The rooting of the universal tree in the so-called "bacterial branch" (**Figures 3–5**) has been often interpreted as suggesting a "prokaryotic phenotype" for LUCA. This is a misleading interpretation that again confuses the phenotypes at the tip and base of a branch. The rooting between a lineage leading to Bacteria and a lineage leading to Archaea and Eukarya is compatible with diverse types of LUCA, including a LUCA with some "eukaryoticlike features" that were lost in Archaea and Bacteria (Forterre, 2013a).

Importantly, rooting of the universal tree in the "bacterial branch" formally requires giving a name to the clade grouping Archaea and Eukarya. Woese (2000) never proposed such a pathway (see text). name, adopting a "gradist" view of life evolution, with the three Domains emerging independently from a "communal LUCA" before the "Darwinian threshold" (Woese, 2000, 2002). In such view, the notion of clade itself cannot be used to group organisms

that diverged at the time of LUCA when no real speciation occurred. I have criticized the Darwinian threshold concept, assuming—with many others—that Darwinian evolution started as soon as biological evolution take off (see Forterre, 2012, and references therein). In particular, extensive genes exchanges that possibly take place at the time of LUCA (but see Poole, 2009) cannot be opposed to Darwinian evolution occurring through variation and selection, since gene transfer only corresponds to a specific type of variation (Forterre, 2012).

I think that it's time now to look back at the universal tree with a cladistics perspective and to propose a name for the clade grouping Archaea and Eukarya. It is challenging to find a common synapomorphy to Archaea and Eukarya that could provide a

good name for the clade corresponding to these two domains. David Prangishvili suggests Arkarya (personal communication), combining the names of the two domains belonging to this clade (**Figure 3**). Notably, universal trees in which Eukarya emerge from within Archaea are often viewed as "two domains trees" *versus* the "three domains tree" of Carl Woese (Gribaldo et al., 2010). However, the new nomenclature proposed here emphasizes that the classical Woese tree is also *stricto sensu*, a two domains tree (Bacteria and Arkarya)!

# **Updated Trees for Everybody**

The backbone of the updated universal trees proposed here (**Figures 3–5**) was selected from the 1990 tree of Woese et al. (1990) as a tribute to Carl Woese and the historical work of the Urbana school (Sapp, 2009; Albers et al., 2013). The relative lengths of the branches linking the three domains together combine features of the rDNA and protein trees. It is indeed puzzling that Archaea and Eukaryotes are very close in trees based on universal protein sequence comparison (Rochette et al., 2014), but are more divergent in those based on rDNA (Pace et al., 1986). The reason for this discrepancy remains unclear and should be worth exploring further. The rather long branches between domains in the trees of **Figures 3–5** also reflect the "three major transformation events" (*sensu* Forterre and Philippe, 1999) that occurred between LUCA and the formation of each domain (Woese, 1998; Forterre and Philippe, 1999).

Evidently, it is not possible to draw a tree including all presently recognized phyla, especially in the bacterial domain, so I made arbitrary choices, and tried to include most well studied bacterial and archaeal phyla, as well as major eukaryotic divisions and/or supergroups. In the case of Archaea, I only indicate the phyla Euryarchaeota, Crenarchaeota, Thaumarchaeota and the candidate phylum "Lokiarchaeota" because other proposed archaeal phyla are represented by a single species and/or their phylum status is controversial or has been refuted by robust phylogenetic and phylogenomic analyses (see below). Although still preliminary, the study of three partial lokiarchaeal genomes has shown that these archaea encode many eukaryotic-like genes absent in Thaumarchaea and are clearly separated from both Thaumarchaeota and Crenarchaeota in phylogenetic analysis (Spang et al., 2015). Furthermore, Lokiarchaeota correspond to a large clade of abundant and diversified uncultivated archaea, previously named deep-sea archaeal group (DSAG), that are widely distributed in both marine and fresh water environments (Jørgensen et al., 2013).

I divide the phylum Euryarchaeota in sub-phylum I (I) and sub-phylum II (II), according to the presence/absence of

DNA gyrase (see below; Forterre et al., 2014b). Dotted lines indicate the endosymbiosis events that had a major impact on the history of life by triggering the emergence of both modern eukaryotes (mitochondria) and Plantae (chloroplasts). In particular, this reminds us that the first mitochondriate eukaryote (FME) emerged after the diversification of alpha proteobacteria, indicating that "modern eukarya" are indeed much more recent than Archaea and Bacteria.

In the tree of **Figure 3**, I use the terms "synkaryote" and "akaryote" (with and without a nucleus, respectively) instead of eukaryotes and prokaryotes (Forterre, 1992; Harish et al., 2013; Penny et al., 2014). This is because the latter terms are the hallmark of the traditional (pre-Woesian) view of the evolution of life from primitive bacteria ("pro" karyotypes) to lower and finally higher eukaryotes (Forterre, 1992; Pace, 2006; Penny et al., 2014).

Some major events that shaped modern domains are indicated, such as the introduction of peptidoglycan (PG) in the lineage leading to Bacteria. The last bacterial and archaeal common ancestors (LBCA and LACA) are colored in pink and red, respectively, to indicate their probable thermophilic and hyperthermophilic nature based on ancestral protein and rRNA sequence reconstruction (Boussau et al., 2008a; Groussin and Gouy, 2011; Groussin et al., 2013). The grouping of hyperthermophiles at the base of the archaeal tree also suggests that LACA was a hyperthermophile (Brochier-Armanet et al., 2011; Petitjean et al., 2015), whereas LUCA was probably a mesophile or a moderate thermophile (Boussau et al., 2008a;

Groussin and Gouy, 2011). Some proposed events are more speculative but supported by theoretical arguments, such as the independent introduction of DNA (blue arrows) from viruses, into the lineages leading to Bacteria and to Archaea/Eukarya (Forterre, 2002) and the thermoreduction (red arrows) at the origin of the modern "akaryotic" phenotype (Forterre, 1995).

Rooting of the domain Eukarya and internal branching in this domain have been adapted from the recent tree of Baldauf and colleagues which is based on a concatenation of mitochondrial proteins rooted with their bacterial homologs (a rather close outgroup compared to Archaea; He et al., 2014). This tree is rooted between Discoba (Jakobida plus Discritata) and all other eukaryotic megagroups. This rooting has been criticized by Derelle et al. (2015) who found distant paralogs in the data set used by He et al. (2014). These authors located the root of the eukaryotic tree between Amorpha and other eukaryotic groups in mitochondrial proteins trees. This rooting corresponds to the previous division between Unikonta and Bikonta originally proposed by (Stechmann and Cavalier-Smith, 2003; Cavalier-Smith, 2010). Derelle et al. (2015) have now suggested naming these two assemblages Optimoda (Amorpha in **Figures 3–5**) and Diphoda.

The position of the root of the domain Eukarya has been constantly changing with further phylogenetic analyses, raising doubt about the possibility of settling this issue using molecular phylogenetic methods based on protein sequences. I thus decided to root this domain between Jakobida and all other eukaryotes in

the universal tree of **Figure 3**, as recently suggested by Kannan et al. (2014), because Jakobida contain large mitochondrial genomes that still encode the bacterial RNA polymerase genes (Burger et al., 2013; Kamikawa et al., 2014; Kannan et al., 2014). The mitochondria of the LECA probably still had this RNA polymerase, which was subsequently replaced with a viral RNA polymerase in all Eukaryotes, except Jakobida (Kannan et al., 2014). These viral RNA polymerases have been recruited from a provirus integrated into the genome of the alphaproteobacterium, which gave rise to mitochondria (Filée and Forterre, 2005). It seems unlikely that this non-orthologous replacement occurred twice independently in the history of mitochondria. Accordingly, the rooting between Jakobida and other eukaryotes is reasonable (more parsimonious) because it requires a single NOR of RNA polymerase in mitochondrial evolution, whereas the rooting between Amorpha and other eukaryotes would require several independent non-orthologous replacements.

Rooting of the domain Bacteria and internal branching in this domain have been adapted from the ribosomal protein trees of Koonin and colleagues (Yutin et al., 2012). These authors have suggested several superphyla beside the previously recognized PVC superphylum, which includes Planctomycetes, Verrucomicrobia and Chlamydiae (Kamke et al., 2014). These putative superphyla are indicated by circles in the tree of **Figure 3**. Branchings within Proteobacteria are drawn according to the ribosomal protein tree of Brochier-Armanet and colleagues (Ramulu et al., 2014). The tree is tentatively rooted between the PVC superphylum and all other Bacteria, according to the basal rooting of Planctomycetes obtained by Brochier and Philippe using slowly evolving positions in ribosomal RNA sequences (Brochier and Philippe, 2002). Bacteroidetes are indicated in the second branching because these Bacteria are grouped with PVC bacteria in the phylogenetic analysis based on ribosomal proteins (Yutin et al., 2012).

The basal position of PVC in the bacterial tree of **Figure 3**, which remains to be confirmed, is appealing because the ancestor of PVC bacteria contained several genes encoding proteins structurally analogous to various eukaryotic coat proteins involved in vesicle and nuclear pore formation (Santarella-Mellwig et al., 2010). These proteins are probably involved in the invagination of the cytoplasmic membrane that led to the formation of the intracellular cytoplasmic membrane (ICM) in most PVC bacteria. This mimics the role of coat proteins in eukaryotes that are involved in the formation of the endoplasmic reticulum and nuclear membranes. The basal position of PVC bacteria suggests a parsimonious scenario in which these proteins were present in LUCA, and later on lost in Archaea and most Bacteria (Forterre and Gribaldo, 2010). However, this scenario is still controversial since it is presently unclear whether the structurally analogous proteins of PVC Bacteria and Eukarya are also homologous (McInerney et al., 2011; Devos, 2012). These proteins are formed by the fusion of two domains (one rich in alpha helices, the other in beta strands) that are each present in the three domains. Accordingly, they can also have originated independently by the fusion of these domains in the branches leading to Eukarya and PVC bacteria.

Archaea have been tentatively rooted in the branch leading to Lokiarchaeota in the tree of **Figure 3**, because this candidate phylum contains most eukaryotic features present in Archaea and branches closer to Eukarya than to other archaea in phylogenetic analyses of universal proteins, even when bacterial proteins are removed from the analysis (Spang et al., 2015, Figure S13D). Previously, the archaeal ribosomal tree was rooted in the branch leading to Thaumarchaeota when eukaryotic proteins were used as outgroup (Brochier-Armanet et al., 2008a). This rooting was also observed in a phylogenetic analysis of the archaeal replicative helicase MCM, which is a good phylogenetic marker for the archaeal domain (Krupovic et al., 2010), and in a phylogeny of five informational proteins present in deeply branching Thaumarchaeota from Kamchatkan thermal springs (Eme et al., 2013). I thus place Thaumarchaeota as the second branch in the archaeal subtree.

Moreira and colleagues, using bacterial proteins (including ribosomal proteins) as outgroup, have recently proposed to root the archaeal tree in the branch leading to Euryarchaeota (Petitjean et al., 2014). As a consequence, they propose to create a new phylum, Proteoarchaeota, grouping Crenarchaeota and Thaumarchaeota, together with the putative phyla Aigarchaeota and Korarchaeota. Proteoarchaea thus corresponds to the previously so-called "TACK superphylum" (Thaumarchaeota, "Aigarchaeota," Crenarchaeota, Korarchaeota). However, as previously mentioned, using the same strategy, Gribaldo and colleagues obtained a root located within Euryarchaeota, more precisely in between subphyla I and II (Raymann et al., 2015). In contrast, their archaeal tree is rooted between "TACK/Proteoarchaeota" and Euryarchaeota when they used eukaryotic proteins as an outgroup. It will be interesting to see if they recover the root in the branch leading to Lokiarchaeota in future analyses using eukaryotic sequences as an outgroup.

Moreira and colleagues argue that eukaryotic proteins cannot be used to root the archaeal tree if Eukarya emerged from within Archaea. However, in the framework of the classical Woese tree, it makes more sense to root the archaeal tree using eukaryotic proteins as outgroup, because these proteins are much more closely related than bacterial proteins to their archaeal orthologs. Notably, the rooting between Lokiarchaeota/Thaumarchaeota and other Archaea, obtained in that case is more parsimonious than the rooting between Euryarchaeota and other Archaea in explaining the presence in Lokiarchaeota/Thaumarchaeota (including "Aigarchaeota," see below) of many eukaryotic features lacking in other Archaea (Brochier-Armanet et al., 2008b; Spang et al., 2010, 2015; Koonin and Yutin, 2014).

Euryarchaeota are divided in two sub-phyla I and II, according to the presence/absence of DNA gyrase, a bacterial DNA topoisomerase that was transferred once in the phylum Euryarchaeota (Raymann et al., 2014). The sub-phylum I Euryarchaeota corresponds to those lacking DNA gyrase and encompasses Thermococcales*, Nanoarchaeum*, and class I methanogens, whereas sub-phylum I corresponds to those containing DNA gyrase and encompasses Archaeoglobales,

Thermoplasmatales, Halobacteriales, and class II methanogens (Forterre et al., 2014b).

Phylogenetic analyses have shown that DNA gyrase has been transferred from Bacteria to Archaea (Raymann et al., 2014). This transfer was an important and unique event that had a critical impact on chromosome structure and patterns of gene expression. Indeed, plasmids from all archaea are relaxed or slightly positively supercoiled, whereas plasmids from member of sub-phylum II Euryarchaeota containing gyrase are negatively supercoiled (Forterre et al., 2014b). Once transferred, DNA gyrase became most likely essential, as demonstrated in the case of Halobacteriales and Methanococcales, because such drastic modification in DNA topology modifies all protein DNA interactions involved in replication and transcription (for review and discussion, see Forterre and Gadelle, 2009). Indeed, to date, the loss of DNA gyrase has not been reported in any organism. Importantly, the phylogeny of archaeal DNA gyrase is fully congruent with the phylogeny of sub-phylum II Euryarchaeota, suggesting that, once transferred to the ancestor of this group, DNA gyrase has co-evolved with sub-phylum II Euryarchaeota (Raymann et al., 2014). Accordingly, considering the importance of DNA gyrase in cell physiology (DNA topology controlling all gene expression patterns) I suggest calling sub-phylum II Euryarchaeota, the neo-euryarchaeota. This name emphasizes the fact that the ancestor of this sub-phylum lived after the formation of the major bacterial phyla, since archaeal DNA gyrases branch within bacterial ones (Raymann et al., 2014).

Since all rooting indicated in the tree of **Figure 3**, as well as most internal nodes within domains, are controversial, I present a second tree (**Figure 4**), in which the information is limited to only that accepted by consensus. Accordingly, each one of the three domains is shown in a radial form without roots and only a few nodes within domains that seem supported by strong phylogenetic analyses are indicated (Brochier-Armanet et al., 2008a; He et al., 2014; Kamke et al., 2014; Ramulu et al., 2014; Spang et al., 2015).

Finally, I also present a third tree in which Eukarya emerged from within Archaea (**Figure 5**). This tree includes the new root proposed by Gribaldo and co-workers for Archaea (Raymann et al., 2015) and shows Lokiarchaeota as sister group of Eukarya (Spang et al., 2015). Notably, if future analyses demonstrate that such a tree is the more likely tree, Archaea will not be a valid taxon anymore, except if one accepts to consider eukaryotes as a particular archaeal phylum (much like *Homo* is a particular lineage of Apes)! In that case, the name Arkarya could be substituted to Archaea. Eukarya will become a particular phylum of Arkarya, beside Euryarkaryota, Crenarkaryota, Thaumarkaryota, and Lokiarkaryota (**Figure 5**).

As can be seen, **Figures 4** and **5** can be easily deduced from **Figure 3**. This indicates that it will be easy to update and modify these trees following the accumulation of new data from comparative genomics and phylogenetic analyses.

# **The Archaeal Tree**

**Figure 6** illustrates a rather detailed, but schematic, archaeal tree as a tribute to this issue devoted to Archaea. This tree has been adapted from the ribosomal protein tree of Brochier-Armanet et al. (2011) and from a recent phylogeny based on the concatenation of 273 proteins conserved in at least 119 archaeal species out of 129 (Petitjean et al., 2015; thereafter called the archaeal protein tree). I also include the recently described candidate phylum "Lokiarchaeota" (corresponding to the DSAG clade) considering its importance for the discussions about the origin of Eukarya. The various roots that have been proposed for the domain Archaea are indicated by orange circles.

Aigarchaeota are included within Thaumarchaeota, because the latter were originally defined as a major archaeal phylum encompassing all archaeal lineages that are sister groups of Crenarchaeota in rDNA analyses (Brochier-Armanet et al., 2011). In the original paper in which we propose this new phylum, we noticed that: "*The diversity of mesophilic crenarchaeota* that we proposed to rename Thaumarchaeota*—based on SSU rRNA sequence is comparable to that of hyperthermophilic Crenarchaeota and Euryarchaeota, which suggests that they represent a major lineage that has equal status to Euryarchaeota and Crenarchaeota*" (Brochier-Armanet et al., 2008a). We also predicted that the mesophily of Thaumarchaeota "*could be challenged by the future identification of non-mesophilic organisms that belong to this phylum*." This suggests considering *candidatus* Caldarchaeum subterraneum as a thermophilic member of the Thaumarchaeota and not as the prototype of a new phylum (Aigarchaeota).  Furthermore, *candidatus* C. subterraneum exhibits all molecular features first used to define the phylum Thaumarchaeota (Brochier-Armanet et al., 2008a; Spang et al., 2010), such as a eukaryotic-like Topo IB, which is absent from all other Archaea (Brochier-Armanet et al., 2008b). Topo IB is absent in Lokiarchaeota, but one must bear in mind that the reconstituted genome is only 92% complete (Spang et al., 2015). The phylum Thaumarchaeota should also include uncultivated archaea from the clade MCG (the Miscellaneous Crenarchaeal Group), since these organisms systematically form monophyletic groups with Thaumarchaea and "Aigarchaeota" in phylogenetic analyses (Spang et al., 2015).

The recently proposed phylum "Geoarchaeota" is included as a sister group of Thermoproteales (without phylum status) as suggested by the analysis of Ettema and co-workers (Guy et al., 2014; Spang et al., 2015). *Thermofilum* always branch very early as a sister group of Thermoproteales. This suggests that *Thermofilum*, as well as Geoarchaeota, could have an order status.

Korarchaeota branch in-between Euryarchaeota and other archaea (Crenarchaeota, Thaumarchaeota) in the ribosomal proteins tree. Unfortunately, this phylum is presently represented by a single species whose genome has been sequenced, *candidatus* Korarchaeum cryptofilum. The genome of *candidatus* K. cryptofilum harbors a mixture of features characteristic of the three other archaeal phyla. This can justify maintaining a phylum status for this group for the moment. More genome sequences of Korarchaeota are nevertheless required to confirm this point.

As previously discussed, Euryarchaeota are divided into two groups depending of the presence of DNA gyrase. Neo-euryarchaeota (sub-phylum II) is a monophyletic group in all phylogenetic analyses. In contrast, sub-phylum I is paraphyletic in the ribosomal and archaeal protein trees (Brochier-Armanet et al., 2011; Petitjean et al., 2015). However, they form a monophyletic assemblage in a tree based on replication proteins (Raymann et al., 2014) and in a recent phylogenomic analysis performed by Makarova et al. (2015) that involved both comparison of multiple phylogenetic trees and a search for putative synapomorphies. I thus decided to favor the monophyly of sub-phylum I in the tree of **Figure 6**.

The close relationship between plasmids of Thermococcales and Methanococcales also suggests that these two orders could be closely related (Soler et al., 2010). It is possible that the emergence of pseudomurein in Methanobacteriales and Methanopyrales allowed these archaea to get rid of mobile elements that used to infect the ancestors of Thermococcales and Methanococcales. Notably, Methanopyrales and Methanobacteriales are monophyletic in the archaeal protein tree, suggesting that the presence of pseudomurein is a synapomorphy for these sister groups (Petitjean et al., 2015). Petitjean and coworkers recently proposed the name Methanomada (superclass) for class I methanogens, that are monophyletic in their protein tree (Petitjean et al., 2015) and in the DNA replication tree (Raymann et al., 2014).

In contrast to class I, the four orders of class II methanogens that are included within neo-euryarchaeota are always paraphyletic in phylogenetic analyses. Methanogens of the recently described order Methanomassiliicoccus form a monophyletic assemblage with Thermoplasmatales, the moderate thermoacidophilic strain *Aciduliprofundum boonei* and several lineages of uncultivated archaea in a ribosomal protein tree (Borrel et al., 2013). The name Diafoarchaea has been proposed for this major subgroup (superclass) of neo-Euryarchaeota (Petitjean et al., 2015).

Altiarchaeales correspond to a recently described mesophilic archaeum, *Candidatus* Altiarchaeum hamiconexum, characterized by fascinating appendages (Hami) that groups with Methanococcales in a ribosomal protein tree, but between Euryarchaeota of sub-phyla I and II in a tree based on several other universal proteins (Probst et al., 2014). Since *Candidatus* A. hamiconexum contain the two DNA gyrase genes, it is located at the base of the neo-euryarchaeota in the tree of **Figure 6**.

Rinke et al. (2013) have recently proposed promoting the nanosized archaea *Nanoarchaeum*, *Parvarchaeum*(ARMAN 4 and 5) and Nanohaloarchaea to phylum level and grouping them with two other putative new phyla of Archaea ("Aenigmarchaeota" and "Diapherotrites") into a new superphylum, called DPANN (Rinke et al., 2013). In their phylogenetic analysis based on 38 "universal proteins," the root of the archaeal tree is located between this putative DPANN superphylum and all other Archaea (see Figure S11 in Rinke et al., 2013). However, their universal protein data set is confusing because it contains eukaryotic proteins of bacterial origin (Williams and Embley, 2014). In the recent tree of Williams and Embley supporting the archaeal origin of eukaryotes, the archaeal tree is rooted between DPANN and Euryarchaeota (see Figure 3 in Williams and Embley, 2014). The basal position of the nanosized archaea in these trees confirms the difficulty of using universal proteins to resolve ancient phylogenies, because

this position is clearly misleading (see below and Petitjean et al., 2014). In the case of nanosized archaea, phylogenetic analyses are even more challenging because their proteins tend to evolve rapidly, producing very long branches in phylogenetic trees. This suggests that these nanosized archaea evolve mainly by streamlining (Brochier-Armanet et al., 2011; Petitjean et al., 2014). They have not been included in the protein tree of Petitjean and coworkers (Petitjean et al., 2015).

Previous phylogenetic and comparative genomic analyses focusing on *Nanoarchaeum equitans* have suggested that this fascinating archaeal symbiont belongs to the Euryarchaeota and could be distant relatives of Thermococcales (Brochier et al., 2005b). This sister relationship was later on supported by the discovery of the shared presence of a tRNA modification protein that was recently transferred from Bacteria to both *N. equitans* and Thermococcales (Urbonavicius et al., 2008). The sisterhood of *N. equitans* and Thermococcales has been observed again in more recent analyses based on ribosomal proteins (Brochier-Armanet et al., 2011) or in the archaeal tree of Moreira and colleagues (Petitjean et al., 2014).

In the tree of **Figure 6**, Thermococcales, *Nanoarchaeum*, *Parvarchaeum*, and Nanohaloarchaea tentatively form a monophyletic group. This is supported by several lines of converging (but weak and controversial) evidences. The grouping of Thermococcales and *Nanoarchaeum* with *Parvarchaeum* (ARMAN 4 and 5) is supported by the phylogeny based on ribosomal proteins (Brochier-Armanet et al., 2011), whereas the grouping of *Nanoarchaeum* and *Parvarchaeum* with Nanohaloarchaea is supported by the phylogeny of DNA replication proteins (Raymann et al., 2014). Interestingly, the grouping of *Parvarchaeum*, *Nanoarchaeum*, and Nanohaloarchaea is supported by the shared presence of an atypical small primase corresponding to the fusion of the two subunits of the *bona fide* archaeal/eukaryal primases, PriS, and PriL (Raymann et al., 2014). It has been suggested that this fusion corresponds to a convergent evolution associated to streamlining (Petitjean et al., 2014). However, this is unlikely because these unusual monomeric primase primases are also highly divergent in terms of amino-acid sequence from the classical archaeal/eukaryal primases and very similar in all nanosized archaea. Alternatively, it has been suggested that this primase has been distributed between nanosized archaea by horizontal gene transfers (Raymann et al., 2014). This seems also unlikely because nanosized archaea live in very different types of environments (high temperature for known *Nanoarchaeum*, high salt for Nanohaloarchaea, high acidity for *Parvarchaeum*). It seems more likely that this primase was acquired by a common ancestor to *Nanoarchaeum*, *Parvarchaeum*, and Nanohaloarchaea from a mobile genetic element (Raymann et al., 2014). It has indeed been shown that some archaeal plasmids encode unusual primases from the PriS/PriL superfamily that are only distantly related to *bona fide* archaeal and eucaryal primases (Lipps et al., 2004; Krupovic et al., 2013; Gill et al., 2014).

The grouping of Nanohaloarchaea with other nanosized archaea, as in **Figure 6**, is especially controversial because they emerge as sister group of Halobacteriales in a ribosomal protein tree (Narasingarao et al., 2012). However, if Nanohaloarchaea are sister group of Halobacteriales, they would be the only members of neo-euryarchaeota lacking DNA gyrase. Nanohaloarchaea might have lost this enzyme during the streamlining process related to their small size. However, the loss of DNA gyrase has not been reported until now in any other free-living organisms. This is why I finally choose to group Nanohaloarchaea with other nanosized archaea in the tree of **Figure 6**. It is possible that the atypical amino acid composition of the nanohaloarchaeal proteome (Narasingarao et al., 2012), linked to salt adaptation, introduces a bias favoring the artificial grouping of Nanohaloarchaea and Haloarchaea. This would also explain why Nanohaloarchaea attracts *Nanoarchaeum* and *Parvarchaeum* away from Thermococcales and closer to Haloarchaea in the DNA replication tree (Raymann et al., 2014).

The nanosized archaeon *Candidatus Micrarchaeum acidophilum* (ARMAN 2) branches together with *Parvarchaeum* (ARMAN 4, 5) in a rDNA tree (Baker et al., 2006) and in the tree of Moreira and coworkers (Petitjean et al., 2014). However, it branches away from *Parvarchaeum* in the ribosomal tree, as an early branching neo-euryarchaeon. The latter position is supported by the presence of DNA gyrase in *Candidatus Micrarchaeum acidophilum* and the absence of the single subunit primase characteristic of other nanosized archaea (Raymann et al., 2014). I thus included *Micrarchaeum* among the superclass Diafoarchaea in the tree of **Figure 6**. Finally, the phylogenetic position and status of "Aenigmarchaeota" and "Diapherotrites" cannot presently be determined because these groups have only been defined from single cell genomic analyses and their genomes are probably incomplete (Petitjean et al., 2014).

Further analyses and (hopefully) many more isolates are clearly required to determine the correct position of the various groups of nanosized archaea in the archaeal tree. From all these considerations, it is clear that the "DPNN superphylum" is an artificial construction. The same is true for the "TACK superphylum" and the candidate phylum "Proteoarchaea" if the root of the archaeal tree is located within Euryarchaeota (Raymann et al., 2015) or between Thaumarchaeota/Lokiarchaeota and other Archaea. As we previously discussed, defining the root of the archaeal tree strongly depends on choosing between different scenarios for the universal tree. Since the rooting of the archaeal tree is still in debate, I would not recommend at the moment to use names such as "Proteoarchaea" or "TACK superphylum" in archaeal phylogeny.

This section on archaeal phylogeny has illustrated the fact that, in addition to the root of the tree itself, several nodes in the archaeal tree are still controversial and require more data and more work to be carried out. These nodes have been marked by circles in blue in the tree of **Figure 6**. Future progress will probably come from the sequencing of more genomes, especially in poorly represented groups and in the many groups that are presently only known from environmental rDNA sequences.

# **Conclusion**

I hope that the universal and archaeal trees proposed here will be useful as new metaphors illustrating the history of life on our planet. From the above review, it should be clear that there is no protein or groups of proteins that can give the real species tree, i.e., allow us to recapitulate safely the exact path of life evolution. In particular, one should be cautious with composite trees based on the concatenation of protein sequences or addition of individual trees. The results obtained should always be compared to the result of individual trees. Martin and colleagues recently reported a lack of correspondence between individual protein trees and the concatenation tree in several datasets of archaeal and bacterial proteins (Thiergart et al., 2014). This can reflect either LGT and/or the absence of real phylogenetic signal. Importantly, this lack of correspondence between individual protein trees and concatenation trees is also observed in the analyses that place Eukarya within Archaea (Cox et al., 2008; Williams and Embley, 2014), raising doubts on results supporting the tree of **Figure 5**. Unfortunately, individual trees are not always available in the supplementary data of published studies (Katz and Grant, 2014; Raymann et al., 2015). In our previous work on archaeal phylogeny, careful analysis of individual trees in parallel to their concatenation was critical to obtain a rather confident tree for Archaea based on ribosomal proteins (Matte-Tailliez et al., 2002) and to find the (probably) correct position of Nanoarchaeota, as sister group of Thermococcales (Brochier et al., 2005b). Notably, to obtain this result in Brochier et al. (2005a), it was necessary to remove the proteins from the large ribosome subunit because several of them exhibit a surprising affinity with their crenarchaeal homologs in individual trees, possibly indicating LGT between *N. equitans* and its host, the crenarchaeon *Ignicoccus*. It is also very important to compare trees obtained with different datasets, such as translation, transcription and DNA replication trees, to pint-point discrepancies and identify their causes (Brochier et al., 2004, 2005a; Raymann et al., 2014).

In summary, it is (and it will be) only possible to draw schematic (theoretical) trees by combining data from multiple phylogenetic protein trees with information deduced from probable synapomorphies. It is the exercise that I have tried to do here in drawing the trees of **Figures 3–6**. These organismal trees, which are supposed to recapitulate the history of ribosome encoding organisms, will probably evolve themselves, with the availability of new genomes (especially from poorly sampled groups), better phylogenetic analyses, and the identification of new synapomorphies defining specific domains, sub-phyla groups and superphyla. For instance, my preliminary analyses of universal proteins sequence alignments indicate that the Lokiarchaeon is probably neither an early braching archaeon nor a missing link between Archaea and Eukarya (see also Nasir et al., 2015). It should be relatively easy to update these trees as new data

# **References**

accumulate in the future and use them in discussions of various controversial scenarios regarding the evolution of ancient life.

# **Acknowledgments**

I thank Mechthild Pohlschroeder and Sonja-Verena Albers for inviting me to draw an updated version of the universal tree of life and David Prangishvili for suggesting the name Arkarya. I am grateful to Sukhvinder Gill for English corrections and a referee for extensive critical analysis. I am supported by an ERC grant from the European Union's Seventh Framework Program (FP/2007-2013)/Project EVOMOBIL—ERC Grant Agreement no.340440.

Filée, J., and Forterre, P. (2005). Viral proteins functioning in organelles: a cryptic origin? *Trends Microbiol.* 13, 510–513. doi: 10.1016/j.tim.2005.08.012

Forterre, P. (1992). Neutral terms. *Nature* 335, 305. doi: 10.1038/355305c0

Forterre, P. (1995). Thermoreduction, a hypothesis for the origin of prokaryotes. *C. R. Acad. Sci. III.* 318, 415–422.

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 Forterre. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Archaeal membrane-associated proteases: insights on Haloferax volcanii and other haloarchaea

# *María I. Giménez\*, Micaela Cerletti and Rosana E. De Castro\**

Instituto de Investigaciones Biológicas, Facultad de Ciencias Exactas y Naturales, Universidad Nacional de Mar del Plata, Consejo Nacional de Investigaciones Científicas y Técnicas, Mar del Plata, Argentina

#### *Edited by:*

Mechthild Pohlschroder, University of Pennsylvania, USA

#### *Reviewed by:*

Kelly Bidle, Rider University, USA Maria-Jose Bonete, University of Alicante, Spain

#### *\*Correspondence:*

María I. Giménez and Rosana E. De Castro, Instituto de Investigaciones Biológicas, Facultad de Ciencias Exactas y Naturales, Universidad Nacional de Mar del Plata, Consejo Nacional de Investigaciones Científicas y Técnicas, Funes 3250 4to nivel, Mar del Plata 7600, Argentina e-mail<EMAIL_ADDRESS><EMAIL_ADDRESS>
The function of membrane proteases range from general house-keeping to regulation of cellular processes. Although the biological role of these enzymes in archaea is poorly understood, some of them are implicated in the biogenesis of the archaeal cell envelope and surface structures. The membrane-bound ATP-dependent Lon protease is essential for cell viability and affects membrane carotenoid content in Haloferax volcanii. At least two different proteases are needed in this archaeon to accomplish the posttranslational modifications of the S-layer glycoprotein. The rhomboid protease RhoII is involved in the N-glycosylation of the S-layer protein with a sulfoquinovose-containing oligosaccharide while archaeosortase ArtA mediates the proteolytic processing coupled-lipid modification of this glycoprotein facilitating its attachment to the archaeal cell surface. Interestingly, two different signal peptidase I homologs exist in H. volcanii, Sec11a and Sec11b, which likely play distinct physiological roles. Type IV prepilin peptidase PibD processes flagellin/pilin precursors, being essential for the biogenesis and function of the archaellum and other cell surface structures in H. volcanii.

**Keywords: archaeal proteolysis, membrane-associated proteases,** *Haloferax volcanii***, cell envelope, S-layer glycoprotein**

# **INTRODUCTION**

Membrane-associated proteases participate in a variety of processes essential for cell physiology including membrane protein quality control, processing of exported and/or membraneanchored polypeptides, regulatory circuits, cell-signaling, the stress response and assembly of cell surface structures (Akiyama, 2009; Dalbey et al., 2011; Schneewind and Missiakas, 2012; Konovalova et al., 2014). Their targets are mainly membrane-bound or secreted proteins which account for 20–30% of total proteins encoded in most genomes (Wallin and von Heijne, 1998) and include membrane receptors, structural proteins, transporters and enzymes such as transferases, oxidoreductases, and hydrolases.

Integral membrane proteases comprise two distinct groups. The first group is represented by peptidases anchored to the cytoplasmic membrane that exert their catalytic activity in an aqueous compartment (cytoplasm, periplasm, or extracellular milieu) either at the aqueous-membrane boundary or after the substrate has been released or extracted from the membrane. Within this category are signal peptidases (SP), site 1 proteases (S1P) or sheddases, signal peptide hydrolases SPPA, HtpX, sortases and the energy-dependent proteases FtsH and LonB. The second group is represented by the so-called intramembrane cleaving proteases (ICliPs) which have their active sites immersed in the hydrophobic environment of the membrane (Wolfe, 2009; Dalbey et al., 2012). This group includes GxGD-aspartyl proteases (eukaryal signal peptide peptidase SPP and presenilin families), rhomboids and site 2 proteases (S2P).

*Archaea*, one of the three domains of life, are widespread in nature but predominate in environments with extreme values of pH, temperature, salt concentration and pressure (Robertson et al., 2005). Studies on archaeal biology are encouraged as they provide the opportunity to better understand cell physiology as well as extend the resources for biotechnology.

The genome sequences of archaea show that these unusual organisms encode a variety of proteolytic enzymes some of which have been characterized (Ward et al., 2002; Maupin-Furlow et al., 2005; De Castro et al., 2006; Ng et al., 2007). Most of the membrane protease families found in bacteria and/or eukaryotic cells also occur in archaea, however, the role of these enzymes in the context of the archaeal cell is poorly understood. In the last decade a number of studies have started to advance the knowledge on this field (see references in **Table 1**). This mini review describes what is known about proteases associated with the cell surface of archaeal cells on the basis of complete genome sequences and biochemical and/or genetic studies. Emphasis will be placed on the proteolytic enzymes affecting the cell envelope and surface structures of the euryarchaeon *Haloferax volcanii* and other haloarchaea. *H. volcanii* grows in a wide range of salinity (1.5–3.5 M NaCl) and is a model organism to study archaeal biology due to a number of advantages including the simplicity of its culture conditions, availability of complete genome sequences and feasibility of its genetic manipulation.

# **MEMBRANE-ASSOCIATED PROTEASES OF** *ARCHAEA*

An overview of the repertoire of membrane proteases that occur in archaeal cells is shown in Table S1 based on *in silico* examination of the complete genome sequences of some representative archaea members. Some protease families are widely represented among archaeal genomes such as HtpX homologs, LonB, SP, and Site 2

**Table 1 |**

**Predicted membrane**

 **proteases of**

**represtenative**

 **archaeal genomes.**

**Table 1 |**

**Continued** (Continued)

proteases (S2Ps) whereas others are restricted to a limited number of organisms (for instance the protease families A5, M10, and PrsW protease). **Table 1** describes the membrane proteases that have been experimentally characterized from the *Archaea* domain. Some of them have been studied in more detail (SPI and TFPP-like SP) and at least a few of their endogenous substrates have been identified (e.g., preflagellins, prepilins, and sugar-binding proteins for TFPP-like peptidases). However, most families have been examined to a limited extent or remain uncharacterized, and their biological relevance and/or targets are unknown (e.g., rhomboids, LonB, CAAX prenyl protease homologs, S2Ps).

The crystal structures of a number of archaeal membrane proteases have been solved (*Methanococcus maripaludis* FlaK; *Thermococcus onnurineus* and *Archaeoglobus fulgidus* LonB proteolytic domains; S2P transmembrane segments (TMSs) core from *Methanococcus jannaschii;* MCMJR1 peptidase from *Methanoculleus marisnigri*) providing valuable structure/function insights on these protease families (see **Table 1** for references).

# **MEMBRANE PROTEASES IMPLICATED IN THE ASSEMBLY OF THE ARCHAEAL CELL ENVELOPE AND SURFACE STRUCTURES**

Probably one of the most distinctive features of archaea is their ability to survive in environments with extremely adverse conditions that are lethal for most life forms. To this end, they have adapted their physiology and cellular structures. One such instance is the cell envelope. The archaeal cell envelope is composed of an atypical cellular membrane constituted by isoprenyl ether glycerol phospholipids surrounded by surface S-layer proteins as the major (or sole) component of the cell wall (Albers and Meyer, 2011). These structures maintain the cellular integrity and functionality as well as serve as a shell to cope with the harsh conditions predominating in their surroundings (Claus et al., 2005).

In addition to the S-layer, archaea show very diverse and complex cell surface structures (reviewed in Lassak et al., 2012). The biogenesis of the appendages composed of bacterial type IV pilin subunits, the pili and the archaeal flagellum or archaellum, has been characterized to some extent. These structures play important roles in cell motility as well as in surface attachment, DNA exchange and cell-cell interaction.

Haloarchaea, a very diverse and probably the best characterized group of archaea, flourish in habitats with high salinity (> 2M NaCl) and intense solar irradiation. In the haloarchaeon *H. volcanii* the structure and maturation of the S-layer glycoprotein as well as the biogenesis of pili and flagella have been examined (Jarrell et al., 2010; Kaminski et al., 2013; Kandiba et al., 2013; Tripepi et al., 2013; Esquivel and Pohlschroder, 2014). The adequate localization and functionality of these structures requires the participation of different families of proteases which are immersed in the context of the cytoplasmic membrane. Below we describe the recent advances on the membrane-associated proteases involved in the processes leading to the assembly of the cell envelope and surface structures in the euryarchaeon *H. volcanii*. The currently available information is summarized in **Figure 1**.

# **PROTEASES INVOLVED IN THE BIOGENESIS OF THE CYTOPLASMIC MEMBRANE AND SECRETION OF PREPROTEINS**

The quality control of membrane proteins is essential for proper cell physiology. In bacteria and eukaryotic organelles a major role in this process is performed by the energy-dependent membrane protease FtsH (Dalbey et al., 2012; Langklotz et al., 2012). Archaea possess only two ATP-dependent proteases: the 20S proteasome (soluble enzyme) and an unusually membrane-bound version of the Lon protease (LonB). The archaeal LonB probably resembles functionally to the FtsH protease which is absent in archaea. LonB has been biochemically and/or structurally characterized in several archaeal members (**Table 1**). In agreement with the genomic prediction, LonB has been immunolocalized in association with the cell membrane in the haloarchaea *Natrialba magadii* and *H. volcanii*. The recombinant protease derived from *N. magadii* (NmLon) showed DNA binding capacity *in vitro*, a feature in common with LonA proteases (Sastre et al., 2011). As FtsH is for *Escherichia coli* (Langklotz et al., 2012), LonB is essential for viability of *H. volcanii* cells. On the other hand suboptimal expression of this protease affects growth rate, cell shape, antibiotic sensitivity, and lipid composition (Cerletti et al., 2014). Also, *H. volcanii* mutant cells deficient in Lon content are more sensitive to puromycin compared to wild type cells suggesting that LonB is involved in the disposal of abnormal proteins. A distinctive feature of haloarchaea is the presence of red membrane-bound carotenoid pigments (C50-bacterioruberins) which serve to protect their macromolecules from the damaging effects of UV light (Khanafari et al., 2010). Interestingly, the cellular content of bacterioruberins dramatically increased in *H. volcanii* mutant cells with a suboptimal Lon concentration while overexpression of this protease rendered the cells colorless (Cerletti et al., 2014). This observation suggests that LonB controls carotenoid biosynthesis in *H. volcanii* probably by degrading enzyme/s involved in this pathway. It is likely that deregulation of the cellular concentration of bacterioruberins and other lipids affects membrane stability contributing to the lethal phenotype of the *lon* knock out mutant.

Signal peptidases are central in the protein secretion process as they remove signal peptides from secretory and membranebound polypeptides. In archaea, type I signal peptidase (SPI), type IV prepilin peptidase (TFPP)-like enzymes and signal peptide peptidase (SPP) have been characterized. A detailed description on the distribution and properties of these enzymes has been previously reported (Ng et al., 2007). SPIs process the majority of pre-proteins that are translocated through the general secretion pathway (Sec), however, whether this enzyme also cleaves Tat signal peptides remains to be demonstrated. Like all members of the SPI family, archaeal SPIs are serine proteases and based on studies performed in SPI from *M. voltae* (Ng and Jarrell, 2003) and *H. volcanii* (Fink-Lavi and Eichler, 2008) the catalytic mechanism of the archaeal SPI homolog seems to rely on a Ser/His/Asp tryad resembling the eukaryotic enzyme. In *H. volcanii* two different SPIs with distinct efficiency for substrate cleavage exist, Sec11a and Sec11b, however, only Sec11b is essential for viability (Fine et al., 2006). It is likely that these enzymes exert different roles and/or cleave distinct substrates *in vivo*.

SPII removes signal peptides from lipoproteins. Although there are numerous proteins in archaea that contain signal peptides with the lipobox motif, including several predicted to be secreted via the Tat pathway, homologs of bacterial SPII have not been identified in archaeal genomes (Giménez et al., 2007). Thus, it has been proposed that a distinct enzyme may exist in archaea to process prelipoproteins (Ng et al., 2007).

### **PROTEASES INVOLVED IN MATURATION OF THE CELL WALL (S-LAYER GLYCOPROTEIN)**

In *H. volcanii* the S-layer glycoprotein is the sole structure that constitutes the cell wall. This protein has been used to examine the molecular/structural adaptations of haloarchaeal proteins to high salt and has served as a model to study protein glycosylation in archaea (Eichler et al., 2013; Jarrell et al., 2014). In haloarchaea, maturation of the S-layer glycoprotein requires at least three different types of posttranslational modifications: glycosylation, proteolytic cleavage and isoprenylation (Konrad and Eichler, 2002; Eichler, 2003). The glycosylation process of the S-layer has been recently reviewed (Eichler et al., 2013).

Sortases are cysteine proteases from Gram-positive bacteria that "sort" proteins to the cell surface by covalently joining them to the cell wall or polymerize pilins to build pili (Proft and Baker, 2009; Clancy et al., 2010; Hendrickx et al., 2011; Spirig et al., 2011). These enzymes modify surface proteins by recognizing and cleaving a sorting signal located either in the N or *C*-terminus of the target protein. Many genomes in bacteria and archaea encode proteins containing a *C*-terminal domain with structural similarity to the *C*-terminus of sortase substrates. These proteins coexist in these genomes with at least one member of the protease families denoted as exosortases (bacteria) or archaeosortases (archaea). Exo and archaeosortases are polytopic membrane proteins with no sequence homology to bacterial sortases. However, they contain the conserved cysteine, arginine, and histidine residues found in the active site of sortases suggesting that they may perform similar functions (Haft et al., 2012). Recently it was reported that *H. volcanii* mutant cells with a deletion in the archaeosortase gene *artA* showed growth defects (which were more evident under low-salt conditions), alterations in cell shape and the S-layer organization, impaired motility and decreased conjugation rates (Abdul Halim et al., 2013). This work demonstrated that ArtA is involved in *C*-terminal processing of the S-layer glycoprotein suggesting that archaeosortases are functional homologs of bacterial sortases. Considering the location of the archaeosortase recognition sequence (PGF) immediately following the TMS of the substrate protein, it was proposed that this enzyme may facilitate the covalent attachment of target proteins (e.g., S-layer glycoprotein) to a membrane lipid in contrast to sortases which attach proteins to the growing cell wall.

Rhomboids are membrane serine proteases involved in regulatory intramembrane proteolysis (RIP) and are conserved in the three domains of life (Lemberg, 2013). The catalytic mechanism of rhomboids relies on a Ser/His dyad located in different TMS of the protease to cleave membrane protein substrates. In eukaryotic cells the functions of this protease family are very diverse and include cell-cell signaling, development, apoptosis, organelle integrity and parasite invasion (reviewed in Freeman, 2014). The relevance of rhomboids in the prokaryotic cell physiology is scarcely understood. In bacteria, rhomboid null mutants show phenotypes that may be related to defective cell envelope and/or cell-surface structures. In *Bacillus subtilis*, a mutant strain in the rhomboid homolog YqgP displayed a slight decrease in glucose uptake and a defect in cell division leading to the formation of filamentous cells (Mesak et al., 2004); *Mycobacterium smegmatis* rhomboid mutants showed reduced capacity for biofilm formation and increased sensitivity to antibiotics (Kateete et al., 2012). So far only TatA, a protein component of the Tat translocon in the pathogenic bacterium *Providencia stuartii,* has been experimentally confirmed as a rhomboid substrate (Stevenson et al., 2007). In this organism the rhomboid protease AarA cleaves an *N*-terminal extension of TatA which in turn allows for secretion of an unknown quorum sensing signal. Archaea appear to encode various sequences for rhomboid proteases (Table S1). In haloarchaea, homologs with various topologies can be found including proteins with six or more TMS as well as unusual rhomboids containing an AN-1 Zn-finger domain at the *N*-terminus. *H. volcanii* has two putative genes for rhomboids, RhoI (nine TMS) and RhoII (six TMS, with *N*-terminal AN-1 Zn finger domain). A knock-out mutant of *rhoII* in *H. volcanii* displayed mild defects in motility and novobiocin sensitivity. This mutant strain was also affected in the glycosylation of the S-layer. In *H. volcanii* wild type cells the S-layer glycoprotein Asn732 is bound to an oligosaccharide containing at least 6 repeating units of sulfoquinovose-hexose (SQ-Hex) while in the mutant strain this residue contained only two SQ-Hex suggesting that RhoII controls (directly or indirectly) the protein glycosylation process in *H. volcanii* (Parente et al., 2014).

#### **PROTEASES INVOLVED IN THE BIOGENESIS OF CELL SURFACE APPENDAGES**

In bacteria, the precursors of type IV pilins and related pseudopilins are processed by a special enzyme belonging to a novel aspartic acid protease family, the type IV prepilin signal peptidase (SPIV/TFPP; Ng et al., 2009). In contrast to SPI and SPII, this enzyme cleaves the signal peptides directly after the *n*-region leaving the *h*-region bound to the mature protein facilitating anchoring/assembly of pilin subunits onto the cell surface (Ng et al., 2007). Archaea encode TFPP-like proteins and they have been studied with regard to their role in the assembly of the structures composing the motility apparatus (**Table 1**). The archaellum is composed of unique proteins that are unrelated to bacterial flagellins. Archaeal preflagellins contain short signal peptides at the *N*-terminus which are similar to those of bacterial type IV pilins, the protein components of pili. These filamentous surface

structures facilitate twitching motility in bacteria. TFPP-like proteases process the signal peptides of archaeal preflagellins. The enzymes present in *M. maripaludis* and *Methanococcus voltae* (FlaK), *Sulfolobus solfataricus,* and *H. volcanii* (PibD) are the most extensively characterized TFPPs of archaea (see references in **Table 1**). FlaK and PibD show some divergences including the length of the signal peptide, key amino acid residues surrounding the cleavage site as well as substrate preference (Ng et al., 2009). PibD from *S. solfataricus* and *H. volcanii* has a broader substrate selection than FlaK, as, in addition to preflagellins, these enzymes can mature prepilins (Albers et al., 2003; Tripepi et al., 2010). *S. solfataricus* PibD also processes certain sugarbinding proteins of the "bindosome," filamentous-like structures that extend from the cell surface (Albers et al., 2003; Szabo et al., 2006).

The *H. volcanii* genome encodes flagellins and contains genes for other type IV pilin-like proteins. Tripepi et al. (2010) showed that deletion of *pibD* disrupted preflagellins processing and prevented maturation of type IV pilin-like proteins. The mutant cells were non-motile and were unable to adhere to a glass surface. These results suggest that PibD is needed for maturation of preflagellins and other type IV pilin-like proteins in *H. volcanii*.

Recently, based on *in vivo* analysis of the catalytic activity of *Sulfolobus acidocaldarius* PibD, TFPPs were renamed as GxHyD group of proteases (rather than DxGD; Henche et al., 2014).

# **CONCLUDING REMARKS**

In prokaryotes the assembly and composition of cell surface structures are essential for the adjustment to the varying conditions of the environment and to interact with their surroundings (e.g., establish cell-cell and/or cell-substrate contacts). In the haloarchaeon *H. volcanii*, several membrane-associated proteases are implicated in different processes (protein secretion, processing and sorting) leading to the biogenesis of the cell wall and extracellular appendages (**Figure 1**), highlighting the importance of these enzymes in the adaptation and interaction of archaea with their environment.

Structural analysis of archaeal membrane proteases (Flak and GxGD proteases) have advanced the knowledge on the catalytic and molecular mechanism of intramembrane cleaving proteases. This will help to understand the mechanism of the eukaryotic homologous enzymes which are implicated in human physiology (regulation of immune response) and/or in the development of diseases (e g. Alzheimer).

There are still many open questions in this field: e.g., endogenous substrates of most membrane proteases are unknown. Efforts should continue to better understand the role of membrane proteases in archaeal physiology.

#### **AUTHOR CONTRIBUTIONS**

All authors made substantial contributions to the acquisition, analysis and interpretation of data for this review. All authors critically reviewed and edited the manuscript, and approved the final version before submission to publication. All authors agree to be accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

#### **ACKNOWLEDGMENTS**

Research at De Castro's laboratory is supported by grants from the National Council of Scientific and Technological Research (CON-ICET) and the National University of Mar del Plata (UNMDP) Argentina. Micaela Cerletti is a PhD student at the UNMDP supported by a research fellow from CONICET; María I. Giménez and Rosana E. De Castro are researchers from CONICET.

#### **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at:http://www.frontiersin.org/journal/10.3389/fmicb.2015.00039/ abstract

#### **REFERENCES**

of Providencia stuartii and gene deletion in *Mycobacterium smegmatis*. *PLoS ONE* 7:e45741. doi: 10.1371/journal.pone.0045741

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 27 November 2014; accepted: 12 January 2015; published online: 06 February 2015.*

*Citation: Giménez MI, Cerletti M and De Castro RE (2015) Archaeal membraneassociated proteases: insights on Haloferax volcanii and other haloarchaea. Front. Microbiol. 6:39. doi: 10.3389/fmicb.2015.00039*

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology.*

*Copyright © 2015 Giménez, Cerletti and De Castro. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited andthatthe original publication inthis journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

**REVIEW ARTICLE** published: 26 November 2014 doi: 10.3389/fmicb.2014.00641

# Biosynthesis of archaeal membrane ether lipids

# *Samta Jain1,2 †, Antonella Caforio1,2 and Arnold J. M. Driessen1,2 \**

<sup>1</sup> Department of Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, Netherlands <sup>2</sup> The Zernike Institute for Advanced Materials, University of Groningen, Groningen, Netherlands

#### *Edited by:*

Sonja-Verena Albers, University of Freiburg, Germany

#### *Reviewed by:*

Dong-Woo Lee, Kyungpook National University, South Korea Jerry Eichler, Ben Gurion University of the Negev, Israel

#### *\*Correspondence:*

Arnold J. M. Driessen, Department of Molecular Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute –The Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 7, 9747 AG Groningen, Netherlands e-mail<EMAIL_ADDRESS>
#### *†Present address:*

Samta Jain, Department of Medicine, Section of Infectious Diseases, Boston University School of Medicine, 02118 Boston, MA, USA

A vital function of the cell membrane in all living organism is to maintain the membrane permeability barrier and fluidity. The composition of the phospholipid bilayer is distinct in archaea when compared to bacteria and eukarya. In archaea, isoprenoid hydrocarbon side chains are linked via an ether bond to the sn-glycerol-1-phosphate backbone. In bacteria and eukarya on the other hand, fatty acid side chains are linked via an ester bond to the sn-glycerol-3-phosphate backbone. The polar head groups are globally shared in the three domains of life.The unique membrane lipids of archaea have been implicated not only in the survival and adaptation of the organisms to extreme environments but also to form the basis of the membrane composition of the last universal common ancestor (LUCA). In nature, a diverse range of archaeal lipids is found, the most common are the diether (or archaeol) and the tetraether (or caldarchaeol) lipids that form a monolayer. Variations in chain length, cyclization and other modifications lead to diversification of these lipids. The biosynthesis of these lipids is not yet well understood however progress in the last decade has led to a comprehensive understanding of the biosynthesis of archaeol. This review describes the current knowledge of the biosynthetic pathway of archaeal ether lipids; insights on the stability and robustness of archaeal lipid membranes; and evolutionary aspects of the lipid divide and the LUCA. It examines recent advances made in the field of pathway reconstruction in bacteria.

#### **Keywords: archaea, ether lipids, isoprenoids, biosynthesis, lipid divide**

**INTRODUCTION**

The "Woesian Revolution" in 1977 defined the three domains of life as the Eukarya, the Bacteria and the Archaea (Woese and Fox, 1977). The archaeal membrane lipid composition is one of the most remarkable feature distinguishing Archaea from Bacteria and Eukarya where the hydrocarbon chain consists of isoprenoid moieties which are ether linked to the enantiomeric glycerol backbone, glycerol-1-phosphate (G1P) in comparison to glycerol-3-phosphate (G3P) of bacteria and eukarya that is ester linked to the fatty acid derived hydrocarbon chain. Polar head groups on the other hand are common in all three domains of life. Other than this core archaeal diether lipid structure, a bipolar tetraether lipid structure is also prevalent in many archaea that span the entire archaeal membrane forming a monolayer (Koga and Morii, 2007). It should be stressed that ether-linked lipids are not unique to archaea *per se*, but are also found in Bacteria and Eukarya, although not ubiquitously distributed and usually only a minor component of the lipid membrane.

The stereo specificity of archaeal lipids and their unique structure was hypothesized to be chemically more stable thereby rendering the organism with the ability to resist and thrive in extreme environmental conditions (Koga, 2012). However, archaea are also found in mesophilic and neutrophilic environment where such a structural role of ether lipids is still not postulated. At the same time, the distinguishing lipid structures have formed the basis to the evolutionary studies describing archaeal and bacterial differentiation. Several models hypothesizing the early evolution of archaeal and bacterial phospholipid biosynthesis were proposed

to answer intriguing questions about the nature of the ancestral membrane lipid composition (Lombard et al., 2012b). Understanding the archaeal lipid biosynthetic pathway is crucial to the above studies.

Decades of studies on the biosynthesis of archaeal lipids have advanced our knowledge on the major enzymatic processes but the pathway is, however still not completely understood. Several enzymes of the pathway have been studied and characterized biochemically but there are also gaps in our understanding of the archaeal lipid biosynthetic pathway and little is known about its regulation. With more genome sequences becoming available, advanced phylogenetic studies have been performed recently (Boucher et al., 2004; Daiyasu et al., 2005; Lombard et al., 2012a; Villanueva et al., 2014) and this helped to more precisely define its evolution. This review will focus on existing knowledge and recent studies on the enzymes of the pathway, the physicochemical properties of archaeal lipids, and the theories on the lipid divide.

# **BIOSYNTHESIS OF ARCHAEAL MEMBRANE LIPIDS ISOPRENOID BUILDING BLOCKS AND CHAIN ELONGATION**

Isoprenoids are ubiquitous to all three domains of life. They are structurally diverse, forming more than 30,000 different compounds in nature ranging from steroids, quinones, carotenoids, and membrane lipids. The building blocks of isoprenoids are universal carbon five subunits called isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) that are isomers. The biosynthetic pathway leading to the synthesis of IPP

and DMAPP vary in different organisms (reviewed in Lombard and Moreira, 2011; Matsumi et al., 2011). To date, three pathways have been reported – 2-*C*-methyl-D-erythritol 4-phosphate/1 deoxy-D-xylulose 5-phosphate pathway (MEP/DOXP pathway) and two mevalonate (MVA) pathways. The MEP pathway genes share no homology to genes of the MVA pathway where pyruvate and glyceraldehyde-3-phosphate molecules are condensed together to form 1-deoxy-D-xylulose-5-phosphate (DXP) which is subsequently converted to IPP and DMAPP by five enzymes (Xue andAhring,2011; **Figure 1**). TheMEP pathway is most common in bacteria although some Firmicutes possess the MVA pathway. The MVA pathway consists of seven enzymatic reactions where two acetyl-CoA molecules are condensed to form acetoacetyl-CoA, which is further condensed to form 3-hydroxy-3-methylglutaryl CoA (HMG-CoA). HMG-CoA undergoes phosphorylation and decarboxylation to form IPP via the formation of MVA (**Figure 1**). The classical MVA pathway is common to eukaryotes while some plants and photosynthetic eukaryotes possessing the MEP pathway in addition (Lombard and Moreira, 2011).

Interestingly, homologs of the last three enzymes of the classical MVA pathway, i.e., phosphomevalonate kinase, diphosphomevalonate decarboxylase and isopentenyl diphosphate isomerase could not be found in the majority of archaea (except in Sulfolobales that have classical MVA pathway; Boucher et al., 2004). This search led to the discovery of the alternate MVA pathway that differs from the classical one in the last three steps (**Figure 1**). The enzyme isopentenyl kinase (IPK) was first discovered in *Methanocaldococcus jannaschii* and found to be conserved in archaea (Grochowski et al., 2006). Its structure was determined (Dellas and Noel, 2010) and IPK enzymes from *Methanothermobacter thermautotrophicus* and *Thermoplasma acidophilum* were characterized biochemically (Chen and Poulter, 2010). In the alternate MVA pathway, phosphomelavonate is decarboxylated to isopentenyl phosphate by a decarboxylase (enzyme yet to be identified in archaea), which is subsequently phosphorylated to IPP by IPK. Furthermore, instead of the typical IDI1 isomerase that performs the last step of the classical MVA pathway, archaea have IDI2 which is not homologous to IDI1 but that performs the same reaction (Matsumi et al., 2011). Interestingly, a decarboxylase enzyme that converts phosphomelavonate to isopentenyl phosphate was found in green non-sulfur bacteria *Roseiflexus castenholzii* along with the presence of IPK enzymes indicating the existence of alternate MVA pathway in organisms other than archaea (Dellas et al., 2013). In general, IPP and DMAPP are synthesized by the MEP pathway in most of the bacteria and by two MVA pathways in eukarya and archaea. The classical MVA pathway of eukaryotes and the alternate MVA pathway of archaea share four of their seven steps.

The isoprenoid building blocks IPP and DMAPP undergo sequential condensation reactions where DMAPP acts as the first allylic acceptor of IPP leading to the formation of a carbon 10 (C10) compound termed geranyl diphosphate (GPP). Further condensation reactions proceed with the addition of IPP molecules where the chain length increases each time by a C5 unit forming farnesyl (C15), geranylgeranyl (C20), farnesylgeranyl (C25) diphosphate etc. This reaction of chain elongation is catalyzed by enzymes belonging to the family of prenyl transferases that are common to all three domains of life (Wang and Ohnuma, 1999; Vandermoten et al., 2009). Depending on the length and geometry of the final molecule, prenyl transferases can have several members in its family. The geometry of the molecule could be *cis* or *trans* and the chain length of the *trans* form generally ranges from C10 (e.g., monoterpenes) to C50 (e.g., Coenzyme Q10) and even longer for the *cis* forms. The chain length found in archaeal membrane lipids is always in the *trans* form and composed mostly of C20 [geranylgeranyl diphosphate (GGPP)] or C25 (farnesylgeranyl diphosphate). Tetraethers are composed of a C40 chain length, the synthesis of which is still unknown (discussed below). The archaeal prenyltransferase enzymes GGPP synthase and farnesylgeranyl diphosphate synthase synthesize specifically C20 or C25 product chain lengths, respectively (Ohnuma et al., 1994; Tachibana et al., 2000; Hemmi et al., 2002; Lai et al., 2009). They belong to short chain *trans* prenyl transferases family (that catalyze reactions ranging from C10–C25). Interestingly, a bifunctional prenyltransferase that catalyzes the synthesis of both C15 and C20 isoprenoids has been characterized from *Thermococcus kodakaraensis* (Fujiwara et al., 2004) and *Methanobacterium thermoautotrophicum*and is considered to be an ancient enzyme (Chen and Poulter, 1993). Multiple sequence alignment of homologues of the family display high sequence similarity with seven conserved regions where region two and seven contain the highly conserved aspartate rich sequences called first aspartate-rich motif (FARM) and second aspartate-rich motif (SARM) domains, respectively. Numerous mutagenesis and structural studies including several members of the family show that the region within the aspartate rich domains are involved in the binding and catalysis of the substrate while the regions flanking these domains are the major determinants of the chain length as they contribute to the size of the active site hydrophobic pocket. For example, GGPP synthase from *Sulfolobus acidocaldarius* could be mutated (at Phe-77 which is fifth amino acid upstream of FARM) to catalyze longer chain length (C30–C50) products (Ohnuma et al., 1996) and farnesyl pyrophosphate (FPP) synthase of *Escherichia coli* could be mutated (at Tyr-79, also 5th amino acid upstream of FARM) to create GGPP synthase (Lee et al., 2005). Physical factors have also been shown to influence the chain length of the product, e.g., the bifunctional enzyme farnesyl diphosphate/GGPP synthase of *Thermococcus kodakaraensis* shows an increase in the FPP/GGPP ratio with the reaction temperature (Fujiwara et al., 2004).

#### **GLYCEROL-1-PHOSPHATE BACKBONE**

The glycerophosphate backbone of archaea has an opposite stereoconfiguration than those of bacteria and eukarya. The archaeal enzyme responsible is G1P dehydrogenase that shares homology with alcohol and glycerol dehydrogenases but no homology to the bacterial/eukaryal G3P dehydrogenase. They belong to two separate families. However, both catalyze the reduction of dihydroxyacetone phosphate (DHAP) using nicotinamide adenine dinucleotide hydrogen (NADH) or nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) as substrate (**Figure 2**; **Table 1**). G1P dehydrogenase uses Zn2<sup>+</sup> for metal ion interaction in its active site (Han and Ishikawa, 2005) and transfers the pro-R hydrogen of NADH in contrast to G3P dehydrogenase that transfers the pro-S hydrogen (Koga et al., 2003); both enzymes bind the nicotinamide ring in an opposite orientation.

G1P dehydrogenase is conserved in archaea. The enzyme has been purified and characterized from *Methanothermobacter thermoautotrophicus* as an octamer (Nishihara and Koga, 1997), *Aeropyrum pernix* as a homodimer (Han et al., 2002) and from *Sulfolobus tokodaii* (Koga et al., 2006). Its activity has been accessed in cell free homogenates of several archaea (Koga and Morii, 2007; Lai et al., 2009).

Until recently it was thought that the stereo specificity is the hallmark of the 'lipid divide' where the G1P backbone is exclusively attributed to archaea. This was challenged by the discovery and characterization of the bacterial G1P dehydrogenase homolog of *Bacillus subtilis* which is annotated as 'AraM' (Guldan et al., 2008). It is also found in other related Gram positive and negative bacteria. Similar to the G1P dehydrogenase of *Aeropyrum*

soluble enzymes of the pathway are colored in blue and the membrane proteins in green. The biosynthetic steps leading to the formation of tetraether lipids is unknown. Archaetidylserine (AS) and saturated AS (sAS) are depicted as an example of polar head

pyrophosphate (DMAPP) are isomers. GGPP, geranylgeranyl diphosphate; G1P, sn-glycerol-1-phosphate; GGGP, 3-Ogeranylgeranyl-sn-glyceryl-1-phosphate; DGGGP, 2,3-bis-O- geranylgeranylsn-glyceryl-1-phosphate.

*pernix*, AraM forms a homodimer and performs G1P dehydrogenase activity. However, the two enzymes have different catalytic efficiencies and AraM is Ni2<sup>+</sup> ion dependent (Han et al., 2002; Guldan et al., 2008). Remarkably, the G1P molecule eventually becomes part of an archaea type ether lipid heptaprenylglyceryl phosphate in *B. subtilis*, the function of which is still unknown (Guldan et al., 2011).

# **ETHER LINKAGES**

The first and second ether bonds between G1P and GGPP is catalyzed by the enzyme geranylgeranylglycerly diphosphate (GGGP) synthase and di-*0*- geranylgeranylglycerly diphosphate (DGGGP) synthase respectively. GGGP synthase is a conserved enzyme found in all archaea except Nanoarchaeota, which is a symbiont and possesses no genes of the lipid biosynthesis pathway

(Podar et al., 2013). It is also found in some bacteria where the polyprenyl diphosphate substrate chain length could vary, e.g., PcrB of *Bacillus subtilis* which is a heptaprenyl diphosphate synthase (Ren et al., 2013). GGGP synthase is a crucial enzyme in the biosynthetic pathway of phospholipid metabolism in archaea as it brings together the three important characteristic features of the archaeal lipid structure – stereoisomeric G1P glycerol backbone and isoprenoid GGPP side chain linking them together via an ether bond (**Figure 2**; **Table 1**). Phylogenetic analysis of the GGGP synthase enzymes distinguishes it into two families, group I and group II, both comprising of archaeal and bacterial sequences. Several enzymes from both the groups have been characterized and a recent study performed the biochemical analysis of 17 members of GGGP synthase family (Peterhoff et al., 2014). The enzymes of group I form dimers (except the monomeric GGGP synthase of *Halobacterium salinarum*) and the group II enzymes are dimeric or hexameric in nature. Both the groups are further subdivided into Ia, Ib, IIa, and IIb with a and b corresponding archaea and bacteria, respectively. Crystal structures of enzymes from all the four groups have been solved. The first crystal structure from the group I GGGP synthase of *Archaeoglobus fulgidus* displays a modified triose phosphate isomerase (TIM)-barrel structure (Payandeh et al., 2006). It forms a dimer bound to the G1P substrate with a central eight-stranded parallel β-barrel and a hydrophobic core surrounded by α-helices (**Figure 3A**). Helix-3 is replaced by a 'strand' which is a novel TIM-barrel modification not observed

previously. The substrate GGPP binds to the deep cleft traversing the top of the β-barrel. There is a 'plug' at the bottom of the barrel and the active site lies at the C-terminal end. The G1P molecule sits near the top inner rim of the barrel and the phosphate group binds to the standard phosphate-binding motif of the TIM-barrel. G1P forms 14 hydrogen bonds within the active site. The (βα)8-barrel fold is found in all the other structures of the GGGP synthases as well with the active site at the C-terminus. The crystal structure of group II archaeal hexameric GGGP synthase of *Methanothermobacter thermoautrophicus* displays a combination of three dimers that resemble the group I dimer (**Figures 3B,C**). In group II, however, the plug of the barrel is longer than in group I and there are 'limiter residues' that restrict the length of hydrophobic pocket to accommodate the polyprenyl diphosphates of a specific length. Interestingly, an aromatic anchor residue is responsible for the hexameric configuration of the enzyme, mutation of which causes it to dimerize without any loss of activity (Peterhoff et al., 2014).

The intrinsic membrane protein DGGGP synthase catalyzes the formation of the second ether bond between the substrate GGGP and GGPP to form DGGGP (**Figure 2**). It belongs to the family of ubiquinone-biosynthetic (UbiA) prenyltransferases, the members of which are responsible for the biosynthesis of respiratory quinones, chlorophyll, heme etc. by transferring a prenyl group to the acceptors that generally have hydrophobic ring structures. DGGGP synthase is divergent among archaea and could not be

identified in the genomes of Thaumarchaeota (Villanueva et al., 2014).

Unlike other enzymes of the pathway, DGGGP synthase has not been well characterized probably due to technical limitations with overexpression of the membrane protein. The DGGGP synthase activity was first found in the membrane fraction of *Methanothermobacter marburgensis* (Zhang and Poulter, 1993). Later the gene was identified in the genome of *Sulfolobus solfataricus* as UbiA-2, cloned in *E. coli* and purified to study the Mg2<sup>+</sup> dependent enzymatic activity using radiolabeled substrates and mass spectrometry (Hemmi et al., 2004). The ratio of the substrates utilized in the reaction was found to be 1:1.1 in a double labeling experiment using [3H]GGPP and [14C]GGGP, respectively. Specificity for GGPP and GGGP was also measured by substituting them with different prenyl substrates, of which none of them were used in the reaction by DGGGP synthase. In another study, DGGGP synthase was shown to accept both the S and R form of GGGP showing that unlike GGGP synthase, it is enantio unselective (Zhang et al., 2006). DGGGP synthase activity of *Archaeoglobus fulgidus* (Lai et al., 2009) and *Methanosarcina acetivorans* (Yokoi et al., 2012) was also observed in *E. coli* when the corresponding genes were expressed along with four previous enzymes of the pathway. However, the expression level of the enzyme was either too low to detect (Lai et al., 2009) or not investigated (Yokoi et al., 2012). In a later study, a higher expression level of DGGGP synthase of *Archaeoglobus fulgidus* was obtained in *E. coli* by changing

the ribosome-binding site and the activity of purified DGGGP synthase was monitored (Jain et al., 2014).

#### **CDP ARCHAEOL FORMATION**

The next step in the archaeal lipid biosynthetic pathway is the activation of DGGGP by cytidine triphosphate (CTP) to form the substrate for polar head group attachment called cytidine diphosphate (CDP)-archaeol (**Figure 2**; **Table 1**). The reaction is brought about by the enzyme CDP-archaeol synthase (CarS), the activity of which was first studied in the membrane fraction of *Methanothermobacter thermoautotrophicus* (Morii et al., 2000). Using various synthetic substrate analog, the activity was found to be specific for unsaturated archaetidic acid with geranylgeranyl chains and did not depend on the stereo specificity or ether/ester bond of the substrate. Minute amount of CDP-archaeol were also detected in growing cells labeled with inorganic 32P. The gene responsible for this activity was only identified in a recent study (Jain et al., 2014). The enzyme CarS is conserved among archaea (except Nanoarchaeota). However, like the enzyme DGGGP synthase, it could not be identified in the families of Thaumarchaeota.

Interestingly, an analogous reaction is found in the bacterial phospholipid biosynthetic pathway where phosphatidic acid is activated by CTP to form CDP diacylglycerol by the enzyme CDP diacylglycerol synthase (CdsA). Although the sequence similarity between CdsA and CarS is very low, hydropathy profile alignment of the two families shows similarity in their secondary structure with overlapping transmembrane segments and cytoplasmic loop regions residing in the C-terminus half. CarS from *Archaeoglobus fulgidus* was expressed and purified from *E. coli*. Similar to CdsA, CarS activity was found to be dependent on Mg2+, both accepts CTP and deoxycytidine triphosphate (dCTP) as substrates and does not utilize adenosine triphosphate (ATP), guanosine triphosphate (GTP), or thymidine triphosphate (TTP) nucleotides in the reaction using substrate DGGGP. However, the two enzymes displayed distinct activity with respect to the lipid substrate specificity where CarS only accepts unsaturated archaetidic acid with geranylgeranyl chains, while CdsA takes phosphatidic acid (Jain et al., 2014).

#### **POLAR HEAD GROUP ATTACHMENT**

The polar head groups serine, ethanolamine, glycerol and *myo*inositol are found in the phospholipids in all three domains of life. The enzymes that catalyze the replacement of the cytidine monophosphate (CMP) entity of CDP-archaeol or CDPdiacylglycerol with a polar head group are homologous and belong to CDP-alcohol phosphatidyltransferase family (Koga, 2011). Archaetidylserine (AS) synthase catalyzes the formation of AS from CDP-archaeol and L-serine (**Figure 2**) and is homologous to bacterial phosphatidylserine (PS) synthase. The enzyme can be classified into two subclasses. Subclass I includes enzymes distributed in Gram-negative bacteria, such as *E. coli* while subclass II enzymes are widespread among Gram-positive bacteria (*B. subtilis*), yeast and archaea. Studies using cell free extracts of *Methanothermobacter thermautotrophicus*, *B. subtilis*, and *E. coli* showed that both the AS and PS synthase from *Methanothermobacter thermautotrophicus* and *B. subtilis* have a broad substrate specificity and can accept lipid derivatives from archaea or bacteria. On the other hand, the *E. coli* PS synthase was specific for bacterial lipid derivatives only (Morii and Koga, 2003).

Archaetidylinositol phosphate (AI) synthase catalyzes the reaction where precursors L-*myo*-inositol-1-phosphate and CDParchaeol are converted to AI phosphate as an intermediate which is further dephosphorylated to AI (Morii et al., 2009). This reaction is similar to the bacterial phosphatidylinositol phosphate (PI) synthase. Similar to AS and PS synthase, the AI and PI synthase show a broad substrate specificity accepting both, archaeal and bacterial lipid derivatives as substrates (Morii et al., 2014). Enzymes homologous to PS decarboxylase and phosphatidylglycerol (PG) synthase have been identified in archaea as AS decarboxylase and archaetidylglycerol (AG) synthase but not yet characterized biochemically (Daiyasu et al., 2005).

#### **SATURATION OF DOUBLE BONDS**

The mature phospholipids of archaea exist in their fully saturated form. The archaeal enzyme digeranylgeranylglycerophospholipid reductase catalyzes the hydrogenation or saturation of the geranylgeranyl chains of unsaturated archaetidic acid (DGGGP) in a stereospecific manner (Xu et al., 2010). It belongs to the geranylgeranyl reductase (GGR) family that includes GGR from plant and prokaryotes that are mainly involved in photosynthesis. Prenyl reductases other than GGRs are also found in all three domains of life and these enzymes catalyze the complete or partial reduction

of isoprenoid compounds like respiratory quinones, tocopherol, dolichol, and other polyprenols (Ogawa et al., 2014).

The structures of the archaeal GGR monomer from *Thermoplasma acidophilum* (Xu et al., 2010) and *Sulfolobus acidocaldarius* (Sasaki et al., 2011) show that they belong to p-hydroxybenzoate hydroxylase (PHBH) superfamily of flavoproteins (**Figure 4A**). The GGR from the thermophilic archaea *Thermoplasma acidophilum* was crystalized in complex with flavin adenine dinucleotide (FAD) where FAD adopts the close confirmation that possibly changes with the binding of the substrate, like in other members of the PHBH family. The reduction of FAD is brought about by either NADH or other reducing agents. Since the protein was overexpressed in *E. coli*, a surrogate lipid-like ligand assigned as phosphatidylglycerol (PGX) was found in the active site forming an imperfect fit to the substrate binding pocket. The lipid binding cavity of GGR is R shaped having two tunnels where the larger tunnel B is more permissive than the smaller tunnel A which is restricted in shape (**Figure 4B**). The *S. acidocaldarius* GGR is structurally similar to GGR from *Thermoplasma acidophilum* in FAD binding and the catalytic region (**Figures 4C,D**) but not in the C-terminal domain which is longer in *S. acidocaldarius* GGR. The conserved sequence motif (YxWxFPx7-8GxG) lies in the large cavity of the catalytic domain and is thought to keep the substrate in position for the reduction reaction as also indicated by mutational studies. Although the enzymes reduce GGGP, they also reduce the double bonds of related compounds like GGGP and GGPP (Sasaki et al., 2011). Another study where the *Methanosarcina acetivorans* GGR was expressed in *E. coli* along with four previous genes of the archaeal lipid biosynthetic pathway, the DGGGP derivative with a fully saturated isoprenoid chain could be obtained (Isobe et al., 2014). Interestingly, the saturation only took place when GGR was coexpressed with a ferredoxin gene found upstream of GGR in the genome of *Methanosarcina acetivorans*, the ferredoxin possibly functioning as a specific electron donor. However, no ferredoxin coexpression was required when the *Methanosarcina acetivorans* GGR was replaced by *S. acidocaldarius* GGR in the same study. Also, the conservation of the ferredoxin gene upstream of GGR in other archaea was not analyzed.

It is not known at what step of the biosynthetic pathway, hydrogenation takes place. However, since that CarS is specific for the unsaturated substrate, saturation probably takes place after theformation of CDP-archaeol. Although the enzyme AS synthase can accept both saturated and unsaturated substrates for catalysis, the detection of unsaturated AS in the cells of *Methanothermobacter thermautotrophicus* suggests that hydrogenation may already take place after the polar head groups are attached (Koga and Morii, 2007).

#### **TETRAETHER FORMATION**

Tetraether (caldarchaeol) lipid structure with varying number (0–8) of cyclopentane moieties are widespread among archaea and a dominating membrane lipid structure in Crenarchaeota and Thaumarchaeota. Euryarchaeota synthesize archaeol or both archaeol and caldarchaeols. On the other hand, Thaumarchaeota have characteristic tetraether lipids with four cyclopentane moieties and a cyclohexane moiety (Villanueva et al., 2014). One of the most intriguing steps in the archaeal biosynthetic pathway is

the tetraether formation. *In vivo* studies suggested that tetraethers are formed from saturated diethers via head to head condensation reaction. Pulse chase and labeling experiment of *Thermoplasma acidophilum* cells with [14C]-MVA showed that the label first incorporates into the archaeol until saturation and only then into caldarchaeol. When an inhibitor of tetraether lipid synthesis (terbinafine) is used, pulse labeling leads to the accumulation of diethers and this phenomenon can be reversed by removal of the inhibitor (Nemoto et al., 2003). However, in another study, radiolabelled archaeol was not incorporated into the tetraethers of *Methanospirillum hungatei* cells and the presence of double bonds was necessary for the incorporation of labeled DGGGP into the tetraethers of *Methanobacterium thermoautotrophicus* cells (Poulter et al., 1988; Eguchi et al., 2000). The enzyme responsible for the formation of the presumed and unusual C–C bond for tetraethers has not been identified and there is no *in vitro* data to support this hypothesis (Koga and Morii, 2007). Recently, an alternative pathway for tetraether and cyclopentane ring formation was hypothesized (Villanueva et al., 2014). A multiple lock and key mechanism was proposed owing to the 'greater functional plasticity' of the enzymes IPP synthase, GGGP synthase, and DGGGP synthase so that they can accommodate prenyl substrates with a ring structure and chain length longer than C20. The cyclopentane rings would be formed early in the pathway before attachment of the glycerol moiety and the C20 geranyl molecules would couple together via a tail-to-tail mechanism to form the C40 phytoene chain by phytoene synthase, an enzyme that is wide spread in archaea. However, both possibilities still need to be experimentally demonstrated.

# **PHYSICOCHEMICAL PROPERTIES OF ARCHAEAL LIPIDS**

**ARCHAEAL MEMBRANE LIPID COMPOSITION – RESPONSE TO STRESS** Within the archaeal lipids there is a great diversity varying in length, composition and configuration of the side chains (**Figure 5**). The most common archaeal core lipid is *sn*-2,3 diphytanylglycerol diether, generally called archaeol, which can undergo several modifications including hydroxylation and condensation. Pioneering studies on archaeal membrane lipid composition and biosynthesis have been performed in early eighties and nineties especially in halophiles using *in vivo* labeling experiments by Kates and colleagues (Kates, 1992, 1993; Kamekura and Kates, 1999) and reviewed in detail (Koga and Morii, 2007). Among others, the studies showed that halophiles are mainly characterized by the phospholipid known as phosphatidylglycerol methyl phosphate (PGP-Me) along with sulfated and desulfated archaeols (Kates, 1993). Archaeol bearing elongated hydrocarbon chains (C20–C25) are found in some methanogens and halobacteriales (**Figure 5A**; Koga et al., 1993; Kamekura and Kates, 1999). A seemingly head-to-head condensation of two diether lipid molecules is one of the most frequent and functionally important structural variations that leads to a glycerol-dialkyglycerol tetraether lipid, known as caldarcheol. It should be stressed, however, that the enzymatic mechanism resulting in this lipid species is entirely unresolved. This core lipid is the most widespread in Archaea, characterized by different modifications depending on the archaeal species. It is in particular abundant among the phyla Euryarcheaota, Creanarchaeota, and Thaumarchaetoa. Up to 8 cyclopentane moieties can be found in lipids of the Thermoplasmatales (**Figure 5B**) and in the Euryarchaeota phylum,

in general (Macalady et al., 2004; UDA et al., 2004; Schouten et al., 2013). Interestingly, the presence of cyclopentane and cyclohexane ring is a distinct feature of Thaumarchaeota leading to a structure known as creanarcheol (**Figure 5C**; Pitcher et al., 2010; Damsté et al., 2012). In some thermoacidophiles and methanogens, a polar head group called nonitol is found which is composed of nine-carbon chain. Recent studies have revealed that the 9-carbon nonitol structure is often found as a polyhydroxylated cyclopentanic form called calditol (Koga and Morii, 2005). Therefore, these structure are now known as glycerol-dialkyl-calditol-tetraether (**Figure 5D**) and are the major component of the membrane of *Sulfolobales* species (Sugai et al., 1995; Untersteller et al., 1999; Gambacorta et al., 2002). The presence of tetraether lipids and the ratio of diether/tetraether lipids vary depending on the archaeal species and also upon growth conditions (Gambacorta et al., 1995). Likewise, there

is also a wide diversity of polar lipids in archaea including phospholipids, glycolipids, phosphoglycolipids, sulpholipids, and aminolipids (Koga and Morii, 2005). The occurrence of different polar head groups depends on the archaeal family and can be used as unique taxonomic marker (Ulrih et al., 2009). Aminolipids, for instance, are prevalent in methanogens and are completely absent in halophiles and thermophiles (Gambacorta et al., 1995).

Bacteria and Eukarya use several mechanisms to maintain the membrane fluidity over a range of temperatures such the regulation of fatty acid composition adapting the degree of branching, saturation and chain length (Zhang and Rock, 2008). The homeoviscous adaption theory states that the lipid compositions in the membrane varies in response to environmental stresses in order to preserve a proper membrane fluidity (Oger and Cario, 2013). However, the exact changes in fatty acid composition in

the membrane upon, for instance, a temperature shift differs from species to species. In *E. coli* the degree of fatty acid unsaturation increases along with a lowering of the temperature while some Bacillus species increase the amount of iso-fatty acids with the growth temperatures. The membrane also has to maintain its permeability barrier and in general it is believed that at the growth temperature, the lipid bilayer is a liquid crystalline state. The phase transition temperature at which the membrane is transferred from the crystalline into the liquid state is considered as a very important characteristic, and depends on the length of the hydrocarbon chains, the degree of saturation and the position of methyl groups; in Bacteria, the transition temperature ranges between −20 to up to 65◦C compared to archaea where the range is much wider between even up to 100◦C, a temperature where some archaea grow (Koga, 2012).

Archaea use different mechanisms to maintain the liquid crystalline phase over the entire growth temperature range. One control mechanism has been reported in the archaeon *Methanococcus jannaschii*, in which case the membrane properties are finely tuned by varying the ratio of diether, macrocyclic diether, and tetraether lipid (Sprott et al., 1991). In contrast, at higher temperatures, hyperthermophilic archaea may incorporate a higher degree of cyclopentane rings in their isoprenoid chain that increase the transition temperature. The presence of such lipid structures in archaea is an indication of a need to preserve the membrane function at the hostile environmental conditions. In particular, the presence of tetraether lipids and the chemically stable ether bonds are major adaptions (Jacquemet et al., 2009). The latter confers resistance to phospholipases attack from other organisms. Despite the general thought that the isoprenoid chains of the ether lipid are involved in thermal resistance, it has been shown that they are not absolutely required for tolerance to high temperature. Archaeol and caldarcheol were also found in the thermophilic methanoarchaeota (65◦C) and in mesophilic species (37◦C; Koga and Nakano, 2008). This suggests that the remarkable properties of the membranes of hyperthermophiles depend not exclusively on the tetraether composition of their lipids but that other aspects are involved as well. When fully stretched, the tetraether lipids span the entire membrane thickness leading to a monolayer, which is believed to stiffen the membrane in the presence of high growth temperatures (Kates, 1992; Jacquemet et al., 2009). This may also protect the membrane from possible lysis at high temperatures (Jacquemet et al., 2009). The presence of cyclic structures, in particular cyclopentene rings (De Rosa et al., 1980), is a hallmark for high growth temperatures and causes an increased membrane packing and thus a reduction in membrane fluidity (Benvegnu et al., 2008). However, also the characteristics of the polar head groups may influence membrane fluidity since the proper balance of negative and positive charges at the membrane surface is essential for its functioning. Therefore, varying the proportion of different polar head groups might be another way to response to external stresses (Oger and Cario, 2013). Furthermore, modification of the polar headgroups with carbohydrates increases hydrogen bonding between the lipids and thus will influence the stability of the membranes. In halophiles, the presence of glycerol methylphosphate attached to the archaeol moiety, contributes to the low membrane permeability under

high salt concentration (Tenchov et al., 2006). A further example is cold adaption in the psychrophile archaeon *Methanococcoides burtonii*, where an increase in the degree of unsaturation in the isoprenoid chains allow growth at low temperatures as they exist in glacial environments (Russell and Nichols, 1999; Oger and Cario, 2013).

#### *IN VITRO* **BASED STUDIES ON THE STABILITY OF ARCHAEAL LIPIDS**

Temperature impacts the membrane properties, influencing the ion permeability and phase behavior. It is believed that the unique membrane organization of archaeal tetraether lipid in a monolayer structure along with the presence of ether bonds renders such membranes, thermal resistance. Therefore, many studies have been conducted to understand the higher thermal stability of archaeal membrane lipids compared to the bacterial phospholipids. By comparing liposomes made of a polar lipid fraction from *S. acidocaldarius* and liposomes prepared from a bacterial lipid (POPC) or a synthetic lipid with a phytanyl chain (DPhPC), the importance of the methyl branched isoprenoid chain in membrane stability has been examined (Chang, 1994). When incubated at 100◦C for approximately half hour, archaeal liposomes showed a very low ion leakage compared to POPC and DPhPC liposomes that collapse after a few minutes incubation at that temperature. Likewise, only very slow release (8–10%) of the fluorophore carboxyfluorescin (CF) was observed with liposomes composed of *S. acidocaldarius* lipids compared to *E. coli* liposomes (50%) over a time interval of 62 days, while liposomes composed of a lipid extractfrom the thermophile bacterium *B. stearothermophilus* showed an intermediate stability (Elferink et al., 1994).

The extremely high heat tolerance of archaeal liposomes open opportunities for biotechnological applications with the ability to be stable even after several autoclaving cycles, which might be exploited for biomedical uses. Autoclaving is a common and effective method for decontamination and the possibility to autoclave the archaeosomes vesicle for the drug delivery gives new prospects for the liposomes formulation. Several studies have been performed regarding the ability of archaeosomes to enhance the immune response when used as novel vaccine and drug delivery system (Sprott et al.,1997; Patel and Sprott,1999; Patel et al.,2000). The superior adjuvant activity of archaeosomes compared to the liposomes evokes sufficient immunostimulation and sustained immune response against cancer or specific infectious diseases. Archaeosomes confer higher stability to the vesicles enhancing the fusion with the immune cells (Krishnan and Sprott, 2008). These properties are dependent on the type of polar head groups and degrees of glycosylation on archaeal lipids (Dicaire et al., 2010; Whitfield et al., 2010). An enhanced cytotoxic cell response was in fact observed in archaeosomes enriched with archaeatidylserine and archaeatidylethanomine due to theirfusogenic nature (Dicaire et al., 2010; Sprott et al., 2012). The effective archaeosomes stability against autoclaving was tested showing a remarkable strength against 2–3 cycles of autoclaving at pH 4.0–10.0 (Brown et al., 2009). Besides the high heat tolerance, archaeal lipids are known to be resistant to conditions of extremely low pH and their low proton permeability contributes to maintaining a constant intracellular pH. Monolayer liposomes reconstituted from the lipid fraction of

*S. acidocaldarius* indeed exhibited very low proton permeability even at higher temperatures (Elferink et al., 1994; Komatsu and Chong, 1998). However, at acidic conditions the archaeosomes appeared less resistant to autoclaving possibly because of a higher protonation of the polar head groups, influencing hydrogenbonds formation among the lipids. Liposomes containing long sugar chains linked to the phospholipids show much lower proton permeability than liposomes composed of lipids with only one sugar unit. The glycolipids amount in the polar lipid fraction of *Thermoplasma acidophilum* increases at lower pH and this seems a general mechanism for acidophiles against the chemically unstable conditions (Shimada et al., 2008). The low permeability of such liposomes can be exploited for drug-delivery with the added value of a high resistance against phospholipase A2, B and pancreatic lipase (Choquet et al., 1994). Due to their high pH tolerance they can easily pass the gastro-intestinal tract without damage. Overall, these studies confirm that the presence of tetraethers in archaeal membranes confers these membranes with a remarkable stability against high temperatures, low pH, and high salt. The degree of hydrocarbon chain saturation, the features of the polar head groups and the presence of cyclopentane ring (Dannenmuller et al., 2000) appear of secondary importance in providing stability.

### **THE LIPID DIVIDE**

#### **PROSPECT OF LUCA WITH MIXED MEMBRANE LIPID COMPOSITION**

During the last decades many theories have been proposed about the origin of life and how the differentiation between the three domains of life occurred. All of these theories accept the existence of the last universal common ancestor (LUCA), also known as LUCAS or cenancestor from which organisms have diverged. Particular attention has been given on the membrane composition of LUCA. Isoprenoids are involved in a wide range of functions, and found both in archaea and bacteria. Whereas fatty acid metabolism is also widely distributed, fatty acid biosynthesis seems underdeveloped in archaea and in some organisms even absent. Therefore, one of the hypothesis is that early life forms were dependent on the presence of membrane lipids with isoprenoid hydrocarbon core. However, the most divergent feature that is at the base of the Lipid Divide is the glycerophosphate backbone. Archaea contains G1P as glycerophosphate moiety while bacteria depend on G3P. These two compounds are synthetized by two different enzymes that are not evolutionary correlated (Koga et al., 1998).

Thus, different hypothesis were suggested to understand the process that brought the ancestor to archaeal, bacterial and eukaryotic organisms. According to Koga et al. (1998) the evolution of the two phospholipid pathways occurred independently leading to the simultaneous appearance of bacterial and archaeal enzymes. This contrast the theory of Martin and Russell (2003), according to whom the two different organisms evolved from an ancestor characterized by iron monosulphide compartment. Another hypothesis (Wächtershäuser, 2003) proposed a three stage process from the pre-cell to the eukaryotic cell. It was suggested that there was a cenancestor with a chemically derived heterochiral membrane containing both the enantiomeric forms of the glycerophosphate backbone, which slowly diverged into a more stable homochiral membrane organism leading to the differentiation

between Bacteria and Archaea. This idea implies that such heterochiral mixed membranes are intrinsically unstable leading to the emergence of chiral selective enzymes and a divide between bacteria and archaea. However, hybrid membranes were tested for their stability using a mixture of egg phosphatidylcholine and extracted lipid from *Sulfolobus solfataricus* (Fan et al., 1995) and a higher stability compared to liposomes reconstituted of only archaeal lipid was observed. Moreover a heterochiral membrane consisting of bacterial G3P lipids and archaeal G1P lipids was analyzed (Shimada and Yamagishi, 2011) and the heterochiral membranes were found to be more stable at higher temperatures compared to liposomes prepared from only the bacterial lipids. Thus, based on these studies, the hypothesis of the existence of an ancestor with both the G1PDH and G3PDH would be possible but there must have been other factors that have driven the segregation into the two different domains. The theory of the coexistence of both the enzymes for the glycerophosphate enantiomers productions can be also extended to the other enzymes involved either in the synthesis of isoprenoid and fatty acid based phospholipids (**Figure 6**). In LUCA, four different membrane lipids may be obtained by the combination of the two glycerophosphate backbones with either isoprenoid or fatty acid hydrocarbon chains (Koga,2011). Environmental pressure and the need to adapt to extreme conditions may have induced archaea to evolve their membranes (Valentine, 2007; Lombard et al., 2012b). In particular, the use of different hydrocarbon chains in response to the growth environment may have induced the segregation in Archaea and Bacteria and resulted in homochiral membrane formation (Koga, 2014).

#### **DIFFERENTIATION OF MEMBRANE LIPIDS IN ARCHAEA AND BACTERIA**

The structural variability found in the membrane lipid composition of archaea and bacteria reflects differences and similarities in the respective biosynthetic pathways. When compared with the well-characterized bacterial ester-lipid biosynthetic pathway, several similarities with the archaeal ether-lipid biosynthesis are evident.

First, the sequence of reactions that yield the final membrane lipid from the building blocks is basically the same even though some enzymes involved in these reactions are equipped with specific features to the lipid of the two different domains. Second, the glycerophosphate backbone in both cases is synthetized by a reduction of DHAP at the 2-OH using NADH as cofactor while the two hydrocarbon chains are linked to the same position on the glycerol moiety. Third, the polar head attachment takes place via a CDP-activated intermediate (Koga, 2014). In particular the peculiar features that distinguish archaeal lipids from bacterial ones occur in the first half of the biosynthetic pathway while the last stages, which involve the replacement of the CDP GROUP with one of the polar groups, is essentially similar in these two domains of life. For the latter reactions, the enzymes belong to the same superfamily and share sequence similarity (Daiyasu et al., 2005; Koga, 2011). On the other hand, the isoprenoid building blocks synthesis differs in Bacteria and in Archaea since it takes place via two different pathway, the DOXP and the alternate MVA pathway, respectively (Koga and Morii, 2007). The other striking difference involved in the lipid divide is the enantiomeric

configuration of the glycerophosphate backbone along with the saturation of the double bonds present on the isoprenoid chains and further modification of the hydrocarbon chains. The remarkable similarities of both biosynthetic pathways are an indication of the existence of a common ancestor with promiscuous enzymatic composition that sufficed for the synthesis of heterochiral membrane lipids.

# **REVERSING EVOLUTION – SYNTHESIS OF ARCHAEAL LIPIDS IN BACTERIA?**

Did a complex heterochiral membrane ever exist? There are several speculations about the membrane lipid composition of LUCA (Lombard et al., 2012b) and one evolutionary way to approach this is to design a cell which has a heterochiral membrane. The prospect of engineering *E. coli* with membranes harboring archaeal lipids has been initiated in several studies. In the first study, five genes were overexpressed in *E. coli*, four from a hyperthermophilic archaeon *Archaeoglobus fulgidus* (G1P dehydrogenase, GGPP synthase, GGGP synthase and DGGGP synthase) and one from *E. coli* (IPP isomerase; Lai et al., 2009). The enzyme IPP isomerase catalyzes the isomerization of IPP and DMAPP. Together with GGPP synthase, the carbon flux could be increased toward the synthesis of GGPP building blocks which had been demonstrated previously for carotenoid production in *E. coli* (Wang et al., 1999). The activity of each enzyme was monitored by different methods. IPP isomerase and GGPP synthase were analyzed by lycopene based

calorimetric assay, G1P dehydrogenase activity was measured spectrophotometrically to detect DHAP dependent NADH oxidation reaction and using radiolabeled substrates, GGGP synthase activity was measured by thin layer chromatography (TLC) and high performance liquid chromatography (HPLC). To detect the formation of DGGGP in *E. coli*, lipids were extracted and dephosphorylated from a 24 h growing culture and analyzed by liquid chromatography mass spectrometry (LC-MS/MS) using electron spray ionisation (ESI-MS). This study showed that indeed the archaeal lipid biosynthesis pathway is functional in *E. coli*. Unlike G1P dehydrogenase and GGGP synthase, DGGGP synthase protein could not be detected and the amount of DGGGP formed was not quantitated.

In another study, the same four genes but now from the methanogen *Methanosarcina acetivorans* were chosen as it grows at 37◦C and are readily overexpressed in *E. coli* (Yokoi et al., 2012). Activity of all the enzymes was monitored in a TLC based assay using radiolabeled substrates. The lipids were extracted from the recombinant *E. coli* cells and unlike in previous study, lipids were not dephosphorylated but analyzed directly using LC/ESI-MS. Interestingly, the result showed that DGGGP synthesized by the archaeal enzymes was further metabolized by *E. coli* endogenous enzymes to form a PG-type derivative of DGGGP which was named DGGGP-Gro. Two speculations were made regarding the endogenous enzymes that might have brought about the reaction. If it emerged from the phospholipid biosynthetic enzymes of *E. coli*, it would require three enzymes to recognize and accept the archaeal substrate, namely CdsA,*sn*-G3P transferase and the phosphatase. However, addition of CTP to their*in vitro*TLC based assay did not increase the amount of product formation and no other polar head group attachment was observed. The other possibility is that the *sn*-1-phosphoglycerol group from the osmoregulated periplasmic glucans was transferred to digeranylgeranylglycerol (DGGGOH) directly by the phosphoglyceroltransferase system. The estimated amount of total archaeal membrane lipids extracted from *E. coli* cells in this study was only 60 μg/g wet cells and at these levels it is not possible to study the influence of archaeal lipids on the physical properties of the cytoplasmic membrane. In a follow up study, GGR and ferredoxin (see section saturation of double bonds) were introduced along with the other genes from *Methanosarcina acetivorans* and expressed in *E. coli*. The formation of DGGGP-Gro was reduced yielding mostly saturated archaetidic acid in *E. coli* (Isobe et al., 2014).

The archaeal lipids have also been synthesized *in vitro* using a set of five purified enzymes, two from bacteria and three from archaea. A mutant of FPP synthase of *E. coli* was used that was shown previously to synthesize GGPP, G1P dehydrogenase was from *B. subtilis*, GGGP synthase from methanogen *Methanococcus maripaludis*, DGGGP synthase and CarS from the hyperthermophilic *Archaeoglobus fulgidus*. All enzymes were purified, and by using substrates IPP, FPP, DHAP, and NADH, the enzymes were shown to be able to synthesize CDP-archaeol in the presence of CTP, Mg2<sup>+</sup> and detergent at 37◦C (Jain et al., 2014). The feasibility of synthesizing archaeal lipids in *E. coli* and *in vitro* are promising first steps toward deciphering the biosynthetic pathway further and eventually understanding the properties of a cell with a heterochiral membrane lipid composition.

#### **FUTURE CHALLENGES**

Although during the last decade, many of the intimate features of the archaeal lipid biosynthesis pathway have been resolved, there are still several important questions that need to be answered. Understanding the mechanism of tetraether formation and identifying the enzyme(s) involved in the reaction requires thorough investigations. *In vitro* analysis of such a reaction(s) would be great advancement in the field. Various other derivatives of diether and tetraether lipids like cyclopentane and macrocyclic ring formation, glycosylation and formation of crenarchaeol are also not well understood. The pathway refractory of archaeal lipid biosynthesis in *E. coli* is currently incomplete where the amount of archaeal lipids formed in comparison to the *E. coli* lipids is very low. Challenging aspect is to modulate and suppress the endogenous pathway and integrate the archaeal lipid biosynthesis pathway in the genome of the host yielding the exclusive formation of these lipid species. Further structural and biochemical analysis of the enzymes of the pathway from different families of archaea would progress the field and bring it in par with the understanding of bacterial phospholipid biosynthesis. Also, regulation of the phospholipid metabolism is poorly understood and could be enhanced through the use of genetic studies, which now became feasible because of the rapid developments in genetic toolbox in archaea. However, to study essential genes using these techniques is still a challenge.

Recent studies have shown that in spite of the uniqueness of the archaeal membrane lipid structure, they are not as distinct as previously thought. The presence of fatty acids and isoprenoids in the three domains of life and the common mode of polar head group attachment in bacteria and archaea, and the presence of homologues of archaeal G1P dehydrogenase and GGGP synthase in bacteria are a few of the similarities. Other question that still remains unanswered is the exact reason why G1P is ether linked to the isoprenoid hydrocarbon chains and G3P is linked via ester bonds to the fatty acid chains; are there still organisms with such mixed membranes? In order to answer such questions, more biochemical and functional investigation are needed on the archaeal lipid biosynthetic pathway along with a deep phylogenetic analysis.

#### **ACKNOWLEDGMENTS**

Samta Jain and Antonella Caforio are supported by the research program of the biobased ecologically balanced sustainable industrial chemistry (BE-BASIC). We thank Ilja Kusters, John van der Oost, Melvin Siliakus and Servè Kengen for fruitful discussions.

#### **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

#### *Received: 26 September 2014; accepted: 06 November 2014; published online: 26 November 2014.*

*Citation: Jain S, Caforio A and Driessen AJM (2014) Biosynthesis of archaeal membrane ether lipids. Front. Microbiol. 5:641. doi: 10.3389/fmicb.2014.00641*

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology.*

*Copyright © 2014 Jain, Caforio and Driessen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Archaeal S-layer glycoproteins: post-translational modification in the face of extremes

# **Lina Kandiba and Jerry Eichler \***

Department of Life Sciences, Ben Gurion University of the Negev, Beersheva, Israel

#### **Edited by:**

Sonja-Verena Albers, University of Freiburg, Germany

**Reviewed by:**

Kelly Bidle, Rider University, USA Reinhard Rachel, University of Regensburg, Germany

#### **\*Correspondence:**

Jerry Eichler, Department of Life Sciences, Ben Gurion University of the Negev, PO Box 653, Beersheva 84105, Israel e-mail<EMAIL_ADDRESS>
Corresponding to the sole or basic component of the surface (S)-layer surrounding the archaeal cell in most known cases, S-layer glycoproteins are in direct contact with the harsh environments that characterize niches where Archaea can thrive. Accordingly, early work examining archaeal S-layer glycoproteins focused on identifying those properties that allow members of this group of proteins to maintain their structural integrity in the face of extremes of temperature, pH, and salinity, as well as other physical challenges. However, with expansion of the list of archaeal strains serving as model systems, as well as growth in the number of molecular tools available for the manipulation of these strains, studies on archaeal S-layer glycoproteins are currently more likely to consider the various post-translational modifications these polypeptides undergo. For instance, archaeal S-layer glycoproteins can undergo proteolytic cleavage, both N- and O-glycosylation, lipidmodification and oligomerization. In this mini-review, recent findings related to the posttranslational modification of archaeal S-layer glycoproteins are considered.

**Keywords: Archaea, lipid modification, post-translational modification, protein glycosylation, S-layer glycoprotein**

Although Archaea are now recognized as denizens of an enormous range of environments, they remain best known in their capacities as extremophiles, namely organisms able to thrive in some of the most physically challenging settings on the planet. In direct contact with these often hostile surroundings, the archaeal cell surface must not only maintain its integrity but also must carry out a variety of normal physiological functions. In Bacteria, the cell boundary consists of membranes and a peptidoglycan-based cell wall together with other polysaccharidebased molecules (e.g., lipopolysaccharide, teichoic acid) and proteins (Braun, 1975; Lugtenberg and Van Alphen, 1983; Raetz et al., 2007), in many cases comprising a surface (S)-layer (Fagan and Fairweather, 2014). By contrast, the cell wall in Archaea tends to be much simpler. Apart from a number of documented examples (König, 2001), the S-layer, in many cases comprising a single protein species but not always (Peters et al., 1995; Grogan, 1996; Veith et al., 2009), corresponds to the sole cell wall structure (Eichler, 2003; Albers and Meyer, 2011). Studies from several groups studying different Archaea have shown that the S-layer glycoprotein is not just a standardized building block used to generate the two-dimensional lattice of the S-layer but rather that S-layer glycoproteins undergo a variety of posttranslational modifications. In this mini-review, recent findings concerning such processing of archaeal S-layer glycoproteins are considered.

#### **DIFFERENCES IN THE SUGAR COATING**

The S-layer glycoprotein of the haloarchaeon *Halobacterium salinarum* offered the first example of *N*-glycosylation in a domain other than the Eukarya (Mescher and Strominger, 1976a). This observation led to a flurry of biochemical activity aimed at describing the composition of *N*-linked glycans decorating the *Hbt. salinarum* S-layer glycoprotein and their biosynthesis (cf. Lechner and Wieland, 1989). However, the lack of sufficient genetic tools for manipulating this and other archaeal species shown to contain glycosylated S-layer proteins (Sumper et al., 1990; Brockl et al., 1991; Karcher et al., 1993) stood in the way of gaining detailed information into such post-translational modification of this protein. Since then, the sequencing of a growing list of archaeal genomes, the development of techniques for manipulating the genetic content of numerous strains and the analytical power of mass spectrometry have been combined to help clear obstacles encountered by earlier studies of S-layer glycoprotein *N*-glycosylation.

Genomic analyses point both to the presence of S-layer glycoproteins and *N*-glycosylation machineries in almost all sequenced Archaea (Magidovich and Eichler, 2009; Albers and Meyer, 2011; Kaminski et al., 2013a). Still, the majority of research on archaeal S-layer glycoprotein *N*-glycosylation to date has focused on *Methanococcus voltae*, *Methanococcus maripaludis*, *Sulfolobus acidocaldarius*, and *Haloferax volcanii* (for recent review, see Jarrell et al., 2014). In each of these species, genes involved in the assembly and attachment of N-linked glycans and often their protein products have been studied. Yet, apart from *S. acidocaldarius*, where *N*-glycosylation is essential for cell survival (Meyer and Albers, 2014), the elimination of such protein processing seemingly has limited impact on the organism (Abu-Qarn and Eichler, 2006; Chaban et al., 2006; VanDyke et al., 2009). As such, one can ask why Archaea devote such a significant number of genes to this post-translational modification. Recent studies on *Hfx. volcanii* S-layer glycoprotein have begun to shed light on this point.

The *Hfx. volcanii* S-layer glycoprotein contains seven putative *N*-glycosylation sites (Sumper et al., 1990).  However, when *Hfx. volcanii* cells are grown in medium containing 1.75 M NaCl ("low salt" conditions) rather than 3.4 M NaCl ("high salt" conditions), Asn-498 is modified by a distinct glycan comprising a sulfated hexose, two hexoses and a rhamnose (Guan et al., 2012). Indeed, the same glycan had been reported earlier as bound to dolichol phosphate in *Hfx. volcanii* grown in the presence of 1.25 M NaCl (Kuntz et al., 1997), the lipid carrier that serves as the platform for *N*-glycan assembly in this and other Archaea (Lechner et al., 1985; Guan et al., 2010; Calo et al., 2011). As such, it would appear that the *Hfx. volcanii* S-layer glycoprotein undergoes differential *N*-glycosylation as a function of environmental salinity. While it remains to be defined how such differential S-layer glycoprotein *N*-glycosylation translates into an appropriate response to changes in surrounding salt levels, the path involved in the biogenesis of the so-called "low-salt" tetrasaccharide has been revealed (Kaminski et al., 2013b). Unexpectedly, the cluster of genes involved does not include an obvious oligosaccharyltransferase, namely that enzyme responsible for transferring a glycan from its lipid carrier to select Asn residues of target proteins (Mohorko et al., 2011). The observation that AglB, the only known archaeal oligosaccharyltransferase (Abu-Qarn and Eichler, 2006; Chaban et al., 2006), is not involved in "low-salt" tetrasaccharide attachment implies the existence of a novel yet undefined enzyme as serving this role (Kaminski et al., 2013b).

It is possible that *N*-glycosylation of the *Hfx. volcanii* S-layer glycoprotein is even more complicated still. It was recently reported that the Asn-732 position is modified by a sulfoquinovose-hexose-based glycan, *N*-linked via a chitobiose core (Parente et al., 2014). Moreover, the composition of this glycan was modified in response to the absence or presence of a membrane-localized rhomboid protease. The presence of such a glycan in *Hfx. volcanii* is surprising, given this organism does not contain a homolog of *S. acidocaldarius* Agl3 (Meyer et al., 2011), a UDP-sulfoquinovose synthase responsible for converting UDP-glucose and sodium sulfite into UDP-sulfoquinovose, the activated form of this sugar that is presumably used in *S. acidocaldarius* and presumably *Hfx. volcanii N*-glycosylation. It should also be noted that Asn-732 is found in the same Cterminal region as a cluster of *O*-glycosylated threonine residues (Sumper et al., 1990) and a lipid anchor (see below). This suggests that post-translational modification of the *Hfx. volcanii* S-layer glycoprotein C-terminal region is a complex event that requires the orchestrated involvement of numerous protein processing pathways.

Unlike *Hfx. volcanii*, which must cope with an environment characterized by molar concentrations of salt, *S. acidocaldarius* is a thermophile that grows optimally at 75–80◦C and pH 2– 3 (Brock et al., 1972). Possibly due to the challenges presented by its surroundings, not only is *N*-glycosylation essential in *S. acidocaldarius* (Meyer and Albers, 2014) but at least one of the glycoproteins comprising the S-layer in this species (SlaA) presents an extremely high number of *N*-glycosylation sites. The 1,395 amino acid-long protein contains 31 potential sites for *N*-glycosylation scattered throughout the polypeptide, translating to an *N*-glycosylation site every 45 residues on average, with the highest densely of such sites being seen in the Cterminal quarter of the protein (Peyfoon et al., 2010). In the region spanning Lys-1004 to Gln-1395, nine of the 11 potential *N*-glycosylation sites were experimentally verified as being charged with a tri-branched hexasaccharide comprising a glucose, a mannose, two *N*-acetylglucosamines and a sulfoquinovose, an unusual sugar routinely found in chloroplasts and photosynthetic bacteria (Zahringer et al., 2000; Peyfoon et al., 2010). Indeed, a tallying of the number of putative *N*-glycosylation sites in 20 different archaeal S-layer glycoproteins reveals that the S-layer glycoproteins of thermo(acido)philes can contain up to 20-fold more such sites than do S-layer glycoproteins in species isolated from other growth conditions (Jarrell et al., 2014). Based on this comparison, it was proposed that such high densities of *N*glycosylation sites reflect the need for a rigid and stable cell wall to cope with the challenges of elevated temperatures and acidity encountered by thermo(acido)philic Archaea.