. (1979) , as modified by Butsch & Bachofen (1984) , and contained the following: NH4Cl (40 mM), MgCl2.6H2O (1·5 mM), nitrilotriacetate (0·15 mM), NaCl (10 mM), KH2PO4 (10 mM), CoCl2.6H2O (1 µM), Na2MoO4.2H2O (1 µM), NiCl2.6H2O (1 µM) and FeCl2.4H2O (25 µM). Cells were grown in chemostat mode in a 2 l bioreactor equipped with controls for temperature, pH and redox potential, as described by Jud et al. (1997) . The temperature was held at 58 °C and the pH at 6·9. The culture was supplied with Na2S (310 mM) at intervals, producing a final concentration of 0·5 mM sulfide in the reactor. The gas supplied was H2/CO2, 80%:20% (v/v); the rate was controlled electronically and kept at 220 ml min-1 (equal to 0·12 vol. per vol. per min). Traces of oxygen were removed by a BASF catalyst, R0–20, sealed in an iron tube in the gas supply line. The sterile media were kept under nitrogen.

Spectroscopic investigations.
The absorption spectra of solutions were obtained with a Uvicon 810 spectrophotometer. Optical measurements of cell suspensions were obtained with an Aminco DW-2 dual-wavelength spectrophotometer using either specially made anaerobic cuvettes with rubber septa or a 30 ml minibioreactor built in our workshop and coupled to the DW-2 optics by light pipes. The spectrophotometer was connected to a computer with an ADALAB A/D converter (Interactive Microware).

Gas analysis.
Hydrogen, methane, oxygen, carbon monoxide and carbon dioxide were quantified by gas chromatography [using a Shimadzu GC-R1A with integrator RPR-G1 and a CSS column Carbosieve S 120/140 (Supelco)] with a TCD detector. The gas was supplied reproducibly to the cuvettes and the minibioreactor through a stainless steel needle as pulses of 10 s at a flow rate of 240 ml min-1 by a computer-controlled valve.

Cell preparations.
Cells were harvested by centrifugation (15 min at 1800 g, 4 °C) and washed twice with growth medium. All steps were performed under anoxic conditions using bottles flushed nine times with nitrogen (cycling between 0·5 and 2 bar). For the experiments, the cell concentration, measured as OD660, was set between 1·5 and 2, which is equivalent to 0·8–1·1 g cells (dry weight) l-1. The suspension was transferred either into an anaerobic cuvette equipped with a valve for gas pulses or into the minibioreactor in an anaerobic box (Forma Scientific 1024). To ensure the complete absence of oxygen, all glassware was kept in the anaerobic box for 24 h prior to the experiments. All manipulations were done under strictly anaerobic conditions, in closed vessels under purified nitrogen or in the anaerobic glove box. If these precautions for anoxic conditions were observed, the cells could be used reproducibly in experiments over a period of at least 4 h.

Measurement of the pHin.
The cuvette or the minibioreactor was stirred and kept at constant temperature under strictly anaerobic conditions during the measurements. The Aminco DW-2 spectrophotometer was used in the dual-wavelength mode, which allows the quantification of small absorption changes in the presence of a large optical background signal (cell scatter). The determination of the relevant wavelengths and the calibration of the intracellular pH (pHin) are described in Results.

Isolation of F420.
F420 was isolated, according to Cheeseman et al. (1972) and Schönheit et al. (1981) , by extracting the compound using 50% (v/v) acetone followed by chromatography twice on QAE-Sephadex A-25.

Other determinations.
The minibioreactor was equipped with electrodes for the continuous measurement of pH, redox potential and Na+ ions (Ingold). The OD600 of the cell suspension was calibrated by using dry-weight determination.


   RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Previous experiments using CF demonstrated a hydrogen-induced change in absorption which was interpreted as a rise in intracellular pH (Bachofen & Butsch, 1986 ). However, the spectrum of CF overlaps with that of the intrinsic electron carrier, F420, and no useful calibration was obtained with CF. Similarly, carboxynaphthofluorescein could not be used satisfactorily as an indicator for the intracellular pH, although it has an absorption maximum shifted 100 nm to longer wavelengths. Factor F420 has a defined absorption spectrum (Fig. 1) that changes upon reduction and also shows a typical fluorescence. This allows methanogens to be easily distinguished from other bacteria in environmental samples (Cheeseman et al., 1972 ). Furthermore, the signal has been found to be proportional to the biomass (Reuter et al., 1986 ). As the absorption spectrum of F420 is also dependent on pH (Cheeseman et al., 1972 ; Eirich et al., 1978 ), it can serve as a pH indicator to measure changes in the pH in the cytoplasm.



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Fig. 1. pH dependence of the absorption spectrum of isolated oxidized F420. Gas phase, N2; F420, 10 µg ml-1. Buffer used: pH 8–9, 50 mM Tris/HCl; below pH 8, 20 mM KH2PO4. Dashed line: F420 reduced with dithionite.

 
Fig. 1 shows absorption spectra in the visible range for the oxidized form of F420 at different pH values. To prevent irreversible degradation, the spectra of oxidized F420 were taken under nitrogen in the absence of an electron donor. A quasi-isosbestic point is obvious at 400 nm and maximum absorption changes are observed at 420 nm. As the spectrum of F420 is also dependent on its redox state, the absorption at the isosbestic point at 400 nm is a measure of the redox state independent of the pH. Full reduction of isolated F420 by dithionite causes the absorption peak to disappear. The change in absorption at 420 nm relative to that at 400 nm correlates well with the pH of the solution. Thus the absorption at 400 nm relative to the value of the fully oxidized state is indicative of the reduction state of F420. To compensate for variations in F420 concentrations in different preparations, both wavelengths, 400 and 420 nm, are related to the absorption at 450 nm as the reference wavelength [ratio (A420-A450)/(A400-A450)]. For isolated F420, the calibration curve is linear between pH 5·8 and pH 7·5. Using the same wavelengths, a calibration curve was obtained for whole cells of M. thermoautotrophicum. Schönheit & Beimborn (1985) and Bachofen & Butsch (1986) demonstrated that no measurable proton potential is present under de-energized conditions in the absence of an electron donor, and that the pHin is then in equilibrium with, and thus equal to, the external pH (pHout). Measurements were taken with cell suspensions equilibrated under nitrogen at medium pH (pHout) values between 5·8 and 7·2. This ratio method makes the pH determinations independent of the intracellular concentration of the marker, F420, and compensates for parallel absorption changes of unknown compounds or variable scattering effects of cell suspensions (Kotyk & Slavik, 1989 ).

The experiment presented in Fig. 2 is an example showing the kinetics of the changes in pHin as calculated from the calibration curve. The rapid alkalinization of the cytoplasm upon the substrate pulse cannot be visualized because bubbles produced during gas injection make optical measurements impossible. Under defined conditions, the duration of the alkalinization until the pHin returned to the initial pH of the medium was highly reproducible within the same cell preparation but it could vary between cell batches. The redox potential in the medium did not change after a hydrogen pulse (not shown). The size of the pH increase in the cytoplasm was dependent on the pHout: it is larger at acid values of the medium pH and becomes smaller towards neutrality. Around neutrality, Schönheit & Beimborn (1985) found, using M. thermoautotrophicum, that they could not measure a {Delta}pH under metabolically active conditions; in a more acidic environment (pH 5), however, the cytoplasm was found to be more alkaline, resulting in a pH gradient of 1–1·3 pH units under energized conditions. This is in close agreement with our observations. In contrast, the {Delta}pH reported by Dybas & Konisky (1992) for Methanococcus voltae under growing conditions was only a few millivolts.



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Fig. 2. Time course of the pHin after a 10 s H2/CO2 (80:20) pulse (black bar) at different medium pH values: A, pH 6·6; B, 6·0. Gas flow rate during pulse, 240 ml min-1; T, 42 °C; mean of five experiments.

 
The duration of the signal change upon hydrogen addition characterizes the length of time for which the cells stay in an energized state, conditions when a pH gradient and a membrane potential are present across the membrane. This is governed by the amount of H2 taken up by the cells, the concentration of H2 reached in the medium after flushing, and (mainly) by the speed at which the electrons are used up in cellular metabolism. The length of the signal is determined from the time immediately after hydrogen injection (pulses of H2/CO2, 80:20, 10 s duration) to the time at which 50% decay is reached; it was strongly dependent on environmental factors such as temperature (Fig. 3). The decay of the hydrogen-induced proton gradient is rapid between 45 °C and the growth temperature of 58 °C, but slows down rapidly at lower temperatures – a temperature-dependence similar to that of the growth rate of the organism.



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Fig. 3. Dependence on temperature of the half-life of the {Delta}pH after a 10 s H2/CO2 (80:20) pulse. Gas flow rate during pulse, 240 ml min-1, pH=6·8.

 
Control experiments with a nitrogen/CO2 mixture (80:20) and with inactivated denatured cells (stirred for 48 h under air, conditions in which F420 is rapidly inactivated and converted to factor390 (Hausinger et al., 1985 ; Schönheit et al., 1981 ) prove that the hydrogen-induced pH changes are driven by the bacterial metabolism.

It has been suggested that F420 is not homogeneously distributed within the cell but is, rather (for functional reasons) concentrated near the membrane at the site of the hydrogenase (Muth, 1988 ). Thus, an averaging method such as the distribution of weak acids and bases (Schönheit & Beimborn, 1985 ) or measurement with homogeneously distributed indicator dyes may not give the same values as a more localized pH indicator (Kotyk & Slavik, 1989 ). The pHin measured by F420 probably does not indicate the mean pH within the cell but, rather, represents the pH found close to the membrane. This would support the suggestion that the pumped protons are held along the membrane by anionic lipids and are not in equilibrium with the cytoplasm (Haines, 1983 ).

The concentration of Na+ ions in the medium, important for the formation of the {Delta}{Psi} in methanogens, was measured simultaneously. It increases when hydrogen is supplied, and the return to the original value is delayed relative to the decay of the {Delta}pH. The decay clearly accelerates after the {Delta}pH has dropped to below approximately 50% of the original size (Fig. 4). At the cell concentration chosen for the experiments, the ratio between the volume of medium and the volume of cells has been estimated to be approximately 600. Thus, an Na+ increase of a few millimoles per litre in the medium upon an H2 pulse would represent a drastic decrease in the ion concentration in the cell. Although most of the Na+ ions may have been bound to cell components, it indicates the formation of a noticeable membrane potential. Indeed, it has been suggested that Na+ and K+ ions are strongly complexed with specific lipids of the cell membrane (Kramer et al., 1988 ). The Na+ efflux is probably the result of a directly coupled Na+ pump, whereas the Na+/H+ antiporter driven by H+ extrusion during methanogenesis acts as a mechanism for regulating the pHin (Deppenmeier et al., 1996 ; Schäfer et al., 1999 ). Because of the presence of CO2 in the gas pulse, the pH of the medium drops slightly and returns slowly (within 30–40 s) to the original value before the pulse (Fig. 4).



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Fig. 4. Time course of intracellular and medium pH, and extracellular Na+ concentration after a 10 s H2/CO2 (80:20) pulse. Gas flow rate during pulse, 240 ml min-1; T, 42 °C; mean of five experiments.

 
Inhibitors of methanogenesis, such as the uncoupler tetrachlorosalicylanilide (TCS), the Na+ ionophore monensin, the inhibitor of the sodium–proton antiporter amiloride (for a review, see Kleyman & Cragoe, 1988 ), and the analogue of Coenzyme M and inhibitor of the methylreductase, bromoethanesulfonic acid (BESA), were tested for effects on the hydrogen-induced pHin changes. With TCS at 10 µM, the formation of a {Delta}pH was completely abolished, which is typical of an uncoupling action. The other reagents altered the reaction kinetics less drastically (Table 1). Monensin at 10 µM reduced the initial {Delta}pH upon the H2 pulse and slowed down the signal recovery after the pulse. BESA showed no effects on the size of the {Delta}pH up to 50 mM, but, again, the decay time also had increased. Amiloride at 10 µM had some effect on the initial size of the {Delta}pH and also retarded the decay kinetics. A prolonged signal duration is indicative of a longer lifetime of the pH gradient. In the presence of amiloride, the Na+/H+ antiporter, and thus the conversion of a {Delta}pH into a {Delta}Na+, is blocked. Similar effects of amiloride were noted by Müller et al. (1987) for Methanosarcina barkeri. Monensin abolishes part of the {Delta}pH, probably as a consequence of the higher Na+ permeability.


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Table 1. Effect of inhibitors on the size and the decay time of the {Delta}pH after a 10 s H2/CO2 (80:20, v/v) pulse

 
Conclusions
Following absorption changes of the endogenous marker F420 at three wavelengths allows the determination of pH changes in the cytoplasm of whole cells of methanogenic bacteria. The {Delta}pH is dependent on the external pH; its decay is determined by the metabolism and affected by external factors. Furthermore, it is influenced by inhibitors of proton and sodium transport.


   ACKNOWLEDGEMENTS
 
We thank the Swiss National Science Foundation for generous support (grant no. 32-27583.89), the unknown reviewers for their work in improving the manuscript, and D. Bollier for the construction of the minibiofermenter.


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
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Dybas, M. & Konisky, J. (1992). Energy transduction in the methanogen Methanococcus voltae is based on a sodium current. J Bacteriol 174, 5575-5583.[Abstract]

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Received 22 May 2000; revised 30 August 2000; accepted 12 September 2000.



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