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Research Article
13 March 2019

Purification of a Crenarchaeal ATP Synthase in the Light of the Unique Bioenergetics of Ignicoccus Species


In this study, the ATP synthase of Ignicoccus hospitalis was purified, characterized, and structurally compared to the respective enzymes of the other Ignicoccus species, to shed light on energy conservation in this unique group of archaea. The crenarchaeal genus Ignicoccus comprises three described species, i.e., I. hospitalis and Ignicoccus islandicus from hot marine sediments near Iceland and Ignicoccus pacificus from a hydrothermal vent system in the Pacific Ocean. This genus is unique among all archaea due to the unusual cell envelope, consisting of two membranes that enclose a large intermembrane compartment (IMC). I. hospitalis is the best studied member of this genus, mainly because it is the only known host for the potentially parasitic archaeon Nanoarchaeum equitans. I. hospitalis grows chemolithoautotrophically, and its sole energy-yielding reaction is the reduction of elemental sulfur with molecular hydrogen, forming large amounts of hydrogen sulfide. This reaction generates an electrochemical gradient, which is used by the ATP synthase, located in the outer cellular membrane, to generate ATP inside the IMC. The genome of I. hospitalis encodes nine subunits of an A-type ATP synthase, which we could identify in the purified complex. Although the maximal in vitro activity of the I. hospitalis enzyme was measured around pH 6, the optimal stability of the A1AO complex seemed to be at pH 9. Interestingly, the soluble A1 subcomplexes of the different Ignicoccus species exhibited significant differences in their apparent molecular masses in native electrophoresis, although their behaviors in gel filtration and chromatography-mass spectrometry were very similar.
IMPORTANCE The Crenarchaeota represent one of the major phyla within the Archaea domain. This study describes the successful purification of a crenarchaeal ATP synthase. To date, all information about A-type ATP synthases is from euryarchaeal enzymes. The fact that it has not been possible to purify this enzyme complex from a member of the Crenarchaeota until now points to significant differences in stability, possibly caused by structural alterations. Furthermore, the study subject I. hospitalis has a particular importance among crenarchaeotes, since it is the only known host of N. equitans. The energy metabolism in this system is still poorly understood, and our results can help elucidate the unique relationship between these two microbes.


ATP synthases are among the most important enzymes of living cells (1). Generally, they function in both directions (synthesizing and hydrolyzing) and therefore are often named ATPases. In this article, we refer to them as ATP synthases, pointing to their function in synthesizing ATP. The term ATPase is used only to describe the A1 subcomplex, which is also able to hydrolyze ATP, or if the function is not clear, as in the case of Nanoarchaeum equitans. The enzymes are found in all three domains of life and exhibit great structural and functional similarity (2). The F-type ATP synthases are found in bacteria, mitochondria, and chloroplasts and are used for ATP synthesis (3), whereas the V-type ATPases from eukaryotes function as ATP-driven ion pumps in vivo (4). The A-type ATP synthases are found in archaea as well as in some bacteria (i.e. Thermus thermophilus and Enterococcus hirae), likely due to horizontal gene transfer (5). To date, all structural information about A-type ATP synthases has been derived from investigations of euryarchaeal enzymes. Possibly best studied are the ATP synthases from Methanosarcina mazei (6, 7), Methanocaldococcus jannaschii (8), and Pyrococcus furiosus (9, 10). Like all members of this class of enzymes, the A-type ATP synthases consist of a soluble A1 subcomplex that contains the catalytic subunits and a membrane-embedded AO subcomplex that is responsible for ion translocation. The proposed subunit composition is A3B3CDE2FH2acx, and the structure and function have been reviewed in detail by Grüber et al. (11).
Gathering information about the ATP synthases in crenarchaeotes has been much more challenging. The genes for the single subunits are scattered across the genome and are not organized in an operon as in Euryarchaeota; therefore, they are harder to identify (12). Attempts to purify the respective enzymes from different crenarchaeotes have not been successful, resulting merely in subcomplexes of various sizes (1316). Thus, the ATP synthases of the crenarchaeotes seem to differ significantly from the euryarchaeal enzymes, concerning their stability and possibly also their structure.
The crenarchaeal genus Ignicoccus comprises three described species (Ignicoccus hospitalis Kin4/IT, Ignicoccus islandicus Kol8T, and Ignicoccus pacificus LPC33T) (17, 18) and is unique among Crenarchaeota genera mostly because of its unusual cell architecture. The cell envelope consists of two membranes instead of a cytoplasmic membrane and an S-layer, as in many other crenarchaeotes (1719). The cytoplasm, with DNA and ribosomes, is surrounded by the inner membrane (IM), followed by a large intermembrane compartment (IMC) and a so-called outer cellular membrane (OCM) (20). I. hospitalis is the best-studied member of this genus, mainly because it is the only known host for Nanoarchaeum equitans and thus is of particular interest. N. equitans cells are tiny cocci, with diameters of 350 to 500 nm, that thrive only in direct contact with I. hospitalis cells (21). It has been shown that lipids, amino acids, and even some proteins are transferred from I. hospitalis to N. equitans (2224). However, the energy metabolism in this intimate association is still poorly understood, although all annotated subunits (A, B, D, a, and c) of a rudimentary ATP synthase/ATPase of N. equitans, except subunit c, were detected in proteome studies (25). Whether this enzyme is an actual ATP synthase capable of synthesizing ATP or merely acts as an ATPase (an ATP-driven proton pump) is still an open question. Recent studies on the A3B3 hexameric complex of N. equitans expressed in Escherichia coli suggest that this complex is probably inactive and N. equitans may not be able to synthesize ATP in vivo (26).
The genome of I. hospitalis encodes all essential subunits of an A-type ATP synthase except for subunit H (25). The localization of the ATP synthase is a unique feature of I. hospitalis. It was shown that the ATP synthase and the sulfur reductase are both located in the OCM. Thus, energy conservation takes place in the IMC (27). With the acetyl coenzyme A (acetyl-CoA) synthetase, an energy-consuming process is located in the IMC as well (20). Whether this particular localization is a feature common to all Ignicoccus species, as proposed by Huber et al. (20), or a unique trait of I. hospitalis, possibly related to its association with N. equitans, is not unambiguously resolved to date.
In order to learn more about the ATP synthase of I. hospitalis, the aim of the present study was the purification and characterization of this enzyme. Furthermore, the ATP synthases of the different Ignicoccus species were compared, to determine possible differences and similarities in their structures and subunit compositions.


Purification of the ATP synthase of I. hospitalis.

Previous attempts by our group to purify the ATP synthase complex in a Tris-based buffer at pH 8.0 did not result in a complete enzyme complex (27, 28). Since preliminary experiments with the hydrogenase complex of I. hospitalis, which is located in the OCM like the ATP synthase, revealed optimal activity of the purified enzyme at pH 8.5 (A. Kletzin, unpublished data), the pH of the buffer was even further elevated to pH 9.0 for the present study.
Approximately 10 g of frozen I. hospitalis cells was resuspended in 40 ml lysis buffer (pH 9.0) (see Materials and Methods) and disrupted by three passages in a French press at 3.5 MPa. Solid components of the lysate were centrifuged down and homogenized in lysis buffer on ice. The pellet was then applied to a sucrose gradient. A distinct band of membranes was visible at approximately 60% sucrose after ultracentrifugation and was extracted from the tube with a syringe. The presence of the ATP synthase in the fractions was verified by ATP hydrolysis activity assays and Western blotting. Membrane proteins were solubilized for 2 h at room temperature with 1 mg n-dodecyl-β-d-maltopyranoside (DDM) per mg protein and then were separated from lipids by ultracentrifugation.
The solubilizate was diluted with AEX-A buffer (pH 9.0) (see Materials and Methods) to a final volume of 50 ml and was purified by anion-exchange chromatography. For elution, the conductivity of the buffers was increased stepwise from 9.0 to 12.5 to 16.0 mS/cm, which was achieved by manual buffer exchange, followed by a gradient to 100% AEX-B buffer. Two protein peaks with ATP hydrolysis activity eluted at conductivities of 12.5 mS/cm and 16 mS/cm (see Fig. S1 in the supplemental material). Fractions of the peak in box 1 were combined and showed a prominent band in clear native electrophoresis (CNE) at an apparent molecular mass of 440 kDa (Fig. S1, inset). This band had been identified previously as the A1 subcomplex of the ATP synthase (27) and therefore was not investigated further. Analysis of the combined fractions of peak 2 resulted in two major bands in CNE, with apparent molecular masses of 440 kDa and 669 kDa (Fig. 1A). The 440-kDa complex showed ATP hydrolysis activity in an in-gel assay (Fig. 1B), representing the A1 subcomplex of the ATP synthase, as described previously (27). The complex contained the subunits A, B, D, and E, as determined by peptide mass fingerprinting (data not shown). The 669-kDa complex was analyzed further; it did not show ATP hydrolysis activity in in-gel assays and showed only low activity in colorimetric cuvette assays. All annotated subunits (Table 1), except for subunit F, of the ATP synthase of I. hospitalis were identified (Fig. 1C) after separation of the proteins in an SDS-PAGE gel and analysis of distinct proteins by peptide mass fingerprinting (see the supplemental material). The fate of the accessory subunit F of the ATP synthase is unclear to date. There were only few other bands, representing mostly hypothetical proteins (e.g., the protein Igni_0533, between 25 and 35 kDa). Therefore, all subunits of the ATP synthase except subunit F eluted collectively from the column, most likely as a coupled complex. As the dissociation into two bands in CNE already indicates, the enzyme complex seems to be highly unstable and dissociates easily into its subcomplexes even during electrophoresis. The same phenomenon occurs during gel filtration of the 669-kDa complex, with two main protein peaks being visible (Fig. 2, arrows 1 and 2). According to their respective protein patterns in CNE (Fig. 2, inset), the peak of higher molecular weight (peak 1) represents the intact enzyme, with an apparent molecular mass of 669 kDa. The peak of lower molecular weight (peak 2) shows the familiar protein band in CNE at an apparent molecular mass of 440 kDa, which represents the A1 subcomplex of the ATP synthase (27). As shown by our data, the protein complex dissociates into its subcomplexes very easily. Due to this instability, it has not been possible to date to obtain meaningful electron micrographs or protein crystals for structure determination. Therefore, there is no molecular or structural explanation for such instability.
FIG 1 Purified ATP synthase of I. hospitalis after anion-exchange chromatography. (A) CNE, with Coomassie blue staining. (B) ATP hydrolysis in-gel assay in a CNE gel. (C) SDS-PAGE, with Coomassie blue staining. Subunits of the ATP synthase identified by peptide mass fingerprinting are labeled in red.
TABLE 1 Subunits of the ATP synthase of I. hospitalis as annotated by Podar et al. (25) and identified by our group (shaded in gray)
SubunitaGene no.Molecular mass (kDa)GenBank accession no.
Difficulties identifying proteins like subunit H using standard algorithms are a well-known problem, due to the low level of conservation (12). Our group was able to identify subunit H in the genome of I. hospitalis during the purification process for the A1 subcomplex (28), by manual sequence comparison after peptide mass fingerprinting.
FIG 2 Size exclusion chromatography of the ATP synthase of I. hospitalis. Blue, UV absorption in arbitrary units; red, ATP hydrolysis activity in arbitrary units. Arrows 1 and 2 mark the two major protein peaks. Inset, CNE of the combined fractions for protein peaks 1 and 2.

pH dependency of ATP hydrolysis activity in I. hospitalis.

Membranes from the sucrose gradient were subjected to activity tests after incubation with Nʹ,Nʹ-dicyclohexylcarbodiimide (DCCD), an inhibitor that is known to affect only the coupled enzyme complex (consisting of A1 and AO subcomplexes) (29, 30). Thus, it was ensured that only samples harboring the coupled enzyme complex (intact ATP synthases) were used to test the pH dependency of ATP hydrolysis activity. Activity reached its maximum at pH 6.5 (Fig. 3). At pH 9.0, where purification was carried out, the activity of the coupled enzyme complex was still 40% of its maximum; at pH 8.0, where standard activity tests were performed, the activity reached 73% of its maximum.
FIG 3 pH dependency of ATP hydrolysis activity of the ATP synthase of I. hospitalis. The specific activity is plotted against pH values. Data are from one experiment, in which the different pH conditions were tested in triplicate.

Comparison of the purified A1 subcomplexes of different Ignicoccus species.

Analogous to the A1 subcomplex of the ATP synthase of I. hospitalis, the respective subcomplexes of I. islandicus and I. pacificus were purified by anion-exchange chromatography at pH 9.0. Specific ATP hydrolysis activity was quantified to 20.7 U/mg (I. hospitalis), 16.3 U/mg (I. islandicus), and 13.8 U/mg (I. pacificus) in a photometric assay. This activity could be inhibited by 1.0 mM diethylstilbestrol (DES) to 24% (I. hospitalis), 25% (I. islandicus), and 35% (I. pacificus) of the initial values. In contrast, 0.25 mM DCCD led to decreases in activity only to 79% (I. hospitalis), 82% (I. islandicus), and 86% (I. pacificus). DES specifically inhibits the activity of the catalytic A1 subcomplex, even when separated from the AO subcomplex (31), while DCCD is known to bind to the c ring of the AO subcomplex and hence affects only the activity of the coupled enzyme (29, 30). Therefore, these results confirm the presence of only the A1 subcomplexes of the respective enzymes.
The three subcomplexes showed very similar behavior in anion-exchange chromatography, leading to elution profiles comparable to that in Fig. S1 in the supplemental material. In CNE, however, the three purified A1 subcomplexes differed significantly in their apparent molecular masses (Fig. 4A and B). The complex from I. hospitalis contained subunits A, B, and E of the soluble A1 part of the ATP synthase, as determined by matrix-assisted laser desorption ionization–time of flight (MALDI-TOF) tandem mass spectrometry (MS/MS) after digestion with trypsin (Fig. 4C, lane 1). The complex from I. islandicus contained the subunits A and B, as well as an aggregate of about 35 kDa. Analysis by MALDI-TOF MS/MS revealed peptide fragments of both subunit F and subunit B. Due to the smaller molecular mass, we expect the aggregate to consist of subunit F and partly degraded subunit B (Fig. 4C, lane 2). In the complex from I. pacificus, however, only subunits A and B could be identified by MALDI-TOF MS/MS (Fig. 4C, lane 3). Accordingly, all three complexes showed a strong reaction with antiserum against the A1 subcomplex of the I. hospitalis ATP synthase (Fig. 4B). To investigate whether the different apparent molecular masses are based on significant differences of surface charges, blue native electrophoresis (BNE) was performed. During BNE, the negatively charged dye Coomassie blue G-250 binds to the proteins without changing their conformation, applying an equal negative charge and thus ensuring that the protein complexes are separated according to their sizes (32). Here, the A1 subcomplexes showed the same difference in apparent molecular masses as in CNE (Fig. 5).
FIG 4 Detection and subunit composition of the A1 subcomplexes of different Ignicoccus species after anion-exchange chromatography (Kin4/I, I. hospitalis; Kol8, I. islandicus; LPC33, I. pacificus). (A) CNE, with Coomassie staining, with the major protein complexes labeled 1 to 3. (B) Immunoblot of a CNE gel using an antibody against the A1 subcomplex of I. hospitalis (dilution, 1:5,000). (C) 2D native SDS-PAGE, with silver staining, of the labeled complexes from panel A. Subunits identified by peptide mass fingerprinting are labeled in red.
FIG 5 BNE of the A1 subcomplexes of different Ignicoccus species, with Coomassie staining (Kin4/I, I. hospitalis; Kol8, I. islandicus; LPC33, I. pacificus). BNE allows the separation of proteins according to their sizes alone, eliminating surface charges using the Coomassie dye.
During gel filtration, the protein complexes with ATP hydrolysis activity eluted at the same elution volume (data not shown). This indicates that the protein complexes indeed had the same molecular masses and the differences in CNE and BNE have other reasons. Native mass spectrometry (MS) data (Fig. S2 to S7) supported this hypothesis. In laser-induced liquid bead ion desorption (LILBID) MS experiments, no differences with respect to the masses of the intact complexes or individual proteins were observed when two different buffers (lysis buffer at pH 9.0 and NH4HCO3 buffer at pH 8.0) were used. The only change observed was the signal-to-noise ratio. For the A1 subcomplex from I. hospitalis, a molecular mass of 392 kDa was determined (Fig. S3). Similarly, the molecular masses of the complexes from I. islandicus and I. pacificus were 392 kDa and 392 kDa, respectively (Fig. S5 and S7). In addition, the complexes seemed to consist of the subunits A, B, E, and F, while the masses of the A1 subcomplexes suggest an A3B3EF stoichiometry. This is consistent with the results from the two-dimensional (2D) SDS-PAGE and analysis by peptide mass fingerprinting (Fig. 4C).

Sequence comparison of the two major subunits, A and B.

As a consequence of the former results, multiple sequence alignments of the subunits A and B were generated using Clustal Omega ( Alignments of amino acid sequences showed very high levels of identity (between 89% and 91% for subunit A and between 92% and 94% for subunit B) among the three Ignicoccus species (Fig. S8 and S9). The preliminary genome sequences of I. islandicus and I. pacificus were kindly provided by Mircea Podar prior to the GenBank release. Genome data from Integrated Microbial Genomes was used, because GenBank annotations are missing for some subunits due to different gene-calling algorithms. According to the sequence data, the proposed molecular masses of subunits A and B are quite similar (subunit A, 66.6 kDa [I. hospitalis], 65.9 kDa [I. islandicus], and 66.2 kDa [I. pacificus]; subunit B, 52.2 kDa [I. hospitalis], 53.3 kDa [I. islandicus], and 52.2 kDa [I. pacificus]). This finding is in accordance with the 2D SDS-PAGE results (Fig. 4C), in which the respective subunits were found at identical apparent molecular weights.


ATP synthase of I. hospitalis.

This paper describes the first purification of a crenarchaeal ATP synthase (with only subunit F missing). Previous attempts at purifying this enzyme complex from members of the Crenarchaeota resulted in much smaller subcomplexes of various subunit compositions. From Sulfolobus acidocaldarius, ATPases consisting of three subunits, with molecular masses of 65, 51, and 20 kDa (33) or 69, 54, and 28 kDa (14), were isolated. Similarly, the purified ATPase from Sulfolobus solfataricus consisted of three subunits (63, 48, and 24 kDa [15]). Purification of the ATP synthase/ATPase of Pyrodictium abyssi resulted in a complex consisting of six subunits, with molecular masses of 67, 51, 41, 26, 22, and 7 kDa (c subunit) (16). Previous purification experiments with the ATP synthase/ATPase of I. hospitalis, carried out at pH 8.0 in Tris buffer, resulted only in a subcomplex with the subunits A, B, E, and F (28). The current results, however, suggest that the ATP synthase of I. hospitalis elutes from the anion-exchange column as a coupled complex when the procedure is carried out under the described conditions at pH 9.0. However, the complex is extremely unstable and easily dissociates into its subcomplexes, as native electrophoresis and gel filtration chromatography experiments suggest. The dissociation seems to be not quantitative, leading to a mixed population of protein complexes of different sizes. Therefore, structural analyses using electron microscopy or X-ray crystallography have not been possible. Attempts to stabilize the complex by chemical fixation using GraFix gradient centrifugation (34, 35) have not been successful. Furthermore, different purification approaches, such as precipitation using polyethylene glycol, did not lead to a more stable ATP synthase complex (data not shown). Overall, purification at an elevated, more alkaline pH seems to be quite favorable for the stability of the I. hospitalis ATP synthase, as we were able to detect all subunits of the enzyme complex in the same fractions. Whether this is true for all crenarchaeotes or is related to the unique cell structure of I. hospitalis cannot be definitively resolved to date.
I. hospitalis has a streamlined genome and is missing many genes, e.g., for metabolic enzymes that are present in most crenarchaeotes (25). It exhibits low variability of metabolic pathways (e.g., no substrate-level phosphorylation) and therefore is a suitable organism for investigations of ATP synthase, as this is its only known mechanism for energy conservation. A typical archaeal ATP synthase has a stoichiometry of A3B3CDE2FH2acx, with subunit c being present in multiple copies (typically 10 to 15 for an 8-kDa subunit) (11). Assuming that in I. hospitalis the enzyme possesses 10 c subunits to form the c ring in the membrane, the whole complex would have a molecular mass of 680 to 690 kDa. This calculated mass is in agreement with the observed apparent molecular mass of about 670 kDa in CNE (Fig. 1A).

Physiology of the IMC.

The results may also shed light on the nature of the IMC. As a space between the IM and the OCM, it was long thought to be mostly empty (not electron dense, except for vesicles [18, 19]). With the findings that ATP synthase and sulfur-hydrogen oxidoreductase (27), as well as acetyl-CoA synthetase (36), are located in/associated with the OCM, it became clear that several essential cellular processes are located in the IMC. Further studies indicate that cytochromes are likely involved in electron transport within the OCM (37). However, there is not much information about the chemical composition (e.g., predominant ions and pH) in the IMC. Preliminary characterization of the hydrogenase subcomplex of sulfur-hydrogen oxidoreductase revealed optimal activity at pH 8.5 (A. Kletzin, personal communication). Additionally, a transmembrane electrochemical ion potential of −180 mV is necessary to drive ATP synthesis (38), which equals a ΔpH value of 3 units. This was confirmed in experiments in which an artificial ΔpH of 2 to 3, applied to I. hospitalis cells, was sufficient to drive ATP synthesis (39). Since the medium of I. hospitalis has a pH of 5.5 to 6.0 (40), a pH of 8.5 to 9.0 in the IMC could be physiological. The results on the pH dependency of ATP hydrolysis activity further support this hypothesis. Although the greatest in vitro ATP hydrolysis activity of I. hospitalis membranes was detected at pH 6.5, there was still significant measurable activity at pH 9.0. Moreover, the curve showed a shift of 0.5 to 1.3 units toward a higher pH, compared to similar investigations of ATP synthases from members of the Euryarchaeota (i.e. Methanosarcina mazei, Methanocaldococcus jannaschii, and Pyrococcus furiosus) (79). Therefore, we hypothesize that the pH in the IMC is indeed somewhat alkaline.
In addition, it seems clear that the IMC is an ATP-rich environment. This could have consequences in terms of the intimate association of I. hospitalis and N. equitans, as the latter possesses only five genes for an ATP synthase/ATPase (A, B, D, a, and c). This may be enough to function as an ATPase but not as an ATP synthase. Therefore, we suggest that N. equitans receives ATP from the IMC of its host I. hospitalis, meeting its energetic demand.

ATP synthases within the genus Ignicoccus.

All described Ignicoccus species exhibit very similar chemolithoautotrophic lifestyles, as well as similar cell architectures (17, 18). However, there are certain differences in the protein composition of the OCM. For example, the protein Ihomp1 is found in abundance in the OCM of I. hospitalis (19, 41) but not in other Ignicoccus species (20). Freeze-etching studies showed protein patterns similar to that of Ihomp1 in the OCM of those species (R. Rachel, unpublished data), but the proteins could not be labeled with a specific antibody against Ihomp1 (20) and the respective genes were not identified in their genomes. Based on the preliminary annotated genomes of I. islandicus and I. pacificus, the species likely both possess a fully functional ATP synthase, although subunit H could not be annotated yet. This is a rather common problem, due to the low level of conservation in this accessory subunit, which hinders the identification by sequence comparison (12). We attempted to purify the intact ATP synthase complexes for both I. pacificus and I. islandicus, for comparison of the whole enzyme complexes. However, the complexes turned out to be even more unstable than the I. hospitalis complex. Therefore, we proceeded to investigate the A1 subcomplexes in detail. The catalytic subunits A and B show an exceptionally high level of identity among the three species (see Fig. S8 and S9 in the supplemental material). A striking difference of more than 100 kDa in the apparent molecular mass of the A1 subcomplexes was observed after CNE (Fig. 4A). The reproduction of this result in BNE, a method in which a negative charge is applied to all protein complexes by the Coomassie dye, eliminated a difference in the surface charge as a possible cause of the observed migration behavior. Moreover, similar distributions of charged and hydrophobic amino acids would be expected, due to the high level of sequence identity of the major subunits A and B. All other studies, including gel filtration and LILBID (native) MS, did not point to an actual difference in size or subunit composition of the enzyme complexes, nor was there evidence that the single subunits varied in their respective molecular masses. Therefore, we have no explanation for the unusual migration behavior of the A1 subcomplexes during native PAGE (CNE and BNE) to date.


Cultivation conditions.

Cells of the type strains of I. hospitalis (Kin4/IT), I. pacificus (LPC33T), and I. islandicus (Kol8T) were grown in anaerobic 0.5× artificial seawater medium at 90°C, as described previously (18, 27). For large-scale cultivation in a 300-liter enamel-protected bioreactor, yeast extract was added to a final concentration of 0.1% (wt/vol). Cells were harvested aerobically by flow-through centrifugation (Padberg, Lahr, Germany), frozen in liquid nitrogen, and stored at −80°C.

Membrane preparation.

Frozen cells (approximately 0.25 g/ml) were resuspended in lysis buffer [25 mM N-(1,1-dimethyl-2-hydroxyethyl)-3-amino-2-hydroxypropanesulfonic acid [AMPSO], 10 mM MgCl2, 0.1 mM phenylmethylsulfonyl fluoride [PMSF] [pH 9.0]] containing 0.1 mg DNase I/ml and were disrupted by three passages in a French press at 3.5 MPa. Solid components of the lysate were centrifuged at 60,000 × g for 1 h at 4°C. After homogenization in lysis buffer on ice, the pellet was applied to a sucrose gradient (20% to 70% [wt/vol] sucrose in lysis buffer for I. hospitalis or 50% to 80% [wt/vol] sucrose in lysis buffer for I. pacificus and I. islandicus). Following ultracentrifugation (210,000 × g for 4 h at 10°C), membranes were extracted from the tube with a syringe. Protein concentrations were determined with a bicinchoninic acid (BCA) protein assay kit (Pierce, USA), according to the manufacturer’s instructions.

Solubilization of membrane proteins.

One milligram DDM per mg protein was added to the membrane fraction, leading to a final concentration of approximately 0.5% (wt/vol). After incubation in 1-ml aliquots for 2 h at room temperature with gentle mixing, lipids were removed by ultracentrifugation (110,000 × g for 1.5 h at 4°C).

Purification of the ATP synthase.

The solubilizate was diluted with AEX-A buffer (25 mM AMPSO, 10 mM MgCl2, 10% [vol/vol] glycerol, 0.05% [wt/vol] DDM, 0.1 mM PMSF [pH 9.0]) to a final volume of 50 ml and was purified by anion-exchange chromatography at 4°C using a HiTrap Q HP column and the ÄKTA fast protein liquid chromatography (FPLC) system (GE Healthcare, UK). Proteins were eluted by increasing the conductivity stepwise from 9.0 to 12.5 to 16.0 mS/cm (adjusted by titration of AEX-A buffer and AEX-B buffer [AEX-A buffer with 1 M NaCl] at 18°C), which was achieved by manual buffer exchange, followed by a gradient to 100% AEX-B buffer. Protein fractions were concentrated if necessary (Vivaspin; Sartorius Stedim Biotech, Germany). For separation of the proteins according to their sizes, a Superdex 200 gel filtration column and AEX-A buffer containing 200 mM NaCl were used.

ATP hydrolysis activity assay.

ATP hydrolysis activity was determined in triplicate in a colorimetric cuvette assay at 90°C, as described previously (9), in slightly modified reaction buffer [100 mM Tris, 100 mM 2-(N-morpholino)ethanesulfonic acid [MES], 10 mM MgCl2, 40 mM K2S2O5, 200 mM KCl [pH 8.0]]. For determination of the pH optima (ranging from pH 6.0 to pH 9.5) 100 mM HEPES was added to the reaction buffer. For inhibition assays, specific inhibitors of the ATP synthase, namely, DES and DCCD, were added to the reaction tube at their respective concentrations (1 mM for DES and 0.25 mM for DCCD) and incubated for 30 min at room temperature prior to the activity assay. The inhibitors were dissolved in 100% ethanol (pro analysis), which can induce an increase of ATP hydrolysis activity. Therefore, controls with the same volume of ethanol were analyzed and used as a baseline.

Electrophoresis, in-gel enzyme activity assay, and Western blotting.

Denaturing electrophoresis was performed as described previously (42). Separation of protein complexes without denaturation was carried out by CNE (43) or BNE (44). For 2D native SDS electrophoresis, distinct bands from the CNE gel were cut out and soaked in 1% (wt/vol) SDS for 20 min. A polyacrylamide gel was poured as described previously (42), and the gel slices were placed in the stacking gel prior to polymerization. An in-gel activity assay was used to detect protein complexes with ATP hydrolysis activity after the CNE (27, 45). Western blot analyses were performed as described previously for the I. hospitalis ATPase subcomplex (27). The generation and specificity of the primary antibody against the A1 subcomplex of I. hospitalis were described previously (27). The primary antibody was used at a dilution of 1:5,000. The secondary antibody was a goat anti-rabbit IgG–DyLight 650 (product no. SA5-10034; Thermo Fisher) and was used at a dilution of 1:2,500.

Peptide mass fingerprinting.

In addition to Western blot analyses, components of the A1 subcomplexes were identified by peptide mass fingerprinting. For this purpose, gels were silver stained using a specific protocol developed for MS (46). Single bands were cut out and destained as described by Gharahdaghi et al. (47). After the gel slices were washed in 50 mM ammonium bicarbonate, 50 mM ammonium bicarbonate-acetonitrile (3:1), 25% acetonitrile, and 50% acetonitrile for 30 min each, they were lyophilized to dryness. Proteins were then digested with trypsin for 16 h at 37°C. Peptides were extracted twice with 100 mM ammonium bicarbonate and once with 100 mM ammonium bicarbonate-acetonitrile (1:1) for 1 h each. The extracts were lyophilized and washed with distilled water. MS was performed by Eduard Hochmuth from the Biochemistry Department of the University of Regensburg, using a 4700 Proteomics MALDI-TOF MS/MS analyzer (Applied Biosystems, Foster City, CA).

Native MS.

In LILBID MS (native MS), droplets are generated by a piezo-driven droplet generator (product no. MD-K-130; Microdrop Technologies GmbH, Norderstedt, Germany). This generator produces droplets of 50-μm diameter, with a frequency of 10 Hz. The droplets are transferred to a vacuum and irradiated with an infrared laser at 2.94 μm, a vibrational absorption wavelength of water, which leads to explosive expansion of the sample droplet. The released ions are accelerated by a pulsed electric field, and the mass of the ions is analyzed by a homebuilt reflectron time-of-flight system. For LILBID MS measurements, the samples were either used “as is” in elution buffer (pH 9.0) or buffer exchanged (where noted), directly before the measurement, into 30 mM NH4HCO3 (pH 8.0) containing 0.03% DDM. The buffer exchange took place in desalting columns (Zeba micro spin desalting columns, product no. 89887; Thermo Scientific), which operate with a 7-kDa cutoff filter. In each case, 5 μl of the sample was used for each measurement. The LILBID instrument was run using standard settings, which have already been reported (48). The presented spectra show the averaged signals of several hundred to a thousand droplets. Data processing was performed using the software Massign (49).
For electrospray ionization (ESI) MS measurements, approximately 7 μl of buffer-exchanged samples were loaded into self-pulled and self-coated glass capillaries for offline operation. The measurements took place on a Synapt G2s instrument (Waters). Quadrupole time of flight (QTOF) MS measurements took place with a QTOF I system (Micromass) incorporating a high-mass upgrade (MSVision) (50). The instrument was further modified in-house to include the aforementioned LILBID source.


This research was funded by the Deutsche Forschungsgemeinschaft.
We thank Rainer Deutzmann and Eduard Hochmuth for MALDI-TOF analyses and Mircea Podar for providing I. islandicus and I. pacificus sequence information prior to release of the whole genomes in GenBank.
L.J.K. designed, performed, and analyzed most of the experiments and wrote the paper. A.W. performed and analyzed alkaline pH experiments. J.W. performed and analyzed experiments on the ATP synthase/ATPase of Ignicoccus islandicus. A.Z. performed and analyzed experiments on the ATP synthase/ATPase of Ignicoccus pacificus. J.H. and N.M. designed and performed LILBID MS and ESI MS experiments and analyzed the data. V.M. analyzed the data and critically revised the manuscript. H.H. designed the study, analyzed the data, and critically revised the manuscript. All authors reviewed the results and approved the final version of the manuscript.

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Boyer PD. 1997. The ATP synthase: a splendid molecular machine. Annu Rev Biochem 66:717–749.
Hilario E, Gogarten JP. 1993. Horizontal transfer of ATPase genes: the tree of life becomes a net of life. Biosystems 31:111–119.
Capaldi RA, Aggeler R. 2002. Mechanism of the F1FO-type ATP synthase, a biological rotary motor. Trends Biochem Sci 27:154–160.
Cipriano DJ, Wang Y, Bond S, Hinton A, Jefferies KC, Qi J, Forgac M. 2008. Structure and regulation of the vacuolar ATPases. Biochim Biophys Acta 1777:599–604.
Müller V, Grüber G. 2003. ATP synthases: structure, function and evolution of unique energy converters. Cell Mol Life Sci 60:474–494.
Wilms R, Freiberg C, Wegerle E, Meier I, Mayer F, Müller V. 1996. Subunit structure and organization of the genes of the A1A0 ATPase from the archaeon Methanosarcina mazei Gö1. J Biol Chem 271:18843–18852.
Pisa KY, Weidner C, Maischak H, Kavermann H, Müller V. 2007. The coupling ion in the methanoarchaeal ATP synthases: H+ vs. Na+ in the A1AO ATP synthase from the archaeon Methanosarcina mazei Gö1. FEMS Microbiol Lett 277:56–63.
Lingl A, Huber H, Stetter KO, Mayer F, Kellermann J, Müller V. 2003. Isolation of a complete A1AO ATP synthase comprising nine subunits from the hyperthermophile Methanococcus jannaschii. Extremophiles 7:249–257.
Pisa KY, Huber H, Thomm M, Müller V. 2007. A sodium ion-dependent A1AO ATP synthase from the hyperthermophilic archaeon Pyrococcus furiosus. FEBS J 274:3928–3938.
Vonck J, Pisa KY, Morgner N, Brutschy B, Müller V. 2009. Three-dimensional structure of A1A0 ATP synthase from the hyperthermophilic archaeon Pyrococcus furiosus by electron microscopy. J Biol Chem 284:10110–10119.
Grüber G, Manimekalai MSS, Mayer F, Müller V. 2014. ATP synthases from archaea: the beauty of a molecular motor. Biochim Biophys Acta 1837:940–952.
Lewalter K, Müller V. 2006. Bioenergetics of archaea: ancient energy conserving mechanisms developed in the early history of life. Biochim Biophys Acta 1757:437–445.
Lübben M, Schäfer G. 1987. A plasma-membrane associated ATPase from the thermoacidophilic archaebacterium Sulfolobus acidocaldarius. Eur J Biochem 164:533–540.
Konishi J, Wakagi T, Oshima T, Yoshida M. 1987. Purification and properties of the ATPase solubilized from membranes of an acidothermophilic archaebacterium, Sulfolobus acidocaldarius. J Biochem 102:1379–1387.
Hochstein LI, Stan-Lotter H. 1992. Purification and properties of an ATPase from Sulfolobus solfataricus. Arch Biochem Biophys 295:153–160.
Dirmeier R, Hauska G, Stetter KO. 2000. ATP synthesis at 100°C by an ATPase purified from the hyperthermophilic archaeon Pyrodictium abyssi. FEBS Lett 467:101–104.
Huber H, Burggraf S, Mayer T, Wyschkony I, Rachel R, Stetter KO. 2000. Ignicoccus gen. nov., a novel genus of hyperthermophilic, chemolithoautotrophic archaea, represented by two new species, Ignicoccus islandicus sp. nov. and Ignicoccus pacificus sp. nov. Int J Syst Evol Microbiol 50:2093–2100.
Paper W, Jahn U, Hohn MJ, Kronner M, Nather DJ, Burghardt T, Rachel R, Stetter KO, Huber H. 2007. Ignicoccus hospitalis sp. nov., the host of ‘Nanoarchaeum equitans.’ Int J Syst Evol Microbiol 57:803–808.
Rachel R, Wyschkony I, Riehl S, Huber H. 2002. The ultrastructure of Ignicoccus: evidence for a novel outer membrane and for intracellular vesicle budding in an archaeon. Archaea 1:9–18.
Huber H, Küper U, Daxer S, Rachel R. 2012. The unusual cell biology of the hyperthermophilic crenarchaeon Ignicoccus hospitalis. Antonie Van Leeuwenhoek 102:203–219.
Huber H, Hohn MJ, Rachel R, Fuchs T, Wimmer VC, Stetter KO. 2002. A new phylum of archaea represented by a nanosized hyperthermophilic symbiont. Nature 417:63–67.
Jahn U, Summons R, Sturt H, Grosjean E, Huber H. 2004. Composition of the lipids of Nanoarchaeum equitans and their origin from its host Ignicoccus sp. strain KIN4/I. Arch Microbiol 182:404–413.
Jahn U, Gallenberger M, Paper W, Junglas B, Eisenreich W, Stetter KO, Rachel R, Huber H. 2008. Nanoarchaeum equitans and Ignicoccus hospitalis: new insights into a unique, intimate association of two archaea. J Bacteriol 190:1743–1750.
Giannone RJ, Huber H, Karpinets T, Heimerl T, Küper U, Rachel R, Keller M, Hettich RL, Podar M, Randau L. 2011. Proteomic characterization of cellular and molecular processes that enable the Nanoarchaeum equitans-Ignicoccus hospitalis relationship. PLoS One 6:e22942.
Podar M, Anderson I, Makarova KS, Elkins JG, Ivanova N, Wall MA, Lykidis A, Mavromatis K, Sun H, Hudson ME, Chen W, Deciu C, Hutchison D, Eads JR, Anderson A, Fernandes F, Szeto E, Lapidus A, Kyrpides NC, Saier MH, Richardson PM, Rachel R, Huber H, Eisen JA, Koonin EV, Keller M, Stetter KO. 2008. A genomic analysis of the archaeal system Ignicoccus hospitalis-Nanoarchaeum equitans. Genome Biol 9:R158.
Mohanty S, Jobichen C, Chichili VPR, Velazquez-Campoy A, Low BC, Hogue CWV, Sivaraman J. 2015. Structural basis for a unique ATP synthase core complex from Nanoarchaeum equitans. J Biol Chem 290:27280–27296.
Küper U, Meyer C, Müller V, Rachel R, Huber H. 2010. Energized outer membrane and spatial separation of metabolic processes in the hyperthermophilic archaeon Ignicoccus hospitalis. Proc Natl Acad Sci U S A 107:3152–3156.
Küper U. 2010. Untersuchungen zur Energiegewinnung des hyperthermophilen, schwefelreduzierenden Archaeons Ignicoccus hospitalis. PhD dissertation. Universität Regensburg, Regensburg, Germany.
Mayer F, Leone V, Langer JD, Faraldo-Gómez JD, Müller V. 2012. A c subunit with four transmembrane helices and one ion (Na+)-binding site in an archaeal ATP synthase: implications for c ring function and structure. J Biol Chem 287:39327–39337.
Pogoryelov D, Krah A, Langer JD, Yildiz Ö, Faraldo-Gómez JD, Meier T. 2010. Microscopic rotary mechanism of ion translocation in the Fo complex of ATP synthases. Nat Chem Biol 6:891–899.
Lemker T, Ruppert C, Stöger H, Wimmers S, Müller V. 2001. Overproduction of a functional A1 ATPase from the archaeon Methanosarcina mazei Gö1 in Escherichia coli. Eur J Biochem 268:3744–3750.
Schägger H, von Jagow G. 1991. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem 199:223–231.
Lübben M, Lünsdorf H, Schäfer G. 1987. The plasma membrane ATPase of the thermoacidophilic archaebacterium Sulfolobus acidocaldarius: purification and immunological relationships to F1-ATPases. Eur J Biochem 167:211–219.
Kastner B, Fischer N, Golas MM, Sander B, Dube P, Boehringer D, Hartmuth K, Deckert J, Hauer F, Wolf E, Uchtenhagen H, Urlaub H, Herzog F, Peters JM, Poerschke D, Lührmann R, Stark H. 2008. GraFix: sample preparation for single-particle electron cryomicroscopy. Nat Methods 5:53–55.
Stark H. 2010. GraFix: stabilization of fragile macromolecular complexes for single particle cryo-EM. Methods Enzymol 481:109–126.
Mayer F, Küper U, Meyer C, Daxer S, Müller V, Rachel R, Huber H. 2012. AMP-forming acetyl coenzyme A synthetase in the outermost membrane of the hyperthermophilic crenarchaeon Ignicoccus hospitalis. J Bacteriol 194:1572–1581.
Naß B, Pöll U, Langer JD, Kreuter L, Küper U, Flechsler J, Heimerl T, Rachel R, Huber H, Kletzin A. 2014. Three multihaem cytochromes c from the hyperthermophilic archaeon Ignicoccus hospitalis: purification, properties and localization. Microbiology 160:1278–1289.
Müller V, Lemker T, Lingl A, Weidner C, Coskun U, Grüber G. 2005. Bioenergetics of archaea: ATP synthesis under harsh environmental conditions. J Mol Microbiol Biotechnol 10:167–180.
Mayer F. 2008. Isolierung und biochemische Charakterisierung von Membran-assoziierten Proteinkomplexen des hyperthermophilen Archaeums Ignicoccus hospitalis. Diplomarbeit. Universität Regensburg, Regensburg, Germany.
Huber H, Stetter KO. 2006. Desulfurococcales, p 52–68. In Dworkin M, Falkow S, Rosenberg E, Schleifer K-H, Stackebrandt E (ed), The prokaryotes, vol 3. Springer, New York, NY.
Burghardt T, Näther DJ, Junglas B, Huber H, Rachel R. 2007. The dominating outer membrane protein of the hyperthermophilic archaeum Ignicoccus hospitalis: a novel pore-forming complex. Mol Microbiol 63:166–176.
Schägger H. 2006. Tricine–SDS-PAGE. Nat Protoc 1:16–22.
Wittig I, Schägger H. 2005. Advantages and limitations of clear-native PAGE. Proteomics 5:4338–4346.
Wittig I, Braun H-P, Schägger H. 2006. Blue native PAGE. Nat Protoc 1:418–428.
Suhai T, Heidrich NG, Dencher NA, Seelert H. 2009. Highly sensitive detection of ATPase activity in native gels. Electrophoresis 30:3622–3625.
Shevchenko A, Wilm M, Vorm O, Mann M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal Chem 68:850–858.
Gharahdaghi F, Weinberg CR, Meagher DA, Imai BS, Mische SM. 1999. Mass spectrometric identification of proteins from silver-stained polyacrylamide gel: a method for the removal of silver ions to enhance sensitivity. Electrophoresis 20:601–605.
Morgner N, Barth H-D, Brutschy B. 2006. A new way to detect noncovalently bonded complexes of biomolecules from liquid micro-droplets by laser mass spectrometry. Aust J Chem 59:109–114.
Morgner N, Robinson CV. 2012. Massign: an assignment strategy for maximizing information from the mass spectra of heterogeneous protein assemblies. Anal Chem 84:2939–2948.
Sobott F, Hernandez H, McCammon MG, Tito MA, Robinson CV. 2002. A tandem mass spectrometer for improved transmission and analysis of large macromolecular assemblies. Anal Chem 74:1402–1407.

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Published In

cover image Journal of Bacteriology
Journal of Bacteriology
Volume 201Number 71 April 2019
eLocator: 10.1128/jb.00510-18
Editor: William W. Metcalf, University of Illinois at Urbana Champaign


Received: 23 August 2018
Accepted: 8 January 2019
Published online: 13 March 2019


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  1. ATP synthase
  2. ATPase
  3. Archaea
  4. Crenarchaea
  5. Ignicoccus
  6. bioenergetics
  7. protein purification
  8. protein stability



Lydia J. Kreuter
Institute for Microbiology and Archaeal Center, Regensburg University, Regensburg, Germany
Present address: Lydia J. Kreuter, Institute of Marine and Environmental Technology, Baltimore, Maryland, USA.
Andrea Weinfurtner
Institute for Microbiology and Archaeal Center, Regensburg University, Regensburg, Germany
Alexander Ziegler
Institute for Microbiology and Archaeal Center, Regensburg University, Regensburg, Germany
Julia Weigl
Institute for Microbiology and Archaeal Center, Regensburg University, Regensburg, Germany
Jan Hoffmann
Institute of Physical and Theoretical Chemistry, Johann Wolfgang Goethe University, Frankfurt am Main, Germany
Nina Morgner
Institute of Physical and Theoretical Chemistry, Johann Wolfgang Goethe University, Frankfurt am Main, Germany
Volker Müller
Department of Molecular Microbiology and Bioenergetics, Johann Wolfgang Goethe University, Frankfurt am Main, Germany
Harald Huber
Institute for Microbiology and Archaeal Center, Regensburg University, Regensburg, Germany


William W. Metcalf
University of Illinois at Urbana Champaign


Address correspondence to Lydia J. Kreuter, [email protected].

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