Voltage-dependent Porin-like Ion Channels in the Archaeon Haloferax volcanii*

(Received for publication, August 7, 1996, and in revised form, September 26, 1996)

Madeleine Besnard , Boris Martinac Dagger and Alexandre Ghazi §

From the Laboratoire des Biomembranes, URA CNRS 1116, Bât. 430, Université Paris-Sud, 91405 Orsay, France and the Dagger  Department of Pharmacology, University of Western Australia, Nedlands WA 6907, Australia

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
Acknowledgments
REFERENCES


ABSTRACT

Membrane vesicles isolated from the cell envelope of the archaebacterium Haloferax volcanii were either reconstituted in giant liposomes and examined by the patch-clamp technique or were fused into planar lipid bilayers. In addition, cell envelope proteins were solubilized by detergent and reconstituted in azolectin liposomes, which were then fused into planar lipid bilayers. Independently of the technique used the predominant channel activity encountered exhibited the following characteristics. Channels were open at all voltages in the range approximately -120 to +120 mV and exhibited frequent fast transitions to closed levels of different amplitudes. At voltages greater than 120 mV the channels tended to close in a manner characterized by large, slow transitions of variable amplitudes. The tendency to close at high membrane potentials was much stronger at one polarity. The channel gating pattern was complex exhibiting a range of subconductances of 10-300 picosiemens in symmetric 100 mM KCl. These electrophysiological characteristics are comparable with those of bacterial and mitochondrial porins, suggesting that the archaeal channels may belong to the general class of porin channels. Some channels showed preference for K+, whereas the others preferred Cl-, suggesting the existence of at least two types of porin-like channels in H. volcanii.


INTRODUCTION

The phylogenetic tree is composed of three domains: Eucarya (eukaryotes), Bacteria (eubacteria), and Archaea (archaebacteria) (1). Archaebacteria comprises several different families of cells adapted to extreme environmental conditions (e.g. temperature, pH, salt concentration). Although eubacteria and archaebacteria are both prokaryotes, it is now generally accepted that archaebacteria are not closer, phylogenetically, either to eubacteria or to eukaryotes (2). Therefore, archaebacteria appear to constitute an intermediary domain between eubacteria and eukaryotes.

Ion channels have been mostly studied in eukaryotic cells (3) and have also been documented in eubacteria. In particular bacterial porins and mechanosensitive ion channels are well described. Porins, large water-filled pores across the outer membrane of Gram-negative bacteria (reviewed in Ref. 4), have also been found in the cell wall of certain Gram-positive bacteria such as mycobacteria (5, 6, 7). More recently, different mechanosensitive ion channels have been described in Gram-negative and Gram-positive bacteria (8, 9, 10, 11, 12, 13, 14, 15, 16, 17). These channels, most likely localized in the plasma membrane, are implicated in osmoregulation (13).

In contrast, the presence of ion channels in archaebacteria has not been documented. A search for ion channels in Archaea thus seems to be of interest not only for the understanding of the phylogeny of ion channels, but also for a better knowledge of the physiology of these unique organisms. For our studies we chose Haloferax volcanii (formerly Halobacterium volcanii), a moderate halophilic archaeon amenable to genetic studies.

We report here the existence of ion channels in the cell envelope of H. volcanii, that with regard to electrophysiological characteristics resemble bacterial porins. This finding indicates the importance of the porin family in the phylogeny of ion channels. Furthermore, it questions our current knowledge of the organization of the cell envelope of archaebacteria.


EXPERIMENTAL PROCEDURES

Cell Growth

H. volcanii (strain NCMB 2012) cells were grown in a medium containing (in mM) NaCl (3350), MgCl2 (170), MgSO4 (200), CaCl2 (6), KCl (26), NaHCO3 (6.6), NaBr (5.4) plus 5 g of yeast extract/liter. In some cases the medium was supplemented with MnCl2 (2.9), ZnSO4 (1.6), Cu SO4 (0.2), FeSO4 (1.5) plus 5 g of tryptone/liter.

Isolation of the Cell Envelope

One liter of cells was grown up to an A600 of 1 and harvested by centrifugation and the pellet resuspended in 10 ml of H20. Addition of EDTA (50 mM final concentration) resulted in lysis of most of the cells. To ensure total lysis, the suspension, supplemented with DNase (20 µg/ml) and MgCl2 (5 mM), was passed through a French press (8000 p.s.i.). The resulting suspension was centrifuged at 10,000 × g to eliminate cell debris. The supernatant was centrifuged for 1 h at 45,000 rpm, using a Ti45 Beckman rotor. The pellet was then resuspended in 10 mM Hepes-NaOH, pH 7.4, plus 5% ethylene glycol. Protein content was measured by the method of Lowry et al. (18). 50-µl aliquots (at 2-5 mg of protein/ml) were stored at -80 °C for further use.

Preparation of Giant Proteoliposomes

Cell envelopes were mixed with azolectin (from soybean, type II-S, Sigma) liposomes at a lipid to protein ratio (w/w) of 50 and fused into giant proteoliposomes by dehydration-rehydration as described previously (11). Rehydration was performed in 100 mM KCl, 10 mM Hepes-KOH, pH 7.4.

Solubilization and Reconstitution of Membrane Proteins into Proteoliposomes

Membrane proteins from cell envelopes were solubilized using octyl beta -D-glucoside; 100 µl of the cell envelope suspension was mixed with 900 µl of the solubilization buffer (10 mM Hepes-KOH, pH 7.4, 300 mM KCl, 1 mM dithiothreitol, 100 mM octyl beta -D-glucoside) and incubated for 20 min. The suspension was then centrifuged at 90,000 rpm for 40 min using a TL100 Beckman ultracentrifuge. The supernatant was added to a 1-ml suspension of asolectin liposomes (1 mg of lipids/ml) in 10 mM Hepes-KOH, pH 7.4, 300 mM KCl and incubated for 20 min before the addition of Bio-Beads SM-2 (Bio-Rad) (160 mg/ml). The suspension was gently agitated for 5 h, the Bio-Beads were discarded, and the suspension was centrifuged for 30 min at 90,000 rpm, using a TL100 Beckman ultracentrifuge. The pellet was then resuspended in 200 µl of 300 mM KCl, 10 mM Hepes-KOH, pH 7.4.

Electrophysiological Recording

Giant proteoliposomes were examined for channel activity using the standard patch-clamp method (19). 2 µl of the giant proteoliposomes suspension was added to the patch-clamp chamber and covered with 2 ml of 100 mM KCl, 10 mM Hepes-KOH, pH 7.4. Patch electrodes were pulled from pyrex capillaries (Corning code 7740) and were not fire-polished. They were filled with 10 mM Hepes-KOH, pH 7.4, 100 mM KCl, 2 mM CaCl2, 5 mM MgCl2. After gigaohm-seal formation on a giant liposome, the patch was excised, and unitary currents were recorded using a Biologic (Claix, France) RK-300 patch-clamp amplifier with a 10-gigaohm feedback resistance. The membrane potential refers to the potential in the bath minus the potential in the pipette.

Planar lipid bilayers were formed across a 250-µm diameter hole by presenting a bubble of asolectin lipids dissolved in n-decane (30 mg/ml) in front of the hole. After membrane formation, the mechanical and electrical stability of the membrane was monitored for 10 min before addition of crude cell envelopes or proteoliposomes to the cis compartment (8 µg protein/ml final concentration in both cases). Fusion to the planar bilayer was induced by imposing a salt gradient between the two chambers (500 mM KCl in the cis compartment versus 100 mM KCl in the trans compartment). The trans compartment was held at virtual ground potential and the membrane current amplified using a homemade device. All solutions were filtered using 0.22-µm Millipore filter.

Electrical recordings from patch-clamp or planar bilayer experiments were stored on a digital audio tape (Biologic DTR 1200 DAT recorder). Records were subsequently filtered at 1 kHz (-3 db point) through a 4-pole Bessel low-pass filter and digitized off-line at 2 kHz on a personal computer. Data were plotted by a Hewlett-Packard Laserjet printer.


RESULTS

Several attempts were made to record channel activities from native membranes of giant protoplasts of H. volcanii. Despite a concentrated effort to make the surface of the giant protoplasts amenable to patch-clamp recording, we were unable to form gigaohm-seal on their surface. Therefore, we decided to record ion channel activities by either reconstituting H. volcanii cell envelope in artificial liposomes or into planar bilayers.

The archaeal membrane vesicles were fused into giant liposomes by dehydration and rehydration (10, 11), and the giant liposomes were then examined by the patch-clamp technique. In parallel experiments the membrane vesicles were fused into planar lipid bilayers. In addition, cell envelope membrane proteins were solubilized using octyl beta -D-glucoside, reconstituted in liposomes that were then fused into planar lipid bilayers. Independently of the technique used the predominant channel activity encountered exhibited the characteristics described below.

Channels were mostly open at all voltages in the range -120 mV to +120 mV and exhibited frequent fast transitions to closed levels of different amplitudes (data not shown). At voltages greater than 120 mV the channels tended to close in a manner characterized by large, slow (seconds) transitions (Figs. 1 and 2). The greater the magnitude of the membrane potential, the shorter the open-time duration following the voltage step. Sporadic fast closures were often superimposed on the slow kinetic closures. For some patches the channels also entered a sustained fast gating mode (millisecond) as shown in Fig. 1.


Fig. 1. Closure of H. volcanii ion channels at positive bilayer potentials. Membrane proteins of the H. volcanii envelope were solubilized in octyl beta -D-glucoside and reconstituted in proteoliposomes that were fused into a planar lipid bilayer. After fusion, symmetrical media (500 mM KCl, 10 mM Hepes-KOH, pH 7.4) were established in the two chambers and the activity recorded. Voltage steps were applied from a holding potential of 0 mV, as indicated on the figure.
[View Larger Version of this Image (16K GIF file)]



Fig. 2. Closure of H. volcanii ion channels at negative bilayer potentials. The bilayer and recording solution were the same as in Fig. 1. Applied voltages are as indicated.
[View Larger Version of this Image (14K GIF file)]


The tendency to close at high membrane potentials was greater at one polarity (positive potential for bilayer experiments and negative potential for patch-clamp experiments), indicating both a preferential insertion of the channels in the lipid bilayer and an asymmetry in the voltage dependence.

Figs. 1 and 2 show recordings from the same bilayer at positive and negative voltages. Upon application of +180 mV across the bilayer the channels reached a closed inactivated state (Fig. 1). At negative potential, full closure of the channels could not be reached, and the time course of closure was much slower (Fig. 2). In most cases the inactivated state was reversible and the channels reopened by returning to 0 mV for several seconds (Fig. 1). However, in a few instances, the channels appeared to be locked in the inactivated state, and reopening could not be obtained at any membrane potential.

In general the channel gating was complex, showing a range of conductances in between 10 and 300 picosiemens in symmetric 100 mM KCl. While this could be attributed in part to a mixture of different channels (see below), in some cases clear cooperative events were observed, indicating that the channels can gate at different conductance states. This is illustrated in Fig. 3, which shows recordings obtained by two successive steps to -160 mV on the same patch. The most frequent transition had a conductance of 80 picosiemens, but a clear cooperative 160-picosiemens transition was also observed. Furthermore, channel gating at a subconductance level of approximately 10 picosiemens was observed.


Fig. 3. Multiplicity of subconductances of H. volcanii ion channels. Cell envelopes were fused into giant liposomes by dehydration-rehydration and studied by patch-clamp methods in symmetrical media (100 mM KCl, 10 mM Hepes-KOH, pH 7.4). The patch was subjected to voltage steps from a holding potential of 0 mV, as indicated in the figure.
[View Larger Version of this Image (22K GIF file)]


The selectivity of the channels was examined in bilayer experiments performed in asymmetric (500/100 mM KCl) and symmetric solution (500 mM KCl). I-V curves for unitary currents were difficult to obtain because of the different conductances encountered at each membrane potential. Instead, we chose to plot the total initial current through a given bilayer obtained during steps to various values of the membrane potential. Between steps the membrane potential was returned to 0 mV to ensure proper reopening of the channels. The current thus corresponded to that flowing through all the fully open channels present in a bilayer. In a series of nine different bilayer experiments, the following reversal potentials (in millivolts) were obtained: +24, +20, +20, +12, -4, -8, -14, -18, -23. The two extreme cases are shown in Fig. 4. These results suggest that at least two different types of porin-like channels with opposite selectivity are present in H. volcanii envelopes.


Fig. 4. Selectivity of H. volcanii channels. Membrane vesicles were fused into planar lipid bilayers, and recordings were performed in 500/100 mM KCl, 10 mM Hepes-KOH, pH 7.4, asymmetrical media (open circle ) (cis versus trans) and 500 mM KCl symmetrical solution (bullet ) after addition of concentrated KCl in the trans chamber. The total initial current through the bilayer obtained during steps to various potentials is plotted against the corresponding membrane potential. Between steps the membrane potential was returned to 0 mV to ensure proper reopening of the channels. Plots A and B are from two different bilayers. The reversal potentials were -23 and +24 mV, respectively.
[View Larger Version of this Image (16K GIF file)]



DISCUSSION

We report here for the first time the existence of ion channels found in the cell envelope of an archaebacterium. The electrophysiological characteristics of these channels, documented by two different techniques, can be summarized as follows: 1) the channels that have large conductances are maximally open at or around 0 mV and close at positive and negative voltages, 2) voltage dependence is asymmetric, 3) the channels exhibit both fast and slow kinetics, and 4) the channels exhibit several subconducting states. The same electrophysiological characteristics can be found in eubacterial porins as documented for OmpF and OmpC (20) and PhoE from Escherichia coli1 and also in mitochondrial porins (reviewed in Ref. 21). This suggests that all these channels may belong to the same family of functionally and/or structurally related membrane pores. The observation that these types of channels are found in all domains of the tree of life indicates the importance of the porin family in the phylogeny of ion channels.

What is the localization of these channels? Due to their characteristics, their presence in the cytoplasmic membrane is unlikely. Although the channels would normally be closed at the high negative membrane potential characteristic of microbial cells, any depolarization would result in the sustained opening of these high conductance channels, probably leading to cell death. Therefore it is likely that these porin-like channels are located in the cell envelope outside the plasma membrane, as in eubacteria.

In eubacteria, porins are located in the outer membrane of Gram-negative bacteria where they allow the diffusion of small hydrophylic compounds through this protection barrier (4). Gram-positive bacteria were long supposed to lack porin channels. However, porin-like channels have been recently identified in the cell wall of Gram-positive bacteria such as Mycobacterium chelonae (5, 6), Mycobacterium smegmatis (7), and Corynebacterium glutamicum (22). These bacteria are known to contain lipids in their cell wall in the form of mycolic acids, and it is possible that mycolic acids and other lipids form part of another bilayer (23), explaining the low permeability of the mycobacterial cell wall (24). Diffusion through this bilayer would be facilitated by porins.

Only one membrane, the plasma membrane, is documented in archaea such as Halobacterium halobium or H. volcanii. The main component of the cell wall is the S-layer formed by a hexagonal arrangement of a glycoprotein (25). A detailed structural study of the S-layer of H. volcanii (26) together with a a previous x-ray study of the envelope of H. halobium (27) led to a model in which the upper external part of the glycoprotein forms a dome-shaped region separated from the cell membrane by spacer elements (26). The protein is thought to be anchored in the plasma membrane by a hydrophobic stretch that is observed near the C terminus of both the H. halobium and H. volcanii glycoprotein sequence (28, 29). The interspace between the plasma membrane and the external part of the S-layer has been considered as an aqueous space analogous to the periplasm of Gram-negative bacteria (26, 27). However, if porins are present in the cell wall, but do not belong to the plasma membrane, they should be located in this interspace, which should then contain a lipid membrane. This membrane should represent in itself a permeability barrier through which movement of solutes would be facilitated by porins. Therefore, the discovery of porins in H. volcanii would suggest a more complex organization for the cell envelope of archaea than previously believed, possibly more similar to that of some Gram-positive eubacteria such as Mycobacteria (30).


FOOTNOTES

*   This work was supported by Australian Research Council Grant A19332733, CNRS, and a grant (to B. M.) from the French Ministry of Higher Education and Research. The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§   To whom correspondence should be addressed. Tel.: 33 1 69 15 71 94; Fax: 33 1 69 85 37 15; E-mail: ghazi{at}psisun.u-psud.fr.
1    Berrier, C., Besnard, M., and Ghazi, A. (1997) J. Membr. Biol., in press.

Acknowledgments

We thank P. Forterre, B. Labedan, and P. Lopez-Garcia for advice, discussions, and the gift of the H. volcanii strain. We also thank C. Berrier and A. C. Le Dain for critical reading of the manuscript.


REFERENCES

  1. Woese, C. R. (1994) Microbiol. Rev. 58, 1-9 [Abstract]
  2. Rowlands, T., Baumann, P., and Jackson, S. P. (1994) Science 264, 1326-1329 [Medline] [Order article via Infotrieve]
  3. Hille, B. (1992) Ionic Channels of Excitable Membranes, 2nd Ed., Sinauer Associates, Inc., Sunderland, MA
  4. Nikaido, H. (1994) J. Biol. Chem. 269, 3905-3908 [Free Full Text]
  5. Trias, J., Jarlier, V., and Benz, R. (1992) Science 258, 1479-1481 [Medline] [Order article via Infotrieve]
  6. Trias, J., and Benz, R. (1993) J. Biol. Chem. 268, 6234-6240 [Abstract/Free Full Text]
  7. Trias, J., and Benz, R. (1994) Mol. Microbiol. 14, 283-290 [Medline] [Order article via Infotrieve]
  8. Martinac, B., Buechner, M., Delcour, A. H., Adler, J., and Kung, C. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 2297-2301 [Abstract]
  9. Zoratti, M., and Petronilli, V. (1988) FEBS Lett. 240, 105-109 [CrossRef][Medline] [Order article via Infotrieve]
  10. Delcour, A. H., Martinac, B., Adler, J., and Kung, C. (1989) Biophys. J. 56, 631-636 [Abstract]
  11. Berrier, C., Coulombe, A., Houssin, C., and Ghazi, A. (1989) FEBS Lett. 259, 27-32 [CrossRef][Medline] [Order article via Infotrieve]
  12. Szabo', I., Petronilli, V., and Zoratti, M. (1992) Biochim. Biophys. Acta 1112, 29-38 [Medline] [Order article via Infotrieve]
  13. Berrier, C., Coulombe, C., Szabo', I., Zoratti, M., and Ghazi, A. (1992) Eur. J. Biochem. 206, 559-565 [Abstract]
  14. Sukharev, S. I., Martinac, B., Arshavsky, V. Y., and Kung, C. (1993) Biophys. J. 65, 1-7
  15. Szabo', I., Petronilli, V., and Zoratti, M. (1993) J. Membr. Biol. 131, 203-218 [Medline] [Order article via Infotrieve]
  16. Sukharev, S. I., Blount, P., Martinac, B., Blattner, F. R., and Kung, C. (1994) Nature 368, 265-268 [CrossRef][Medline] [Order article via Infotrieve]
  17. Berrier, C., Besnard, M., Ajouz, B., Coulombe, A., and Ghazi, A. (1996) J. Membr. Biol. 151, 175-187 [CrossRef][Medline] [Order article via Infotrieve]
  18. Lowry, D. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265-275 [Free Full Text]
  19. Hamill, O. P., Marty, A., Neher, E., Sakmann, B., and Sigworth, F. J. (1981) Pfluegers Arch. 391, 85-100 [Medline] [Order article via Infotrieve]
  20. Berrier, C., Coulombe, A., Houssin, C., and Ghazi, A. (1992) FEBS Lett. 306, 251-256 [CrossRef][Medline] [Order article via Infotrieve]
  21. Benz, R. (1994) Biochim. Biophys. Acta 1197, 167-196 [Medline] [Order article via Infotrieve]
  22. Niederweiss, M., Maier, E., Lichtinger, T., Benz, R., and Krämer, R. (1995) J. Bacteriol. 177, 5716-5718 [Abstract]
  23. Nikaido, H., Kim, S. H., and Rosenberg, E. Y. (1993) Mol. Microbiol. 8, 1025-1030 [Medline] [Order article via Infotrieve]
  24. Jarlier, V., and Nikaido, H. (1990) J. Bacteriol. 172, 1418-1423 [Medline] [Order article via Infotrieve]
  25. Lechner, J., and Wieland, F. (1989) Annu. Rev. Biochem. 58, 173-194 [CrossRef][Medline] [Order article via Infotrieve]
  26. Kessel, M., Widhaber, I., Cohen, S., and Baumeister, W. (1988) EMBO J. 7, 1549-1554
  27. Blaurock, A., Stoeckenius, W., Oesterhelt, D., and Scherphof, G. (1976) J. Cell Biol. 71, 1-22 [Abstract]
  28. Sumper, M., Berg, E., Mengele, R., and Strobel, I. (1990) J. Bacteriol. 172, 7111-7118 [Medline] [Order article via Infotrieve]
  29. Lechner, J., and Sumper, M. (1987) J. Biol. Chem. 262, 9724-9729 [Abstract/Free Full Text]
  30. Brennan, P. J., and Nikaido, H. (1995) Annu. Rev. Biochem. 64, 29-63 [CrossRef][Medline] [Order article via Infotrieve]

©1997 by The American Society for Biochemistry and Molecular Biology, Inc.