(Table S5). Members of this family typically repress transcription from their own promotor, but activate transcription from a divergent promotor. The products of these activated genes perform a wide range of functions [130] while their activation is mediated by co-inducers, such as extracellular signaling molecules, metabolites, or ions. Other important one-component systems in C. metallidurans were GntR (n = 43), TetR (n = 35), AraC (n = 25), MarR (n = 16), and IclR (n = 15) (Table S5), all of which contribute to the transcriptional control of stress responses, carbon metabolism, catabolic pathways, multidrug efflux pumps, biosynthesis of antibiotics, response to osmotic stress and toxic chemicals, differentiation processes, and virulence and pathogenicity.

In terms of one-component metalloregulatory proteins in CH34, three families MerR (n = 10), ArsR (n = 6), and Fur (n = 2) deserve our attention. Although these regulators are evolutionary related in their DNA binding properties, their mode of action is different [131]. While MerR-type regulators nearly exclusively function as transcriptional activators, ArsR-type regulators generally act as repressors that upon metal binding dissociate from the DNA giving way to the RNA polymerase (coined as transcriptional derepression). Fur proteins although being typical repressors can also activate gene expression, either directly, or indirectly via the Fur-dependent repression of a small regulatory antisense RNA. Importantly, there is a wide range in metal selectivity within these three families. Metal sensing in MerR regulators is governed by the effector binding domain in their C-terminal region. Structural diversity in this domain allows MerR family proteins to sense not only various metal ions, e.g., Zn²⁺ by ZntR, Au⁺ by GolS, Cd²⁺ by CadR, Pb²⁺ by PbrR, or Hg²⁺ by MerR, but also oxidative stress, e.g., by SoxR via an [2Fe-2S]²⁺ cluster (reviewed in [131]). Preferential sensing by MerR regulators of one metal ion over another, or monovalent ions over divalent ions, is defined by the positioning of Cys and His but also other conserved residues, mainly in the DNA binding loop. The stringency of this preference probably varies among MerR regulators as some respond to more than one metal. In strain CH34 there are four MerR proteins and three PbrR proteins encoded by the *mer* and *pbr* loci (regulators are underlined in Table 5), and one CupR (Rmet_3523) which responds to Cu⁺ and Au⁺ [35]. The remaining MerR-type regulators in CH34, including SoxR (Rmet_4538), are not physically linked to any structural HMR gene.

The metal sensing in ArsR regulators in contrast is not defined by a single metal binding pocket but is distributed over distinct locations of the repressor involving a number of alpha helices containing key Cys or His residues [131]. Members of this family sense, next to As(III) and Sb(III), a wide variety of divalent metal ions including Zn²⁺, Ni²⁺, Co²⁺, Pb²⁺, and Cd²⁺. Some ArsR family members display sensitivity to multiple metal ions and are able to respond to either monovalent or multivalent ions, probably as a result of evolutionary adaptations in their protein structure particularly in the α3N and α5 helical regions. The CH34 genome enlists only one bona fide ArsR regulator (Rmet_0333 of the *ars* locus on CHR1; Table 5). This repressor has been shown to dissociate from its operator DNA in a metal and valence qualifying manner [94], with three groups of heavy metals displaying decreasing degrees of ArsR dissociation: As(III)/Bi(III)/Co²⁺/Cu²⁺/Ni²⁺; Cd²⁺; and Pb²⁺/Zn²⁺. No ArsR dissociation was observed in the presence of As(V). The other five ArsR family proteins of CH34 appear to be unlinked to any HMR gene cluster except Rmet_5676 which is located adjacent to the defective *nim* locus on CHR2 (Table 5). The two Fur proteins in CH34 are discussed in the next section on siderophores. Besides the MerR, ArsR, and Fur families, no other structural families of one-

component metal responsive regulators are currently recognized in the CH34 genome. Although also LysR, TetR, and MarR families reportedly contain members that are directly involved in metal homeostasis or metal-regulated responses [131], only very few well-documented cases are known sofar. Nonetheless, these three families are represented in CH34 by a total of a few hundred proteins so it may be worthwhile to look also within this pool for further candidate metal response regulators.

Among two-component systems the OmpR system constituted the largest group (n = 33) while the LuxR family (best known for its quorum sensing regulators) also was well represented (n = 41) (Table S5). According to the MiST2 database [132], there are 56 two-component pairs of adjacent genes in the CH34 genome. Several of those have been implicated in HMR, notably two pairs of copRS and czcRS, and one pair each of agtRS, hmzRS, zniRS, and zneRS (underlined in Table 5). From MaGe data and the DBD and MiST2 databases we discovered that the second replicon of C. metallidurans (CHR2) carries proportionally almost twice the number of regulatory genes as compared to the main replicon CHR1. This enrichment in regulatory genes agrees well with the earlier notion that CHR2 carries many functions relevant to the survival and responsive behavior of C. metallidurans CH34 (Figure 2).

Clearly, the wide diversity of sigma factors and transcriptional regulators in C. metallidurans CH34 points to intricate regulatory mechanisms for activating genes in response to environmental stimuli. For instance, microarray analyses of genome-wide gene expression studies on strain CH34 pointed to layered multiple responses when cells are challenged with heavy metals [10]. Similar complex expression patterns were shown for CH34 cells grown under other harsh conditions, including simulated microgravity, growth on basalt, and spaceflight (data not shown). To unravel these regulatory mechanisms and better understand the molecular basis of C. metallidurans' ability to survive prolonged periods of stress, we now are focussing on reconstructing the regulatory networks by systematically identifying regulatory motifs in the upstream regions of stress-induced and co-regulated genes.

## Siderophores

At neutral pH and oxic conditions, bacteria cannot readily utilize low-solubility Fe(III) minerals. Accordingly, they synthesise and secrete soluble iron-complexing compounds, called siderophores. C. metallidurans CH34 grown under iron-limiting conditions produces a unique phenolate-type siderophore containing neither hydroxamate nor catecholate groups [133]. This siderophore, originally referred to as “alcaligen E”, eventually was classified as a polycarboxylate staphyloferrin B [134] because it resembled the siderophores typically produced by staphylococci. It since was recognized that other organisms produce staphyloferrin B, such as R. solanacearum [135]. Notably, C. metallidurans staphyloferrin B can interact with Cd²⁺ thereby decreasing the metal's bioavailability and hence lowering its cellular toxicity in the bacterium. Also, siderophore-producing C. metallidurans tested in bioaugmentation-assisted phytoextraction increased, by a factor of 5, the accumulation of Cr(VI) in plant shoots [136]. Future studies on siderophore production in C. metallidurans CH34 might thus open up exciting lines of research in heavy metal tolerance and bioremediation.

Genes for staphyloferrin B biosynthesis (Rmet_1109-1118) and two genes encoding ferric/ferrichrome siderophore TonB-dependent receptors BfrC and FcuA (Rmet_1104 and Rmet_1108, respectively) are located on CHR1 within a 52 kb region that lacks synteny with Cpin, Ceut, and Cai. Although a tRNA-Ser gene precedes this region, we could not identify any tyrosine-based site-

<figure><img src="image_21.png" /></figure>
specific recombinases, flanking IS elements, or direct repeats; hence, we did not classify it as a genomic island (see our discussion on Mobile Genetic Elements). It includes genes encoding a siderophore synthase decarboxylase (ssd, Rmet_1110), an aldolase (Rmet_1111), staphyloferrin B biosynthesis proteins (Rmet_1112, Rmet_1113, and Rmet_1115), a permease of the Major Facilitator Superfamily (MSF) (Rmet_1114), an ornithine cyclodeaminase (ocd, Rmet_1116), needed to biosynthesize alcaligin [137], a cysteine synthase (cysK, Rmet_1117), and the staphyloferrin B receptor (aleB, Rmet_1118). This core set of genes (Rmet_1110 to Rmet_1117) displays high synteny with the staphyloferrin B biosynthesis regions of *Ralstonia solanacearum* GMI1000 and *Staphylococcus aureus* MW2 [135] and of all other *S. aureus* strains present in the MaGe database. In addition, *R. solanacearum* GMI1000 and *C. metallidurans* CH34 share the *aleB* gene (Rmet_1118) and a gene for the sigma-19 transcription factor FecI (RpoI; Rmet_1120). The insertion of a mini-transposon Tn5 into *aleB* decreased iron uptake three to four times in iron-starved cells compared to the wild type strain CH34 [133]. The *C. metallidurans* CH34 genome lacks the *entA*, *entB*, *entC*, *entD*, *entF*, and *entS* genes required for enterobactin biosynthesis but seems to contain an *entE* gene (Rmet_3777) on CHR2.

The regulation of staphyloferrin B biosynthesis is controlled by two systems: A. *fecRI* regulon located immediately upstream of *aleB*, and the *fur* gene distant from *aleB*. The regulator *FecR* (Rmet_1119) is a sensor for the presence of iron dicitrate in the periplasm and regulates its transport by inactivating *FecI* (Rmet_1120). The Fur protein (for ferric uptake regulation) tightly represses staphyloferrin B biosynthesis under iron-replete conditions [138]. In both *C. metallidurans* and *R. solanacearum*, a gene (Rmet_2977) encoding the global virulence regulator *PhcA* is located next to the *fur* gene (Rmet_2976). In *R. solanacearum*, *PhcA* negatively regulates staphyloferrin B biosynthesis [135]. Furthermore, the *phcA* gene of *C. metallidurans* CH34 can complement a *phcA*⁻ mutant of *R. solanacearum* [139]. Interestingly, two Fur-like regulators, Rmet_0128 and Rmet_5746, respectively, were found on CHR1 and CHR2.

Besides the *aleB* locus, the genome of *C. metallidurans* CH34 contains additional genes that are probably involved in binding siderophores. These are Rmet_0837 and Rmet_4617 encoding hydroxamate-type ferrisiderophore receptors, Rmet_4496 and Rmet_4497 encoding ferric siderophore receptors, Rmet_5806 and Rmet_5807 encoding *FecA*-type ferric citrate outer membrane receptors, a small locus Rmet_2277/8/9 encoding a complete TonB-ExbB-ExbD type energy-transducing complex. Supplementary genes encode a TonB-like protein (Rmet_3055) and ExbB/ExbD homologues (Rmet_0536/7). The presence of multiple siderophore binding and translocation systems is consistent with the observation that *C. metallidurans* CH34 employs various ferric ion chelators for iron uptake, including desferriferrioxamine B, D2 and E, desferriferricrocin, desferriferrichrysin, desferriferrichrome A, and ornibactin [133]. In addition, its genome encodes proteins involved in iron storage (*bfr*, Rmet_0195, Rmet_0248) and soluble ferrous Fe(II) transport (*feoB*, Rmet_5890 and Rmet_5890). Table S6 offers an overview of CH34 genes implemented in iron uptake and -metabolism.

Signal transduction

Like many other environmental bacteria, *C. metallidurans* CH34
deploys early warning systems and response mechanisms to cope
with many different processes such as growth and development,
metabolic regulation, and sensing, and to react swiftly to various
stresses. In bacteria, the linkage of input signals to output responses
entails a cascade of protein modifications, basically of two modes:

One-component and two-component systems (denominated as
1CST and 2CST, respectively). In 1CST systems, the input- and
output-domains are fused in a single protein, whereas in 2CST
systems, the sensory- and receiver-domains lie on two different
proteins (reviewed by Galperin and Gomelsky [140]). The latter
has the advantage that the sensory component can be membrane-
associated to receive *extracellular* stimuli while the receiver domain
can be part of the actual regulator, often acting at the level of
transcription. Until recently, 2CST systems were thought to be the
primary mode of signal transduction in prokaryotes, but it became
evident that 1CST systems carry out much of the signal
transduction [128]. The input and output domains of these
systems display a remarkable diversity, are widely distributed
across species, and are mainly involved in DNA binding and the
intracellular detection of environmental stimuli [128]. To survey
signal transduction proteins in *C. metallidurans* and related
organisms we used the *Microbial Transduction System* (MiST)
database [141] (http://genomics.ornl.gov/mist/) that holds the
predicted signal transduction proteins for nearly 750 bacterial and
archaeal organisms, using Pfam and SMART reference domains
(added note: an extended database MiST2 was very recently
established at http://mistdb.com/ [132]). These data demonstrat-
ed that all four *Cupriavidus* species contained approximately the
same proportional number of proteins putatively involved in signal
transduction (approx. 9–10% of their proteome) (Table S7). The
same proportion was evident in *D. acidovorans*, *Janthinobacterium*, and
*B. xenovorans*, while *B. petrii*, *R. solanacearum*, and *H. arsenicoxydans*
displayed slightly lower values (8.3-, 8.2- and 7.0%, respectively).
In contrast, and in line with its deterministic lifestyle, the
intracellular symbiont *P. necessarius* STIR1, whose single replicon
displays good synteny with *C. metallidurans* CHR1 (see section on
Phylogeny), deploys only 1.6% of its protein complement for signal
transduction. For all the organisms we compared, there were two-
to three-fold more predicted proteins involved in 1CST systems
than in 2CST systems (Table S7) (the only exception being *B. petrii*
that has a slightly higher preference for 1CST systems). Also, this
comparison demonstrated that the second replicon of all
organisms with a multipartite genome (i.e. the four *Cupriavidus*
species, *R. solanacearum*, and *B. xenovorans*) displayed an enrichment
on their second replicon for proteins involved in signal
transduction (data not shown), once more underpinning the
importance of this replicon for adaptation and survival.

According to MiST data, protein domains commonly used for
signal transduction in *C. metallidurans* CH34 were Bug (123),
LysR_substrate (122), Response_reg (98), HATPase_c (72), HisKA
(55), HAMP (45), TetR_N (36), GGDEF (30), PAS (30), FCD (24),
EAL (23), MCPsignal (16), Sigma54_activation (16), and GAF (10)
(domain names are in Pfam notation with numbers in para-
ntheses). Most of these domains are integrated as signal receivers
into transcriptional regulators (e.g. LysR_substrate, TetR_N,
FCD) whether in 2CST or 1CST format, while others have more
specific functions suggesting a role in chemotaxis (e.g. Respon-
se_reg, MCPsignal), His-Asp phosphorelay (HisKA, HAMP,
HATPase_c), cyclic di-GMP turnover (GGDEF, EAL, GAF), or
solute binding (e.g. Bug). Undoubtedly, many output domains
constitute DNA binding motifs for the transcriptional regulation of
genes. These domains typically have helix-turn-helix (e.g. HTH_1,
IclR) or winged helix (e.g. Trans_reg_C, MarR) structures (Table
S7, not all data shown).

In general, the distribution of signal transduction domains is the
same across the four *Cupriavidus* species (Table S7) and is not very
different from their distributions in the other free living species.
However, we noted an important deviation for the Bug domain
that occurs at high frequency (>100) for all four *Cupriavidus* species

<figure><img src="image_22.png" /></figure>
specific recombinases, flanking IS elements, or direct repeats; hence, we did not classify it as a genomic island (see our discussion on Mobile Genetic Elements). It includes genes encoding a siderophore synthase decarboxylase (ssd, Rmet_1110), an aldolase (Rmet_1111), staphyloferrin B biosynthesis proteins (Rmet_1112, Rmet_1113, and Rmet_1115), a permease of the Major Facilitator Superfamily (MSF) (Rmet_1114), an ornithine cyclodeaminase (ocd, Rmet_1116), needed to biosynthesize alcaligin [137], a cysteine synthase (cysK, Rmet_1117), and the staphyloferrin B receptor (aleB, Rmet_1118). This core set of genes (Rmet_1110 to Rmet_1117) displays high synteny with the staphyloferrin B biosynthesis regions of *Ralstonia solanacearum* GMI1000 and *Staphylococcus aureus* MW2 [135] and of all other *S. aureus* strains present in the MaGe database. In addition, *R. solanacearum* GMI1000 and *C. metallidurans* CH34 share the *aleB* gene (Rmet_1118) and a gene for the sigma-19 transcription factor FecI (RpoI; Rmet_1120). The insertion of a mini-transposon Tn5 into *aleB* decreased iron uptake three to four times in iron-starved cells compared to the wild type strain CH34 [133]. The *C. metallidurans* CH34 genome lacks the *entA*, *entB*, *entC*, *entD*, *entF*, and *entS* genes required for enterobactin biosynthesis but seems to contain an *entE* gene (Rmet_3777) on CHR2.

The regulation of staphyloferrin B biosynthesis is controlled by two systems: A. *fecRI* regulon located immediately upstream of *aleB*, and the *fur* gene distant from *aleB*. The regulator *FecR* (Rmet_1119) is a sensor for the presence of iron dicitrate in the periplasm and regulates its transport by inactivating *FecI* (Rmet_1120). The Fur protein (for ferric uptake regulation) tightly represses staphyloferrin B biosynthesis under iron-replete conditions [138]. In both *C. metallidurans* and *R. solanacearum*, a gene (Rmet_2977) encoding the global virulence regulator *PhcA* is located next to the *fur* gene (Rmet_2976). In *R. solanacearum*, *PhcA* negatively regulates staphyloferrin B biosynthesis [135]. Furthermore, the *phcA* gene of *C. metallidurans* CH34 can complement a *phcA*⁻ mutant of *R. solanacearum* [139]. Interestingly, two Fur-like regulators, Rmet_0128 and Rmet_5746, respectively, were found on CHR1 and CHR2.

Besides the *aleB* locus, the genome of *C. metallidurans* CH34 contains additional genes that are probably involved in binding siderophores. These are Rmet_0837 and Rmet_4617 encoding hydroxamate-type ferrisiderophore receptors, Rmet_4496 and Rmet_4497 encoding ferric siderophore receptors, Rmet_5806 and Rmet_5807 encoding *FecA*-type ferric citrate outer membrane receptors, a small locus Rmet_2277/8/9 encoding a complete TonB-ExbB-ExbD type energy-transducing complex. Supplementary genes encode a TonB-like protein (Rmet_3055) and ExbB/ExbD homologues (Rmet_0536/7). The presence of multiple siderophore binding and translocation systems is consistent with the observation that *C. metallidurans* CH34 employs various ferric ion chelators for iron uptake, including desferriferrioxamine B, D2 and E, desferriferricrocin, desferriferrichrysin, desferriferrichrome A, and ornibactin [133]. In addition, its genome encodes proteins involved in iron storage (*bfr*, Rmet_0195, Rmet_0248) and soluble ferrous Fe(II) transport (*feoB*, Rmet_5890 and Rmet_5890). Table S6 offers an overview of CH34 genes implemented in iron uptake and -metabolism.

Signal transduction

Like many other environmental bacteria, *C. metallidurans* CH34
deploys early warning systems and response mechanisms to cope
with many different processes such as growth and development,
metabolic regulation, and sensing, and to react swiftly to various
stresses. In bacteria, the linkage of input signals to output responses
entails a cascade of protein modifications, basically of two modes:

One-component and two-component systems (denominated as
1CST and 2CST, respectively). In 1CST systems, the input- and
output-domains are fused in a single protein, whereas in 2CST
systems, the sensory- and receiver-domains lie on two different
proteins (reviewed by Galperin and Gomelsky [140]). The latter
has the advantage that the sensory component can be membrane-
associated to receive *extracellular* stimuli while the receiver domain
can be part of the actual regulator, often acting at the level of
transcription. Until recently, 2CST systems were thought to be the
primary mode of signal transduction in prokaryotes, but it became
evident that 1CST systems carry out much of the signal
transduction [128]. The input and output domains of these
systems display a remarkable diversity, are widely distributed
across species, and are mainly involved in DNA binding and the
intracellular detection of environmental stimuli [128]. To survey
signal transduction proteins in *C. metallidurans* and related
organisms we used the *Microbial Transduction System* (MiST)
database [141] (http://genomics.ornl.gov/mist/) that holds the
predicted signal transduction proteins for nearly 750 bacterial and
archaeal organisms, using Pfam and SMART reference domains
(added note: an extended database MiST2 was very recently
established at http://mistdb.com/ [132]). These data demonstrat-
ed that all four *Cupriavidus* species contained approximately the
same proportional number of proteins putatively involved in signal
transduction (approx. 9–10% of their proteome) (Table S7). The
same proportion was evident in *D. acidovorans*, *Janthinobacterium*, and
*B. xenovorans*, while *B. petrii*, *R. solanacearum*, and *H. arsenicoxydans*
displayed slightly lower values (8.3-, 8.2- and 7.0%, respectively).
In contrast, and in line with its deterministic lifestyle, the
intracellular symbiont *P. necessarius* STIR1, whose single replicon
displays good synteny with *C. metallidurans* CHR1 (see section on
Phylogeny), deploys only 1.6% of its protein complement for signal
transduction. For all the organisms we compared, there were two-
to three-fold more predicted proteins involved in 1CST systems
than in 2CST systems (Table S7) (the only exception being *B. petrii*
that has a slightly higher preference for 1CST systems). Also, this
comparison demonstrated that the second replicon of all
organisms with a multipartite genome (i.e. the four *Cupriavidus*
species, *R. solanacearum*, and *B. xenovorans*) displayed an enrichment
on their second replicon for proteins involved in signal
transduction (data not shown), once more underpinning the
importance of this replicon for adaptation and survival.

According to MiST data, protein domains commonly used for
signal transduction in *C. metallidurans* CH34 were Bug (123),
LysR_substrate (122), Response_reg (98), HATPase_c (72), HisKA
(55), HAMP (45), TetR_N (36), GGDEF (30), PAS (30), FCD (24),
EAL (23), MCPsignal (16), Sigma54_activation (16), and GAF (10)
(domain names are in Pfam notation with numbers in para-
ntheses). Most of these domains are integrated as signal receivers
into transcriptional regulators (e.g. LysR_substrate, TetR_N,
FCD) whether in 2CST or 1CST format, while others have more
specific functions suggesting a role in chemotaxis (e.g. Respon-
se_reg, MCPsignal), His-Asp phosphorelay (HisKA, HAMP,
HATPase_c), cyclic di-GMP turnover (GGDEF, EAL, GAF), or
solute binding (e.g. Bug). Undoubtedly, many output domains
constitute DNA binding motifs for the transcriptional regulation of
genes. These domains typically have helix-turn-helix (e.g. HTH_1,
IclR) or winged helix (e.g. Trans_reg_C, MarR) structures (Table
S7, not all data shown).

In general, the distribution of signal transduction domains is the
same across the four *Cupriavidus* species (Table S7) and is not very
different from their distributions in the other free living species.
However, we noted an important deviation for the Bug domain
that occurs at high frequency (>100) for all four *Cupriavidus* species

<figure><img src="image_22.png" /></figure>
but is far less prevalent in either *R. solanacearum* or *B. xenovorans* (respectively, 2 and 13 Bug domains). The Bug domain, named after the *Bordetella pertussis bug* gene identified by Antoine *et al.* [142], was reported as the first member of a large family of periplasmic solute-binding receptors present in in great numbers (up to 181 per genome) in *Bordetella* species, with much lower numbers (1 to 11) in some other bacterial genomes [47]. The only exception in their study was *C. metallidurans* for which 102 Bug homologs were predicted. Typically, Bug proteins are highly conserved in size (median size 325 residues), contain an identifiable signal-sequence, and share well-conserved carboxylate-binding motifs that form a characteristic Venus-fly-trap fold [143,144]. This latter characteristic and the fact that the family member TctC, a periplasmic citrate-binding protein of *Salmonella typhimurium*, is part of a novel transport-system TctABC, led researchers to classify many members of this family to the tripartite tricarboxylate transporter (TTT) family (listed as TC 2.A.80 in the Transport Classification Database; http://www.tcdb.org). Nonetheless, although Bug proteins clearly form a distinct family of solute receptors, their sequences have diverged widely and their genes often are not linked to transport-related operons. Antoine and co-workers [47,145] posited that these bug genes may be used by other transport systems, or involved in other metabolic activities or other pathways activated under stress. In this context it is interesting to note that many of the bug genes in CH34 seem linked or are in the close vicinity of transporter and regulatory genes. We also find it intriguing that *C. metallidurans*, and the other *Cupriavidus* species, harbour so many bug genes: Out of 200 genomes analysed by Antoine *et al.* [47], only *Bordetella* species and *C. metallidurans* were listed with a prodigious number of bug genes. We might argue that *Cupriavidus* species are free-living organisms wherein the use of solute receptors for specialised strategies of adaptation and survival evolved, but then, we question why so few other free-living organisms failed to develop this approach? We surveyed for the Bug domain in the 1,216 complete and draft, bacterial and archaeal genomes currently in the MiST2 database [132] and found 2,377 proteins with Bug domains. Outside the genera *Bordetella* and *Cupriavidus* only four genomes had a comparable count: *Delftia acidovorans* SPH-1 (159), *Variovorax paradoxus S110* (137), *Verminophrobacter eiseniae* EF01-2 (131), and *Polaromonas* sp. JS666 (97). The only other genomes with a large count (>40) were *Diaphorobacter* sp. TPSY (45), *Acidovorax avenae* ssp. *citrulli* (55), *Acidovorax* JS42 (46), and *Polaromonas naphtalenivorans* CJ2 (41). To our surprise, all organisms that showed an overrepresentation of bug genes belonged to the same beta-proteobacterial order of *Burkholderiales* (Figure 5; Table S8). We propose to call this group of prokaryotes the “Bug Receptor Bacteria” or BuRBs. Such overrepresentation may be a recent innovation in bacteria, or may be more useful to those with a particular lifestyle and habitat. Whether the propensity to utilise large numbers of Bug-domain containing receptors arose within the genus *Bordetella* and was radiated out to closely related genera, or whether it has originated elsewhere within the order of *Burkholderiales* is an open question. It is useful here to mention that *B. petrii*, which carries 107 bug genes, is not host-restricted like the other *Bordetella* species. It is an environmental species typically obtained from river sediment, grass root, and polluted soil [146,147] and has the capacity to degrade aromatic compounds and detoxify heavy metals [148]. However, *B. petrii* strains subsequently were isolated from patients with mandibular osteomyelitis and chronic suppurative mastoiditis [149,150]; thus, this species appears to be an opportunistic pathogen after all. It also contains many mobile genetic elements [148], some strikingly resembling those found in CH34 [59], that may have played an intermediate role in the evolution of BuRBs

## Beta-proteobacteria

Burkholderiales

Alcaligenaceae

*Bordetella*

*B. bronchiseptica* RB50 (183)

*B. parapertussis* 12822 (140)

*B. pertussis* Tohama I (79)

*B. petrii* DSM 12804 (107)

*B. avium* 197N (26)

Burkholderiaceae

Burkholderia

*B. xenovorans* LB400 (13)

Ralstonia

*R. solanacearum* GMI1000 (2)

Polymucleobacter

*P. necessarius* subsp. asymbioticus QLW (16)

*P. necessarius* subsp. necessarius STIR1 (8)

Cupriavidus

*C. metallidurans* CH34 (123)

*C. eutropha* H16 (155)

*C. pinatubonensis* JMP134 (131)

*C. taiwanensis* (103)

Comamonadaceae

Acidovorax

*A. citrulli* AAC00-1 (55)

*Acidovirax* sp. JS42 (46)

Delftia

*D. acidovorans* (159)

Diaphorobacter

*Diaphorobacter* sp. TPSY (45)

Polaromonas

*Polaromonas* sp. JS666 (97)

*P. naphtalenivorans* CJ2 (41)

Variovorax

*V. paradoxus* S110 (137)

Verminephrobacter

*V. eisiniae* EF01-2 (131)

Oxalobacteraceae

Herminiimonas

Herminiimonas arsenicoxydans (5)

Janthinobacterium

Janthinobacterium sp. Marseille (10)

**Figure 5. Distribution of the Bug domain.** Overrepresentation of 'Bug' domains in the family *Burkholderiales* according to MIST data (http://mistdb.com/), with numbers in parentheses. Organisms encoding more than 40 Bug domain-containing proteins are in bold. Only relevant taxa and species are shown (see text). Detailed data are in Table S8.

doi:10.1371/journal.pone.0010433.g005

and the spread of the Bug domain. We speculate that BuRBs evolved many Bug-domain proteins as periplasmic receptors to expand their cellular capacity to bind and transport, via the Venus fly trap fold, a wide range of environmental carboxylated compounds. They might include aminoacids (glutamate, aspartate), other forms of C-skeletons useful for growth (oxalate, malate, succinate, lactate, citrate), humic-acid derivatives from soil, metal-complex carboxylated acids in polluted environments, or plant-root-exuded carboxylates naturally accumulating in the rhizosphere.

Another unusual feature of signal transduction in the *C. metallidurans* CH34 genome is the presence of an intact phosphorelay two-component system entailing a multidomain histidine-kinase sensor protein BvgS1 (Rmet_5710) and a response-transcriptional activator BvgA (Rmet_5714). This system BvgSA controls, via the transduction of environmental signals, the expression of numerous virulence- and colonization-factors in pathogenic *Bordetella* species [50,151] and resembles, respectively, the EvgSA and RocSA systems in *E. coli* and *P. aeruginosa*. *E. coli* contains at least 37 EvgA-activated genes, including stationary-

<figure><img src="image_23.png" /></figure>
but is far less prevalent in either *R. solanacearum* or *B. xenovorans* (respectively, 2 and 13 Bug domains). The Bug domain, named after the *Bordetella pertussis bug* gene identified by Antoine *et al.* [142], was reported as the first member of a large family of periplasmic solute-binding receptors present in in great numbers (up to 181 per genome) in *Bordetella* species, with much lower numbers (1 to 11) in some other bacterial genomes [47]. The only exception in their study was *C. metallidurans* for which 102 Bug homologs were predicted. Typically, Bug proteins are highly conserved in size (median size 325 residues), contain an identifiable signal-sequence, and share well-conserved carboxylate-binding motifs that form a characteristic Venus-fly-trap fold [143,144]. This latter characteristic and the fact that the family member TctC, a periplasmic citrate-binding protein of *Salmonella typhimurium*, is part of a novel transport-system TctABC, led researchers to classify many members of this family to the tripartite tricarboxylate transporter (TTT) family (listed as TC 2.A.80 in the Transport Classification Database; http://www.tcdb.org). Nonetheless, although Bug proteins clearly form a distinct family of solute receptors, their sequences have diverged widely and their genes often are not linked to transport-related operons. Antoine and co-workers [47,145] posited that these bug genes may be used by other transport systems, or involved in other metabolic activities or other pathways activated under stress. In this context it is interesting to note that many of the bug genes in CH34 seem linked or are in the close vicinity of transporter and regulatory genes. We also find it intriguing that *C. metallidurans*, and the other *Cupriavidus* species, harbour so many bug genes: Out of 200 genomes analysed by Antoine *et al.* [47], only *Bordetella* species and *C. metallidurans* were listed with a prodigious number of bug genes. We might argue that *Cupriavidus* species are free-living organisms wherein the use of solute receptors for specialised strategies of adaptation and survival evolved, but then, we question why so few other free-living organisms failed to develop this approach? We surveyed for the Bug domain in the 1,216 complete and draft, bacterial and archaeal genomes currently in the MiST2 database [132] and found 2,377 proteins with Bug domains. Outside the genera *Bordetella* and *Cupriavidus* only four genomes had a comparable count: *Delftia acidovorans* SPH-1 (159), *Variovorax paradoxus S110* (137), *Verminophrobacter eiseniae* EF01-2 (131), and *Polaromonas* sp. JS666 (97). The only other genomes with a large count (>40) were *Diaphorobacter* sp. TPSY (45), *Acidovorax avenae* ssp. *citrulli* (55), *Acidovorax* JS42 (46), and *Polaromonas naphtalenivorans* CJ2 (41). To our surprise, all organisms that showed an overrepresentation of bug genes belonged to the same beta-proteobacterial order of *Burkholderiales* (Figure 5; Table S8). We propose to call this group of prokaryotes the “Bug Receptor Bacteria” or BuRBs. Such overrepresentation may be a recent innovation in bacteria, or may be more useful to those with a particular lifestyle and habitat. Whether the propensity to utilise large numbers of Bug-domain containing receptors arose within the genus *Bordetella* and was radiated out to closely related genera, or whether it has originated elsewhere within the order of *Burkholderiales* is an open question. It is useful here to mention that *B. petrii*, which carries 107 bug genes, is not host-restricted like the other *Bordetella* species. It is an environmental species typically obtained from river sediment, grass root, and polluted soil [146,147] and has the capacity to degrade aromatic compounds and detoxify heavy metals [148]. However, *B. petrii* strains subsequently were isolated from patients with mandibular osteomyelitis and chronic suppurative mastoiditis [149,150]; thus, this species appears to be an opportunistic pathogen after all. It also contains many mobile genetic elements [148], some strikingly resembling those found in CH34 [59], that may have played an intermediate role in the evolution of BuRBs

## Beta-proteobacteria

Burkholderiales

*Alcaligenaceae*

*Bordetella*

*B. bronchiseptica* RB50 (183)

*B. parapertussis* 12822 (140)

*B. pertussis* Tohama I (79)

*B. petrii* DSM 12804 (107)

*B. avium* 197N (26)

*Burkholderiaceae*

*Burkholderia*

*B. xenovorans* LB400 (13)

*Ralstonia*

*R. solanacearum* GMI1000 (2)

*Polymucleobacter*

*P. necessarius* subsp. asymbioticus QLW (16)

*P. necessarius* subsp. necessarius STIR1 (8)

*Cupriavidus*

*C. metallidurans* CH34 (123)

*C. eutropha* H16 (155)

*C. pinatubonensis* JMP134 (131)

*C. taiwanensis* (103)

*Comamonadaceae*

*Acidovorax*

*A. citrulli* AAC00-1 (55)

*Acidovirax* sp. JS42 (46)

*Delftia*

*D. acidovorans* (159)

*Diaphorobacter*

*Diaphorobacter* sp. TPSY (45)

*Polaromonas*

*Polaromonas* sp. JS666 (97)

*P. naphtalenivorans* CJ2 (41)

*Variovorax*

*V. paradoxus* S110 (137)

*Verminephrobacter*

*V. eisiniae* EF01-2 (131)

*Oxalobacteraceae*

*Herminiimonas*

*Herminiimonas* arsenicoxydans (5)

*Janthinobacterium*

*Janthinobacterium* sp. Marseille (10)

**Figure 5. Distribution of the Bug domain.** Overrepresentation of 'Bug' domains in the family *Burkholderiales* according to MIST data (http://mistdb.com/), with numbers in parentheses. Organisms encoding more than 40 Bug domain-containing proteins are in bold. Only relevant taxa and species are shown (see text). Detailed data are in Table S8.

doi:10.1371/journal.pone.0010433.g005

and the spread of the Bug domain. We speculate that BuRBs evolved many Bug-domain proteins as periplasmic receptors to expand their cellular capacity to bind and transport, via the Venus fly trap fold, a wide range of environmental carboxylated compounds. They might include aminoacids (glutamate, aspartate), other forms of C-skeletons useful for growth (oxalate, malate, succinate, lactate, citrate), humic-acid derivatives from soil, metal-complex carboxylated acids in polluted environments, or plant-root-exuded carboxylates naturally accumulating in the rhizosphere.

Another unusual feature of signal transduction in the *C. metallidurans* CH34 genome is the presence of an intact phosphorelay two-component system entailing a multidomain histidine-kinase sensor protein BvgS1 (Rmet_5710) and a response-transcriptional activator BvgA (Rmet_5714). This system BvgSA controls, via the transduction of environmental signals, the expression of numerous virulence- and colonization-factors in pathogenic *Bordetella* species [50,151] and resembles, respectively, the EvgSA and RocSA systems in *E. coli* and *P. aeruginosa*. *E. coli* contains at least 37 EvgA-activated genes, including stationary-

<figure><img src="image_23.png" /></figure>
phase acid resistance genes and multidrug efflux pump genes [152], while *P. aeruginosa* RocA activates fimbrial adhesin genes for forming biofilms [153]. The BvgA transcriptional activator is well conserved in the four *Cupriavidus* species (63–68% aa identity) and *B. petrii* (58% aa identity), and displays a good similarity to EvgA and RocA, and to the BvgA regulators of pathogenic *Bordetella* species (39–45% aa identity) (all identities with minLrap >0.95). Likewise, the length and sequence of the *C. metallidurans* BvgS₁ protein is conserved to EvgS and RocS, as well as to the BvgS sensory proteins of pathogenic *Bordetella* species (29–33% aa identity, minLrap 1.0). By contrast, the BvgS equivalents of *B. petrii* and the three other *Cupriavidus* strains are devoid of the large periplasmic domain spanning nearly 400 residues (Figure 6). Thus, only the BvgSA system of *C. metallidurans* is fully conserved in respect to the *E. coli* EvgSA, *P. aeruginosa* RocSA, and BvgSA systems of pathogenic *Bordetella*. Adding to the complexity of the BvgS structures in *C. metallidurans* and *B. petrii*, both have a second Bvg histidine-kinase sensor. It is called BvgS₂ (encoded by Rmet_5724 and Bpet4472) and lacks the periplasmic domain. Also, in both organisms the phosphotransfer domain HPt, which is

in pathogenic *Bordetella* part of the BvgS sensor, is an independent protein (Rmet_5713 and Bpet4470, respectively) (Figure 6). Certainly, the region spanning *bvgS*₁ (Rmet_5710) to *bvgS*₂ (Rmet_5724) has undergone multiple insertions and DNA rearrangements, proof of which is a nearby IS1088 copy (Rmet_5718) inserted into a defective serine protease gene (the IS element is indicated with a blue rectangle in the +1 reading frame of Figure 6).

## Capsular proteins

Immediately downstream of the *bvg* gene region (Rmet_5710-24; see above) there is a stretch of genes *kpsMTED-kdsAB-kpsFCS* putatively involved in synthesizing and transporting capsular polysaccharides (genes Rmet_5729 through Rmet_5737) (Figure 6). The *kdsA* and *kdsB* genes (Rmet_5733-34) located between these *kps* loci encode enzymes for the biosynthesis of 2-keto-3-deoxyoctulosonic acid (KDO), a common component of (lipo)polysaccharides. A similar gene cassette is present in the *C. taiwanensis* genome as a cluster of nine genes (RALTA_B1091 through RALTA_B1100). Such gene casettes appear to be

<figure><img src="image_24.png" /></figure>

**Figure 6.** The *bvgSA* and *kps* loci of *C. metallidurans* CH34. Genomic region on the CHR2 replicon of *C. metallidurans* CH34 covering the *bvgSA* locus encoding a master virulence regulon (Rmet_5710, _5714, _5724) and the *kps* locus for synthesis and transport of capsular polysaccharides (Rmet_5729 to _5737). Relevant synteny maps are given for 11 organisms. Abbreviations: Cpin, *C. pinatubonensis* JMP134; Ceut, *C. eutrophus* H16; Ctai, *C. taiwanensis*; Bpet, *B. petrii* DSM 12804; Bbro, *Bordetella bronchiseptica* RB50; Paer, *Pseudomonas aeruginosa* PAO1; Bpar, *B. parapertussis* 12822; Bper, *B. pertussis* Tohama I; Ecol-A, avian pathogenic *Escherichia coli* (APEC01); Ecol-U, uriopathogenic *E. coli* (UTI89); Ecol-K, *E. coli* K12. A nearby IS1088 element (Rmet_5718) is indicated with a blue rectangle in the +1 reading frame (see text).
doi:10.1371/journal.pone.0010433.g006

<figure><img src="image_25.png" /></figure>
phase acid resistance genes and multidrug efflux pump genes [152], while *P. aeruginosa* RocA activates fimbrial adhesin genes for forming biofilms [153]. The BvgA transcriptional activator is well conserved in the four *Cupriavidus* species (63–68% aa identity) and *B. petrii* (58% aa identity), and displays a good similarity to EvgA and RocA, and to the BvgA regulators of pathogenic *Bordetella* species (39–45% aa identity) (all identities with minLrap >0.95). Likewise, the length and sequence of the *C. metallidurans* BvgS₁ protein is conserved to EvgS and RocS, as well as to the BvgS sensory proteins of pathogenic *Bordetella* species (29–33% aa identity, minLrap 1.0). By contrast, the BvgS equivalents of *B. petrii* and the three other *Cupriavidus* strains are devoid of the large periplasmic domain spanning nearly 400 residues (Figure 6). Thus, only the BvgSA system of *C. metallidurans* is fully conserved in respect to the *E. coli* EvgSA, *P. aeruginosa* RocSA, and BvgSA systems of pathogenic *Bordetella*. Adding to the complexity of the BvgS structures in *C. metallidurans* and *B. petrii*, both have a second Bvg histidine-kinase sensor. It is called BvgS₂ (encoded by Rmet_5724 and Bpet4472) and lacks the periplasmic domain. Also, in both organisms the phosphotransfer domain HPt, which is

in pathogenic *Bordetella* part of the BvgS sensor, is an independent protein (Rmet_5713 and Bpet4470, respectively) (Figure 6). Certainly, the region spanning *bvgS*₁ (Rmet_5710) to *bvgS*₂ (Rmet_5724) has undergone multiple insertions and DNA rearrangements, proof of which is a nearby IS1088 copy (Rmet_5718) inserted into a defective serine protease gene (the IS element is indicated with a blue rectangle in the +1 reading frame of Figure 6).

## Capsular proteins

Immediately downstream of the *bvg* gene region (Rmet_5710-24; see above) there is a stretch of genes *kpsMTED-kdsAB-kpsFCS* putatively involved in synthesizing and transporting capsular polysaccharides (genes Rmet_5729 through Rmet_5737) (Figure 6). The *kdsA* and *kdsB* genes (Rmet_5733-34) located between these *kps* loci encode enzymes for the biosynthesis of 2-keto-3-deoxyoctulosonic acid (KDO), a common component of (lipo)polysaccharides. A similar gene cassette is present in the *C. taiwanensis* genome as a cluster of nine genes (RALTA_B1091 through RALTA_B1100). Such gene cassettes appear to be

<figure><img src="image_24.png" /></figure>

**Figure 6.** The *bvgSA* and *kps* loci of *C. metallidurans* CH34. Genomic region on the CHR2 replicon of *C. metallidurans* CH34 covering the *bvgSA* locus encoding a master virulence regulon (Rmet_5710, _5714, _5724) and the *kps* locus for synthesis and transport of capsular polysaccharides (Rmet_5729 to _5737). Relevant synteny maps are given for 11 organisms. Abbreviations: Cpin, *C. pinatubonensis* JMP134; Ceut, *C. eutrophus* H16; Ctai, *C. taiwanensis*; Bpet, *B. petrii* DSM 12804; Bbro, *Bordetella bronchiseptica* RB50; Paer, *Pseudomonas aeruginosa* PAO1; Bpar, *B. parapertussis* 12822; Bper, *B. pertussis* Tohama I; Ecol-A, avian pathogenic *Escherichia coli* (APEC01); Ecol-U, uri pathogenic *E. coli* (UTI89); Ecol-K, *E. coli* K12. A nearby IS1088 element (Rmet_5718) is indicated with a blue rectangle in the +1 reading frame (see text).
doi:10.1371/journal.pone.0010433.g006

<figure><img src="image_25.png" /></figure>
organised in three regions in analogy with the group II/III capsular polysaccharide gene clusters of K-type *Escherichia coli* [154,155] in which a serotype-specific region (SSR) is flanked by conserved *kps* genes encoding a transport machinery for bringing polysaccharides to the cell surface. This entails a heterodimeric ABC transporter KpsMT, a multimeric KpsD outer membrane porter, and a periplasmic connector KpsE [156,157]. The function of the other Kps proteins KpsF, KpsC, and KpsS is less clear but they were implicated in lipid metabolism, translocation and surface expression of capsule polymers, and determining chain length. We point out that the *C. metallidurans* *kpsF* gene (*Rmet_5735*), encoding a D-arabinose 5-phosphate isomerase, has a duplicate copy on *CHR1* as gene *Rmet_0307*. Likewise, *kdsA* and *kdsB* have duplicate copies on *CHR1*, respectively, as *Rmet_1053* and *Rmet_0533*. The genomes of *C. eutrophus* H16 and *C. pinatubonensis* JMP134 do not carry a *kps-SSR-kps* cassette. In fact, of all the species sofar discussed in this paper, only pathogenic *Bordetella* species and *B. petrii* contain a similar cassette *kpsMT-kpsE/wcbD-SSR-kpsCS/wcbAO* although they all seem to lack the *kpsD* gene encoding the outer membrane porter (in these species it may have been replaced by an unknown alternative protein). Interestingly, the plant symbiont *Sinorhizobium meliloti* produces a K-antigen that is exported by a typical *kpsMTED-SSR-kpsCS* module. This module is encoded in both *S. meliloti* strains Rm41 and 2101 by two separate *rkp* loci of which one is sited on the chromosome and the other one on the symbiotic megaplasmid [158,159] (not shown).

Although there is no direct evidence for an K-antigen polysaccharide (KPS) in *C. metallidurans* CH34 or *C. taiwanensis*, such a Kps 'secretin' complex may be used to secrete other types of exopolysaccharides, lipo-oligosaccharides, or short-chained signaling molecules. In this context, the presence of a full set of *pelABCDEFG* genes (*Rmet_4155* through *Rmet_4561*) in the *C. metallidurans* CH34 captured our interest. This *pel* operon first was described in *P. aeruginosa* PA14 as producing a glucose-rich biofilm matrix [160]; it later was identified in *P. aeruginosa* PAK and *R. solanacearum* GM1000 [161]. Comparative analysis of related genomes via MaGe synteny maps (data not shown) failed to detect a *pel* locus in *C. pinatubonensis* JMP134; however, both *C. eutrophus* H16 and *C. taiwanensis* carry a full set of seven *pel* genes. Other genomes listed in the MaGe database with a complete set are those of three other *R. solanacearum* strains (Molk2, Ipo1609, and UW551), *R. picketti* 12D, *B. xenovorans* LB400, *Pseudomonas fluorescens* Pf5, and *D. acidovorans*. The *pel* operon encodes sugar-processing enzymes, such as oligogalacturonide lyase (PeIA), two glycosyl-transferases (PelC and PelF), and a sucrose synthase (PeLE), along with proteins involved in cellular signalling or transmembrane transport (i.e., PelD, a cyclic-di-GMP signal receptor and PelG, a putative 'flippase'). Implication of the *pel* locus in biofilm formation was apparant by a variety of *pel* mutants of *P. aeruginosa* strains and *R. solanacearum* [160-162]. Immediately upstream of the *C. metallidurans* CH34 *pel* locus we identified three transposase (*tnpA*) genes belonging to different families of insertion elements namely *Rmet_4154* (*ISRme8*), *Rmet_4153* (*ISRme1*), and *Rmet_4152* (*ISRme5*) indicating a hotspot for insertion.

In searching for additional genes that might be involved in capsular protein synthesis and transport, we found two more regions of interest. The first is a large cluster of 20 genes (*Rmet_2711* through *Rmet_2731* situated on *CHR1*) clearly involved in lipopolysaccharide biosynthesis although the exact role of most proteins remains enigmatic. Based on the patched synteny with other genomes, seemingly this region, which is highly conserved compared to *C. taiwanensis*, to a lesser extent to the other *Cupriavidus* and *Ralstonia* species, and very intermittantly with

the genomes of any other of the species in the MaGe database, displays an amalgam of genes encoding secretion proteins and sugar-processing enzymes, possibly of different origins, that may or may not act in concert. A second cluster of nearly 30 genes (*Rmet_5832* through *Rmet_5860* situated in *CHR2*) exhibits a similar mosaic structure with many genes being hypothetical or conserved hypothetical while others generally seem to be involved in transport, signal transduction, or sugar processing activities. Some genes are worth mentioning, for instance *Rmet_5835* and *Rmet_5843*, encoding proteins that respectively display a fair-to-good aa identity with the *P. aeruginosa* PsIA (32%) and PsIB (43%) proteins. Psl proteins in *P. aeruginosa* are implicated in the synthesis of a mannose-rich component of the biofilm matrix [162]. Just downstream of these genes, on the opposite strand of *CHR2*, lies a small cluster of three genes *epsAPB* (*Rmet_5851-5850-5849*) encoding proteins involved in exporting exopolysaccharides; this locus is, on the same strand, immediately preceded by four genes encoding glycosyltransferases (*Rmet_5853* through *Rmet_5856*), and two more genes encoding sugar-processing enzymes mannose epimerase (*Rmet_5859*) and mannose dehydratase (*Rmet_5860*). We believe that these two large regions (in addition to the better defined *pel* locus) deserve special attention in regard to the *kps-SSR-kps* cassette or alternative transport systems encoded by the *C. metallidurans* CH34 genome, particularly in view of the organism's association with biofilms [14,39], and the role of exopolysaccharides in bacterial complexation of metals [163].

## Motility, chemotaxis, biofilms, and Q-sensing

*C. metallidurans* displays flagella uniformly distributed over the cell surface [34] allowing it to swarm and swim. Most of the *flh*, *fli*, and *flg* genes necessary for synthesizing flagella are grouped within four gene clusters: A *flh* locus *Rmet_3698-3703*, two *fli* loci *Rmet_5252-66* and *Rmet_5298-5303*, and the *flg* locus represented by *Rmet_3731-44*. In addition, the CH34 genome encodes *cpa*, *tad*, and *pil* loci essential for synthesizing type IV pili (Tfp), allowing cells to move across solid surfaces via a mode of twitching. These include the *pilVWX* genes (*Rmet_0189/91*), various *cpa* and *tad* genes within the region *Rmet_0639* through *Rmet_0697*, *pilTU* genes (*Rmet_2935/6*), the *pilBCD* genes (*Rmet_3108-10*), the *pilMNOPQ* genes (*Rmet_3268-72*), and the *cpaBCD-tadABCD* genes (*Rmet_3653-59*). The CH34 genome also contains many genes required for the biogenesis of fimbriae important for cell adhesion, colonization, and biofilm formation, including the *fimAC-htrE* genes *Rmet_0574/5/6*, *FimA*-type adhesin genes *Rmet_1667*, *Rmet_4250*, and *Rmet_4961*, and a *fim* locus *Rmet_4250-64*. Our genome survey shows that more than 150 genes may be involved in generating or controlling flagella, pili, and fimbriae. Most occur within the above loci, or are situated within a large cluster of 18 chemotaxis (*che*), motility (*mot*) and flagellar (*flh*, *fli*) genes (*Rmet_3678* through *Rmet_3695*); others are part of smaller sets of genes encoding sensory- or regulatory-proteins.

One major mechanism of regulating motility in strain CH34 is through the action of a Quorum Sensing (QS) system encompassing the *phcBSRQ* operon (*Rmet_2949-52*) and the *phcA* gene (*Rmet_2977*) [139]. This QS or cell-to-cell communication system was first discovered in *R. solanacearum* and is neither related to the classical LuxIR system utilizing acyl-homoserine lactone (AHL) autoinducers nor the LuxS system with autoinducer-2. In *R. solanacearum*, PhcB directs the synthesis of the diffusible signal molecule 3-OH-palmitic acid methyl ester (3-OH-PAME). Typically, with higher cell densities also 3-OH-PAME levels increase to an extent that it saturates the membrane-bound sensor PhcS, effectively lifting the PhcR-mediated repression of the PhcA regulator. All five *phc* genes of this system are conserved in CH34,

<figure><img src="image_26.png" /></figure>
organised in three regions in analogy with the group II/III capsular polysaccharide gene clusters of K-type *Escherichia coli* [154,155] in which a serotype-specific region (SSR) is flanked by conserved *kps* genes encoding a transport machinery for bringing polysaccharides to the cell surface. This entails a heterodimeric ABC transporter KpsMT, a multimeric KpsD outer membrane porter, and a periplasmic connector KpsE [156,157]. The function of the other Kps proteins KpsF, KpsC, and KpsS is less clear but they were implicated in lipid metabolism, translocation and surface expression of capsule polymers, and determining chain length. We point out that the *C. metallidurans* *kpsF* gene (*Rmet_5735*), encoding a D-arabinose 5-phosphate isomerase, has a duplicate copy on *CHR1* as gene *Rmet_0307*. Likewise, *kdsA* and *kdsB* have duplicate copies on *CHR1*, respectively, as *Rmet_1053* and *Rmet_0533*. The genomes of *C. eutrophus* H16 and *C. pinatubonensis* JMP134 do not carry a *kps-SSR-kps* cassette. In fact, of all the species sofar discussed in this paper, only pathogenic *Bordetella* species and *B. petrii* contain a similar cassette *kpsMT-kpsE/wcbD-SSR-kpsCS/wcbAO* although they all seem to lack the *kpsD* gene encoding the outer membrane porter (in these species it may have been replaced by an unknown alternative protein). Interestingly, the plant symbiont *Sinorhizobium meliloti* produces a K-antigen that is exported by a typical *kpsMTED-SSR-kpsCS* module. This module is encoded in both *S. meliloti* strains Rm41 and 2101 by two separate *rkp* loci of which one is sited on the chromosome and the other one on the symbiotic megaplasmid [158,159] (not shown).

Although there is no direct evidence for an K-antigen polysaccharide (KPS) in *C. metallidurans* CH34 or *C. taiwanensis*, such a Kps 'secretin' complex may be used to secrete other types of exopolysaccharides, lipo-oligosaccharides, or short-chained signaling molecules. In this context, the presence of a full set of *pelABCDEFG* genes (*Rmet_4155* through *Rmet_4561*) in the *C. metallidurans* CH34 captured our interest. This *pel* operon first was described in *P. aeruginosa* PA14 as producing a glucose-rich biofilm matrix [160]; it later was identified in *P. aeruginosa* PAK and *R. solanacearum* GM1000 [161]. Comparative analysis of related genomes via MaGe synteny maps (data not shown) failed to detect a *pel* locus in *C. pinatubonensis* JMP134; however, both *C. eutrophus* H16 and *C. taiwanensis* carry a full set of seven *pel* genes. Other genomes listed in the MaGe database with a complete set are those of three other *R. solanacearum* strains (Molk2, Ipo1609, and UW551), *R. picketti* 12D, *B. xenovorans* LB400, *Pseudomonas fluorescens* Pf5, and *D. acidovorans*. The *pel* operon encodes sugar-processing enzymes, such as oligogalacturonide lyase (PeIA), two glycosyl-transferases (PelC and PelF), and a sucrose synthase (PeLE), along with proteins involved in cellular signalling or transmembrane transport (i.e., PelD, a cyclic-di-GMP signal receptor and PelG, a putative 'flippase'). Implication of the *pel* locus in biofilm formation was apparent by a variety of *pel* mutants of *P. aeruginosa* strains and *R. solanacearum* [160-162]. Immediately upstream of the *C. metallidurans* CH34 *pel* locus we identified three transposase (*tnpA*) genes belonging to different families of insertion elements namely *Rmet_4154* (*ISRme8*), *Rmet_4153* (*ISRme1*), and *Rmet_4152* (*ISRme5*) indicating a hotspot for insertion.

In searching for additional genes that might be involved in capsular protein synthesis and transport, we found two more regions of interest. The first is a large cluster of 20 genes (*Rmet_2711* through *Rmet_2731* situated on *CHR1*) clearly involved in lipopolysaccharide biosynthesis although the exact role of most proteins remains enigmatic. Based on the patched synteny with other genomes, seemingly this region, which is highly conserved compared to *C. taiwanensis*, to a lesser extent to the other *Cupriavidus* and *Ralstonia* species, and very intermittently with

the genomes of any other of the species in the MaGe database, displays an amalgam of genes encoding secretion proteins and sugar-processing enzymes, possibly of different origins, that may or may not act in concert. A second cluster of nearly 30 genes (*Rmet_5832* through *Rmet_5860* situated in *CHR2*) exhibits a similar mosaic structure with many genes being hypothetical or conserved hypothetical while others generally seem to be involved in transport, signal transduction, or sugar processing activities. Some genes are worth mentioning, for instance *Rmet_5835* and *Rmet_5843*, encoding proteins that respectively display a fair-to-good aa identity with the *P. aeruginosa* PsIA (32%) and PsIB (43%) proteins. Psl proteins in *P. aeruginosa* are implicated in the synthesis of a mannose-rich component of the biofilm matrix [162]. Just downstream of these genes, on the opposite strand of *CHR2*, lies a small cluster of three genes *epsAPB* (*Rmet_5851-5850-5849*) encoding proteins involved in exporting exopolysaccharides; this locus is, on the same strand, immediately preceded by four genes encoding glycosyltransferases (*Rmet_5853* through *Rmet_5856*), and two more genes encoding sugar-processing enzymes mannose epimerase (*Rmet_5859*) and mannose dehydratase (*Rmet_5860*). We believe that these two large regions (in addition to the better defined *pel* locus) deserve special attention in regard to the *kps-SSR-kps* cassette or alternative transport systems encoded by the *C. metallidurans* CH34 genome, particularly in view of the organism's association with biofilms [14,39], and the role of exopolysaccharides in bacterial complexation of metals [163].

## Motility, chemotaxis, biofilms, and Q-sensing

*C. metallidurans* displays flagella uniformly distributed over the cell surface [34] allowing it to swarm and swim. Most of the *flh*, *fli*, and *flg* genes necessary for synthesizing flagella are grouped within four gene clusters: A *flh* locus *Rmet_3698-3703*, two *fli* loci *Rmet_5252-66* and *Rmet_5298-5303*, and the *flg* locus represented by *Rmet_3731-44*. In addition, the CH34 genome encodes *cpa*, *tad*, and *pil* loci essential for synthesizing type IV pili (Tfp), allowing cells to move across solid surfaces via a mode of twitching. These include the *pilVWX* genes (*Rmet_0189/91*), various *cpa* and *tad* genes within the region *Rmet_0639* through *Rmet_0697*, *pilTU* genes (*Rmet_2935/6*), the *pilBCD* genes (*Rmet_3108-10*), the *pilMNOPQ* genes (*Rmet_3268-72*), and the *cpaBCD-tadABCD* genes (*Rmet_3653-59*). The CH34 genome also contains many genes required for the biogenesis of fimbriae important for cell adhesion, colonization, and biofilm formation, including the *fimAC-htrE* genes *Rmet_0574/5/6*, *FimA*-type adhesin genes *Rmet_1667*, *Rmet_4250*, and *Rmet_4961*, and a *fim* locus *Rmet_4250-64*. Our genome survey shows that more than 150 genes may be involved in generating or controlling flagella, pili, and fimbriae. Most occur within the above loci, or are situated within a large cluster of 18 chemotaxis (*che*), motility (*mot*) and flagellar (*flh*, *fli*) genes (*Rmet_3678* through *Rmet_3695*); others are part of smaller sets of genes encoding sensory- or regulatory-proteins.

One major mechanism of regulating motility in strain CH34 is through the action of a Quorum Sensing (QS) system encompassing the *phcBSRQ* operon (*Rmet_2949-52*) and the *phcA* gene (*Rmet_2977*) [139]. This QS or cell-to-cell communication system was first discovered in *R. solanacearum* and is neither related to the classical LuxIR system utilizing acyl-homoserine lactone (AHL) autoinducers nor the LuxS system with autoinducer-2. In *R. solanacearum*, PhcB directs the synthesis of the diffusible signal molecule 3-OH-palmitic acid methyl ester (3-OH-PAME). Typically, with higher cell densities also 3-OH-PAME levels increase to an extent that it saturates the membrane-bound sensor PhcS, effectively lifting the PhcR-mediated repression of the PhcA regulator. All five *phc* genes of this system are conserved in CH34,

<figure><img src="image_26.png" /></figure>
the other three *Cupriavidus* species, and *R. pickettii*. However, while 3-OH PAME could be extracted and purified from *R. solanacearum*, this was not the case for CH34, indicating that 3-OH PAME-like activity in CH34 may result from a structural variant of this molecule [139]. In the plant-pathogenic bacterium *R. solanacearum* activation of the virulence regulator PhcA is followed by an enhanced production of exoenzymes and extracellular polysaccharides leading to severely decreased motility, presumably in preparation for host colonization [164]. This is not the case in strain CH34 where PhcA activation actually stimulates motility [139]. Likewise, the inactivation of *phcA* in *R. solanacearum* increased siderophore production tenfold [135] but not in *C. metallidurans* CH34 where no affect on siderophore production was seen [139]. Possibly, the Phc regulatory system may connect to different sets of genes in *C. metallidurans* compared to *R. solanacearum*; this is not unduly surprising bearing in mind the fundamental differences between their ecology and physiology. Interestingly, CH34 does possess an AHL acylase (Rmet_4680), which enables CH34 to modulate AHL levels and thereby AHL-dependent regulatory systems of other members in a multispecies microbial community.

## Heterotrophic carbon metabolism

*C. metallidurans* CH34 cannot grow on glucose, fructose, or galactose [9] because it lacks adequate glucose uptake systems and does not possess a functional 6-phosphofructokinase. Consequently, energy is not produced through the Embden-Meyerhof-Parnas pathway, but by assimilating acidic sugar-derivatives such as D-gluconate, acetate, pyruvate, lactate, L-malate, benzoate, 4-hydroxybenzoate, and various dicarboxylic acids (C₄ to C₁₁) [9]. CH34 can derive glucose-6-phosphate from gluconeogenesis (see below) or via the phosphorylation of cytoplasmic glucose by glucokinase (Rmet_5799). It then is further converted to 6-phosphogluconate (6GP) in a two-step reaction involving the *pgl* and *zuf* genes (encoded by Rmet_5800 and Rmet_5801, respectively). The 6GP is metabolized through the alternative glycolytic Entner-Doudoroff (ED) pathway involving a 6-phosphogluconate dehydratase (EDD; Rmet_5802) and a 2-keto-3-deoxy-6-phosphogluconate aldolase (EDA; Rmet_4768), thus generating two pyruvate molecules that finally are processed to acetyl-CoA. In many prokaryotes, this acetyl-CoA is fully oxidized to CO₂ via citrate, D-isocitrate, 2-ketoglutarate, succinate, fumarate, malate, and oxalacetate in the tri carboxic-acid (TCA) cycle, that is, the Krebs cycle. However, *C. metallidurans* apparently also can use the glyoxylate bypass in which D-isocitrate is cleaved by isocitrate lyase (AceA) (Rmet_1385, Rmet_4737) into succinate and glyoxylate. As in full TCA cycles, succinate is converted to fumarate by succinate dehydrogenase (succinate:quinone reductase, SQR), a multimeric enzyme encoded by four genes *sdhCDAB* (Rmet_2483/4/5/6) and further processed to malate by fumarase FumA (Rmet_2273, Rmet_5752). In parallel, glyoxylate condenses with acetyl-CoA yielding malate via malate synthase (AceB) (Rmet_1390). Malate finally is changed into oxaloacetate via the malate dehydrogenase enzyme MDH (Rmet_2489, Rmet_5095), completing a full cycle in which acetate is again transformed to citrate by citrate synthase CitA (Rmet_2481, Rmet_4144, Rmet_4268, Rmet_4946, and Rmet_5380) and citrate to D-isocitrate via aconitase AcnA (Rmet_1585, Rmet_2492, Rmet_4240, and Rmet_5296). We note that D-isocitrate, as for full TCA cycles, is transfigured into 2-ketoglutarate by isocitrate dehydrogenase ICDH (Rmet_2895, Rmet_3729). The carbon skeleton of 2-ketoglutarate then is used, depending on cellular ammonia levels and energy status, by either one of the two enzymes glutamate dehydrogenase GDH (Rmet_1181) or gluta-

mate synthase (GOGAT) (GltAB encoded by Rmet_3262/3) to generate glutamate that in turn feeds the nitrogen metabolic pathways. We offer three important observations here. First, although the *C. metallidurans* CH34 genome encodes the E1, E2, and E3 components of an α-ketoglutarate dehydrogenase (KGDH) complex (Rmet_2048 through Rmet_2050), their role in the TCA cycle for converting 2-ketoglutarate to succinate via succinyl-CoA is undecided. Possibly, an alternative route is followed wherein the gene products of Rmet_2049 and Rmet_2050 actually form a ketoglutarate decarboxylase (KGD) complex that changes 2-ketoglutarate into the intermediate succinate semialdehyde followed by further conversion to succinate by a succinate-semialdehyde dehydrogenase (SSDH) encoded by the *gabD* genes (Rmet_1898, Rmet_3961, and Rmet_4942). Interestingly, these SSDH enzymes may have a role in 4-hydroxybutyric acid catabolism, as was reported for *C. eutrophus* H16 [165]. Second, neither succinate nor oxaloacetate are produced via a reductive TCA cycle as the essential enzymes are missing (see further). And third, most TCA enzymes in *C. metallidurans* CH34 appear to have multiple isoenzymes whose genes are scattered throughout both replicons CHR1 and CHR2 (i.e., we discerned at least five citrate synthase genes, four aconitase genes, three genes for SSDH, and two genes each for ICDH, MDH, fumerase, and isocitrate lyase).

None of the sequenced *Cupriavidus* strains contains a gene for 6-phosphogluconate dehydrogenase (6-PGDH), basically ruling out the oxidative branch of the pentose-phosphate pathway. Nonetheless, all essential genes for gluconeogenesis, in which glucose is generated from non-sugar carbon substrates (such as pyruvate, lactate, oxaloacetate, and glycerol) and glucogenic amino-acids (primarily alanine and glutamine) are present in *C. metallidurans*. They include genes to transform malate via oxaloacetate (Rmet_2489, Rmet_2750) or via pyruvate (Rmet_0878, Rmet_1453) to phosphoenolpyruvate that in turn is processed to fructose-1,6-biphosphate through a cascade of enzymatic actions by, for example, phosphopyruvate hydratase (ENO; Rmet_1055, Rmet_1176), phosphoglycerate mutase (PGM; Rmet_0251, Rmet_0420), phosphoglycerate kinase (PGK; Rmet_0501, Rmet_1516), glyceraldehyde-3-phosphate dehydrogenase (GAP; Rmet_1096, Rmet_1515, Rmet_2979), and fructose-1,6-biphosphate aldolase (FBA; Rmet_0503, Rmet_1492, Rmet_1518)]. The resulting fructose-1,6-biphosphate is further converted to fructose-6-phosphate by fructose-bisphosphatase (FBP; Rmet_0875, Rmet_1507, Rmet_1511) and finally into β-D-glucose-6-phosphate by glucose 6-phosphate isomerase (GPI; Rmet_1879). Without exception, CHR1 encompasses all the genes required for gluconeogenesis, some of which also are involved in the reductive pentose-phosphate pathway (Calvin-Benson-Bassham cycle).

Under aerobic conditions, *C. metallidurans* CH34 potentially can degrade benzoate and 4-hydroxybenzoate to ketoadipate via catechol and protocatechuate, respectively, as it has all the genes for both ortho-cleavage pathways (e.g., the *benABCDM* and *catABCD* genes Rmet_4875 through Rmet_4885, *pcaA* gene Rmet_4270, *pcaBCDHGKQ* genes Rmet_4011 through Rmet_4016, and *pobAR* genes Rmet_4017/8). The ketoadipate thus formed then is processed to acetyl-CoA by the products of the *pcaIJF* genes (Rmet_3667/8/9). In strain CH34, benzoate might be degraded anaerobically via the central metabolite benzoyl-CoA (instead of catechol; see Harwood *et al.* [166]) in an alternate pathway requiring the products of Rmet_1220 through Rmet_1225. Therein, the benzoyl-CoA reductase BoxAB (Rmet_1220/1) will open the benzene ring, followed by β-oxidation of the resulting product (oxidation occurs at the β-carbon C3) leading to 3-ketoadipyl-CoA, and finally to succinyl-

<figure><img src="image_27.png" /></figure>
the other three *Cupriavidus* species, and *R. pickettii*. However, while 3-OH PAME could be extracted and purified from *R. solanacearum*, this was not the case for CH34, indicating that 3-OH PAME-like activity in CH34 may result from a structural variant of this molecule [139]. In the plant-pathogenic bacterium *R. solanacearum* activation of the virulence regulator PhcA is followed by an enhanced production of exoenzymes and extracellular polysaccharides leading to severely decreased motility, presumably in preparation for host colonization [164]. This is not the case in strain CH34 where PhcA activation actually stimulates motility [139]. Likewise, the inactivation of *phcA* in *R. solanacearum* increased siderophore production tenfold [135] but not in *C. metallidurans* CH34 where no affect on siderophore production was seen [139]. Possibly, the Phc regulatory system may connect to different sets of genes in *C. metallidurans* compared to *R. solanacearum*; this is not unduly surprising bearing in mind the fundamental differences between their ecology and physiology. Interestingly, CH34 does possess an AHL acylase (Rmet_4680), which enables CH34 to modulate AHL levels and thereby AHL-dependent regulatory systems of other members in a multispecies microbial community.

## Heterotrophic carbon metabolism

*C. metallidurans* CH34 cannot grow on glucose, fructose, or galactose [9] because it lacks adequate glucose uptake systems and does not possess a functional 6-phosphofructokinase. Consequently, energy is not produced through the Embden-Meyerhof-Parnas pathway, but by assimilating acidic sugar-derivatives such as D-gluconate, acetate, pyruvate, lactate, L-malate, benzoate, 4-hydroxybenzoate, and various dicarboxylic acids (C₄ to C₁₁) [9]. CH34 can derive glucose-6-phosphate from gluconeogenesis (see below) or via the phosphorylation of cytoplasmic glucose by glucokinase (Rmet_5799). It then is further converted to 6-phosphogluconate (6GP) in a two-step reaction involving the *pgl* and *zuf* genes (encoded by Rmet_5800 and Rmet_5801, respectively). The 6GP is metabolized through the alternative glycolytic Entner-Doudoroff (ED) pathway involving a 6-phosphogluconate dehydratase (EDD; Rmet_5802) and a 2-keto-3-deoxy-6-phosphogluconate aldolase (EDA; Rmet_4768), thus generating two pyruvate molecules that finally are processed to acetyl-CoA. In many prokaryotes, this acetyl-CoA is fully oxidized to CO₂ via citrate, D-isocitrate, 2-ketoglutarate, succinate, fumarate, malate, and oxalacetate in the tri carboxic-acid (TCA) cycle, that is, the Krebs cycle. However, *C. metallidurans* apparently also can use the glyoxylate bypass in which D-isocitrate is cleaved by isocitrate lyase (AceA) (Rmet_1385, Rmet_4737) into succinate and glyoxylate. As in full TCA cycles, succinate is converted to fumarate by succinate dehydrogenase (succinate:quinone reductase, SQR), a multimeric enzyme encoded by four genes *sdhCDAB* (Rmet_2483/4/5/6) and further processed to malate by fumarase FumA (Rmet_2273, Rmet_5752). In parallel, glyoxylate condenses with acetyl-CoA yielding malate via malate synthase (AceB) (Rmet_1390). Malate finally is changed into oxaloacetate via the malate dehydrogenase enzyme MDH (Rmet_2489, Rmet_5095), completing a full cycle in which acetate is again transformed to citrate by citrate synthase CitA (Rmet_2481, Rmet_4144, Rmet_4268, Rmet_4946, and Rmet_5380) and citrate to D-isocitrate via aconitase AcnA (Rmet_1585, Rmet_2492, Rmet_4240, and Rmet_5296). We note that D-isocitrate, as for full TCA cycles, is transfigured into 2-ketoglutarate by isocitrate dehydrogenase ICDH (Rmet_2895, Rmet_3729). The carbon skeleton of 2-ketoglutarate then is used, depending on cellular ammonia levels and energy status, by either one of the two enzymes glutamate dehydrogenase GDH (Rmet_1181) or gluta-

mate synthase (GOGAT) (GltAB encoded by Rmet_3262/3) to generate glutamate that in turn feeds the nitrogen metabolic pathways. We offer three important observations here. First, although the *C. metallidurans* CH34 genome encodes the E1, E2, and E3 components of an α-ketoglutarate dehydrogenase (KGDH) complex (Rmet_2048 through Rmet_2050), their role in the TCA cycle for converting 2-ketoglutarate to succinate via succinyl-CoA is undecided. Possibly, an alternative route is followed wherein the gene products of Rmet_2049 and Rmet_2050 actually form a ketoglutarate decarboxylase (KGD) complex that changes 2-ketoglutarate into the intermediate succinate semialdehyde followed by further conversion to succinate by a succinate-semialdehyde dehydrogenase (SSDH) encoded by the *gabD* genes (Rmet_1898, Rmet_3961, and Rmet_4942). Interestingly, these SSDH enzymes may have a role in 4-hydroxybutyric acid catabolism, as was reported for *C. eutrophus* H16 [165]. Second, neither succinate nor oxaloacetate are produced via a reductive TCA cycle as the essential enzymes are missing (see further). And third, most TCA enzymes in *C. metallidurans* CH34 appear to have multiple isoenzymes whose genes are scattered throughout both replicons CHR1 and CHR2 (i.e., we discerned at least five citrate synthase genes, four aconitase genes, three genes for SSDH, and two genes each for ICDH, MDH, fumerase, and isocitrate lyase).

None of the sequenced *Cupriavidus* strains contains a gene for 6-phosphogluconate dehydrogenase (6-PGDH), basically ruling out the oxidative branch of the pentose-phosphate pathway. Nonetheless, all essential genes for gluconeogenesis, in which glucose is generated from non-sugar carbon substrates (such as pyruvate, lactate, oxaloacetate, and glycerol) and glucogenic amino-acids (primarily alanine and glutamine) are present in *C. metallidurans*. They include genes to transform malate via oxaloacetate (Rmet_2489, Rmet_2750) or via pyruvate (Rmet_0878, Rmet_1453) to phosphoenolpyruvate that in turn is processed to fructose-1,6-biphosphate through a cascade of enzymatic actions by, for example, phosphopyruvate hydratase (ENO; Rmet_1055, Rmet_1176), phosphoglycerate mutase (PGM; Rmet_0251, Rmet_0420), phosphoglycerate kinase (PGK; Rmet_0501, Rmet_1516), glyceraldehyde-3-phosphate dehydrogenase (GAP; Rmet_1096, Rmet_1515, Rmet_2979), and fructose-1,6-biphosphate aldolase (FBA; Rmet_0503, Rmet_1492, Rmet_1518)]. The resulting fructose-1,6-biphosphate is further converted to fructose-6-phosphate by fructose-bisphosphatase (FBP; Rmet_0875, Rmet_1507, Rmet_1511) and finally into β-D-glucose-6-phosphate by glucose 6-phosphate isomerase (GPI; Rmet_1879). Without exception, CHR1 encompasses all the genes required for gluconeogenesis, some of which also are involved in the reductive pentose-phosphate pathway (Calvin-Benson-Bassham cycle).

Under aerobic conditions, *C. metallidurans* CH34 potentially can degrade benzoate and 4-hydroxybenzoate to ketoadipate via catechol and protocatechuate, respectively, as it has all the genes for both ortho-cleavage pathways (e.g., the *benABCDM* and *catABCD* genes Rmet_4875 through Rmet_4885, *pcaA* gene Rmet_4270, *pcaBCDHGKQ* genes Rmet_4011 through Rmet_4016, and *pobAR* genes Rmet_4017/8). The ketoadipate thus formed then is processed to acetyl-CoA by the products of the *pcaIJF* genes (Rmet_3667/8/9). In strain CH34, benzoate might be degraded anaerobically via the central metabolite benzoyl-CoA (instead of catechol; see Harwood *et al.* [166]) in an alternate pathway requiring the products of Rmet_1220 through Rmet_1225. Therein, the benzoyl-CoA reductase BoxAB (Rmet_1220/1) will open the benzene ring, followed by β-oxidation of the resulting product (oxidation occurs at the β-carbon C3) leading to 3-ketoadipyl-CoA, and finally to succinyl-

<figure><img src="image_27.png" /></figure>
CoA and acetyl-CoA. Five more genes of this operon (Rmet_1226 through Rmet_1230) probably encode an ABC-like transport system of benzoate.

Similarly, β-oxidation may play an important role in the formation of acetyl-CoA from dicarboxylated compounds because reportedly, *C. metallidurans* CH34 grows on a range of dicarboxy-lated compounds [9]. However, without experimental evidence, it is difficult to predict the genes and enzymes that would be involved in these conversions in CH34. An interesting report by Harrison and Harwood [167] implicated the *pimFABCDE* operon for the β-oxidative degradation of C~7~-C~14~ dicarboxylic- and fatty-acids in *Rhodopseudomonas palustris*. This operon encodes a set of acyl-CoA ligase, enoyl-CoA hydratase-, acyl-CoA dehydrogenase-, and acyl-CoA transferase-enzymes, and is accompanied by a IclR-type regulatory gene, and several genes for ABC-like transport. Our search through the MaGe annotated data of CH34 yielded more than a hundred of these enzymes. Though most of them might degrade fatty acids, we believe that in CH34 many others are recruited to specifically degrade dicarboxylates.

## Aerobic- and anaerobic-respiration

The citric-acid cycle is followed by oxidative phosphorylation in which NADH, FADH~2~, and GTP are reduced by cytochromes and other terminal oxidases in the cell's inner membrane, while protons are expelled into the periplasm. The resulting proton gradient across the membrane then drives ATP synthesis by the conversion of ADP (ATPase; α, β, γ, δ, ε, a, b, and c units encoded by Rmet_3493 through Rmet_3500). A key locus in this aerobic energy metabolism is a 14-gene cluster located on CHR1 (Rmet_0927 through Rmet_0940) encoding a large membrane-bound complex of NADH dehydrogenase (NADH:quinone oxidoreductase) that is highly conserved within the four sequenced *Cupriavidus* strains, and in other closely related species. According to BioCyc pathway analysis, alternative NADH dehydrogenases may exist in *C. metallidurans*. These are encoded by Rmet_2623 as a single-subunit FAD-binding NDHase, by Rmet_0177 as a NAD-dependent epimerase/dehydratase, and by Rmet_4310 as a FAD-dependent pyridine nucleotide-disulphide oxidoreductase. We also discerned the Rmet_4300 gene that apparently codes for a NuoF homologue (NADH ubiquinone oxidoreductase, chain F). Other membrane-associated protein complexes that may be important to the electron transport chain in strain CH34 are succinate:quinone reductase (SQR; see previous section on the TCA cycle), a *bc*~f~-type cytochrome complex (encoded by Rmet_3228/9/30), a *cbb*~3~-type cytochrome c oxidase (encoded by *cco* genes Rmet_2041/2/3/4), two cytochrome *bd*-type quinol oxidases CydAB (encoded by the genepairs Rmet_4955/6 and Rmet_5232/3), a *aa*~3~-type oxidase encoded by the *cta*-gene cluster (Rmet_0261 through Rmet_0271), a *bb*~3~-type cytochrome oxidase encoded by the *cox*-gene cluster (Rmet_3955/6/7/8/9), and three quinol oxidases encoded by the clusters *cyo*~1~ (Rmet_0948/49/50/51), *cyo*~2~ (Rmet_5608/09/10/11), and *cyo*~3~ (Rmet_5790/1/2/3).

Under anoxic conditions, *C. metallidurans* CH34 may resort to alternate modes of energy generation by using electron acceptors other than oxygen. Thus, the bacterium reduces nitrate [32], which is reflected by three loci encoding distinct types of nitrate reductases: a cytoplasmic assimilatory enzyme (NAS; encoded by the *nasAB* genes Rmet_4820/1), a membrane-bound quinol-oxidizing enzyme (NAR; encoded by the *narGHJ1* genes Rmet_2074 through Rmet_2077, accompanied by two ABC-type transporter-genes Rmet_2072/3), and a periplasmic enzyme (NAP; encoded by the *napEDABC* genes Rmet_4061 through Rmet_4065). Probably, the NAS-, NAP-, and NAR- systems have different physiological roles depending on metabolic conditions.

Consequently, their pathways may be interconnected to foster rapid adaptation to changing nitrogen and/or oxygen conditions, a big asset for survival in nature.

Although strain CH34 reportedly does not reduce nitrite [32], it contains two genes Rmet_4018/9 encoding the large- and small-subunits of a putative assimilatory nitrite-reductase (ANR). These genes, preceded by a nitrate transporter gene (Rmet_4817), lie immediately upstream of the NAS genes (Rmet_4820/1). Theoretically, full denitrification by the successive reduction of nitrate to molecular nitrogen seems possible in CH34 because of three key genes. They are the *nirS* gene (Rmet_3172) encoding a NO-forming nitrite-reductase (NNR), the *norB* gene (Rmet_3167) encoding a nitric-oxide reductase (NOR), and a *nosZ* gene (Rmet_4917) coding for a nitrous-oxide reductase (NOS). These three are centered among additional genes that might be required for functional denitrification: (i) The *nirENJ* (Rmet_3168/69/70) and *nirCFGH* genes (Rmet_2079/80/81, RALME2203), (ii) the *norR* gene (Rmet_3166) encoding a $\sigma^{54}$ dependent transcriptional regulator, and, (iii) the NOS accessory genes *nosLYFDR* (Rmet_4912/3/4/5/6) and *nosCX* (Rmet_4918/9). A regulatory two-component system NarX-NarL that may be important for nitrate- or nitrite- controlled transcription is encoded by Rmet_2086/7.

In view of the ability of strain CH34 to metabolize anaerobically and its potential for full denitrification, we note that it has all the genes for the three known classes of ribonucleotide reductase (RNRase): NrdAB (Rmet_3087/8), NrdJ (Rmet_2130), and NrdDG (Rmet_5137, Rmet_5139), along with NrdR (Rmet_2681) as a transcriptional repressor of these genes. According to the RNRase database (http://rnrdb.molbio.su.se/) less than 8% of all bacteria have all three classes of RNRase: *C. eutrophus* H16 and *C. metallidurans* CH34 are among the bacteria with these genes, but *C. pinatubonensis* JMP134 and *C. taiwanensis* lack the *nrdDG* genes. Our identification of *nrdDG* genes in strain CH34 is important because the NrdDG enzyme, which operates only under anaerobic conditions, is essential for complete denitrification in *C. eutrophus* H16 [168]. As the latter's *nrdDG* genes lie on the pHG1 plasmid, a pHG1-free mutant strain grows very poorly on nitrate and does not achieve full denitrification. This defect in the mutant is countered by adding cobalamin (VitB~12~) to the medium [169], most likely because the coenzyme B~12~-dependent ClassII RNRase, NrdJ, assumes the role of NrdDG, while at the same time *C. eutrophus* H16 readily takes up and assimilates cobalamin. Likewise, *C. metallidurans* CH34 encodes most, if not all genes for the uptake and assimilation of cobalamin (see next section). Together, these facts strongly indicate that anaerobic metabolism, with nitrate as the terminal electron-acceptor, runs through very similar pathways in strains H16 and CH34.

## Vitamin biosynthesis

*C. metallidurans* CH34 has multiple genes for assimilating a variety of B family vitamins, or their *de novo* production, including thiamin (VitB1), riboflavin (VitB2), pantothenic acid (VitB5), pyridoxal 5'-phosphate (VitB6), biotin (VitB7), folic acid (VitB9), and cobalamin (VitB12). These vitamins, important cofactors for synthesizing fatty acids, proteins, carbohydrates, and branched-chain amino acids, aid in electron-relay systems and energy production. In our genomic survey for vitamin-related genes we did not find any that are relevant to the biosynthetic pathways of retinol (VitA), niacin (VitB3), ascorbic acid (VitC), calciferol (VitD), tocopherol (VitE), or phylloquinone (VitK).

Thiamin (VitB1) biosynthesis is likely governed in *C. metallidurans* CH34 by the *thiCOSGED* locus (Rmet_0162/6, Rmet_0169). This set of genes is needed to generate the thiazole- and pyrimidine-

<figure><img src="image_28.png" /></figure>
CoA and acetyl-CoA. Five more genes of this operon (Rmet_1226 through Rmet_1230) probably encode an ABC-like transport system of benzoate.

Similarly, β-oxidation may play an important role in the formation of acetyl-CoA from dicarboxylated compounds because reportedly, *C. metallidurans* CH34 grows on a range of dicarboxy-lated compounds [9]. However, without experimental evidence, it is difficult to predict the genes and enzymes that would be involved in these conversions in CH34. An interesting report by Harrison and Harwood [167] implicated the *pimFABCDE* operon for the β-oxidative degradation of C~7~-C~14~ dicarboxylic- and fatty-acids in *Rhodopseudomonas palustris*. This operon encodes a set of acyl-CoA ligase, enoyl-CoA hydratase-, acyl-CoA dehydrogenase-, and acyl-CoA transferase-enzymes, and is accompanied by a IclR-type regulatory gene, and several genes for ABC-like transport. Our search through the MaGe annotated data of CH34 yielded more than a hundred of these enzymes. Though most of them might degrade fatty acids, we believe that in CH34 many others are recruited to specifically degrade dicarboxylates.

## Aerobic- and anaerobic-respiration

The citric-acid cycle is followed by oxidative phosphorylation in which NADH, FADH~2~, and GTP are reduced by cytochromes and other terminal oxidases in the cell's inner membrane, while protons are expelled into the periplasm. The resulting proton gradient across the membrane then drives ATP synthesis by the conversion of ADP (ATPase; α, β, γ, δ, ε, a, b, and c units encoded by Rmet_3493 through Rmet_3500). A key locus in this aerobic energy metabolism is a 14-gene cluster located on CHR1 (Rmet_0927 through Rmet_0940) encoding a large membrane-bound complex of NADH dehydrogenase (NADH:quinone oxidoreductase) that is highly conserved within the four sequenced *Cupriavidus* strains, and in other closely related species. According to BioCyc pathway analysis, alternative NADH dehydrogenases may exist in *C. metallidurans*. These are encoded by Rmet_2623 as a single-subunit FAD-binding NDHase, by Rmet_0177 as a NAD-dependent epimerase/dehydratase, and by Rmet_4310 as a FAD-dependent pyridine nucleotide-disulphide oxidoreductase. We also discerned the Rmet_4300 gene that apparently codes for a NuoF homologue (NADH ubiquinone oxidoreductase, chain F). Other membrane-associated protein complexes that may be important to the electron transport chain in strain CH34 are succinate:quinone reductase (SQR; see previous section on the TCA cycle), a *bc*~f~-type cytochrome complex (encoded by Rmet_3228/9/30), a *cbb*~3~-type cytochrome c oxidase (encoded by *cco* genes Rmet_2041/2/3/4), two cytochrome *bd*-type quinol oxidases CydAB (encoded by the genepairs Rmet_4955/6 and Rmet_5232/3), a *aa*~3~-type oxidase encoded by the *cta*-gene cluster (Rmet_0261 through Rmet_0271), a *bb*~3~-type cytochrome oxidase encoded by the *cox*-gene cluster (Rmet_3955/6/7/8/9), and three quinol oxidases encoded by the clusters *cyo*~1~ (Rmet_0948/49/50/51), *cyo*~2~ (Rmet_5608/09/10/11), and *cyo*~3~ (Rmet_5790/1/2/3).

Under anoxic conditions, *C. metallidurans* CH34 may resort to alternate modes of energy generation by using electron acceptors other than oxygen. Thus, the bacterium reduces nitrate [32], which is reflected by three loci encoding distinct types of nitrate reductases: a cytoplasmic assimilatory enzyme (NAS; encoded by the *nasAB* genes Rmet_4820/1), a membrane-bound quinol-oxidizing enzyme (NAR; encoded by the *narGHJ1* genes Rmet_2074 through Rmet_2077, accompanied by two ABC-type transporter-genes Rmet_2072/3), and a periplasmic enzyme (NAP; encoded by the *napEDABC* genes Rmet_4061 through Rmet_4065). Probably, the NAS-, NAP-, and NAR- systems have different physiological roles depending on metabolic conditions.

Consequently, their pathways may be interconnected to foster rapid adaptation to changing nitrogen and/or oxygen conditions, a big asset for survival in nature.

Although strain CH34 reportedly does not reduce nitrite [32], it contains two genes Rmet_4018/9 encoding the large- and small-subunits of a putative assimilatory nitrite-reductase (ANR). These genes, preceded by a nitrate transporter gene (Rmet_4817), lie immediately upstream of the NAS genes (Rmet_4820/1). Theoretically, full denitrification by the successive reduction of nitrate to molecular nitrogen seems possible in CH34 because of three key genes. They are the *nirS* gene (Rmet_3172) encoding a NO-forming nitrite-reductase (NNR), the *norB* gene (Rmet_3167) encoding a nitric-oxide reductase (NOR), and a *nosZ* gene (Rmet_4917) coding for a nitrous-oxide reductase (NOS). These three are centered among additional genes that might be required for functional denitrification: (i) The *nirENJ* (Rmet_3168/69/70) and *nirCFGH* genes (Rmet_2079/80/81, RALME2203), (ii) the *norR* gene (Rmet_3166) encoding a $\sigma^{54}$ dependent transcriptional regulator, and, (iii) the NOS accessory genes *nosLYFDR* (Rmet_4912/3/4/5/6) and *nosCX* (Rmet_4918/9). A regulatory two-component system NarX-NarL that may be important for nitrate- or nitrite- controlled transcription is encoded by Rmet_2086/7.

In view of the ability of strain CH34 to metabolize anaerobically and its potential for full denitrification, we note that it has all the genes for the three known classes of ribonucleotide reductase (RNRase): NrdAB (Rmet_3087/8), NrdJ (Rmet_2130), and NrdDG (Rmet_5137, Rmet_5139), along with NrdR (Rmet_2681) as a transcriptional repressor of these genes. According to the RNRase database (http://rnrdb.molbio.su.se/) less than 8% of all bacteria have all three classes of RNRase: *C. eutrophus* H16 and *C. metallidurans* CH34 are among the bacteria with these genes, but *C. pinatubonensis* JMP134 and *C. taiwanensis* lack the *nrdDG* genes. Our identification of *nrdDG* genes in strain CH34 is important because the NrdDG enzyme, which operates only under anaerobic conditions, is essential for complete denitrification in *C. eutrophus* H16 [168]. As the latter's *nrdDG* genes lie on the pHG1 plasmid, a pHG1-free mutant strain grows very poorly on nitrate and does not achieve full denitrification. This defect in the mutant is countered by adding cobalamin (VitB~12~) to the medium [169], most likely because the coenzyme B~12~-dependent ClassII RNRase, NrdJ, assumes the role of NrdDG, while at the same time *C. eutrophus* H16 readily takes up and assimilates cobalamin. Likewise, *C. metallidurans* CH34 encodes most, if not all genes for the uptake and assimilation of cobalamin (see next section). Together, these facts strongly indicate that anaerobic metabolism, with nitrate as the terminal electron-acceptor, runs through very similar pathways in strains H16 and CH34.

## Vitamin biosynthesis

*C. metallidurans* CH34 has multiple genes for assimilating a variety of B family vitamins, or their *de novo* production, including thiamin (VitB1), riboflavin (VitB2), pantothenic acid (VitB5), pyridoxal 5'-phosphate (VitB6), biotin (VitB7), folic acid (VitB9), and cobalamin (VitB12). These vitamins, important cofactors for synthesizing fatty acids, proteins, carbohydrates, and branched-chain amino acids, aid in electron-relay systems and energy production. In our genomic survey for vitamin-related genes we did not find any that are relevant to the biosynthetic pathways of retinol (VitA), niacin (VitB3), ascorbic acid (VitC), calciferol (VitD), tocopherol (VitE), or phylloquinone (VitK).

Thiamin (VitB1) biosynthesis is likely governed in *C. metallidurans* CH34 by the *thiCOSGED* locus (Rmet_0162/6, Rmet_0169). This set of genes is needed to generate the thiazole- and pyrimidine-

<figure><img src="image_28.png" /></figure>
moieties of the vitamin as well as to control the condensation reaction of both moieties to form thiamin phosphate. Additional genes involved in thiamin biosynthesis are *dxs* (Rmet_2615), *icsS* (Rmet_1025), and *thiL* (Rmet_3047) encoding an oxidase and catalytic enzymes. Two more kinases, ThiF and ThiK, usually are involved in thiamin synthesis but we identified only three putative *thiF* genes (Rmet_0249 and Rmet_2639, both located on CHR1, and RALMEp10091, present on the pMOL28 plasmid); we did not recognize the *thiK* gene.

The biosynthesis of riboflavin (VitB2) in CH34 might be linked to the genes *ribDC* (Rmet_2688/9), *rib(BA1)*E (Rmet_2693/4), *ribA1* (Rmet_0194), *ribF* (Rmet_2884), *ribA2* (Rmet 4572), and *ribB* (Rmet_4758). No other *rib* genes were detected.

Besides its involvement in energy metabolism, pantothenic acid (VitB5) is regarded as a crucial component of coenzyme A biosynthesis, starting from β-alanine and L-valine. In *C. metallidurans* CH34, the synthesis of VitB5 could be connected to genes *ilvE* (Rmet_0493), *panB* (Rmet_2774), *panC* (Rmet_2917), and two copies of *panE* (Rmet_4536, Rmet_5770).

The biosynthesis of pyridoxal 5'-phosphate (VitB6) in this strain proceeds via two complementary pathways. The biosynthetic pathway involves the genes *pdxC* (*serC*; Rmet_0715), *pdxA* (present in two copies Rmet_0438, Rmet_2590), a gene encoding an FMN-binding protein (Rmet_0820), *pdxJ* (Rmet_2415), *dxs* (Rmet_2615; also involved in VitB1 synthesis, see above), *pdxH* (Rmet_2642), and a putative pyridoxine oxidase gene (Rmet_4050). We did not detect the genes involved in generating 2-oxo-3-hydroxy-4-phosphobutanoate, an early step in VitB6 synthesis and the actual substrate of the SerC catalytic action. The second pathway, the salvage pathway, basically converts pyridoxine, pyridoxamine, or pyridoxal, into pyridoxal-5'-phosphate, the active form of VitB6. This involves only two genes *pdxH* (Rmet_2642) and *pdxK* (Rmet_4114). We note that the former also is part of the *de novo* biosynthetic pathway (above).

The *bioAFDB* locus (Rmet_0114/5/6/7) in CH34 controls the production of biotin (VitB7). There are slight variations in the pathway, depending on whether pimelate or pimeloyl-CoA is the starting substrate; nonetheless, the geneproducts of all four genes are involved in both. Pimeloyl-CoA may be derived from the conversion of aromatic compounds via the intermediates benzoyl-CoA and 3-hydroxypimelyl-CoA, while the direct use of pimelate necessitates a fifth gene, *pauA* (pimelic acid-utilizing A; Rmet_2598). The *pauA* gene codes for a 6-carboxyhexanoate-CoA ligase (often referred to as a 'pimeloyl-CoA synthetase'), essentially synthesising pimeloyl-CoA from pimelate and coenzyme A.

The variant pathways and intermediate reactions leading to the formation of folate (VitB9) from chorismate and GTP are complex and behind the scope of our genome study. We identified several genes that might be involved in folate biosynthesis in *C. metallidurans* CH34; they are scattered throughout the genome and linkage between individual ones is blurred. These genes are *folB* (Rmet_0183), *folD* (Rmet_1192), *purT* (Rmet_1820), *folP* (Rmet_2187), *folC* (Rmet_2463), *folA* (Rmet_2569), *purN* (Rmet_2878), *folK* (Rmet_2915), *trpG* (Rmet_3179), and a putative *folE* (Rmet_3990). We failed to identify a hydrolase gene for converting 7,8-dihydroneopterin triphosphate into 7,8-dihydro-D-neopterin.

There have been extensive studies on cobalamin, a vitamin of a particular interest in public health and medical sciences, and produced only by microorganisms [170]: indeed, its biosynthesis is one of the best known vitamin pathways to date [171]. Two pathways for corrin ring formation were described in bacteria, differing from each other in the way cobalt is incorporated [172].

In *C. metallidurans* CH34, the presence of a sirohydrochlorin cobaltochelatase gene *cbiX* (Rmet_2810) suggests that this organism synthesises cobalamin anaerobically, thereby including an early insertion step of cobalt. Genes involved in the late stage of the biosynthesis are grouped in a large operon of 13 genes (entailing *cbi*, *cob* and *btu* genes Rmet_2777 through Rmet_2789) with a cobalamin riboswitch we identified at the far end (Rmet_R0047). This entire operon, which according to the MaGe interface displays good synteny with all *Cupriavidus* and *Ralstonia* genomes, contains all the essential genes for producing adenosylcobalamin from cobyrinic acid (including those controlling corrinoid transport, viz. *btuB*, *btuC*, *btuD*, and *btuF*). Only the *bluB* gene (Rmet_2213), coding for a cobalt reductase, is at a different location on CHR1. Whilst the early and late stages of the biosynthesis generally are covered by these listed genes, other genes involved in the intermediate corrin biosynthesis in *C. metallidurans* await elucidation.

## Autotrophic metabolism

In the absence of heterotrophic carbon sources, *C. metallidurans* CH34 can switch to an autotrophic lifestyle by fixing and reducing carbon dioxide for synthesizing organic molecules. Generally, autotrophic prokaryotes have four different biochemical routes to accomplish this (reviewed by Atomi [173]): (i) The reductive pentose-phosphate pathway; (ii) the reductive acetyl-CoA pathway; (iii) the 3-hydroxypropionate cycle; and, (iv) the reductive tricarboxylic-acid (rTCA) cycle.

The *C. metallidurans* CH34 genome encodes all enzymes necessary for the 13 biochemical reactions of the reductive pentose phosphate pathway, also known as the Calvin-Benson-Bassham (CBB) cycle. Figure S6 is a schematic view of the *cbb* loci in CH34, using Tabita *et al.* 's gene notations [174]. We note especially that this region contains some IS elements (marked in red by their *tnpA* transposase genes in Figure S6), including two exterior IS*1071* elements, Rmet_1491 and Rmet_1544. This may explain the earlier observed loss of autotrophic growth in CH34 under mutagenic stress [22] since the IS*1071*-mediated excision of this region would remove, amongst other genes, the two structural genes *cbbL* (Rmet_1501) and *cbbS* (Rmet_1500) of RubisCo (ribulose 1-5 bifosfaat carboxylase oxygenase) as well as of the accessory genes *cbbQ* (Rmet_1499) and *cbbO* (Rmet_1498) needed to assemble RubisCo.

Several regulators control the CBB cycle. First, there are three copies of *cbbR*, a LysR-type transcriptional activator of *cbb* gene expression. Two copies juxtapose the central *cbb* gene clusters, *cbbR1* (Rmet_1502) and *cbbR2* (Rmet_1508), while a third copy, *cbbR3* (Rmet_4365), is located on the second replicon *CHR2* (unlike other *cbb* genes that all are sited on CHR1). Furthermore, the *C. metallidurans* CH34 genome harbours a set of *regA* (Rmet_0135) and *regB* (Rmet_0136) genes probably regulating carbon fixation via the CBB-cycle, as in other organisms [173,175]. Remarkably, three non-essential *cbb*-genes appear to be transcriptionally linked to the *cbb* operon: *cbbY* (Rmet_1517), *cbbZ1* (Rmet_1514), and *cbbZ2* (Rmet_3177). Under certain conditions, autotrophic bacteria carrying these exceptional *cbb* genes display a phosphoglycolate phosphatase (PGP) enzyme activity [176] that may help cells to cope with toxic levels of phosphoglycolate accumulated by the oxygenase reaction of RubisCo whilst the carbon skeleton of phosphoglycolate is salvaged as glycolate for further use in glycine synthesis. We did not detect a *cbbX* gene in the *C. metallidurans* CH34 genome.

Besides the CBB cycle, *C. metallidurans* CH34 apparently has functional genes to fix carbon dioxide via the reductive acetyl

<figure><img src="image_29.png" /></figure>
moieties of the vitamin as well as to control the condensation reaction of both moieties to form thiamin phosphate. Additional genes involved in thiamin biosynthesis are *dxs* (Rmet_2615), *icsS* (Rmet_1025), and *thiL* (Rmet_3047) encoding an oxidase and catalytic enzymes. Two more kinases, ThiF and ThiK, usually are involved in thiamin synthesis but we identified only three putative *thiF* genes (Rmet_0249 and Rmet_2639, both located on CHR1, and RALMEp10091, present on the pMOL28 plasmid); we did not recognize the *thiK* gene.

The biosynthesis of riboflavin (VitB2) in CH34 might be linked to the genes *ribDC* (Rmet_2688/9), *rib(BA1)*E (Rmet_2693/4), *ribA1* (Rmet_0194), *ribF* (Rmet_2884), *ribA2* (Rmet 4572), and *ribB* (Rmet_4758). No other *rib* genes were detected.

Besides its involvement in energy metabolism, pantothenic acid (VitB5) is regarded as a crucial component of coenzyme A biosynthesis, starting from β-alanine and L-valine. In *C. metallidurans* CH34, the synthesis of VitB5 could be connected to genes *ilvE* (Rmet_0493), *panB* (Rmet_2774), *panC* (Rmet_2917), and two copies of *panE* (Rmet_4536, Rmet_5770).

The biosynthesis of pyridoxal 5'-phosphate (VitB6) in this strain proceeds via two complementary pathways. The biosynthetic pathway involves the genes *pdxC* (*serC*; Rmet_0715), *pdxA* (present in two copies Rmet_0438, Rmet_2590), a gene encoding an FMN-binding protein (Rmet_0820), *pdxJ* (Rmet_2415), *dxs* (Rmet_2615; also involved in VitB1 synthesis, see above), *pdxH* (Rmet_2642), and a putative pyridoxine oxidase gene (Rmet_4050). We did not detect the genes involved in generating 2-oxo-3-hydroxy-4-phosphobutanoate, an early step in VitB6 synthesis and the actual substrate of the SerC catalytic action. The second pathway, the salvage pathway, basically converts pyridoxine, pyridoxamine, or pyridoxal, into pyridoxal-5'-phosphate, the active form of VitB6. This involves only two genes *pdxH* (Rmet_2642) and *pdxK* (Rmet_4114). We note that the former also is part of the *de novo* biosynthetic pathway (above).

The *bioAFDB* locus (Rmet_0114/5/6/7) in CH34 controls the production of biotin (VitB7). There are slight variations in the pathway, depending on whether pimelate or pimeloyl-CoA is the starting substrate; nonetheless, the geneproducts of all four genes are involved in both. Pimeloyl-CoA may be derived from the conversion of aromatic compounds via the intermediates benzoyl-CoA and 3-hydroxypimelyl-CoA, while the direct use of pimelate necessitates a fifth gene, *pauA* (pimelic acid-utilizing A; Rmet_2598). The *pauA* gene codes for a 6-carboxyhexanoate-CoA ligase (often referred to as a 'pimeloyl-CoA synthetase'), essentially synthesising pimeloyl-CoA from pimelate and coenzyme A.

The variant pathways and intermediate reactions leading to the formation of folate (VitB9) from chorismate and GTP are complex and behind the scope of our genome study. We identified several genes that might be involved in folate biosynthesis in *C. metallidurans* CH34; they are scattered throughout the genome and linkage between individual ones is blurred. These genes are *folB* (Rmet_0183), *folD* (Rmet_1192), *purT* (Rmet_1820), *folP* (Rmet_2187), *folC* (Rmet_2463), *folA* (Rmet_2569), *purN* (Rmet_2878), *folK* (Rmet_2915), *trpG* (Rmet_3179), and a putative *folE* (Rmet_3990). We failed to identify a hydrolase gene for converting 7,8-dihydroneopterin triphosphate into 7,8-dihydro-D-neopterin.

There have been extensive studies on cobalamin, a vitamin of a particular interest in public health and medical sciences, and produced only by microorganisms [170]: indeed, its biosynthesis is one of the best known vitamin pathways to date [171]. Two pathways for corrin ring formation were described in bacteria, differing from each other in the way cobalt is incorporated [172].

In *C. metallidurans* CH34, the presence of a sirohydrochlorin cobaltochelatase gene *cbiX* (Rmet_2810) suggests that this organism synthesises cobalamin anaerobically, thereby including an early insertion step of cobalt. Genes involved in the late stage of the biosynthesis are grouped in a large operon of 13 genes (entailing *cbi*, *cob* and *btu* genes Rmet_2777 through Rmet_2789) with a cobalamin riboswitch we identified at the far end (Rmet_R0047). This entire operon, which according to the MaGe interface displays good synteny with all *Cupriavidus* and *Ralstonia* genomes, contains all the essential genes for producing adenosylcobalamin from cobyrinic acid (including those controlling corrinoid transport, viz. *btuB*, *btuC*, *btuD*, and *btuF*). Only the *bluB* gene (Rmet_2213), coding for a cobalt reductase, is at a different location on CHR1. Whilst the early and late stages of the biosynthesis generally are covered by these listed genes, other genes involved in the intermediate corrin biosynthesis in *C. metallidurans* await elucidation.

## Autotrophic metabolism

In the absence of heterotrophic carbon sources, *C. metallidurans* CH34 can switch to an autotrophic lifestyle by fixing and reducing carbon dioxide for synthesizing organic molecules. Generally, autotrophic prokaryotes have four different biochemical routes to accomplish this (reviewed by Atomi [173]): (i) The reductive pentose-phosphate pathway; (ii) the reductive acetyl-CoA pathway; (iii) the 3-hydroxypropionate cycle; and, (iv) the reductive tricarboxylic-acid (rTCA) cycle.

The *C. metallidurans* CH34 genome encodes all enzymes necessary for the 13 biochemical reactions of the reductive pentose phosphate pathway, also known as the Calvin-Benson-Bassham (CBB) cycle. Figure S6 is a schematic view of the *cbb* loci in CH34, using Tabita *et al.* 's gene notations [174]. We note especially that this region contains some IS elements (marked in red by their *tnpA* transposase genes in Figure S6), including two exterior IS*1071* elements, Rmet_1491 and Rmet_1544. This may explain the earlier observed loss of autotrophic growth in CH34 under mutagenic stress [22] since the IS*1071*-mediated excision of this region would remove, amongst other genes, the two structural genes *cbbL* (Rmet_1501) and *cbbS* (Rmet_1500) of RubisCo (ribulose 1–5 bifosfaat carboxylase oxygenase) as well as of the accessory genes *cbbQ* (Rmet_1499) and *cbbO* (Rmet_1498) needed to assemble RubisCo.

Several regulators control the CBB cycle. First, there are three copies of *cbbR*, a LysR-type transcriptional activator of *cbb* gene expression. Two copies juxtapose the central *cbb* gene clusters, *cbbR1* (Rmet_1502) and *cbbR2* (Rmet_1508), while a third copy, *cbbR3* (Rmet_4365), is located on the second replicon *CHR2* (unlike other *cbb* genes that all are sited on CHR1). Furthermore, the *C. metallidurans* CH34 genome harbours a set of *regA* (Rmet_0135) and *regB* (Rmet_0136) genes probably regulating carbon fixation via the CBB-cycle, as in other organisms [173,175]. Remarkably, three non-essential *cbb*-genes appear to be transcriptionally linked to the *cbb* operon: *cbbY* (Rmet_1517), *cbbZ1* (Rmet_1514), and *cbbZ2* (Rmet_3177). Under certain conditions, autotrophic bacteria carrying these exceptional *cbb* genes display a phosphoglycolate phosphatase (PGP) enzyme activity [176] that may help cells to cope with toxic levels of phosphoglycolate accumulated by the oxygenase reaction of RubisCo whilst the carbon skeleton of phosphoglycolate is salvaged as glycolate for further use in glycine synthesis. We did not detect a *cbbX* gene in the *C. metallidurans* CH34 genome.

Besides the CBB cycle, *C. metallidurans* CH34 apparently has functional genes to fix carbon dioxide via the reductive acetyl

<figure><img src="image_29.png" /></figure>
coenzyme A pathway (also termed the Wood-Ljungdahl pathway). One key biochemical reaction in this pathway is the transition between carbon dioxide and carbon monoxide mediated by a metalloenzyme, CO dehydrogenase (CODH). This enzyme is a dimeric heterotrimer consisting of a small, medium, and large subunit representing respectively, an iron-sulfur protein (CoxS), a flavoprotein (CoxM), and a molybdoprotein (CoxL). Two forms of aerobic CO-DH are known: Form I was characterized explicitly for its ability to oxidize carbon monoxide, but the function of form II CODH, distinct from form I but phylogenetically closely related, remains unclear [177]. Strain CH34 carries the genes *coxSLM* (Rmet_0365 – Rmet_0363) and the accessory genes *coxDEGI* (Rmet_0362 – Rmet_0359). The order of the CODH genes in this operonic structure and the presence of an AYRGAGR motif in CoxL (encoded by Rmet_0364) strongly suggest that the *C. metallidurans* 'CODH is of form II. Although the ability of *C. metallidurans* CH34 to undertake substantial oxidation of carbon monoxide still needs to be tested, CODH-II-encoding *cox* genes were found in *Bradyrhizobium japonicum* USDA 110, an organism relying solely on CODH-II for CO oxidation and hence, growth on carbon monoxide as a sole carbon- and energy- source [178].

Based on our genomic survey and supported by KEGG analysis, *C. metallidurans* CH34 lacks the cycle-specific malonyl-CoA reductase (EC:4.1.1.9), an essential enzyme for the 3-hydroxypropionate cycle [179,180], and hence, cannot employ this pathway to assimilate carbon dioxide. Likewise, the *C. metallidurans* CH34 genome does not code for the key enzymes of a rTCA cycle, i.e., ATP-citrate lyase and 2-oxoglutarate synthase nor does it encode enzymes for alternative formation of oxaloacetate, for example, via the combined action of citryl-CoA synthetase and citryl-CoA lyase, thus effectively ruling out the rTCA-cycle as a route for fixing carbon dioxide.

From this genomic information it appears that *C. metallidurans* CH34 relies primarily on a RubisCO Type I enzyme for fixing carbon dioxide as part of a complete CBB-cycle. Although an intact CODH-II based system is present, it is yet unclear whether strain CH34 can use CO as a sole energy and carbon source.

## Sulfur oxidation (sulfur autotrophy)

*C. metallidurans* CH34 has the genetic capacity to oxidize inorganic sulfur compounds by the action of sulfur oxidizing genes (*sox*) and sulfur oxygenase reductase genes (*sor*) (Figure S7A) [181]. The first set of genes comprises the *soxBWXAζIFE* gene cluster situated on CHR1 as Rmet_3417 through Rmet_3427. Basically, sulfur respiration requires only five *sox* genes whose products form three key periplasmic protein complexes: SoxYZ, a sulfur carrier protein, SoxXA, a c-type cytochrome complex, and SoxB, a sulfate thiol hydrolase. In this system, the oxidation of thiosulfate (*S*₂*O*₃²⁻⁻) to sulfate entails the oxidative linkage, mediated by SoxXA, of thiosulfate to the conserved C-terminal cysteine of SoxY, followed by hydrolysis of the terminal sulfone (*SO*₃⁻⁻) group by SoxB (Figure S7B). The SoxXA complex then covalently binds other pathway intermediates to the SoxY cysteine thus forming extended chains of sulfane (*S*⁻⁻) groups preceding the terminal sulfone group (*SO*₃⁻⁻) [182]. These sulfane chains are broken by the iterated actions of the SoxB hydrolase and a periplasmic sulfur dehydrogenase, SoxCD. Although *C. metallidurans* CH34 lacks the *soxCD* genes, this would not impair its ability to oxydise thiosulfate to sulfate in the first step of the cycle. However, fewer electrons would be liberated compared to the electron generation of complete *sox* systems.

In *C. metallidurans* CH34, the likely route for sulfide (*HS*⁻) oxidation would be via the *soxE* and *soxF* gene products, so forming a typical c-type flavocytochrome complex in which *SoxF* functions as a sulfide dehydrogenase (Figure S7C). Lastly, the *C. metallidurans* *sox* gene cluster includes two genes of unknown function coding for a conserved hypothetical protein (Rmet_3421), putatively exported, and an unknown c-type cytochrome precursor (Rmet_3424).

*C. metallidurans* probably oxidizes sulfites (*SO*₃⁻⁻) through the action of the *sorA* and *sorB* genes (present on CHR2 as Rmet_4891 and Rmet_4892) (Figure S7D) that are orthologues of the *C. eutrophus* H16 *sorA* and *sorB* genes encoding, respectively, a periplasmic sulfite dehydrogenase and a membrane-bound c-type cytochrome [71].

These routes of sulfur oxidation (*SoxB*-XA-YZ, *SoxEF*, and *SorAB*) in combination with an autotrophic lifestyle would enable *C. metallidurans* to use reduced sulfur compounds as electron donors, while employing carbon dioxide as an electron acceptor, and hence it could function as a true chemolithotrophic sulfur-oxidizing bacterium. Such a sulfur-based growth is sustainable only when sufficient amounts of thiosulfate ions are present. A biological source for them is from the fermentation of taurine (2-aminoethanesulfonate), together with the potential fermentation of other organosulfonates in anoxic environments [183]. Interestingly, *C. metallidurans* thrives not only on thiosulfate but also on a wide range of aromatic sulfonates, alkanesulfonates, and sulfate esters [184]. Therefore, it is not surprising that the *C. metallidurans* CH34 genome encodes alkanesulfonate- and taurine-specific ABC transporter systems, such as SsuABC (Rmet_1370/2/3) and TauABC (Rmet_2857/8/9), an alkanesulfate monooxygenase SsuD (Rmet_1371), at least six copies of taurine dioxygenase TauD (encoded by Rmet genes _4136, _4409, _4633, _5119, _5292, and _5823), a reductase-ferredoxin AsfA/AsfB (Rmet_4334/5), and a putative NAD(P)H-dependent FMN reductase SsuE (Rmet_5075) that with AsfA/AsfB may form an electron chain for aromatic desulfonation. We did not identify other genes for enzymatic desulfonation from the *C. metallidurans* CH34 genome data.

## Hydrogen oxidation (hydrogenotrophy)

*C. metallidurans* CH34 can gain its energy from oxidising hydrogen gas (*H*₂) into electrons (e⁻) and protons (*H*⁺). The underlying enzymatic engine has two parts: A periplasmic membrane-bound hydrogenase (MBH), and a cytosolic soluble hydrogenase (SH). This allows the bacterium to switch between an *H*₂-driven energy supply and a classic lithotrophic growth mode. In contrast to the *H*₂-oxidising *C. eutrophus* H16, the expression of MBH and SH in *C. metallidurans* CH34 is not controlled by an *H*₂-sensor in the form of a third hydrogenase. Therefore, the nearby concentration of hydrogen gas does not regulate the lithotrophic growth of CH34. Nonetheless, the lack of a specific *H*₂-sensor-hydrogenase does not entail the constitutive expression of the energy-related hydrogenases, such as in the absence of *H*₂. Rather, both the hydrogenases and the RubisCo enzyme (see autotrophic metabolism) are regulated by the carbon source afforded by the growth substrate [9].

All the genes needed to form and activate MBH and SH are grouped in two large gene clusters situated on CHR1 (Figure S8A). The first cluster, governing MBH, harbors the hydrogenase pleiotropic operon encompassing six genes hypE₁D₁C₁F₁B₁A₁ (Rmet_1281 – Rmet_1286), followed by a set of genes required for hydrogen oxidation, hoxVTRQOLMζGK (Rmet_1287 through Rmet_1293, Rmet_1295, and Rmet_1297-98). This entire cluster is located on genomic island CMGI-2. A second cluster, governing

<figure><img src="image_30.png" /></figure>
coenzyme A pathway (also termed the Wood-Ljungdahl pathway). One key biochemical reaction in this pathway is the transition between carbon dioxide and carbon monoxide mediated by a metalloenzyme, CO dehydrogenase (CODH). This enzyme is a dimeric heterotrimer consisting of a small, medium, and large subunit representing respectively, an iron-sulfur protein (CoxS), a flavoprotein (CoxM), and a molybdoprotein (CoxL). Two forms of aerobic CO-DH are known: Form I was characterized explicitly for its ability to oxidize carbon monoxide, but the function of form II CODH, distinct from form I but phylogenetically closely related, remains unclear [177]. Strain CH34 carries the genes *coxSLM* (Rmet_0365 – Rmet_0363) and the accessory genes *coxDEGI* (Rmet_0362 – Rmet_0359). The order of the CODH genes in this operonic structure and the presence of an AYRGAGR motif in CoxL (encoded by Rmet_0364) strongly suggest that the *C. metallidurans* 'CODH is of form II. Although the ability of *C. metallidurans* CH34 to undertake substantial oxidation of carbon monoxide still needs to be tested, CODH-II-encoding *cox* genes were found in *Bradyrhizobium japonicum* USDA 110, an organism relying solely on CODH-II for CO oxidation and hence, growth on carbon monoxide as a sole carbon- and energy- source [178].

Based on our genomic survey and supported by KEGG analysis, *C. metallidurans* CH34 lacks the cycle-specific malonyl-CoA reductase (EC:4.1.1.9), an essential enzyme for the 3-hydroxypropionate cycle [179,180], and hence, cannot employ this pathway to assimilate carbon dioxide. Likewise, the *C. metallidurans* CH34 genome does not code for the key enzymes of a rTCA cycle, i.e., ATP-citrate lyase and 2-oxoglutarate synthase nor does it encode enzymes for alternative formation of oxaloacetate, for example, via the combined action of citryl-CoA synthetase and citryl-CoA lyase, thus effectively ruling out the rTCA-cycle as a route for fixing carbon dioxide.

From this genomic information it appears that *C. metallidurans* CH34 relies primarily on a RubisCO Type I enzyme for fixing carbon dioxide as part of a complete CBB-cycle. Although an intact CODH-II based system is present, it is yet unclear whether strain CH34 can use CO as a sole energy and carbon source.

## Sulfur oxidation (sulfur autotrophy)

*C. metallidurans* CH34 has the genetic capacity to oxidize inorganic sulfur compounds by the action of sulfur oxidizing genes (*sox*) and sulfur oxygenase reductase genes (*sor*) (Figure S7A) [181]. The first set of genes comprises the *soxBWXAζIFE* gene cluster situated on CHR1 as Rmet_3417 through Rmet_3427. Basically, sulfur respiration requires only five *sox* genes whose products form three key periplasmic protein complexes: SoxYZ, a sulfur carrier protein, SoxXA, a c-type cytochrome complex, and SoxB, a sulfate thiol hydrolase. In this system, the oxidation of thiosulfate (*S*₂*O*₃²⁻⁻) to sulfate entails the oxidative linkage, mediated by SoxXA, of thiosulfate to the conserved C-terminal cysteine of SoxY, followed by hydrolysis of the terminal sulfone (*SO*₃⁻⁻) group by SoxB (Figure S7B). The SoxXA complex then covalently binds other pathway intermediates to the SoxY cysteine thus forming extended chains of sulfane (*S*⁻⁻) groups preceding the terminal sulfone group (*SO*₃⁻⁻) [182]. These sulfane chains are broken by the iterated actions of the SoxB hydrolase and a periplasmic sulfur dehydrogenase, SoxCD. Although *C. metallidurans* CH34 lacks the *soxCD* genes, this would not impair its ability to oxydise thiosulfate to sulfate in the first step of the cycle. However, fewer electrons would be liberated compared to the electron generation of complete *sox* systems.

In *C. metallidurans* CH34, the likely route for sulfide (*HS*⁻) oxidation would be via the *soxE* and *soxF* gene products, so forming a typical c-type flavocytochrome complex in which *SoxF* functions as a sulfide dehydrogenase (Figure S7C). Lastly, the *C. metallidurans* *sox* gene cluster includes two genes of unknown function coding for a conserved hypothetical protein (Rmet_3421), putatively exported, and an unknown c-type cytochrome precursor (Rmet_3424).

*C. metallidurans* probably oxidizes sulfites (*SO*₃⁻⁻) through the action of the *sorA* and *sorB* genes (present on CHR2 as Rmet_4891 and Rmet_4892) (Figure S7D) that are orthologues of the *C. eutrophus* H16 *sorA* and *sorB* genes encoding, respectively, a periplasmic sulfite dehydrogenase and a membrane-bound c-type cytochrome [71].

These routes of sulfur oxidation (*SoxB*-XA-YZ, *SoxFE*, and *SorAB*) in combination with an autotrophic lifestyle would enable *C. metallidurans* to use reduced sulfur compounds as electron donors, while employing carbon dioxide as an electron acceptor, and hence it could function as a true chemolithotrophic sulfur-oxidizing bacterium. Such a sulfur-based growth is sustainable only when sufficient amounts of thiosulfate ions are present. A biological source for them is from the fermentation of taurine (2-aminoethanesulfonate), together with the potential fermentation of other organosulfonates in anoxic environments [183]. Interestingly, *C. metallidurans* thrives not only on thiosulfate but also on a wide range of aromatic sulfonates, alkanesulfonates, and sulfate esters [184]. Therefore, it is not surprising that the *C. metallidurans* CH34 genome encodes alkanesulfonate- and taurine-specific ABC transporter systems, such as SsuABC (Rmet_1370/2/3) and TauABC (Rmet_2857/8/9), an alkanesulfate monooxygenase SsuD (Rmet_1371), at least six copies of taurine dioxygenase TauD (encoded by Rmet genes _4136, _4409, _4633, _5119, _5292, and _5823), a reductase-ferredoxin AsfA/AsfB (Rmet_4334/5), and a putative NAD(P)H-dependent FMN reductase SsuE (Rmet_5075) that with AsfA/AsfB may form an electron chain for aromatic desulfonation. We did not identify other genes for enzymatic desulfonation from the *C. metallidurans* CH34 genome data.

## Hydrogen oxidation (hydrogenotrophy)

*C. metallidurans* CH34 can gain its energy from oxidising hydrogen gas (*H*₂) into electrons (e⁻) and protons (*H*⁺). The underlying enzymatic engine has two parts: A periplasmic membrane-bound hydrogenase (MBH), and a cytosolic soluble hydrogenase (SH). This allows the bacterium to switch between an *H*₂-driven energy supply and a classic lithotrophic growth mode. In contrast to the *H*₂-oxidising *C. eutrophus* H16, the expression of MBH and SH in *C. metallidurans* CH34 is not controlled by an *H*₂-sensor in the form of a third hydrogenase. Therefore, the nearby concentration of hydrogen gas does not regulate the lithotrophic growth of CH34. Nonetheless, the lack of a specific *H*₂-sensor-hydrogenase does not entail the constitutive expression of the energy-related hydrogenases, such as in the absence of *H*₂. Rather, both the hydrogenases and the RubisCo enzyme (see autotrophic metabolism) are regulated by the carbon source afforded by the growth substrate [9].

All the genes needed to form and activate MBH and SH are grouped in two large gene clusters situated on CHR1 (Figure S8A). The first cluster, governing MBH, harbors the hydrogenase pleiotropic operon encompassing six genes hypE₁D₁C₁F₁B₁A₁ (Rmet_1281 – Rmet_1286), followed by a set of genes required for hydrogen oxidation, *hoxVTRQOLMζGK* (Rmet_1287 through Rmet_1293, Rmet_1295, and Rmet_1297-98). This entire cluster is located on genomic island CMGI-2. A second cluster, governing

<figure><img src="image_30.png" /></figure>
the SH, contains one partial *hox* gene denoted Δ*hoxV* (Rmet_1534), nine intact *hox* genes *hoxFUTHWINXA* (Rmet_1522 through Rmet_1527, Rmet_1533, and Rmet_1541-42), and six genes *hypA*2*B*2*F*2*C*2*D*2*E*2 (Rmet_1535 through Rmet_1540) all on genomic island CMGI-3. The latter group of *hyp* genes (index 2) appears to have arisen from a reverse duplication of the *hyp* operon of the first *hyp* cluster (index 1) on CMGI-2. We note that the *hyp* and *hox* genes for SH are located, together with the *cbb* genes, on a region flanked by IS elements (Figures S6 and S8A) and that the IS-mediated excission of this region would lead to the loss of RubisCo and the entire SH system, thereby impairing not only autotrophic growth but causing the loss of hydrogenotrophic growth as well. Indeed, stress-induced mutants of CH34 lacking this genomic region are unable to utilize hydrogen gas as a source of energy [6].

Besides these hydrogenase gene clusters on CHR1, there are six *hydrogenase 'four' (hyf)* genes grouped together on CHR2 as *hyfBCEFGI* (Rmet_4666 through Rmet_4671). This resembles an incomplete operon that, in *E. coli* K12, exists as a 10-gene cluster encoding a multimeric hydrogenase complex. Presumably, this complex interacts with formate dehydrogenase FdhF to form a hydrogenlyase system cleaving formate to dihydrogen and carbon dioxide [185]. The same six *hyf* genes also occur on the first replicon CHR1 of *C. eutrophus* H16 as genes H16-A2196 through H16-A2201 [45,169]. However, because of the lack of at least four subunit-encoding genes, and the structural differences in the CH34 and H16 Hyf proteins compared to the *E. coli* K12 Hyf units, it is very unlikely that the *hyf* genes in either strain are involved in any aspect of lithotrophic growth.

The *C. metallidurans* MBH is a typical [Ni-Fe]-hydrogenase [186] composed of three subunits (Figure S8B): HoxG, holding the active hydrogen-splitting Ni-Fe centre; HoxK, an electron-transferring subunit containing two 4Fe-4S clusters; and HoxZ, a membrane-bound di-heme cytochrome b. The HoxK signal sequence contains the *SRRSFMK* sequence recognized by the Twin arginine translocation (Tat) system. Consequently, MBH is exported from the cytoplasm to the periplasmic space, to where its enzymatic activity is limited. The *C. metallidurans* SH is classified as a cytoplasmic bidirectional or hydrogen-evolving [Ni-Fe]-hydrogenase [186,187] consisting of five subunits HoxHUFIK, with HoxY holding the *e*-transferring 4Fe-4S cluster. While the reducing powers of MBH reside solely under the form of reduced ubiquinone (ubiquinol), the SH can directly reduce NAD⁺ to NADH (Figure S8B). Because SH acts as a bidirectional enzyme complex, it reverses its activity with increasing NADH to NAD⁺ ratios, effectively acting as a “redox valve”. Accordingly, SH is an important factor in protecting the electron transfer chain from becoming hyper-reductive. The fact that *C. metallidurans* CH34 can utilize H₂ instantaneously as a reducing power and hence as an alternative source of energy when nutrients are scarce indicates that chemolithoautotrophic growth is an integral part of the organism's lifestyle.

We point out that, although *C. eutrophus* H16 is known as the model 'knallgas' bacterium, the MBH of *C. metallidurans* CH34 appears to be far more efficient than its counterpart in *C. eutrophus* H16 since it can oxidise H₂ gas at concentrations as low as 3% (v/v) and this under atmospheric conditions (i.e., at 20.9% v/v oxygen) [188]. The remarkable features of the *C. metallidurans* MBH (high catalytic activity, H₂ affinity, and O₂ tolerance) might be valuable in the emerging field of microbial fuel-cell technology [189,190]. Further, the oxygen tolerance of MBH may account for the ability of *C. metallidurans* CH34 to grow autotrophically, even in high-oxygen atmospheres [9].

## Organoautotrophy and degradation of aromatics

C. metallidurans CH34 can utilize various recalcitrant organics as sole source of carbon and energy, including volatile compounds such as 1-propanol, 2-propanol, 1-butanol, 2-butanol, and acetone, as well as the monoaromatic hydrocarbons benzene, toluene, o-xylene, and phenol. The CH34 genome encodes at least 18 alcohol dehydrogenases (ADHs) and at least 10 aldehyde dehydrogenases (ALDHs). This multitude of fermentation enzymes may seem unusual for an organism that typically follows respiratory metabolism. However, as discussed in earlier separate sections, under limiting oxygen and carbon conditions CH34 will degrade short-chain fatty acids, dicarboxylated compounds, or energy-storage products, such as polyhydroxybutyrate (PHB). This may engender elevated levels of acetyl-CoA while simultaneously, fewer terminal electron acceptors are available, thus giving rise to partially oxidized toxic products including alcohols and aldehydes. Hence, the primary role of the multiple ADHs and ALDHs in CH34 would be cellular detoxification. Further accumulation of acetyl-CoA may initiate the condensation of two acetyl-CoA molecules to acetoacetyl-CoA by the action of an acetyl-CoA acetyltransferase (AAT; EC 2.3.1.9). This compound then may be converted by an acetyl-CoA:acetoacetate CoA transferase (CoAT; EC 2.8.3.8) to acetoacetate, which may have various metabolic fates. However, some bacteria change acetoacetate further into acetone via acetoacetate decarboxylase (ADC; EC 4.1.1.4). The CH34 genome carries the genes for ten AATs very similar in length and sequence (though some may have alternate functions in the cell, for example, the β-oxidation of fatty acids and dicarboxylates, or PHB metabolism) and has two CoATases encoded by two independent *atoAD* loci (Rmet_1153/4 and Rmet_5059/60). However, we detected no ADC in the CH34 sequence data (by BlastP analysis against the ADC sequence of *Clostridium acetobutylicum* ATCC 824, a model organism for bacterial solventogenesis). Hence, it is doubtful that CH34 produces acetone directly from acetoacetate through decarboxylation.

We were very surprised to learn that the CH34 genome encodes an acetone carboxylase (ACX; EC 6.4.1.6) (encoded by Rmet_4105/6/7), an (α₂β₂γ₂) multimeric enzyme involved in acetone metabolism and identified in only very few bacteria [191,192]. The ACX catalytic reaction basically is the reverse of the ADC reaction in that it carboxylates acetone to generate acetoacetate that is metabolized to acetoacetyl-CoA and further to acetyl-CoA for oxidation in the TCA cycle. The *acxABC* genes in *C. metallidurans* CH34 encoding the three subunits of ACX seemingly are regulated by the AcxR transcriptional activator, whose gene (Rmet_4104) immediately precedes *acxABC* and lays in the opposite orientation, closely resembling the *acxRABC* operon of *Xanthobacter autotrophicus* Py2 [192]. Both in *X. autothropicus* and *C. metallidurans* the AcxR protein contains a SELFGXXXGAFTGA motif that is typical for σ⁵⁴ subunit binding regulators [193], and there is a clearly identifiable σ⁵⁴ promoter sequence TGGCATGNNNTTGC in the *acxR* intergenic region. Importantly, proteomic studies demonstrated that the growth of strain CH34 on acetone or 2-propanol as the sole carbon source induces ACX expression (C. Rosier, pers. comm.). A survey of CH34-related genomes using the MaGe interface reveals that the *C. pinatubonensis* JMP134 megaplasmid displays an *acxRABC* operon almost identical to the CH34 *acx* locus in organization and sequence (Ceut_C6170/72/73/74; with aa sequence identities of 89-93% between the respective ACX subunits and 71% identity between the AcxR proteins). None of the other organism listed in Table 2 appears to harbor an *acxRABC* locus. According to current MaGe data, only *Azorhizobium caulinodans* ORS 571 carries an

<figure><img src="image_31.png" /></figure>
the SH, contains one partial *hox* gene denoted Δ*hoxV* (Rmet_1534), nine intact *hox* genes *hoxFUTHWINXA* (Rmet_1522 through Rmet_1527, Rmet_1533, and Rmet_1541-42), and six genes *hypA*2*B*2*F*2*C*2*D*2*E*2 (Rmet_1535 through Rmet_1540) all on genomic island CMGI-3. The latter group of *hyp* genes (index 2) appears to have arisen from a reverse duplication of the *hyp* operon of the first *hyp* cluster (index 1) on CMGI-2. We note that the *hyp* and *hox* genes for SH are located, together with the *cbb* genes, on a region flanked by IS elements (Figures S6 and S8A) and that the IS-mediated excision of this region would lead to the loss of RubisCo and the entire SH system, thereby impairing not only autotrophic growth but causing the loss of hydrogenotrophic growth as well. Indeed, stress-induced mutants of CH34 lacking this genomic region are unable to utilize hydrogen gas as a source of energy [6].

Besides these hydrogenase gene clusters on CHR1, there are six *hydrogenase 'four' (hyf)* genes grouped together on CHR2 as *hyfBCEFGI* (Rmet_4666 through Rmet_4671). This resembles an incomplete operon that, in *E. coli* K12, exists as a 10-gene cluster encoding a multimeric hydrogenase complex. Presumably, this complex interacts with formate dehydrogenase FdhF to form a hydrogenlyase system cleaving formate to dihydrogen and carbon dioxide [185]. The same six *hyf* genes also occur on the first replicon CHR1 of *C. eutrophus* H16 as genes H16-A2196 through H16-A2201 [45,169]. However, because of the lack of at least four subunit-encoding genes, and the structural differences in the CH34 and H16 Hyf proteins compared to the *E. coli* K12 Hyf units, it is very unlikely that the *hyf* genes in either strain are involved in any aspect of lithotrophic growth.

The *C. metallidurans* MBH is a typical [Ni-Fe]-hydrogenase [186] composed of three subunits (Figure S8B): HoxG, holding the active hydrogen-splitting Ni-Fe centre; HoxK, an electron-transferring subunit containing two 4Fe-4S clusters; and HoxZ, a membrane-bound di-heme cytochrome b. The HoxK signal sequence contains the SRRSFMK sequence recognized by the Twin arginine translocation (Tat) system. Consequently, MBH is exported from the cytoplasm to the periplasmic space, to where its enzymatic activity is limited. The *C. metallidurans* SH is classified as a cytoplasmic bidirectional or hydrogen-evolving [Ni-Fe]-hydrogenase [186,187] consisting of five subunits HoxHUFIK, with HoxY holding the e⁻ transferring 4Fe-4S cluster. While the reducing powers of MBH reside solely under the form of reduced ubiquinone (ubiquinol), the SH can directly reduce NAD⁺ to NADH (Figure S8B). Because SH acts as a bidirectional enzyme complex, it reverses its activity with increasing NADH to NAD⁺ ratios, effectively acting as a “redox valve”. Accordingly, SH is an important factor in protecting the electron transfer chain from becoming hyper-reductive. The fact that *C. metallidurans* CH34 can utilize H₂ instantaneously as a reducing power and hence as an alternative source of energy when nutrients are scarce indicates that chemolithoautotrophic growth is an integral part of the organism's lifestyle.

We point out that, although *C. eutrophus* H16 is known as the model 'knallgas' bacterium, the MBH of *C. metallidurans* CH34 appears to be far more efficient than its counterpart in *C. eutrophus* H16 since it can oxidise H₂ gas at concentrations as low as 3% (v/v) and this under atmospheric conditions (i.e., at 20.9% v/v oxygen) [188]. The remarkable features of the *C. metallidurans* MBH (high catalytic activity, H₂ affinity, and O₂ tolerance) might be valuable in the emerging field of microbial fuel-cell technology [189,190]. Further, the oxygen tolerance of MBH may account for the ability of *C. metallidurans* CH34 to grow autotrophically, even in high-oxygen atmospheres [9].

## Organoautotrophy and degradation of aromatics

C. metallidurans CH34 can utilize various recalcitrant organics as sole source of carbon and energy, including volatile compounds such as 1-propanol, 2-propanol, 1-butanol, 2-butanol, and acetone, as well as the monoaromatic hydrocarbons benzene, toluene, o-xylene, and phenol. The CH34 genome encodes at least 18 alcohol dehydrogenases (ADHs) and at least 10 aldehyde dehydrogenases (ALDHs). This multitude of fermentation enzymes may seem unusual for an organism that typically follows respiratory metabolism. However, as discussed in earlier separate sections, under limiting oxygen and carbon conditions CH34 will degrade short-chain fatty acids, dicarboxylated compounds, or energy-storage products, such as polyhydroxybutyrate (PHB). This may engender elevated levels of acetyl-CoA while simultaneously, fewer terminal electron acceptors are available, thus giving rise to partially oxidized toxic products including alcohols and aldehydes. Hence, the primary role of the multiple ADHs and ALDHs in CH34 would be cellular detoxification. Further accumulation of acetyl-CoA may initiate the condensation of two acetyl-CoA molecules to acetoacetyl-CoA by the action of an acetyl-CoA acetyltransferase (AAT; EC 2.3.1.9). This compound then may be converted by an acetyl-CoA:acetoacetate CoA transferase (CoAT; EC 2.8.3.8) to acetoacetate, which may have various metabolic fates. However, some bacteria change acetoacetate further into acetone via acetoacetate decarboxylase (ADC; EC 4.1.1.4). The CH34 genome carries the genes for ten AATs very similar in length and sequence (though some may have alternate functions in the cell, for example, the β-oxidation of fatty acids and dicarboxylates, or PHB metabolism) and has two CoATases encoded by two independent *atoAD* loci (Rmet_1153/4 and Rmet_5059/60). However, we detected no ADC in the CH34 sequence data (by BlastP analysis against the ADC sequence of *Clostridium acetobutylicum* ATCC 824, a model organism for bacterial solventogenesis). Hence, it is doubtful that CH34 produces acetone directly from acetoacetate through decarboxylation.

We were very surprised to learn that the CH34 genome encodes an acetone carboxylase (ACX; EC 6.4.1.6) (encoded by Rmet_4105/6/7), an (α₂β₂γ₂) multimeric enzyme involved in acetone metabolism and identified in only very few bacteria [191,192]. The ACX catalytic reaction basically is the reverse of the ADC reaction in that it carboxylates acetone to generate acetoacetate that is metabolized to acetoacetyl-CoA and further to acetyl-CoA for oxidation in the TCA cycle. The *acxABC* genes in *C. metallidurans* CH34 encoding the three subunits of ACX seemingly are regulated by the AcxR transcriptional activator, whose gene (Rmet_4104) immediately precedes *acxABC* and lays in the opposite orientation, closely resembling the *acxRABC* operon of *Xanthobacter autotrophicus* Py2 [192]. Both in *X. autothropicus* and *C. metallidurans* the AcxR protein contains a SELFGXXXGAFTGA motif that is typical for σ⁵⁴ subunit binding regulators [193], and there is a clearly identifiable σ⁵⁴ promoter sequence TGGCATGNNNTTGC in the *acxR* intergenic region. Importantly, proteomic studies demonstrated that the growth of strain CH34 on acetone or 2-propanol as the sole carbon source induces ACX expression (C. Rosier, pers. comm.). A survey of CH34-related genomes using the MaGe interface reveals that the *C. pinatubonensis* JMP134 megaplasmid displays an *acxRABC* operon almost identical to the CH34 *acx* locus in organization and sequence (Ceut_C6170/72/73/74; with aa sequence identities of 89-93% between the respective ACX subunits and 71% identity between the AcxR proteins). None of the other organism listed in Table 2 appears to harbor an *acxRABC* locus. According to current MaGe data, only *Azorhizobium caulinodans* ORS 571 carries an

<figure><img src="image_31.png" /></figure>
*acxRABC* locus closely resembling the CH34 *acx* locus (with 62-71% sequence identity between the respective ACX subunits and 41% sequence identity between the AcxR proteins) albeit that its *acrR* gene is oriented in the same direction as the *acxABC* genes. We also detected *acx* loci in *Mesorhizobium loti* MAFF 303099 and *Sinerhizobium loti* 1021 with 37-40% sequence identity of their ACX subunits to the respective ACX subunits of CH34. However, the *M. loti* and *S. loti* loci do not code for an AcxR-type σ<sup>54</sup> dependent transcriptional activator, but for a LuxR/FixJ type transcriptional regulator.

Bacteria can produce acetone through the dehydrogenation of 2-propanol (iso-propanol) or via the oxidation of 2-nitropropane by a 2-nitropropane dioxygenase (NPD; EC 1.3.11.32). Among the 18 ADH genes we detected in CH34, the most likely candidate for an isopropanol dehydrogenase (iPDH) is Rmet_5645 because its product is highly similar in length and sequence to the multifunctional ADH of *C. eutrophus* H16 (H16_A0757) [194]. In addition, the CH34 genome contains six NPD encoding genes (Rmet_0222, Rmet_2083, Rmet_2605, Rmet_3674, Rmet_5434, and Rmet_5534). The production of acetone by CH34 would explain the presence of an ACX because this way cells can utilize it for growth and keep (toxic) acetone levels low.

Benzene, toluene (methylbenzene), ethylbenzene, and xylene (dimethylbenzene) jointly are known as BTEX compounds, with phenol as a major chemical intermediate. In *C. metallidurans* CH34 aerobic degradation of BTEX involves ring hydroxylation by one of three different multicomponent monooxygenases and proceeds via typical *meta*-cleavage pathways initiated via catechol decyclation by a 2,3-dioxygenase (in contrast to 1,2-dioxygenase *ortho*-cleavage pathways that cleave the catechol ring between the two hydroxyl groups). Bacterial multicomponent monooxygenases (BMMs), transcribed from single operons encoding four to six subunits, are classified in six distinct groups [195]. Although the genomes of aromatic-degrading strains only rarely carry more than one BMM [195], Cafaro et al. [196] suggested that multiple BMMs in the same system may be beneficiary in terms of BMM specificity and efficiency of substrate utilization. We identified two Group 1 (phenol hydroxylases; PHs) and one Group 2 (toluene-benzene monooxygenases; TOMOs) BMMs in two different loci of the CH34 genome. The first locus (Rmet_1305 to Rmet_1331) is part of the CMGI-2 genomic island and contains the genes for two BMMs: a phenol hydroxylase, encoded by six genes Rmet_1326 to Rmet_1331, and a benzene-toluene monooxygenase, encoded by six genes Rmet_1311 to Rmet_1316. All genes are present for the *meta*-cleavage of catechol (by an 2,3-dioxygenase encoded by Rmet_1324), and its subsequent enzymatic processing (by the products of genes Rmet_1317 to Rmet_1322) to the final product acetyl-CoA. The Rmet_1305 gene (*patR*) was identified as a σ<sup>54</sup>-dependent Fis-type transcriptional regulator. This entire region is in high synteny with the aromatic degradation gene clusters in the related genomes of *C. pinatubonensis* JMP134 (Reut_B5671 through Reut_B5693) and *Ralstonia pickettii* 12D (Rpic_5178 through Rpic_5201). The second locus (Rmet_1781 to Rmet_1788) encodes a second phenol hydroxylase (PH) also consisting of six subunits (Rmet_1782/3/4/5/6/7), with Rmet_1788 encoding the Fis-type regulator PoxR. However, gene Rmet_1781 encodes a 1,2-dioxygenase with high similarity (64% aa sequence identity) to the 1,2-dioxygenase for *ortho*-cleavage of benzoate (encoded by Rmet_4881; see 'Heterotrophic carbon metabolism' in this discussion). This eight-gene cassette, including a 1,2-dioxygenase gene, also occurs in strain JMP134 (as Reut_A1698 to Reut_A1705) and in *Wautersia numadzuensis* strain TE26 (as the *phtRABCDEF-catA* locus; [197]). The very high sequence similarities between the eight corresponding gene products of the CH34,

JMP134, and TE26 loci (82-99% sequence identity) point to their acquisition via gene transfer as one cassette.

The combination of a phenol hydroxylase gene cluster with a 1,2-dioxygenase gene seems to be very rare and confined to the above-mentioned strains. As far as we are aware, *P. putida* strain GB1 is the only other organism with a similar locus (as evidenced by MaGe data) although the 1,2-dioxygenase gene is not located downstream of the hydroxylase genes, but on the other side of the regulatory gene. Also, the sequence similarity between the corresponding hydroxylase subunits is much lower (48-68% sequence identity). Notomista et al. [195] postulated that the modularity of multiple BMMs in the same system may foster the invention of new mixed pathways, or even hybrid BMMs with novel substrate specificities. Here, it might be valuable to explore systematically the aromatic-substrate specificity of *C. metallidurans* CH34, as was done recently for *C. pinatubonensis* JMP134 [198]. Only then can we correlate aromatics degradation with pathway predictions and determine the true catabolic potential of CH34.

## Polyester synthesis PHA/PHB

Bacteria produce polyhydroxybutyrate (PHB), a polyester polymer belonging to the family of polyhydroxyalkanoates (PHAs), as a carbon- and energy-reserve to be metabolized after exhausting all other carbon resources (reviewed by Lenz and Marchessault [199]). The bacterial synthesis of PHAs, such as PHB, has gathered renewed interest because their physical properties can be remarkably similar to those of the plastic polypropylene, and because, unlike polypropylene, it is rapidly biodegradable (hence the name “bioplastic” or “green plastic”). Although there are various bacterial pathways to create PHAs [200], the key model one presently used is that from *C. eutrophus* H16 which encompasses only three enzymes: A PHA synthase (or polymerase) encoded by *phaC*; an acetoacetyl-CoA reductase encoded by *phaB*; and, a β-ketothiolase encoded by *phaA*. The synthesis of PHB has been studied most extensively in *C. eutrophus* H16 because it can generate PHB solely using simple carbon substances, such as gluconate, via the Entner-Doudoroff (ED) pathway [45]. Additional genes playing a role in PHB synthesis are *phaR* encoding a transcriptional regulator, and *phaP* encoding a phasin acting on the surface of PHA granules. Phasins inhibit the coalescence of the granules to regulate their number, and may take part in mobilizing the storage polymer. Lastly, several *phaZ* paralogs encode the intracellular depolymerases needed for PHB homeostasis and degradation.

Like its relative *C. eutrophus* H16, *C. metallidurans* CH34 synthesizes PHB and probably accumulates it in the form of intracellular storage granules. Interestingly, PHB synthesis in CH34 is initially (i.e., after 24 hours of growth) enhanced under simulated microgravity [41]. The CH34 genome has three copies of *phaC*, one copy each of *phaA* and *phaP*, three copies of *phaB*, and one regulatory gene *phaR* (for Rmet numbers see Table S9). These PHB synthesis genes mainly are organized into two cassettes: *phaC1-phaA-phaB1-phaR* (Rmet_1356-1359) located on the CHR1, and *phaC3-phaB3* (Rmet_5122-5123) located on CHR2. All the other genes are scattered throughout the genome (Table S9). We also found isologs of *phaA* (*bktB*, *atoB*) encoding additional acetoacetyl-CoA thiolases but these were annotated as having a different substrate specificity than PhaA (Table S9). For intracellular PHB depolymerization, we identified three *phaZ* genes on the *C. metallidurans* CH34 genome: One on CHR1 and two on CHR2. These *phaZ* genes correspond with the *phaZ1*, *phaZ2* and *phaZ3* genes of *C. eutrophus* H16 identified by York et al. [201]. In addition, the CHR1 replicon carries a gene for a PHB-oligomer hydrolase (designated by *phaY* or Rmet_1960) that is an ortholog of

<figure><img src="image_32.png" /></figure>
acxRABC locus closely resembling the CH34 *acx* locus (with 62-71% sequence identity between the respective ACX subunits and 41% sequence identity between the AcxR proteins) albeit that its *acrR* gene is oriented in the same direction as the *acxABC* genes. We also detected *acx* loci in *Mesorhizobium loti* MAFF 303099 and *Sinerhizobium loti* 1021 with 37-40% sequence identity of their ACX subunits to the respective ACX subunits of CH34. However, the *M. loti* and *S. loti* loci do not code for an AcxR-type σ⁵⁴ dependent transcriptional activator, but for a LuxR/FixJ type transcriptional regulator.

Bacteria can produce acetone through the dehydrogenation of 2-propanol (iso-propanol) or via the oxidation of 2-nitropropane by a 2-nitropropane dioxygenase (NPD; EC 1.3.11.32). Among the 18 ADH genes we detected in CH34, the most likely candidate for an isopropanol dehydrogenase (iPDH) is Rmet_5645 because its product is highly similar in length and sequence to the multifunctional ADH of *C. eutrophus* H16 (H16_A0757) [194]. In addition, the CH34 genome contains six NPD encoding genes (Rmet_0222, Rmet_2083, Rmet_2605, Rmet_3674, Rmet_5434, and Rmet_5534). The production of acetone by CH34 would explain the presence of an ACX because this way cells can utilize it for growth and keep (toxic) acetone levels low.

Benzene, toluene (methylbenzene), ethylbenzene, and xylene (dimethylbenzene) jointly are known as BTEX compounds, with phenol as a major chemical intermediate. In *C. metallidurans* CH34 aerobic degradation of BTEX involves ring hydroxylation by one of three different multicomponent monooxygenases and proceeds via typical *meta*-cleavage pathways initiated via catechol decyclation by a 2,3-dioxygenase (in contrast to 1,2-dioxygenase *ortho*-cleavage pathways that cleave the catechol ring between the two hydroxyl groups). Bacterial multicomponent monooxygenases (BMMs), transcribed from single operons encoding four to six subunits, are classified in six distinct groups [195]. Although the genomes of aromatic-degrading strains only rarely carry more than one BMM [195], Cafaro et al. [196] suggested that multiple BMMs in the same system may be beneficiary in terms of BMM specificity and efficiency of substrate utilization. We identified two Group 1 (phenol hydroxylases; PHs) and one Group 2 (toluene-benzene monooxygenases; TOMOs) BMMs in two different loci of the CH34 genome. The first locus (Rmet_1305 to Rmet_1331) is part of the CMGI-2 genomic island and contains the genes for two BMMs: a phenol hydroxylase, encoded by six genes Rmet_1326 to Rmet_1331, and a benzene-toluene monooxygenase, encoded by six genes Rmet_1311 to Rmet_1316. All genes are present for the *meta*-cleavage of catechol (by an 2,3-dioxygenase encoded by Rmet_1324), and its subsequent enzymatic processing (by the products of genes Rmet_1317 to Rmet_1322) to the final product acetyl-CoA. The Rmet_1305 gene (*patR*) was identified as a σ⁵⁴-dependent Fis-type transcriptional regulator. This entire region is in high synteny with the aromatic degradation gene clusters in the related genomes of *C. pinatubonensis* JMP134 (Reut_B5671 through Reut_B5693) and *Ralstonia pickettii* 12D (Rpic_5178 through Rpic_5201). The second locus (Rmet_1781 to Rmet_1788) encodes a second phenol hydroxylase (PH) also consisting of six subunits (Rmet_1782/3/4/5/6/7), with Rmet_1788 encoding the Fis-type regulator PoxR. However, gene Rmet_1781 encodes a 1,2-dioxygenase with high similarity (64% aa sequence identity) to the 1,2-dioxygenase for *ortho*-cleavage of benzoate (encoded by Rmet_4881; see 'Heterotrophic carbon metabolism' in this discussion). This eight-gene cassette, including a 1,2-dioxygenase gene, also occurs in strain JMP134 (as Reut_A1698 to Reut_A1705) and in *Wautersia numadzuensis* strain TE26 (as the *phtRABCDEF-catA* locus; [197]). The very high sequence similarities between the eight corresponding gene products of the CH34,

JMP134, and TE26 loci (82-99% sequence identity) point to their acquisition via gene transfer as one cassette.

The combination of a phenol hydroxylase gene cluster with a 1,2-dioxygenase gene seems to be very rare and confined to the above-mentioned strains. As far as we are aware, *P. putida* strain GB1 is the only other organism with a similar locus (as evidenced by MaGe data) although the 1,2-dioxygenase gene is not located downstream of the hydroxylase genes, but on the other side of the regulatory gene. Also, the sequence similarity between the corresponding hydroxylase subunits is much lower (48-68% sequence identity). Notomista et al. [195] postulated that the modularity of multiple BMMs in the same system may foster the invention of new mixed pathways, or even hybrid BMMs with novel substrate specificities. Here, it might be valuable to explore systematically the aromatic-substrate specificity of *C. metallidurans* CH34, as was done recently for *C. pinatubonensis* JMP134 [198]. Only then can we correlate aromatics degradation with pathway predictions and determine the true catabolic potential of CH34.

## Polyester synthesis PHA/PHB

Bacteria produce polyhydroxybutyrate (PHB), a polyester polymer belonging to the family of polyhydroxyalkanoates (PHAs), as a carbon- and energy-reserve to be metabolized after exhausting all other carbon resources (reviewed by Lenz and Marchessault [199]). The bacterial synthesis of PHAs, such as PHB, has gathered renewed interest because their physical properties can be remarkably similar to those of the plastic polypropylene, and because, unlike polypropylene, it is rapidly biodegradable (hence the name “bioplastic” or “green plastic”). Although there are various bacterial pathways to create PHAs [200], the key model one presently used is that from *C. eutrophus* H16 which encompasses only three enzymes: A PHA synthase (or polymerase) encoded by *phaC*; an acetoacetyl-CoA reductase encoded by *phaB*; and, a β-ketothiolase encoded by *phaA*. The synthesis of PHB has been studied most extensively in *C. eutrophus* H16 because it can generate PHB solely using simple carbon substances, such as gluconate, via the Entner-Doudoroff (ED) pathway [45]. Additional genes playing a role in PHB synthesis are *phaR* encoding a transcriptional regulator, and *phaP* encoding a phasin acting on the surface of PHA granules. Phasins inhibit the coalescence of the granules to regulate their number, and may take part in mobilizing the storage polymer. Lastly, several *phaZ* paralogs encode the intracellular depolymerases needed for PHB homeostasis and degradation.

Like its relative *C. eutrophus* H16, *C. metallidurans* CH34 synthesizes PHB and probably accumulates it in the form of intracellular storage granules. Interestingly, PHB synthesis in CH34 is initially (i.e., after 24 hours of growth) enhanced under simulated microgravity [41]. The CH34 genome has three copies of *phaC*, one copy each of *phaA* and *phaP*, three copies of *phaB*, and one regulatory gene *phaR* (for Rmet numbers see Table S9). These PHB synthesis genes mainly are organized into two cassettes: *phaC1-phaA-phaB1-phaR* (Rmet_1356-1359) located on the CHR1, and *phaC3-phaB3* (Rmet_5122-5123) located on CHR2. All the other genes are scattered throughout the genome (Table S9). We also found isologs of *phaA* (*bktB*, *atoB*) encoding additional acetoacetyl-CoA thiolases but these were annotated as having a different substrate specificity than PhaA (Table S9). For intracellular PHB depolymerization, we identified three *phaZ* genes on the *C. metallidurans* CH34 genome: One on CHR1 and two on CHR2. These *phaZ* genes correspond with the *phaZ1*, *phaZ2* and *phaZ3* genes of *C. eutrophus* H16 identified by York et al. [201]. In addition, the CHR1 replicon carries a gene for a PHB-oligomer hydrolase (designated by *phaY* or Rmet_1960) that is an ortholog of

<figure><img src="image_32.png" /></figure>
the earlier identified 3-hydroxybutyrate-oligomer hydrolase (3HBH) of C. eutrophus H16 [202-204]. The 3HBH enzyme in C. eutrophus H16 degrades the trimeric products formed by intracellular PHB depolymerases [205]; based on the very high aa similarity (63%), PhaY seems to have the same function in C. metallidurans. Lastly, a gene fragment (Rmet_1670) located adjacent to the phaC2 gene (Rmet_1671) is predicted by the MaGe system to encode a defective γ-aminobutyrate transporter, indicating that these genes might be remnants of an obsolete PHB biosynthesis operon.

## DNA repair genes and stress

The robustness of C. metallidurans CH34 under a wide variety of harsh environmental conditions does not rest just on its efficient transport and detoxification of harmful substances, ions or molecules, but also greatly depends on safeguarding its genetic material from irreversible damage. Generally, genome integrity is maintained by a number of DNA-repair mechanisms, including excision repair, direct repair, and recombinational repair. The CH34 genome contains DNA-repair genes for base-excision repair (alkA, mutY, tag, ung, fpg, nth, xthA) (all Rmet numbers are given in Table S10) and the complete pathway for nucleotide-excision repair (mfd, uvrA, uvrB, uvrC, uvrD). Direct DNA repair in strain CH34 is mediated by photoreactivation (phr), oxidative demethylation (alkB1, alkB2), and alkyltransferases activity (ada, ogt1, ogt2, ogt3). Its genome does not seem to carry a RecBCD nor a RecFOR pathway to repair single-strand gaps and double-strand DNA breaks by homologous recombination. The lack of these genes was previously postulated to allow the transformation in CH34 of large linear dsDNA fragments [206]. Nonetheless, strain CH34 has a functionally analogous system in the form of AddAB that is prevalent among β- and α-proteobacteria and firmicutes [207]. The CH34 genome carries the genes encoding the Holliday-junction protein complex RuvAB, the resolvase RuvC, a dsDNA exonuclease SbcD, and the mismatch-repair proteins MutS and MutL. These genes *ruvAB*, *ruvC*, *sbcD*, and *mutSL* often co-occur within genomes [207]. Interestingly, CH34 does not contain a *mutH* gene encoding a dGATC endonuclease but a *dam* homologous gene is present (Rmet_5483, located on CHR2). Dam affects the transposition frequency of several transposons and IS elements (Tn10, Tn5, Tn903, IS3, and possibly others), and modulates gene expression by the adenine methylation of DNA regulatory sites [208]. The presence of Dam in C. metallidurans CH34 might well be significant in view of the complex transcriptional networks and the large number of mobile genetic elements intrinsic to its flexible, fluid genome.

In addition, CH34 harbors the main genes of the RpoH regulon (heat shock) and stringent response (*relA* and *spoT*), the main regulatory genes of the SOS response (*lexA* and *recA*), and the oxidative-stress-response regulators, OxyR and SoxR (all Rmet numbers in Table S10). C. metallidurans CH34, however, lacks a SoxS homolog and does not carry any typical SoxR binding site, as indicated by a genome-scale pattern search using the SoxR binding site consensus sequence [209] and the Regulatory Sequence Analysis Tools webservice (RSAT) [210] (results not shown). Therefore, the involvement of SoxR-mediated regulation of the oxidative-stress response is unresolved. Finally, CH34 harbors 15 proteins belonging to the universal stress protein (UspA) superfamily (Table S11). Generally, UspA proteins are induced in response to a variety of starvation and stress conditions and have partially overlapping but distinct biological functions [211]. Eleven genes encode small proteins harboring one Usp

domain, two encode intermediate proteins with one Usp domain (*uspA13*, *uspA14*), and two encode large proteins consisting of two Usp domains (*uspA6*, *uspA11*). The *uspA11* gene is present in 2 copies and located on transposon Tn6050. All but five (*uspA3*, *uspA4*, *uspA13*, *uspA14*, *uspA15*) contain the G-2X-G-9X-[S/T] motif that is shared by many members of the UspA family [212].

## Conclusion

We have embarked on an exciting journey through the genome of a versatile soil bacterium, C. metallidurans strain CH34, best known for its tolerance to a wide range of heavy metals and representative for a species that is mainly found in areas linked to mining sites and metallurgical or chemical industries. Genome data show that this genome underwent numerous genetic adaptations in its evolutionary history by continuously gathering novel genes and pathways, and by rearranging and combining them in new clusters useful for its survival. This continuous acquisition of genes, and clusters of genes, has led to the co-occurrence of many complementary systems either involved in heavy-metal resistance or other types of stress, requiring intricate fine-tuning of cellular responses on many different levels. Accordingly, its complex genome boasts a vast network of regulatory proteins, including 18 sigma factors and hundreds of transport, signal sensory, and transduction systems. Although we only have just begun to understand this complexity, genome data presented here will now allow us to systematically link whole genome expression studies with high quality annotation data that are publicly available via the MaGe system. In addition, the comparison of the C. metallidurans CH34 genome with the genomes of three other Cupriavidus strains, each from a species with a particular lifestyle and habitat, will offer many clues on their physiology, metabolic capacities, and distinct interactions with their environment.

## Materials and Methods

### C. metallidurans strain CH34 isolation and propagation

Strain CH34 of C. metallidurans was isolated previously as described [3]. This strain is the type strain of the species (currently holding at least 17 strains [32]), and was deposited into the BCCM/LMG public strain collection (http://bccm.belspo.be/) as strain LMG1195. The bacterium was routinely grown at 28-30°C in rich Luria-Bertani medium or in Tris-salt mineral medium with gluconate as sole carbon source (MM284). The latter medium especially was used for its growth in the presence of heavy metals [9].

### Sequencing, assembly, and gap closure

The complete genome sequence of C. metallidurans strain CH34 was determined using the whole-genome shotgun method at the Joint Genome Institute (JGI), USA, using a combination of three randomly sheared libraries that contain inserts in the 3kb and 8kb (plasmids), and 40 kb (fosmids) size range. All general aspects of library construction and sequencing performed at the JGI are described at http://www.jgi.doe.gov/. These libraries were sequenced to a total depth of 9.9X, and all reads assessed for quality. Before assembling the reads, all were trimmed by removing vector-specific sequences. Libraries were assembled using the Paracel Genome Assembler (PGA; http://www.paracel.com/). Possible faulty assemblies, resulting from large repeat elements such as IS elements, were corrected by verifying the correspondence between the left- and right-direct repeat (i.e., of the IS element) via PCR amplification and sequence analysis of the predicted product. Gaps between contigs were closed by using the

<figure><img src="image_33.png" /></figure>
the earlier identified 3-hydroxybutyrate-oligomer hydrolase (3HBH) of C. eutrophus H16 [202-204]. The 3HBH enzyme in C. eutrophus H16 degrades the trimeric products formed by intracellular PHB depolymerases [205]; based on the very high aa similarity (63%), PhaY seems to have the same function in C. metallidurans. Lastly, a gene fragment (Rmet_1670) located adjacent to the phaC2 gene (Rmet_1671) is predicted by the MaGe system to encode a defective γ-aminobutyrate transporter, indicating that these genes might be remnants of an obsolete PHB biosynthesis operon.

## DNA repair genes and stress

The robustness of C. metallidurans CH34 under a wide variety of harsh environmental conditions does not rest just on its efficient transport and detoxification of harmful substances, ions or molecules, but also greatly depends on safeguarding its genetic material from irreversible damage. Generally, genome integrity is maintained by a number of DNA-repair mechanisms, including excision repair, direct repair, and recombinational repair. The CH34 genome contains DNA-repair genes for base-excision repair (alkA, mutY, tag, ung, fpg, nth, xthA) (all Rmet numbers are given in Table S10) and the complete pathway for nucleotide-excision repair (mfd, uvrA, uvrB, uvrC, uvrD). Direct DNA repair in strain CH34 is mediated by photoreactivation (phr), oxidative demethylation (alkB1, alkB2), and alkyltransferases activity (ada, ogt1, ogt2, ogt3). Its genome does not seem to carry a RecBCD nor a RecFOR pathway to repair single-strand gaps and double-strand DNA breaks by homologous recombination. The lack of these genes was previously postulated to allow the transformation in CH34 of large linear dsDNA fragments [206]. Nonetheless, strain CH34 has a functionally analogous system in the form of AddAB that is prevalent among β- and α-proteobacteria and firmicutes [207]. The CH34 genome carries the genes encoding the Holliday-junction protein complex RuvAB, the resolvase RuvC, a dsDNA exonuclease SbcD, and the mismatch-repair proteins MutS and MutL. These genes *ruvAB*, *ruvC*, *sbcD*, and *mutSL* often co-occur within genomes [207]. Interestingly, CH34 does not contain a *mutH* gene encoding a dGATC endonuclease but a *dam* homologous gene is present (Rmet_5483, located on CHR2). Dam affects the transposition frequency of several transposons and IS elements (Tn10, Tn5, Tn903, IS3, and possibly others), and modulates gene expression by the adenine methylation of DNA regulatory sites [208]. The presence of Dam in C. metallidurans CH34 might well be significant in view of the complex transcriptional networks and the large number of mobile genetic elements intrinsic to its flexible, fluid genome.

In addition, CH34 harbors the main genes of the RpoH regulon (heat shock) and stringent response (*relA* and *spoT*), the main regulatory genes of the SOS response (*lexA* and *recA*), and the oxidative-stress-response regulators, OxyR and SoxR (all Rmet numbers in Table S10). C. metallidurans CH34, however, lacks a SoxS homolog and does not carry any typical SoxR binding site, as indicated by a genome-scale pattern search using the SoxR binding site consensus sequence [209] and the Regulatory Sequence Analysis Tools webservice (RSAT) [210] (results not shown). Therefore, the involvement of SoxR-mediated regulation of the oxidative-stress response is unresolved. Finally, CH34 harbors 15 proteins belonging to the universal stress protein (UspA) superfamily (Table S11). Generally, UspA proteins are induced in response to a variety of starvation and stress conditions and have partially overlapping but distinct biological functions [211]. Eleven genes encode small proteins harboring one Usp

domain, two encode intermediate proteins with one Usp domain (*uspA13*, *uspA14*), and two encode large proteins consisting of two Usp domains (*uspA6*, *uspA11*). The *uspA11* gene is present in 2 copies and located on transposon Tn6050. All but five (*uspA3*, *uspA4*, *uspA13*, *uspA14*, *uspA15*) contain the G-2X-G-9X-[S/T] motif that is shared by many members of the UspA family [212].

## Conclusion

We have embarked on an exciting journey through the genome of a versatile soil bacterium, C. metallidurans strain CH34, best known for its tolerance to a wide range of heavy metals and representative for a species that is mainly found in areas linked to mining sites and metallurgical or chemical industries. Genome data show that this genome underwent numerous genetic adaptations in its evolutionary history by continuously gathering novel genes and pathways, and by rearranging and combining them in new clusters useful for its survival. This continuous acquisition of genes, and clusters of genes, has led to the co-occurrence of many complementary systems either involved in heavy-metal resistance or other types of stress, requiring intricate fine-tuning of cellular responses on many different levels. Accordingly, its complex genome boasts a vast network of regulatory proteins, including 18 sigma factors and hundreds of transport, signal sensory, and transduction systems. Although we only have just begun to understand this complexity, genome data presented here will now allow us to systematically link whole genome expression studies with high quality annotation data that are publicly available via the MaGe system. In addition, the comparison of the C. metallidurans CH34 genome with the genomes of three other Cupriavidus strains, each from a species with a particular lifestyle and habitat, will offer many clues on their physiology, metabolic capacities, and distinct interactions with their environment.

## Materials and Methods

### C. metallidurans strain CH34 isolation and propagation

Strain CH34 of C. metallidurans was isolated previously as described [3]. This strain is the type strain of the species (currently holding at least 17 strains [32]), and was deposited into the BCCM/LMG public strain collection (http://bccm.belspo.be/) as strain LMG1195. The bacterium was routinely grown at 28-30°C in rich Luria-Bertani medium or in Tris-salt mineral medium with gluconate as sole carbon source (MM284). The latter medium especially was used for its growth in the presence of heavy metals [9].

### Sequencing, assembly, and gap closure

The complete genome sequence of C. metallidurans strain CH34 was determined using the whole-genome shotgun method at the Joint Genome Institute (JGI), USA, using a combination of three randomly sheared libraries that contain inserts in the 3kb and 8kb (plasmids), and 40 kb (fosmids) size range. All general aspects of library construction and sequencing performed at the JGI are described at http://www.jgi.doe.gov/. These libraries were sequenced to a total depth of 9.9X, and all reads assessed for quality. Before assembling the reads, all were trimmed by removing vector-specific sequences. Libraries were assembled using the Paracel Genome Assembler (PGA; http://www.paracel.com/). Possible faulty assemblies, resulting from large repeat elements such as IS elements, were corrected by verifying the correspondence between the left- and right-direct repeat (i.e., of the IS element) via PCR amplification and sequence analysis of the predicted product. Gaps between contigs were closed by using the

<figure><img src="image_33.png" /></figure>
Consed editing software [213], custom primer walks, and/or PCR
amplification. The final assembly consisted of 184,165 Sanger
reads (i.e., using conventional dye-terminator chemistry). Of those,
3,752 were finishing reads used to correct errors in assembling the
draft. The estimated error rate of the completed genome sequence
was 1 in 2.5 Mb, or less. The final assembly contained two
chromosomes (3,928,089 bp and 2,580,084 bp in size) and two
plasmids (233,720 bp and 171,459 bp in size). We verified the
consistency of the chromosome whole assembly by comparing the
genome sequence map with a genetic experimental map obtained
by conjugation [22]. The sequence and annotation data of the four
replicons of *C. metallidurans* CH34 were deposited in the NCBI
database (http://www.ncbi.nlm.nih.gov/Genbank/) under the
GenBank accession numbers NC_007971 (for pMOL30),
NC_007972 (for pMOL28), NC_007973 (for CHR1), and
NC_007974 (for CHR2).

Structural and functional genome annotation

Prediction of ORFs and other genome features (RNA encoding
genes, riboswitches, ribosome binding sites, signal sequences, etc.)
as well as functional predictions were done by the MaGe platform
[48]. This system allowed the manual curation of all genes and
their functions. MaGe is also interlinked to metabolic pathway
tools such as KEGG and BioCyc, displays synteny maps for easy
genome comparison, and has an exploration interface to query
the underlying SQL databases. All *C. metallidurans* CH34 project
data are freely available at https://www.genoscope.cns.fr/agec/
mage/.

Circular plots

Circular maps for the two main replicons CHR1 and CHR2
(Figure 1) were drawn using the Circos software [214]. We
calculated the GC content and GC-skew data employing the
gc2viz.pl and gcskew2viz.pl scripts from the GenomeViz package
[215].

Identification of *ori* and *ter* regions

Regions representing the putative origin of replication (ori) and
termination (ter) were predicted by Ori-Finder software [216] that
combines gene finding, detection of base composition asymmetry
(e.g. GC-skew), and analysis of DnaA-box distribution. These
regions were scrutinised for additional features and repetitive
elements (Figure S1).

Distribution of functional classes

Functional classes were classified in COGs (Clusters of
Orthologous Groups) by the COGnitor software imbedded into
the MaGe system. All COG tables were retrieved from MaGe via
the Export pages and the derived CDS numbers were used to
calculate the normalised distribution of functional content over the
two main replicons (Table S1; Figure 2).

Clustering

Hierarchical clustering was performed via the heatmap.2 function
of the gplots library in R (version 2.8.0) using single linkage
clustering. For clustering of the different replicons (Figure 3;
Figure S3), we normalized the obtained numbers of orthologs
shared between two replicons according to the actual size of the
replicon, i.e., the distance scores used in hierarchical clustering
were calculated as [1 – number of orthologs/size of the smallest
genome] [217]. For the clustering of different RpoD proteins
(Figure S5) we used the distance measure [100 – the percentage of
protein similarity].

Genome comparative analysis

Whole genome alignments of the four *Cupriavidus* and the *R. solanacearum* genomes (Figure 4; Figure S2) were performed using the anchor-alignment software Murasaki (http://murasaki.dna.bio.keio.ac.jp/) [218]. Genome comparisons between *C. metallidurans* CH34 and *P. necessarius* STIR were done using either the LinePlot tool of MaGe [48] or the PROmer module of the MUMmer v3.0 package [219].

Clustering of protein families (paralogy)

To classify protein sequence sets into families according to
sequence similarity we used the GeneRAGE algorithm [56]. This
algorithm constructs a binary matrix holding all similarity
relationships from an all-to-all protein sequence comparison
performed with BLAST v2.0 [220] and deploys the CAST
algorithm [221] to filter sequences for low-complexity regions. The
matrix then is processed for symmetry and transitivity relationships
using successive rounds of the Smith-Waterman dynamic
programming algorithm [222]. In this way, false relationships
within the matrix are detected, as well as multi-domain protein
families are detected. We have used a cut-off E-value threshold of
10<sup>-10</sup> for all BLAST comparisons, and the default Z-score values
for symmetrification and multi-domain detection (10 and 3,
respectively).

Identification of orthologs

Protein sequence similarity was determined using BLAST (v2.0)
software [220]. Proteins between different organisms were defined
as orthologs when (i) they displayed a minimal percentage
sequence identity of 35%; (ii) they are each other's reciprocal best
BLAST hit; and, (iii) the sequence identity was measured over at
least 80% of the smallest protein.

**Supporting Information**

**Figure S1** Origins of replication for the main replicons (chromosomes) CHR1 and CHR2 of *C. metallidurans* CH34 (panels A and B, respectively). (A) DnaA-binding boxes with consensus sequence TTATCCACA are given as blocked arrows. Dark shaded boxes have a perfect match with the consensus sequence, while lighter shaded boxes have one or two mismatches (marked in red in their respective sequences). (B) Putative RepA-binding sites (see text) are indicated as ovals. Blue shaded ovals indicate putative RepA-binding sites on the opposite strand.

Found at: doi:10.1371/journal.pone.0010433.s001 (0.05 MB PDF)

**Figure S2** Genome dynamics in the second chromosome of *Cupriavidus* and *Ralstonia*. Nucleotide based comparison of the CHR2 replicons of four *Cupriavidus* species and *R. solanacearum* GMI1000 (denoted as Rsol) using the anchor-allignment software Murasaki (http://murasaki.dna.bio.keio.ac.jp/). Scale in Mb is shown on top. Abbreviations: Cmet, *C. metallidurans* CH34; Cpin, *C. pinatubonensis*, JMP134; Ceut, *C. eutrophus*; Ctai, *C. taiwanensis*. Found at: doi:10.1371/journal.pone.0010433.s002 (1.86 MB TIF)

**Figure S3** Similarity matrix between CH34 replicons and replicons of related organisms using protein based orthology data. Protein contents of all replicons for four *Cupriavidus* species and related organisms were compared by scoring the reciprocal best BlastP hits (values in lower part of the matrix) and subsequent cluster analysis of normalised values (upper part of the matrix) (see Methods). Most analysed replicons broadly fall into two large clusters A and B, with main chromosomes grouping tightly together, while three plasmids group together in cluster C owing to

<figure><img src="image_34.png" /></figure>
Consed editing software [213], custom primer walks, and/or PCR
amplification. The final assembly consisted of 184,165 Sanger
reads (i.e., using conventional dye-terminator chemistry). Of those,
3,752 were finishing reads used to correct errors in assembling the
draft. The estimated error rate of the completed genome sequence
was 1 in 2.5 Mb, or less. The final assembly contained two
chromosomes (3,928,089 bp and 2,580,084 bp in size) and two
plasmids (233,720 bp and 171,459 bp in size). We verified the
consistency of the chromosome whole assembly by comparing the
genome sequence map with a genetic experimental map obtained
by conjugation [22]. The sequence and annotation data of the four
replicons of *C. metallidurans* CH34 were deposited in the NCBI
database (http://www.ncbi.nlm.nih.gov/Genbank/) under the
GenBank accession numbers NC_007971 (for pMOL30),
NC_007972 (for pMOL28), NC_007973 (for CHR1), and
NC_007974 (for CHR2).

Structural and functional genome annotation

Prediction of ORFs and other genome features (RNA encoding
genes, riboswitches, ribosome binding sites, signal sequences, etc.)
as well as functional predictions were done by the MaGe platform
[48]. This system allowed the manual curation of all genes and
their functions. MaGe is also interlinked to metabolic pathway
tools such as KEGG and BioCyc, displays synteny maps for easy
genome comparison, and has an exploration interface to query
the underlying SQL databases. All *C. metallidurans* CH34 project
data are freely available at https://www.genoscope.cns.fr/agec/
mage/.

Circular plots

Circular maps for the two main replicons CHR1 and CHR2
(Figure 1) were drawn using the Circos software [214]. We
calculated the GC content and GC-skew data employing the
gc2viz.pl and gcskew2viz.pl scripts from the GenomeViz package
[215].

Identification of ori and ter regions

Regions representing the putative origin of replication (ori) and
termination (ter) were predicted by Ori-Finder software [216] that
combines gene finding, detection of base composition asymmetry
(e.g. GC-skew), and analysis of DnaA-box distribution. These
regions were scrutinised for additional features and repetitive
elements (Figure S1).

Distribution of functional classes

Functional classes were classified in COGs (Clusters of
Orthologous Groups) by the COGnitor software imbedded into
the MaGe system. All COG tables were retrieved from MaGe via
the Export pages and the derived CDS numbers were used to
calculate the normalised distribution of functional content over the
two main replicons (Table S1; Figure 2).

Clustering

Hierarchical clustering was performed via the heatmap.2 function
of the gplots library in R (version 2.8.0) using single linkage
clustering. For clustering of the different replicons (Figure 3;
Figure S3), we normalized the obtained numbers of orthologs
shared between two replicons according to the actual size of the
replicon, i.e., the distance scores used in hierarchical clustering
were calculated as [1 – number of orthologs/size of the smallest
genome] [217]. For the clustering of different RpoD proteins
(Figure S5) we used the distance measure [100 – the percentage of
protein similarity].

Genome comparative analysis

Whole genome alignments of the four *Cupriavidus* and the *R. solanacearum* genomes (Figure 4; Figure S2) were performed using the anchor-alignment software Murasaki (http://murasaki.dna.bio.keio.ac.jp/) [218]. Genome comparisons between *C. metallidurans* CH34 and *P. necessarius* STIR were done using either the LinePlot tool of MaGe [48] or the PROmer module of the MUMmer v3.0 package [219].

Clustering of protein families (paralogy)

To classify protein sequence sets into families according to
sequence similarity we used the GeneRAGE algorithm [56]. This
algorithm constructs a binary matrix holding all similarity
relationships from an all-to-all protein sequence comparison
performed with BLAST v2.0 [220] and deploys the CAST
algorithm [221] to filter sequences for low-complexity regions. The
matrix then is processed for symmetry and transitivity relationships
using successive rounds of the Smith-Waterman dynamic
programming algorithm [222]. In this way, false relationships
within the matrix are detected, as well as multi-domain protein
families are detected. We have used a cut-off E-value threshold of
10<sup>-10</sup> for all BLAST comparisons, and the default Z-score values
for symmetrification and multi-domain detection (10 and 3,
respectively).

Identification of orthologs

Protein sequence similarity was determined using BLAST (v2.0)
software [220]. Proteins between different organisms were defined
as orthologs when (i) they displayed a minimal percentage
sequence identity of 35%; (ii) they are each other's reciprocal best
BLAST hit; and, (iii) the sequence identity was measured over at
least 80% of the smallest protein.

**Supporting Information**

**Figure S1** Origins of replication for the main replicons (chromosomes) CHR1 and CHR2 of *C. metallidurans* CH34 (panels A and B, respectively). (A) DnaA-binding boxes with consensus sequence TTATCCACA are given as blocked arrows. Dark shaded boxes have a perfect match with the consensus sequence, while lighter shaded boxes have one or two mismatches (marked in red in their respective sequences). (B) Putative RepA-binding sites (see text) are indicated as ovals. Blue shaded ovals indicate putative RepA-binding sites on the opposite strand.

Found at: doi:10.1371/journal.pone.0010433.s001 (0.05 MB PDF)

**Figure S2** Genome dynamics in the second chromosome of *Cupriavidus* and *Ralstonia*. Nucleotide based comparison of the CHR2 replicons of four *Cupriavidus* species and *R. solanacearum* GMI1000 (denoted as Rsol) using the anchor-allignment software Murasaki (http://murasaki.dna.bio.keio.ac.jp/). Scale in Mb is shown on top. Abbreviations: Cmet, *C. metallidurans* CH34; Cpin, *C. pinatubonensis*, JMP134; Ceut, *C. eutrophus*; Ctai, *C. taiwanensis*. Found at: doi:10.1371/journal.pone.0010433.s002 (1.86 MB TIF)

**Figure S3** Similarity matrix between CH34 replicons and replicons of related organisms using protein based orthology data. Protein contents of all replicons for four *Cupriavidus* species and related organisms were compared by scoring the reciprocal best BlastP hits (values in lower part of the matrix) and subsequent cluster analysis of normalised values (upper part of the matrix) (see Methods). Most analysed replicons broadly fall into two large clusters A and B, with main chromosomes grouping tightly together, while three plasmids group together in cluster C owing to

<figure><img src="image_34.png" /></figure>
a backbone of essential genes (see also text). General information on all those species is given in Table 2. The genome of *Escherichia coli* K-12 served as a reference and outlier. Abbreviations: MPL, megaplasmid; Cmet, *C. metallidurans* CH34; Cpin, *C. pinatubonensis* JMP134; Ceut, *C. eutrophus* H16; Ctai, *C. taiwanensis*; Ecol, *Escherichia coli* K-12; Bxen, *Burkholderia xenovorans* LB400; Bpet, *Bordetella petrii*; Jmar, *Janthinobacterium* sp. Marseille; Hars, *Hermi- niimonas arsenicoxydans*; Daci, *Delftia acidovorans* SPH-1; Rsol, *Ralstonia solanacearum* GMI1000; and Pnec, *Polynucleobacter necessarius* STIR1. Replicons are denoted by their commonly known name (e.g., pRALTA) or subindexed (e.g., Cmet_CHR1).

Found at: doi:10.1371/journal.pone.0010433.s003 (2.72 MB TIF)

**Figure S4** Protein-based synteny between the *C. metallidurans* CH34 chromosomes and the genome of *P. necessarius* STIR1, an obligate symbiont of freshwater protozoans. (A) Synteny displays were generated by the LinePlot tool of the MaGe system using a synton size (S) of 3 genes. Both species belong to the family of *Burkholderiaceae* and a plausible reason for the high synteny between CH34-CHR1 and the STIR1 genome would be the preservation of essential genes during genomic reduction in the STIR1 symbiont. Genes indicated on the positive (+) and negative (-) strand are rRNA (blue), tRNA (green), and transposons (pink). Strand conservations and strand inversions between de compared replicons are colored green and red, respectively. (B) Dot plot analysis of MUMs (maximal unique matching subsequences) between the main chromosome of *C. metallidurans* CH34 and the *P. necessarius* STIR1 genome. MUMs between both forward strands are shown in red and MUMs between the forward and reverse strands are shown in blue. The analysis was performed at protein level (using the six reading frames) deploying the PROmer module of MUMmer software with default parameters.

Found at: doi:10.1371/journal.pone.0010433.s004 (0.18 MB PDF)

**Figure S5** Sigma-70 sequence similarity heatmap based on pairwise Smith-Waterman alignments of RpoD sequences. While the RpoD1 protein of CH34 (Rmet_2606 sited on CHR1) groups tightly with RpoD orthologs from the other three *Cupriavidus* species, the second sigma-70 factor RpoD2 (Rmet_4661 sited on CHR2) does not group with any of the RpoD orthologues in *Cupriavidus* but displays more significant BlastP hits with the RpoD proteins of *Bordetella* and *Burkholderia* species. All gene notations are from GenBank. Abbreviations: Rmet, *C. metallidurans* CH34; Bxe, *B. xenovorans* LB400; Reut, *C. pinatubonensis* JMP134; RALTA, *C. taiwanensis*; H16, *C. eutrophus* H16; Bpet, *B. petrii*; Daci, *D. acidovorans* SPH-1; Pnec, *P. necessarius* STIR1; mma, *Janthinobacterium* sp. Marseille; HEAR, *H. arsenicoxydans*; RSc, *R. solanacearum* GMI1000; b3067 denotes the RpoD protein sequence of *E. coli* str. K-12 substr. MG1655.

Found at: doi:10.1371/journal.pone.0010433.s005 (0.73 MB PDF)

**Figure S6** Schematic view of the *cbb* loci in *C. metallidurans* CH34. Loci involved in the Calvin-Benson-Bassham cycle (reductive pentose phosphate pathway) are dispersed over the CH34 genome although most are within the Rmet_1491 through Rmet_1544 region. Note that IS-mediated excision of this region (or parts thereof) would lead to the impairment of autotrophic growth in CH34, a phenomenon that can be observed when strain CH34 is under mutagenic stress (see text).

Found at: doi:10.1371/journal.pone.0010433.s006 (0.07 MB PDF)

**Figure S7** Genes and pathways for sulfur oxidation (sulfur autotrophy) in *C. metallidurans* CH34. (A) Genetic loci encompass-

ing sulfur oxidizing genes (*sox*) and sulfur oxygenase reductase genes (*sor*). Note that strain CH34 lacks the *soxCD* genes for a periplasmic sulfur dehydrogenase which is not essential for the oxidation of thiosulfate although fewer electrons are generated (see text and next panel). (B) Proposed pathway for the oxidation of thiosulfate ($S_2O_3^{2-}$) to sulfate ($SO_4^{2-}$) by the combined action of the sulfur carrier protein SoxYZ, a cytochrome c-type complex SoxXA, and the sulfate thiol hydrolase SoxB. (C) Proposed pathway for the oxidation of sulfide ($HS^-$) by the SoxEF c-type flavocytochrome complex in which SoxF acts as a sulfide dehydrogenase. (D) Proposed pathway for the oxidation of sulfite ($SO_3^{2-}$) to sulfate ($SO_4^{2-}$) through the action of a periplasmic sulfite dehydrogenase, SorA, and a membrane-bound c-type cytochrome, SorB.

Found at: doi:10.1371/journal.pone.0010433.s007 (0.05 MB XLS)

**Figure S8** Genetic loci in *C. metallidurans* CH34 for the oxidation of hydrogen. (A) *hox* and *hyp* genes needed to form and activate the two hydrogenases of CH34. The first cluster of genes governs the membrane-bound hydrogenase (MBH) and is sited on CHR1 as part of the CMGI-2 genomic island. The second gene cluster encodes the cytoplasmic hydrogenase (SH) and is sited on CHR2 as part of the genomic island CMGI-3. (B) Multimeric composition of the MBH and SH metalloproteins and their cellular location in *C. metallidurans* CH34. While the MBH enzyme is transloacted to the periplasmic space and transfers electrons in one direction, the SH remains inside the cytoplasm and is able to transfer electrons in both directions, depending on NADH to NAD⁺ ratios in the cell. As such, the SH enzyme effectively acts as a 'redox valve'.

Found at: doi:10.1371/journal.pone.0010433.s008 (0.06 MB XLS)

**Table S1** Functional classification. Normalised distribution of functional content over the two large replicons of *C. metallidurans* CH34, based on COG assignments. These data were used for colorcoding in Figure 1 and the drawing of Figure 2.

Found at: doi:10.1371/journal.pone.0010433.s009 (0.08 MB PDF)

**Table S2** Novel HMR genes in *Cupriavidus metallidurans* CH34. Correspondance between HMR loci reported here with HMR loci reported in an earlier overview [8]. Novel genes are given in bold letter type.

Found at: doi:10.1371/journal.pone.0010433.s010 (0.02 MB XLS)

**Table S3** Efflux and transport proteins in *C. metallidurans* CH34. Transport proteins predicted by TransAAP analysis for the four *Cupriavidus* species by submitting the genomes of these species to the TransportDB website (http://www.membranetransport.org/). Only five classes were considered (indicated with blue bars). Transporters that are not present in *C. metallidurans* CH34 but are present in one or more of the other *Cupriavidus* strains are shaded in gray. Reversely, transporters that are present in CH34 but are not in either of the three other strains are shaded in green.

Found at: doi:10.1371/journal.pone.0010433.s011 (0.03 MB XLS)

**Table S4** Sigma factors of *C. metallidurans* CH34. The 17 sigma factors of *C. metallidurans* CH34 and their orthologs in related organisms (as listed in Table 2) and *E. coli* K-12. Note that some organisms, including strain CH34, possess multiple sigma-70 factors (RpoD proteins). All gene notations are from GenBank (and are as in Figure S5).

<figure><img src="image_35.png" /></figure>
a backbone of essential genes (see also text). General information on all those species is given in Table 2. The genome of *Escherichia coli* K-12 served as a reference and outlier. Abbreviations: MPL, megaplasmid; Cmet, *C. metallidurans* CH34; Cpin, *C. pinatubonensis* JMP134; Ceut, *C. eutrophus* H16; Ctai, *C. taiwanensis*; Ecol, *Escherichia coli* K-12; Bxen, *Burkholderia xenovorans* LB400; Bpet, *Bordetella petrii*; Jmar, *Janthinobacterium* sp. Marseille; Hars, *Hermi- niimonas arsenicoxydans*; Daci, *Delftia acidovorans* SPH-1; Rsol, *Ralstonia solanacearum* GMI1000; and Pnec, *Polynucleobacter necessarius* STIR1. Replicons are denoted by their commonly known name (e.g., pRALTA) or subindexed (e.g., Cmet_CHR1).

Found at: doi:10.1371/journal.pone.0010433.s003 (2.72 MB TIF)

**Figure S4** Protein-based synteny between the *C. metallidurans* CH34 chromosomes and the genome of *P. necessarius* STIR1, an obligate symbiont of freshwater protozoans. (A) Synteny displays were generated by the LinePlot tool of the MaGe system using a synton size (S) of 3 genes. Both species belong to the family of *Burkholderiaceae* and a plausible reason for the high synteny between CH34-CHR1 and the STIR1 genome would be the preservation of essential genes during genomic reduction in the STIR1 symbiont. Genes indicated on the positive (+) and negative (-) strand are rRNA (blue), tRNA (green), and transposons (pink). Strand conservations and strand inversions between de compared replicons are colored green and red, respectively. (B) Dot plot analysis of MUMs (maximal unique matching subsequences) between the main chromosome of *C. metallidurans* CH34 and the *P. necessarius* STIR1 genome. MUMs between both forward strands are shown in red and MUMs between the forward and reverse strands are shown in blue. The analysis was performed at protein level (using the six reading frames) deploying the PROmer module of MUMmer software with default parameters.

Found at: doi:10.1371/journal.pone.0010433.s004 (0.18 MB PDF)

**Figure S5** Sigma-70 sequence similarity heatmap based on pairwise Smith-Waterman alignments of RpoD sequences. While the RpoD1 protein of CH34 (Rmet_2606 sited on CHR1) groups tightly with RpoD orthologs from the other three *Cupriavidus* species, the second sigma-70 factor RpoD2 (Rmet_4661 sited on CHR2) does not group with any of the RpoD orthologues in *Cupriavidus* but displays more significant BlastP hits with the RpoD proteins of *Bordetella* and *Burkholderia* species. All gene notations are from GenBank. Abbreviations: Rmet, *C. metallidurans* CH34; Bxe, *B. xenovorans* LB400; Reut, *C. pinatubonensis* JMP134; RALTA, *C. taiwanensis*; H16, *C. eutrophus* H16; Bpet, *B. petrii*; Daci, *D. acidovorans* SPH-1; Pnec, *P. necessarius* STIR1; mma, *Janthinobacterium* sp. Marseille; HEAR, *H. arsenicoxydans*; RSc, *R. solanacearum* GMI1000; b3067 denotes the RpoD protein sequence of *E. coli* str. K-12 substr. MG1655.

Found at: doi:10.1371/journal.pone.0010433.s005 (0.73 MB PDF)

**Figure S6** Schematic view of the *cbb* loci in *C. metallidurans* CH34. Loci involved in the Calvin-Benson-Bassham cycle (reductive pentose phosphate pathway) are dispersed over the CH34 genome although most are within the Rmet_1491 through Rmet_1544 region. Note that IS-mediated excision of this region (or parts thereof) would lead to the impairment of autotrophic growth in CH34, a phenomenon that can be observed when strain CH34 is under mutagenic stress (see text).

Found at: doi:10.1371/journal.pone.0010433.s006 (0.07 MB PDF)

**Figure S7** Genes and pathways for sulfur oxidation (sulfur autotrophy) in *C. metallidurans* CH34. (A) Genetic loci encompass-

ing sulfur oxidizing genes (*sox*) and sulfur oxygenase reductase genes (*sor*). Note that strain CH34 lacks the *soxCD* genes for a periplasmic sulfur dehydrogenase which is not essential for the oxidation of thiosulfate although fewer electrons are generated (see text and next panel). (B) Proposed pathway for the oxidation of thiosulfate ($S_2O_3^{2-}$) to sulfate ($SO_4^{2-}$) by the combined action of the sulfur carrier protein SoxYZ, a cytochrome c-type complex SoxXA, and the sulfate thiol hydrolase SoxB. (C) Proposed pathway for the oxidation of sulfide ($HS^-$) by the SoxEF c-type flavocytochrome complex in which SoxF acts as a sulfide dehydrogenase. (D) Proposed pathway for the oxidation of sulfite ($SO_3^{2-}$) to sulfate ($SO_4^{2-}$) through the action of a periplasmic sulfite dehydrogenase, SorA, and a membrane-bound c-type cytochrome, SorB.

Found at: doi:10.1371/journal.pone.0010433.s007 (0.05 MB XLS)

**Figure S8** Genetic loci in *C. metallidurans* CH34 for the oxidation of hydrogen. (A) *hox* and *hyp* genes needed to form and activate the two hydrogenases of CH34. The first cluster of genes governs the membrane-bound hydrogenase (MBH) and is sited on CHR1 as part of the CMGI-2 genomic island. The second gene cluster encodes the cytoplasmic hydrogenase (SH) and is sited on CHR2 as part of the genomic island CMGI-3. (B) Multimeric composition of the MBH and SH metalloproteins and their cellular location in *C. metallidurans* CH34. While the MBH enzyme is transloacted to the periplasmic space and transfers electrons in one direction, the SH remains inside the cytoplasm and is able to transfer electrons in both directions, depending on NADH to NAD⁺ ratios in the cell. As such, the SH enzyme effectively acts as a 'redox valve'.

Found at: doi:10.1371/journal.pone.0010433.s008 (0.06 MB XLS)

**Table S1** Functional classification. Normalised distribution of functional content over the two large replicons of *C. metallidurans* CH34, based on COG assignments. These data were used for colorcoding in Figure 1 and the drawing of Figure 2.

Found at: doi:10.1371/journal.pone.0010433.s009 (0.08 MB PDF)

**Table S2** Novel HMR genes in *Cupriavidus metallidurans* CH34. Correspondance between HMR loci reported here with HMR loci reported in an earlier overview [8]. Novel genes are given in bold letter type.

Found at: doi:10.1371/journal.pone.0010433.s010 (0.02 MB XLS)

**Table S3** Efflux and transport proteins in *C. metallidurans* CH34. Transport proteins predicted by TransAAP analysis for the four *Cupriavidus* species by submitting the genomes of these species to the TransportDB website (http://www.membranetransport.org/). Only five classes were considered (indicated with blue bars). Transporters that are not present in *C. metallidurans* CH34 but are present in one or more of the other *Cupriavidus* strains are shaded in gray. Reversely, transporters that are present in CH34 but are not in either of the three other strains are shaded in green.

Found at: doi:10.1371/journal.pone.0010433.s011 (0.03 MB XLS)

**Table S4** Sigma factors of *C. metallidurans* CH34. The 17 sigma factors of *C. metallidurans* CH34 and their orthologs in related organisms (as listed in Table 2) and *E. coli* K-12. Note that some organisms, including strain CH34, possess multiple sigma-70 factors (RpoD proteins). All gene notations are from GenBank (and are as in Figure S5).

<figure><img src="image_35.png" /></figure>