Molecular Microbiology (2014) 92(3), 625–639 ■

doi:10.1111/mmi.12583 First published online 8 April 2014

BarR, an Lrp-type transcription factor in Sulfolobus acidocaldarius, regulates an aminotransferase gene in a β-alanine responsive manner

Han Liu,1 Alvaro Orell,2† Dominique Maes,3 Marleen van Wolferen,2 Ann-Christin Lindås,4 Rolf Bernander,4 Sonja-Verena Albers,2 Daniel Charlier1 and Eveline Peeters1* 1 Research Group of Microbiology and 3Structural Biology Brussels, Department of Bio-engineering Sciences, Vrije Universiteit Brussel, Pleinlaan 2, B-1050 Brussels, Belgium. 2 Molecular Biology of Archaea, Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Strasse, 35043 Marburg, Germany. 4 Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, Svante Arrhenius v. 20C, SE-10691 Stockholm, Sweden.

Summary In archaea, nothing is known about the β-alanine degradation pathway or its regulation. In this work, we identify and characterize BarR, a novel Lrp-like transcription factor and the first one that has a nonproteinogenic amino acid ligand. BarR is conserved in Sulfolobus acidocaldarius and Sulfolobus tokodaii and is located in a divergent operon with a gene predicted to encode β-alanine aminotransferase. Deletion of barR resulted in a reduced exponential growth rate in the presence of β-alanine. Furthermore, qRT-PCR and promoter activity assays demonstrated that BarR activates the expression of the adjacent aminotransferase gene, but only upon β-alanine supplementation. In contrast, auto-activation proved to be β-alanine independent. Heterologously produced BarR is an octamer in solution and forms a single complex by interacting with multiple sites in the 170 bp long intergenic region separating the divergently transcribed genes. In vitro, DNA binding is specifically responsive Accepted 15 March, 2014. *For correspondence. E-mail espeeter@ vub.ac.be; Tel. (+32) 2 6291343; Fax (+32) 2 6291345. †Present address: Molecular Microbiology of Extremophiles Research Group, Centre for Genomics and Bioinformatics, Faculty of Sciences, Universidad Mayor, Camino la Pirámide 5750, Santiago, Chile.

© 2014 John Wiley & Sons Ltd

to β-alanine and site-mutant analyses indicated that β-alanine directly interacts with the ligand-binding pocket. Altogether, this work contributes to the growing body of evidence that in archaea, Lrp-like transcription factors have physiological roles that go beyond the regulation of α-amino acid metabolism.

Introduction Microorganisms constantly need to adapt the activity of metabolic pathways to environmental, and more specifically nutritional, conditions. Sophisticated regulation mechanisms exist on transcriptional, post-transcriptional and post-translational levels. A common regulation strategy in prokaryotes is the use of single-component transcription factors (TFs) that control gene expression during transcription initiation in response to signals such as smallmolecule ligands, which can be either metabolites or molecules taken up from the environment. Archaea are endowed with a eukaryote-like basal transcription machinery that is regulated by bacterial-like TFs (Aravind and Koonin, 1999; Bell et al., 2001). Past efforts have led to the in silico identification of putative TF genes in archaeal genomes (Charoensawan et al., 2010; Pérez-Rueda and Janga, 2010); however, the physiological function and signal response mechanisms of most of these TFs are still poorly understood. One of the most abundant TF families identified in archaeal genomes is the Leucine-responsive Regulatory Protein (Lrp) family, also named FFRP or AsnC (Pérez-Rueda and Janga, 2010). Lrp-like proteins consist of an N-terminal DNA-binding domain with a helix–turn– helix (HTH) motif and a C-terminal domain with a Regulation of Amino acid Metabolism (RAM) ligand binding domain characterized by an αβ sandwich fold (Brinkman et al., 2003; Peeters and Charlier, 2010). They typically bind and respond to small-molecule ligands. In this work, we identify an Lrp-like TF that is responsive to β-alanine (3-aminopropanoic acid), a naturally occurring β-amino acid found in all biological systems. While β-alanine is not incorporated in proteins, it is an important

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Fig. 1. Genomic co-association of putative β-alanine degradation genes and lrp-like genes. A. Illustration of the β-alanine degradation pathway. B. Genetic organization of clusters containing an lrp-like gene (depicted in red), a potential aminotransferase gene (depicted in blue) and/or a potential semialdehyde dehydrogenase gene (depicted in green). Numbers below arrows represent gene numbers in the respective organism. Genes that are predicted to encode a β-alanine aminotransferase or malonate semialdehyde dehydrogenase functioning in the β-alanine degradation pathway according to the KEGG database (Wixon and Kell, 2000) are indicated with an asterisk.

metabolite required for biosynthesis of coenzyme A (CoA) in microorganisms and plants. CoA is essential in numerous metabolic pathways such as the tricarboxylic acid (TCA) cycle, β-oxidation and fatty acid biosynthesis (Spry et al., 2008). In archaea, the CoA biosynthetic pathway is distinct from that in bacteria and eukaryotes as phosphorylation of pantoate, performed by pantoate kinase (PoK), precedes condensation with β-alanine, performed by phosphopantothenate synthetase (PPS), instead of pantoate being converted to pantothenate before phosphorylation (Yokooji et al., 2009; Atomi et al., 2013). PoK and PPS have been characterized genetically and biochemically in the archaeon Thermococcus kodakarensis, and genome sequence analyses indicate that they exist in most archaeal species, including Sulfolobus (Yokooji et al., 2009; Ishibashi et al., 2012; Tomita et al., 2012). An alternative metabolic fate of β-alanine is its degradation, thereby recovering and conserving energy under the form of acetyl-CoA. In this pathway, malonate semialdehyde is formed by transamination of pyruvate to L-alanine and malonate semialdehyde is then converted to acetyl-CoA and CO2 in an oxidative decarboxylation step (Fig. 1A).

The two enzymes responsible for β-alanine catabolism, β-alanine aminotransferase and malonate semialdehyde dehydrogenase, have been characterized in Pseudomonas fluorescens (Hayaishi et al., 1961). Archaea require β-alanine for CoA biosynthesis and are capable of synthesizing it de novo by decarboxylation of aspartate (Tomita et al., 2014). However, the β-alanine degradation pathway and its regulation is as yet uncharacterized in archaea. Here, we report the characterization of an Lrp-like TF in the crenarchaeota Sulfolobus acidocaldarius and Sulfolobus tokodaii that regulates a juxtaposed gene encoding a putative β-alanine aminotransferase in response to the presence of exogenous β-alanine.

Results Identification of an lrp-like gene co-occurring with putative β-alanine degradation genes An lrp-like gene, which is conserved in S. acidocaldarius and S. tokodaii (Saci_2136 and ST1115 respectively), is © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

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Fig. 2. Growth curves of MW001 and MW001ΔbarR in the presence of exogenous β-alanine. Cells were grown in Brock medium supplemented with 0.2 (w/v)% casamino acids, 0.2 (w/v)% sucrose, 20 μg ml−1 uracil and different concentrations of β-alanine: 0 mM (A), 1 mM (B) and 10 mM (C).

located in a divergent operon with an adjacent aminotransferase (at) gene (Saci_2137 and ST1114; arCOG00916). In S. tokodaii, this gene cluster is additionally flanked by an aldehyde dehydrogenase gene (ST1116; arCOG01252). A comparative genomic analysis has furthermore revealed the existence of gene clusters comprising an lrp-like gene with a similar genomic environment, albeit in different gene syntenies, in a variety of archaeal and bacterial species (Fig. 1B). Depending on the exact substrates, aminotransferase and aldehyde dehydrogenase enzymes could act together in butanoate metabolism [using γ-aminobutyrate (GABA) as a substrate], L-valine or β-alanine catabolism. According to the KEGG database (Wixon and Kell, 2000), most of the aminotransferase and aldehyde dehydrogenase genes associated with lrp-like genes shown in Fig. 1A, including those in S. acidocaldarius and S. tokodaii, are predicted to function in the β-alanine degradation pathway (EC2.6.1.18/EC2.6.1.19/EC2.6.1.55 and EC1.2.1.18 for β-alanine transaminase and malonate semialdehyde dehydrogenase respectively). Since TF genes occur commonly in a divergent operon with their target gene(s), we speculate that the Lrp-like TFs encoded by Saci_2136 and ST1115 regulate the expression of the juxtaposed aminotransferase gene. Given the experimental observations presented below, we designate this TF beta-alanine responsive regulator (BarR). The S. tokodaii and S. acidocaldarius BarR orthologues (named St-BarR and Sa-BarR respectively) are highly similar in both the DNA-binding and ligand-binding domain and have an overall sequence identity of 73% (Fig. S1). Besides St-BarR, S. tokodaii harbours Grp, a glutamineresponsive regulator (Kumarevel et al., 2008) that exhibits © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

69% sequence identity with St-BarR and Sa-BarR, suggesting that the TF gene has undergone a duplication in S. tokodaii. Growth characteristics of an Sa-barR gene disruption mutant In an effort to examine the physiological role of BarR, we have constructed an in-frame markerless Sa-barR disruption mutant strain of S. acidocaldarius MW001. To this end, a ‘pop-in pop-out’ strategy was applied based on uracil auxotrophy (Wagner et al., 2012). The complete absence of the Sa-barR gene and its protein product in the deletion strain (designated MW001ΔbarR) was confirmed by PCR and sequencing analysis of the genomic DNA and by Western blotting using anti-BarR antibodies (Fig. S2). The MW001 and MW001ΔbarR strains did not exhibit significantly different growth rates in rich medium (Fig. 2A) and in rich medium to which 10 mM of L-valine or GABA was added (Fig. S3). However, in rich medium supplemented with 10 mM β-alanine MW001ΔbarR exhibited a slower growth rate than the isogenic wild-type strain (Fig. 2C). Noteworthy, the presence of β-alanine resulted in lower cell densities (Fig. 2) and in all growth conditions, the MW001ΔbarR strain reached somewhat higher cell densities than MW001 (Figs 2 and S3). Neither MW001 nor MW001ΔbarR cells grew in minimal medium supplemented with β-alanine as a sole carbon and energy source (data not shown). Taken together, these results indicate that Sa-BarR has a regulatory role in β-alanine metabolism and not in butanoate metabolism or valine degradation.

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Sa-BarR activates Saci_2137 expression in the presence of exogenous β-alanine To determine whether or not Saci_2137 is a regulatory target of Sa-BarR, we performed quantitative RT-PCR (qRT-PCR) and analysed the effect of Sa-barR deletion on the expression of Saci_2137 (Fig. 3A). We did not observe any difference in normal growth conditions. In contrast, upon addition of 10 mM β-alanine to the growth medium, Saci_2137 expression displayed a significant lower expression in the MW001ΔbarR strain when compared with MW001. This observation implies that the expression of Saci_2137 is activated in a Sa-BarRmediated manner, but only in the presence of β-alanine in the growth medium. The intergenic non-coding region between the divergently transcribed Sa-barR and Saci_2137 genes has a length of 170 bp and as a consequence, the oppositely oriented promoters of both genes are located relatively close to each other. We employed a LacS reporter system assaying expression under control of the full-length intergenic region (PbarR-2137) and a truncated part containing only the Saci_2137 promoter region (P2137) (Fig. 3B and C). The results of these assays demonstrated a β-alanineinduced activation [PbarR-2137: 2.4-fold (± 0.64, P < 0.05) and P2137: 2.7-fold (± 1.36, P < 0.05)] for both reporter gene constructs. Therefore, this underscores that BarR activates Saci_2137 expression and suggests that the minimal cis-elements that drive this activation are located close to the Saci_2137 promoter.

Sa-BarR performs an auto-activation We also questioned whether or not Sa-BarR performs an autoregulation. In contrast to Saci_2137, the endogenous expression level of Sa-barR was insensitive to the addition of β-alanine to the growth medium (Fig. 3A). Likewise, Sa-BarR protein levels did not vary significantly when cells were grown in the presence or absence of exogenous β-alanine, as observed by Western blotting (Fig. S4). Also in reporter gene assays, expression did not change significantly when comparing growth conditions with and without exogenous β-alanine (Fig. 3B and C). Independent of genetic background or growth condition, LacS was expressed at a much lower level upon gene fusion with a truncated part containing only the Sa-barR promoter region (PbarR) than with the full-length intergenic region (P2137-barR). This observation suggests that additional sequence elements located upstream of position −109 are required for efficient transcription of Sa-barR. In both growth conditions, a reporter gene fusion with P2137barR resulted in significantly higher expression when comparing MW001 to MW001ΔbarR [without β-alanine: 6.2fold (± 1.95, P < 0.01) and +10 mM β-alanine: 2.9-fold

(± 0.30, P < 0.01)]. Taken together, we can conclude that BarR performs a β-alanine-independent auto-activation. Recombinantly purified Sa-BarR and St-BarR proteins form octamers in vitro Heterologous overexpression of Sa-barR and St-barR in E. coli did not yield soluble protein, but instead resulted in large amounts of inclusion bodies. Therefore, we developed a procedure to purify carboxy-terminally His-tagged BarR protein by affinity chromatography from washed and denatured inclusion bodies, followed by an on-column refolding step. Hence, we obtained highly pure preparations of the 17.7 kDa Sa-BarR and St-BarR proteins (Fig. S5). To analyse quaternary structure of the purified refolded proteins, we applied them to gel filtration chromatography (Fig. 4A and B). Both St-BarR and Sa-BarR eluted at a volume corresponding to about 140 kDa, representing an octameric association state (a BarR octamer has a theoretical molecular weight of 142 kDa). Using the octameric crystal structure of the paralogous Grp protein of S. tokodaii (Kumarevel et al., 2008), a structural homology model of St-BarR was built in the SWISS-MODEL workspace (Arnold et al., 2006) of which we can assume that it is a good representation of the relevant assembly of Sa/StBarR protein in solution (Fig. 4C). Based on the observations described above, we hypothesize that β-alanine is a putative ligand of BarR and we repeated the gel filtration experiments in the presence of 5 mM β-alanine (Fig. 4A and B). In these conditions, Sa-BarR eluted again as a homogenous population of octameric proteins. In contrast, the presence of β-alanine induced a small fraction of St-BarR to dissociate into oligomeric forms with lower molecular weight (p4 and p5), possibly corresponding to dimers and/or tetramers. Sa-BarR is associated with the Sa-barR/Saci_2137 intergenic region in vivo To discern whether or not the Sa-BarR-mediated activation of Saci_2137 and auto-activation of Sa-barR are direct regulatory processes, we tested the in vivo association of the protein with the intergenic region containing promoter regions of both genes. Therefore, we performed chromatin immunoprecipitation (ChIP) in which the Sa-BarR-specific enrichment of genomic DNA spanning the Sa-barR-Saci_2137 intergenic region was tested with quantitative PCR (qPCR) (Fig. 5A). A significant enrichment of 25- and 31-fold was observed upon growth in absence and presence of exogenous 10 mM β-alanine, respectively, indicating that Sa-BarR is bound in vivo to the intergenic region not only in the presence but also in the absence of exogenous β-alanine. © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

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Fig. 3. Effects of Sa-barR deletion and exogenous β-alanine on Saci_2137 and Sa-barR gene expression. A. qRT-PCR analysis in different genetic backgrounds (MW001 (WT) versus MW001xΔbarR (Δ)) and different growth conditions (absence (− β-Ala) versus presence (+ β-Ala) of 10 mM β-alanine in the growth medium). Relative gene expression ratios were normalized against the expression of Saci_0691 and Saci_1336. Values represent means of biological triplicate measurements. B. Schematic overview of the notation of the different reporter gene constructed in the promoter activity assays. Promoters are represented by black and grey rectangles for the factor B recognition element (BRE) and TATA box respectively. Lengths of the promoter regions fused to β-glycosidase (lacS) are indicated on the top scheme. C. Promoter activity assays for Saci_2137 and Sa-barR promoters performed in different genetic backgrounds (MW001 (WT) and MW001ΔbarR (KO)) and different growth conditions (absence (− β-Ala) and presence (+ β-Ala) of 10 mM β-alanine in the growth medium). LacS activities (Miller units), which are a measure of promoter activities, were determined using the ortho-nitrophenyl-β-D-galactopyranoside (ONPG) method. Values represent means of biological triplicates.

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Characteristics of the interaction between BarR and the intergenic barR/at control region DNA in vitro Electrophoretic mobility shift assays (EMSAs) were performed to monitor the interaction between refolded recombinant BarR protein and DNA comprising the barR/at control region (Fig. 5B and C). Sa-BarR and St-BarR formed with high affinity a single specific complex with their cognate DNA fragments (both with an apparent equilibrium dissociation constant (KD) of about 200 nM), confirming the specificity of the association observed with ChIP. Furthermore, the observed binding demonstrates that the refolded protein preparations are active and contain biologically relevant assembly forms. For both proteins, we also tested binding to subfragments containing only part of the intergenic region, either the barR or at promoter region (Fig. 5B and C). In the case of Sa-BarR, these subfragments, which were designed to have a similar length, did not result in any observable binding, demonstrating that interactions are required in both the Sa-barR and Saci_2137 promoter regions for stable complex formation (Fig. 5B). In contrast, St-BarR exhibited binding to the PbarR subfragment, albeit with a twofold lower binding affinity than to the full-length fragment (Fig. 5C). Binding to the PST1114 subfragment displayed complex formation with a very low binding affinity. Notably, complexes formed with either the full-length fragment or one of the subfragments displayed a similar relative mobility, indicating that binding stoichiometry is identical. We did not perform in vivo experiments with S. tokodaii and thus, it is unknown if ST1114 transcription is upregulated in S. tokodaii cells upon addition of β-alanine to the medium as is the case for Saci_2137 in S. acidocaldarius. Identification of BarR binding sites in the operator DNA

Fig. 4. Quaternary structure analysis of St/Sa-BarR protein. A. Gel filtration elution profile of Sa-BarR performed in running buffer (20 mM Na2HPO4, 150 mM NaCl, pH 7.4) with or without 5 mM β-alanine. Blue dextran (bd), was injected concomitantly with Sa-BarR protein to determine V0. The p3 and p4 peak maxima correspond to a calculated molecular weight of 143 kDa. An SDS PAGE analysis of the peak fractions is shown in the inset (MWL = molecular weight ladder). B. Gel filtration elution profile of St-BarR with following calculated molecular weights: 140 kDa (p1), 137 kDa (p3), 74 kDa (p4) and 50 kDa (p5). C. Homology model of octameric St-BarR structure created with SWISS-MODEL using the Grp crystal structure (PDB: 2EZW) (Kumarevel et al., 2008) as a template. Each subunit is coloured differently.

To identify the interaction zones of St/Sa-BarR binding in the barR/at intergenic region, we performed ‘in gel’ Cu-phenanthroline (Cu-OP) footprinting with the full-length DNA fragments (Fig. 6A and B). Comparison of the cleavage patterns of free and bound DNA revealed that Sa-BarR binding to its homologous operator DNA yields protection in two distinct zones, one encompassing P2137 and the other encompassing PbarR. On the other hand, the St-BarR footprinting experiment demonstrated only clear protection in a region overlapping PbarR. These results support the previous observations in the subfragment EMSAs that Sa-BarR binding strictly depends on all binding sites within the Sa-barR and Saci_2137 promoter regions, while St-BarR binding occurs with significant higher affinity in the St-barR promoter region than in the ST1114 promoter region. Given the high-affinity nature of binding between St-BarR and PbarR, we further analysed this interaction with © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

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Fig. 5. Binding of BarR to the intergenic divergent barR/at promoter region. A. ChIP experiment to analyse in vivo association of Sa-BarR to the Sa-barR/Saci_2137 intergenic region in growth conditions without β-alanine (− β-Ala) or with 10 mM β-alanine (+ β-Ala). Quantification of the fold-enrichment ratio in ChIP DNA versus input DNA was performed with qPCR using the Saci_0691 genomic region for normalization. As a negative control, enrichment of part of the ORF of Saci_1336, a region that is presumably not bound by Sa-BarR, was also quantified. A mock control experiment was carried out without antibodies. Data represent the mean of triplicate measurements. B. Electrophoretic mobility shift assays (EMSAs) of the interaction between Sa-BarR and different fragments encompassing either the complete intergenic Sa-barR/Saci_2137 control region or parts of it. The inset provides a schematic overview of the used fragments. Used protein concentrations are mentioned above each lane. The position of bands corresponding to free DNA (F) and bound DNA (B) is indicated. C. EMSAs of the interaction between St-BarR and different fragments encompassing either the complete or partial intergenic St-barR/ST1114 control region.

a variety of footprinting and high-resolution binding interference techniques using a 160 bp PbarR subfragment (Figs 6C and S6). Whereas a continuous footprint protection zone of about 70 bp was observed with both ‘in gel’ Cu-OP and DNase I footprinting, the location of strong binding interference effects defined two distinct interaction regions. St-BarR-induced DNA deformations occur in a zone separating these two regions, as demonstrated by the observation of multiple footprinting hyperreactivity and reverse binding interference effects. Each of the two contact zones, named site A and site B, in the St-barR promoter region comprises a highly similar 15 bp semi-palindromic sequence and it can be assumed that these sequences represent the recognized St-BarR binding motif (Fig. 6B and C). Furthermore, a single occurrence of a similar sequence can be discerned in the © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

ST1114 promoter region (site C; Fig. 6B). Whereas no protection was evident at this site, it is not excluded that transient contacts contribute to complex formation (given the observation that binding affinity to the full-length operator fragment exceeded that of the PbarR subfragment). The consensus sequence of the St-BarR binding motif, based on the forward and reverse sequences of the three sites, is depicted as a sequence logo (Fig. 6D). Whereas site B and C are well conserved in S. acidocaldarius, site A is very poorly conserved and lacks resemblance to the binding motif (Fig. 6B). Site A and site B border the predicted barR promoter on both sides, an operator architecture that is typical for repressors. Nevertheless, as shown above Sa-BarR activates expression of its own gene, which is supported by the observation of simultaneous binding of St-BarR and the

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Fig. 6. Identification and high-resolution contact probing of BarR DNA binding sites. A. ‘In gel’ Cu OP footprinting analysis of the interaction between Sa-BarR and the Sa-barR/Saci_2137 control region (top panel) and of the interaction between St-BarR and the St-barR/ST1114 control region (bottom panel). Top and bottom strands of the probes were labelled for the Sa-BarR and St-BarR experiment respectively. The direction of migration is indicated with an arrow. Populations that were excised from the EMSA gel are indicated as F (free DNA), B (bound DNA) and I (input DNA), the latter corresponding to free DNA in the no-protein lane. A+G and C+T represent sequencing ladders. The position of promoter elements (dark blue = BRE; light blue = TATA box) and of protected regions (red line) is indicated below the autoradiographs. B. Sequence alignment of the divergent intergenic barR/at region for S. tokodaii and S. acidocaldarius with indication of (putative) BarR binding sites and promoter elements. Experimentally confirmed binding sites are coloured dark green and in silico predicted binding sites light green. Translational start codons are boxed. Protected regions identified in the experiment shown in panel A are indicated with a red line. C. Summary of high-resolution contact probing of the interaction between St-BarR and the St-barR promoter, using a truncated fragment comprising only the St-barR control region. Original autoradiographs are shown in Supplementary Information (Fig. S6). Protein-DNA interaction effects are symbolized as follows: orange line = protection zone identified with ‘in gel’ Cu OP footprinting; orange ball-and-stick symbol = hyperreactivity effect in ‘in gel’ Cu OP footprinting; red-coloured sequence = protection zone identified with DNase I footprinting; red ball-and-stick symbol = hyperreactivity effect in DNase I footprinting; open green rectangle = weak effects in depurination binding interference; filled green rectangle = strong effect in depurination binding interference; green triangle = reverse effect in depurination binding interference; open blue rectangle = weak effect in depyrimidation binding interference; filled blue rectangle = strong effect in depyrimidation binding interference; blue triangle = reverse effect in depyrimidation binding interference; open purple diamond = weak effect in premethylation binding interference; filled purple diamond = strong effect in premethylation binding interference. The translational start codon of St-barR is depicted in bold. D. Sequence logo, created with web logo (Crooks et al., 2004), representing the DNA-binding specificity of St-BarR, based on the three identified binding sites shown in panel B.

basal TFs TATA-binding protein (TBP) and transcription factor B (TFB) (Fig. S7). It appears that St-BarR presence on the promoter even stimulates the binding of TBP/TFB. β-Alanine is the specific ligand of BarR To investigate the hypothesis that β-alanine is the specific ligand of BarR, postulated based on the observed transcription regulation in response to the presence of β-alanine in the medium, we tested the effect of adding different concentrations of this amino acid to EMSA reaction mixtures (Fig. 7A). β-alanine clearly exerts an effect on DNA binding but unexpectedly, this effect consists of the disruption of St/Sa-BarR protein-DNA complexes. The St-BarR-DNA interaction displayed a higher sensitivity to β-alanine than the Sa-BarR-DNA interaction with inhibition coefficients (IC50) of 0.5 and 4 μM respectively. None of the 20 α-amino acids including L-alanine affected complex formation (data not shown), demonstrating a high specificity to β-alanine. Previously, it has been demonstrated that two β4 residues in Lrp-like proteins, corresponding to Ser132 and Thr134 in St-BarR (see Fig. S1), are crucial for the formation of the ligand binding pocket and for the interaction with the COO− group of the ligand, which are invariably amino acids, in this pocket (Okamura et al., 2007). Alanine substitution of Ser132 or Thr134 in St-BarR almost completely alleviated the response to β-alanine, monitored at the level of protein-DNA complex formation (Fig. 7A), thereby demonstrating that β-alanine binds into the ligand binding pocket to exert its effect. Using the homology model of the St-BarR structure, we have modelled the conformation of β-alanine into the binding pocket (Fig. 7B). In this model, the carboxyl group of β-alanine is positioned similar to that of glutamine in Grp-Gln and is predicted to establish multiple interactions © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

with both backbone and side-chain atoms of Ser132, Ser133 and Thr134. Also Gly99 in the loop between β2 and β3 could interact with the carboxyl group of β-alanine, while the ammonia group of the Cβ of β-alanine might exhibit multiple interactions with Tyr73, Tyr77 and Asn102 of St-BarR. All Grp residues that interact directly with the ligand are either conserved or similar (a serine instead of threonine at position 132) in St/Sa-BarR except for an Asn residue in the loop between β2 and β3, which is a Thr residue in St/Sa-BarR (Thr101/Thr103; see Fig. S1).

Discussion In this study, we have demonstrated that BarR activates a gene encoding a putative β-alanine aminotransferase and that the TF is specifically responsive to β-alanine both in vitro and in vivo. Thus far, all identified ligands of characterized bacterial and archaeal Lrp-like TFs are classical α-amino acids (Brinkman et al., 2003; Peeters and Charlier, 2010) and hence, we identify for the first time a non-proteinogenic amino acid as a ligand for an Lrp-like TF. This enforces our previously stated hypothesis that certain proteins of the archaeal class of Lrp-type TFs might respond to other metabolic intermediates (Peeters et al., 2009). The β4 residues Ser132 and Thr134 in St-BarR are highly conserved as Thr or Ser in all Lrp-like proteins and considered to be markers of effector regulation as they are crucial for the formation of the ligand binding pocket (Thaw et al., 2006; Shrivastava and Ramachandran, 2007; Kumarevel et al., 2008). In vitro, β-alanine dissociates BarR-DNA complexes with a low IC50 and this effect is almost completely abolished by alanine substitution of either Ser132 or Thr134. This indicates that β-alanine specifically binds into the ligand binding pocket of the

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Fig. 7. Identification of β-alanine as the specific ligand of BarR. A. EMSA showing the effect of β-alanine on DNA binding of different BarR protein variants. The position of free (F) and complexed (B) DNA is indicated. The reaction mixtures contained 400 nM Sa-BarR, 110 nM St-BarR or 165 nM St-BarRS132A or St-BarR T134A protein (when present, as indicated with ‘+’); the total applied β-alanine concentrations are given. B. Illustration of the ligand binding pocket of BarR in a homology model based on the Grp-Gln structure. Chains belonging to different dimers are depicted in different colours (light blue or green) with indication of the β-strands β1 and β4 respectively. The modelled β-alanine molecule is indicated in white, and is superimposed over the L-glutamine molecule (grey) that is positioned similarly as in the homologous Grp-Gln crystal structure. Residues that are important for formation of the ligand binding pocket are depicted in stick models and labelled. Where relevant, atoms are coloured by atom type (nitrogen: blue, oxygen: red). The predicted atomic distances between β-alanine ligand and St-BarR residues are: 2.57 Å between O and O(S132), 3.41 Å between O and OG(S132), 3.23 Å between O and N(S133), 3.28 Å between O and OG1(T134), 3.10 Å between OXT and OG1(T134), 2.52 Å between OXT and N(T134), 2.92 Å between OXT and N(G99), 2.92 Å between N and OH(Y73), 3.23 Å between N and OH(Y77) and 3.91 Å between N and OD1(N102).

protein to elicit its effect. A structural homology model predicts indeed that Ser132 and Thr134 interact with the β-alanine carboxyl group (Fig. 7B). All ligand-interacting residues of Grp are conserved in BarR, except for an Asn that is a Thr in BarR (Fig. S1) and which undergoes a drastic conformational change upon ligand binding in Grp thereby switching from being located inside to outside the ligand binding pocket (Kumarevel et al., 2008). Possibly, the corresponding Thr residue is a determinant of β-alanine specificity of BarR.

St/Sa-BarR protein forms octamers, which is a common oligomeric state of Lrp-like TFs. To convert ligand binding into a regulatory response, structural changes take place in Lrp proteins, which can be either a transition between different association states (Chen and Calvo, 2002; Okamura et al., 2007; Yokoyama et al., 2007; Shrivastava et al., 2009) or very subtle conformational changes, in accordance with an allosteric regulation process. The latter is the case for Grp, in which glutamine binding induces conformational changes only in the neighbourhood of the © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

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ligand binding pocket (Kumarevel et al., 2008). A similar mechanism is plausible to occur upon β-alanine binding in Sa-BarR and St-BarR. Although for the latter, a small fraction of lower-molecular weight species was detected in the presence of β-alanine, the majority of the protein retained the octameric state. Possibly, β-alanine binding destabilizes dimer–dimer interactions within the St-BarR octamer through small conformational changes. Although this is the opposite effect as observed for the homologous Grp, in which binding of glutamine stabilizes dimer–dimer interactions (Kumarevel et al., 2008), it is conceivable since the effect on DNA binding in vitro is also opposite: while ligand binding reduces DNA binding affinity for BarR, it enhances binding affinity for Grp. A single BarR-DNA complex is formed in EMSAs probing the intergenic region of the divergent barR/at operon, which is most likely containing a single BarR octamer. Two/three partially conserved semi-palindromic binding motifs were identified by a combination of experimental observations and sequence searches, and extend over a region of 109 and 113 bp respectively (Fig. 6B). Since DNA wrapping is a common phenomenon observed for Lrp-like proteins and several structural models of Lrp-DNA complexes have been proposed in which a DNA region of about 110 bp is wrapped around an octameric protein (Leonard et al., 2001; Thaw et al., 2006; de los Rios and Perona, 2007; Yokoyama et al., 2007), we propose that a similar octameric BarR-DNA complex is formed. Protein-induced DNA deformations are indeed supported by the observation of hyperreactivity sites and reverse effects in the footprinting and binding interference experiments (Fig. 6). Envisaging the establishment of weak transient interactions without sequence specificity in the region between site B and site C would uncover a situation with 4 regularly spaced binding sites in which each site interacts with one of the dimeric peripheral facing DNA-binding domains [for St-BarR: 15 bp site (A) + 17 bp linker + 15 bp site (B) + 18 bp linker + 15 bp site + 18 bp linker + 15 bp site (C); for Sa-BarR: 15 bp site (A) + 16 bp linker + 15 bp site (B) + 16/17 bp linker + 15 bp site + 16/17 bp linker + 15 bp site (C)]. The lack of observing additional interactions in footprinting experiments could be explained by them being weak and transient. We also did not observe footprint protection for St-BarR binding upstream of the ST1114 promoter, although interactions do take place in that region since binding affinity is higher for the full-length fragment than for the truncated fragment containing only the PbarR region (Fig. 5C). Notably, while Sa-BarR is unable to bind in vitro with any of the truncated fragments containing only one part of the intergenic region, this is not the case in vivo: both PbarR and P2137 convey Sa-BarR-mediated activation (Fig. 3C). We propose that Sa-BarR exerts activation of both divergent genes from within the same octameric complex. © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

In case of the aminotransferase gene, the operator sequence lies upstream of the BRE element, which is an ideal location for transcription activation (Peeters et al., 2013). In contrast, operator sites border PbarR on both sides (Fig. 6). The observed auto-activation could be explained by a mechanism in which TBP and/or TFB binding is stimulated (Fig. S7). In a model in which BarR binds to the other side of the DNA helix than the basal TFs, this stimulation is more likely to occur through BarRinduced conformational changes in the promoter DNA than through protein–protein interactions. Whereas auto-activation occurs ligand-independent, target activation is induced by β-alanine. Curiously, while β-alanine abrogates DNA binding of Sa-BarR with a linear DNA probe in vitro, this is clearly not the case in vivo, as observed by ChIP-qPCR. This discrepancy could be explained by the presence of additional stabilizing factors, such as other TFs or proteins interacting with Sa-BarR, DNA interaction sites outside the intergenic region probed in EMSAs or chromatin structure elements that are absent in the naked DNA. Both effects, disruption of DNA binding in vitro and activation in vivo, could be explained by the above-proposed allosteric mechanism of β-alanine inducing subtle conformational changes in the protein and thus affecting the alignment of different subunits and domains. In vivo, these changes do not affect overall complex stoichiometry, as evidenced by similar levels of binding observed in ChIP-qPCR, but might affect interaction with the basal transcription machinery at P2137, not PbarR. The proposal that Saci_2137 and ST1115 encode enzymes of the β-alanine degradation pathway has been solely put forward based on sequence analysis, and the enzymes still await experimental characterization. Nevertheless, the β-alanine-dependent activation that we describe here argues for the Saci2137 aminotransferase indeed catalysing the first step of β-alanine degradation. Although β-alanine is important as precursor of CoA, it appears logical that the degradation pathway is induced in conditions of excess β-alanine, thereby generating energy useful to the cell. Also in P. fluorescens, β-alanine breakdown is induced when the cells are grown in conditions with β-alanine as sole energy and carbon source (Hayaishi et al., 1961); however, the underlying regulatory mechanism is unknown. Although MW001ΔbarR exhibits a slower growth rate in the presence of 10 mM β-alanine, it eventually outgrows the WT strain (Fig. 2). Possibly, a lack in activation of β-alanine degradation could be beneficial to the cells during stationary phase as it results in a larger intracellular β-alanine pool available for CoA biosynthesis. Another observation is that the presence of 10 mM β-alanine in the medium causes both the WT and mutant strain to display slow exponential growth rates and to yield low cell densities in stationary

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phase. We speculate that efficient uptake and consequent high intracellular concentration of β-alanine could cause toxicity due to osmotic effects, metabolic imbalances and/or competition in the uptake of α-amino acids. BarR is present in the two Sulfolobus species studied in this work but not in related Sulfolobus solfataricus and Sulfolobus islandicus strains. While these organisms harbour genes encoding a putative β-alanine aminotransferase (Sso2720 in S. solfataricus P2 and SiRe_2479 in S. islandicus REY15A, having respectively 74% and 75% sequence identity with Saci_2137) and a putative malonate semialdehyde dehydrogenase (Sso1218 in S. solfataricus P2 and SiRe_0376 in S. islandicus REY15A, having respectively 79% and 80% sequence identity with ST1116), they are embedded in a different genomic context as compared with S. tokodaii/S. acidocaldarius and nothing is known about their regulation. It is unknown which gene in S. acidocaldarius encodes a malonate semialdehyde dehydrogenase responsible for the second step in the β-alanine degradation pathway. In contrast, we have shown that gene clusters containing BarR homologues and homologues of putative β-alanine degradation enzymes occur in several other archaeal and some bacterial phyla (Fig. 1). This suggests that the β-alanine dependent regulation by an Lrp-like regulator as described in this work is not restricted to Sulfolobus and that it is more common than previously thought that Lrp-like TFs are involved in the regulation of central cellular metabolism and not only in amino acid metabolism.

Experimental procedures Strains and growth conditions Sulfolobus acidocaldarius DSM639 was cultivated aerobically at 75°C in Brock basal salts medium (Brock et al., 1972) supplemented with 0.2 (w/v)% tryptone and 0.2 (w/v)% D-xylose and adjusted to pH 3.5 with sulphuric acid. The uracil auxotrophic strain S. acidocaldarius MW001 (Wagner et al., 2012) was grown in the same medium supplemented with 10 μg ml−1 uracil. S. tokodaii strain 7 (DSM 16993) was grown in the same basic medium, but supplemented with 0.1 (w/v) % yeast extract and adjusted to pH 3.0. For growth tests, MW001 and MW001ΔbarR strains were cultivated in Brock medium supplemented with 0.2 (w/v)% casamino acids, 0.2 (w/v)% sucrose, 20 μg ml−1 uracil and, where appropriate, β-alanine at the indicated concentration. For growth on plates, Brock medium was solidified by adding 0.6 (w/v)% gelrite, 10 mM MgCl2 and 3 mM CaCl2. Plates were then incubated for 5–10 days at 75°C. Escherichia coli DH5α was used for propagation of plasmid DNA constructs. E. coli BL21(DE3) was used for heterologous overexpression of proteins. Microbial growth was followed by measuring the optical density at 600 nm (OD600). Genotypes of used strains are given in Table S1.

Construction of a barR deletion mutant A suicide disruption vector for Sa-barR (Saci_2136) gene deletion (pSVA406xΔbarR) was constructed by cloning 800 bp up- and downstream flanking regions of Sa-barR (using S. acidocaldarius DSM639 genomic DNA) into plasmid pSVA406, after fusing them by overlap PCR. An overview of all plasmids used in this work is given in Table S2. For all cloning purposes in this work, genomic DNA was extracted from 2 ml of culture and purified by magnetic bead purification with a QuickPick SML gDNA kit (Bio-Nobile). The sequences of the disruption vector and of all other constructs prepared in this work were verified by DNA sequencing. Methylation and transformation of the plasmid in strain MW001 were performed according to the procedure described before (Wagner et al., 2012), as were selection and screening of transformants. This resulted in an in-frame deletion mutant (MW001ΔbarR), leaving 15 bp of the barR gene intact (Fig. S2). All primer sequences are given in Table S3.

Western blotting Purified recombinant Sa-BarR (see below) was used to immunize rabbits for the production of polyclonal anti-SaBarR antibodies (Innovagen), which were affinity-purified before use. Cell-free extracts were prepared by harvesting cells at an OD600 of 0.4 as described before (Lassak et al., 2013). After separating cell-free extracts on SDS-PAGE, proteins were blotted on a PVDF membrane (Life Technologies) employing a mini-PROTEAN system (Bio-Rad) and immunodetection was performed using a WesternBreeze kit (Life Technologies) with a CDP-Star chemiluminescent substrate followed by exposure to an X-ray sensitive film or a CCD camera.

qRT-PCR For qRT-PCR analysis, total RNA was isolated from MW001 and MW001ΔbarR strains at an OD600 of 0.3 using an RNeasy mini kit (Qiagen) and residual DNA was removed by treatment with TURBO DNase (Ambion) according to manufacturer’s instructions. First-strand cDNA was synthesized from 1 μg RNA with a SuperScript III First-Strand Synthesis Super Mix kit (Invitrogen). Primers (Table S3) were designed with Primer3 Plus software (Rozen and Skaletsky, 2000). Real-time qPCR was carried out in a Bio-Rad iCycler as described before (Nguyen Duc et al., 2013). Biological triplicates were assayed and normalization was done with respect to the expression of reference genes Saci_0691 (encoding RNA polymerase subunit A) and Saci_1336 (encoding TBP). These genes were selected as having the most stable expression in the different growth conditions assayed here from a set of potential reference genes (Saci_0691, Saci_0693, Saci_0900, Saci_1191, Saci_1336 and Saci_1892) using BestKeeper software (Pfaffl et al., 2004).

Promoter activity assays Reporter gene constructs were prepared by fusing full-length and truncated versions of the barR-Saci_2137 intergenic © 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

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region in both orientations (Fig. 3B) to the S. solfataricus β-glycosidase (lacS) reporter gene in the E. coli/S. acidocaldarius shuttle vector pSVA1450 (Henche et al., 2012). SacII and NcoI restriction sites were used for cloning and all constructs were sequenced and methylated prior to transforming them into MW001 and MW001ΔbarR. Transformants were grown in the above-mentioned growth conditions and cells were collected at an OD600 of 0.4. LacS activity of the cell extracts was determined using the ortho-nitrophenyl-β-Dgalactopyranoside (ONPG) method as described previously (Jonuscheit et al., 2003; Peng et al., 2009). Total protein content was determined using a MicroBCA Protein Assay Kit (Pierce). One specific unit is defined as 1 mmol orthonitrophenyl produced per minute per mg total protein. Biological triplicates were assayed and data were corrected for background values obtained for MW001 and MW001ΔbarR transformed with pSVA1614, a promoterless control plasmid. Statistical analysis was performed with GraphPad Prism.

Chromatin immunoprecipitation The preparation of cross-linked and sheared chromatin was done as described before (Nguyen Duc et al., 2012). Chromatin immunoprecipitation (ChIP) was performed with polyclonal anti-BarR rabbit antibodies using Dynabeads M-280 sheep anti-rabbit IgG beads (Life Technologies) as described previously (Smollett et al., 2012). As a control, we also prepared an antibody-free mock sample. Subsequently, ChIP samples were purified with the iPure DNA extraction kit (Diagenode) and analysed by qPCR. Primers (Table S3) were designed with Primer3 software (Rozen and Skaletsky, 2000) and enrichment of the Sa-barR/Saci_2137 intergenic region was quantified by using a presumably unbound genomic region (part of the open reading frame (ORF) of Saci_0691, RNA polymerase subunit A) for normalization and comparing ChIP DNA to input DNA (sample taken before ChIP). As a negative control, enrichment of part of the ORF of Saci_1336 (encoding TBP) was also quantified.

repeating the 15000-g centrifugation. Inclusion body proteins were denatured by resuspending the pellet in buffer A [50 mM Na2HPO4 (pH 7.4), 0.5 M NaCl, 8 M urea, 0.4 M L-arginine, 20 mM imidazole] and incubating the suspension at room temperature during 2 h. Following a centrifugation at 25 000 g during 20 min, the supernatant was loaded on a 5 ml HisTrap HP column (GE Healthcare) and using an AKTA FPLC system (GE Healthcare) two subsequent linear gradients were set to perform on-column refolding and elution of BarR protein: a gradient between buffer A and B1 [50 mM Na2HPO4 (pH 7.4), 0.5 M NaCl, 0.4 M L-arginine, 20 mM imidazole] and a gradient between buffer B1 and B2 [50 mM Na2HPO4 (pH 7.4), 0.5 M NaCl, 0.4 M L-arginine, 0.5 M imidazole]. Finally, fractions containing BarR protein (Fig. S5) were pooled and dialysed into storage buffer [20 mM Tris (pH 8.0), 50 mM NaCl, 0.4 mM EDTA, 1 mM DTT, 1 mM MgCl2, 12.5% (v/v) of glycerol]. To examine the quaternary structure of St/Sa-BarR proteins, 0.5 mg of protein was loaded on a Superdex 200 gel filtration column (GE Healthcare) with as a mobile phase buffer 20 mM Na2HPO4 (pH 7.4), 150 mM NaCl. The following proteins were used for the molecular weight standard curve: RNase T1 (11 kDa), chymotrypsin (25 kDa), ovalbumin (43 kDa), albumin (66 kDa) and alcohol dehydrogenase (147 kDa).

Protein-DNA interaction assays Radiolabelled DNA was prepared by PCR, with one of the two oligonucleotides 5′-end-labelled with 32P using [γ-32P]-ATP (Perkin Elmer) and T4 polynucleotide kinase (Thermo Scientific). PCRs were performed using Taq DNA polymerase (Ready Mix, Sigma-Aldrich) and either genomic DNA or plasmid DNA as a template. Labelled DNA fragments were purified by polyacrylamide gel electrophoresis prior to subsequent experiments. EMSAs (Peeters et al., 2013), ‘in gel’ Cu-OP footprinting (Peeters et al., 2004), DNase I footprinting (Enoru-Eta et al., 2000) and depurination, depyrimidation and premethylation binding interference (Wang et al., 1998) was performed as described previously. All binding reactions contained an excess non-specific competitor DNA.

Recombinant protein overexpression and purification

Bioinformatic analyses and protein structure modelling

The Sa-barR and St-barR coding regions were cloned into pET24a vector using NdeI and XhoI restriction sites, thereby fusing a His-tag encoding sequence to the C-terminal end of the ORF. Site-directed mutagenesis of St-barR, yielding vectors for overexpression of St-BarRS132A and St-BarRT134A, was performed by overlap PCR mutagenesis. All resulting overexpression plasmids were transformed into E. coli BL21(DE3) and proteins were overexpressed with induction at an OD600 of 0.6 by adding 5 μM isopropyl-β-D-1thiogalactopyranoside (IPTG), followed by overnight growth. Cells were harvested by centrifugation and resuspended into a buffer containing 50 mM Tris (pH 7.5) and 10 mM EDTA. Subsequently, the cell suspension was sonicated during 15 min at 20% of the maximal amplitude (750 Watt VibraCell sonicator, Bioblock Sciences). After centrifugation during 15 min at 15 000 g, the pellet, containing BarR-rich inclusion bodies, was collected and washed twice with washing buffer [50 mM Tris (pH 7.5), 10 mM EDTA, 5 mM DTT, 2% Triton X-100, 0.5 M NaCl], thereby collecting the pellet each time by

Comparative genomic analyses were performed with Absynte (Despalins et al., 2011) in expert mode, with the Saci2136, Saci2137, ST1114, ST1115 or ST1116 protein sequence as input. Functional predictions of genes were made using the KEGG database (Wixon and Kell, 2000). T-coffee (Notredame et al., 2000) was used for protein sequence alignments. A homology model of the three-dimensional structure of St-BarR was generated using the SWISS-MODEL workspace (Arnold et al., 2006), with either the native Grp structure (PDB: 2EZW) or complexed Grp-Gln structure (PDB: 2E7X) as a template (Kumarevel et al., 2008). β-Alanine has been modelled into the ligand binding pocket using the program Wincoot (Emsley and Cowtan, 2004).

© 2014 John Wiley & Sons Ltd, Molecular Microbiology, 92, 625–639

Acknowledgements We gratefully acknowledge the scientific achievements and life of Rolf Bernander, who sadly passed away just before

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submission of this paper. This work was supported by the Research Council of the Vrije Universiteit Brussel, the Vlaamse Gemeenschapscommissie and by the Research Foundation Flanders (FWO-Vlaanderen). H.L. received a PhD scholarship from the China Scholarship Council-Vrije Universiteit Brussel (CSC-VUB) and E.P. is a postdoctoral fellow of the Research Foundation Flanders (FWO-Vlaanderen). R.B and A.C.L. were supported by a Swedish Research Council grant 621-2010-5551. M.v.W. was supported by DFG grant AL1206/3-1, A.O. and S.V.A. received intramural funds from the Max Planck Society. We declare that we have no conflicts of interest.

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