Regulation of alkane degradation pathway by a TetR ...

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Journal Code: MMI Article No: MMI13232 Page Extent: 22

Toppan Best-set Premedia Limited Proofreader: Emily Delivery Date: 13 Oct 2015 Copyeditor: Mildred

Molecular Microbiology (2015) ■

doi:10.1111/mmi.13232

Regulation of alkane degradation pathway by a TetR family repressor via an autoregulation positive feedback mechanism in a Gram-positive Dietzia bacterium

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Jie-Liang Liang,1 Yong Nie,1 Miaoxiao Wang,1 Guangming Xiong,2 Yi-Ping Wang,3 Edmund Maser2 and Xiao-Lei Wu1* 1 Department of Energy and Resources Engineering, College of Engineering, Peking University, Beijing, 100871, China. 2 Institute of Toxicology and Pharmacology for Natural Scientists, University Medical School, Schleswig-Holstein, Campus Kiel, Kiel, 24105, Germany. 3 State Key Laboratory of Protein and Plant Gene Research, College of Life Sciences, Peking University, Beijing, 100871, China.

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Summary n-Alkanes are ubiquitous in nature and serve as important carbon sources for both Gram-positive and Gramnegative bacteria. Hydroxylation of n-alkanes by alkane monooxygenases is the first and most critical step in n-alkane metabolism. However, regulation of alkane degradation genes in Gram-positive bacteria remains poorly characterized. We therefore explored the transcriptional regulation of an alkB-type alkane hydroxylase-rubredoxin fusion gene, alkW1, from Dietzia sp. DQ12-45-1b. The alkW1 promoter was characterized and so was the putative TetR family regulator, AlkX, located downstream of alkW1 gene. We further identified an unusually long 48 bp inverted repeat upstream of alkW1 and demonstrated the binding of AlkX to this operator. Analytical ultracentrifugation and microcalorimetric results indicated that AlkX formed stable dimers in solution and two dimers bound to one operator in a positive cooperative fashion characterized by a Hill coefficient of 1.64 (± 0.03) [kD = 1.06 (± 0.16) μM, kD′ = 0.05 (± 0.01) μM]. However, the DNA-binding affinity was disrupted in the presence of long-chain fatty acids (C10–C24), suggesting that AlkX can sense the concentrations of n-alkane degradation metabolites. A model was there-

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Accepted 25 September, 2015. *For correspondence. E-mail [email protected]; Tel. and Fax +86-10-62759047.

© 2015 John Wiley & Sons Ltd

fore proposed where AlkX controls alkW1 expression in a metabolite-dependent manner. Bioinformatic analysis revealed that the alkane hydroxylase gene regulation mechanism may be common among Actinobacteria.

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Introduction

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Alkanes are ubiquitous in nature and produced by many organisms. Microorganisms, including cyanobacteria, reportedly produce high proportions of heptadecane (Winters et al., 1969; Mcinnes et al., 1980), which makes n-alkanes available in natural habitats dominated by cyanobacteria (Shiea et al., 1990). Plant epidermal cells produce cuticular waxes to protect against water loss and pathogen infection (Samuels et al., 2008). These waxes are complex mixtures of straight-chain aliphatics with lengths of C20–C60 and typically constitute 20–60% of the cuticle mass (Heredia, 2003). Insects use the hydrocarbon backbone to produce pheromones (Tillman et al., 1999). Human skin cells excrete n-alkanes to form a waterproof barrier (Elias et al., 1990; Küster et al., 1991). Global transportation throughout the ages has slowly spread alkanes to most parts of the earth, aquatic via terrestrial and the surface via subsurface, although in low concentrations (Schirmer et al., 2010). The ubiquitous presence of alkanes has led to the natural existence of numerous and diverse alkane-degrading microorganisms, including both Gram-positive and Gram-negative bacteria and even Archaea (Bertrand et al., 1990; van Beilen et al., 2003; Wentzel et al., 2007; Tapilatu et al., 2010). Microbial degradation processes have accomplished natural alkane attenuation and shifted the compositions of crude oil constituents in oil reservoirs (Voordouw, 2011; CastorenaCortés et al., 2012), ultimately contributing to crude oil formation and global carbon cycling, as well as shaping the microbial community in the biosphere. Moreover, degradation of alkanes in plants and human skin may cause a loss of host defense, and subsequently result in microbial infection in alkane-producing host. Indeed, some bacterial strains, such as Pseudomonas, Mycobacterium and Dietzia, can degrade n-alkanes and are well-known opportunistic pathogens in humans (Cole et al., 1998; Alonso

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et al., 1999; Koerner et al., 2009). In addition, microbial n-alkane degradation is also central to industrial applications such as biodegradation and microbial enhanced oil recovery (Brown, 2010). A large number of Gram-positive and Gram-negative bacteria degrade n-alkanes of various chain lengths. These bacteria include Acinetobacter (Sakai et al., 1994; Throne-Holst et al., 2007; Sun et al., 2012), Alcanivorax (van Beilen et al., 2004; Schneiker et al., 2006), Bacillus (Kato et al., 2001), Burkholderia (Yuste et al., 2000), Pseudomonas (Smits et al., 2002), Rhodococcus (Whyte et al., 2002), Dietzia (Bihari et al., 2011; Wang et al., 2011; Procópio et al., 2012), Gordonia (Kotani et al., 2003), Geobacillus (Feng et al., 2007) and Amycolicicoccus (Nie et al., 2013). Alkane hydroxylases are reportedly the key and rate-limiting enzymes for aerobic alkane degradation. Among the three major alkane hydroxylases functionally identified thus far, the best characterized is integralmembrane non-heme diiron monooxygenase (AlkB), which degrades middle- and long-chain n-alkanes and is found with high sequence diversity in both Gram-positive and Gram-negative bacteria (van Beilen et al., 2002; 2003; Smits et al., 2002; Kuhn et al., 2009). Indeed, many diverse alkB genes have recently been detected and are widely distributed in microorganisms and environments (Nie et al., 2014). In contrast to the intensely investigated functions of the AlkB hydroxylases, the mechanism of alkane hydroxylase regulation has been clarified most thoroughly in the OCT plasmid of Pseudomonas putida GPo1 (Rojo, 2009), which contains two gene clusters, alkBFGHJKL and alkST (van Beilen et al., 1994; 2001). In the presence of inducers such as n-alkanes, the LuxR family activator AlkS induces the expression of both gene clusters, whereas when inducers are absent, AlkS binds to its own promoter and represses its expression (Canosa et al., 2000). The only related study reported thus far in Grampositive alkane-degrading bacteria identified the transcriptional start point and described promoter-inducing activity in Rhodococcus sp. SCP1 (Cappelletti et al., 2011). However, an investigation of 137 environmental metagenomes reveals that Actinobacteria-related alkB genes are significantly enriched and abundant in terrestrial habitats (Nie et al., 2014). This result and the abundance of Actinobacteria in terrestrial environment (Venter et al., 2004; Giovannoni and Stingl, 2005) suggest the importance of Gram-positive bacterial n-alkane degradation, at least in terrestrial environment. A search of the GenBank database revealed a genetic arrangement, i.e., a putative TetR family regulator (TFR) gene located downstream of alkB, in the genomes of Gram-positive bacteria such as Prauserella rugosa NRRL B-2295, Mycobacterium tuberculosis H37Rv (Whyte et al., 2002), Nocardioides sp. strain CF8 (Hamamura et al., 2001), Gordonia

sp. Socg (Lo Piccolo et al., 2011) and a few Rhodococcus strains (Na et al., 2005; McLeod et al., 2006; Sekine et al., 2006). This finding might imply a link between TFRs and AlkB in Gram-positive bacterial alkane degradation. TFRs compose the third most common transcriptional regulator family in bacteria and regulate a wide range of cellular activities, including antibiotic production, multidrug resistance, amino acid metabolism, osmotic stress, pathogenicity and development (Ramos et al., 2005; Yu et al., 2010b). However, whether TFRs can regulate alkB expression is unknown. The present study showed for the first time the regulation mode of n-alkane metabolism in a Gram-positive bacterium, Dietzia sp. DQ12-45-1b. The putative TFR, AlkX (with the GenBank accession number being AEM66515.1), represses the transcription of the alkB-like gene, alkW1, by cooperatively binding to its promoter. The mechanism seems common in Dietzia and may be widely used by other Actinobacteria such as Rhodococcus and Mycobacterium based on the bioinformatics analysis. This repression was diminished by fatty acids, which are the intermediates of n-alkane degradation, via an autoregulatory positive feedback mechanism.

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Results

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AlkX is a repressor of alkW1 and n-alkane releases the repressive effect

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We annotated a putative TetR family transcriptional regulatory gene (alkX) located 19 bp downstream of an alkBtype alkane hydroxylase-rubredoxin fusion gene (alkW1) with the same orientation on the chromosome of Dietzia sp. DQ12-45-1b (Nie et al., 2011). Sequence analyses revealed that AlkX harbors a putative helix–turn–helix (HTH) domain (PF00440), the sequence of which places it in the TFR family of DNA binding proteins. Reverse transcription polymerase chain reaction (PCR) analysis showed that alkX and alkW1 were cotranscribed in Dietzia sp. DQ12-45-1b cells grown with n-hexadecane as sole carbon source (Fig. S1). In this way, alkW1 and alkX formed an operon involved in n-alkane oxidation. To investigate the possible regulatory role of AlkX on AlkW1 expression in Dietzia sp. DQ12-45-1b, we constructed an enhanced green fluorescent proteinkanamycin resistance (EGFP-Kan) gene insertion mutant (designated DQ4502; Fig. 1A, Table S1) and analyzed the expression level of AlkW1 in both the wild-type strain and alkX mutant. The AlkW1 expression level detected in the alkX mutant was similar to that in the wild-type strain induced by n-hexadecane. However, when glucose was used as sole carbon source, the AlkW1 expression level was much higher than that detected in the wild-type strain (Fig. 1B). When complemented with plasmid pXL1817

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© 2015 John Wiley & Sons Ltd, Molecular Microbiology

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AlkX regulates alkane hydroxylase gene expression 3

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Fig. 1. AlkX represses the expression of alkW1 in vivo. A. Schematic diagram of mono-homologous recombination to construct the alkX mutant. Fusion PCR was performed to synthesize a DNA fragment for the fusion of an EGFP gene, a Kan gene and a 400 bp partial alkX gene fragment. After the linear DNA fragment was introduced into Dietzia sp. DQ12-45-1b, mono-homologous recombination of alkX occurred in the strain. B. Western blot results for the wild-type and mutant strains cultured in minimal medium with glucose or n-hexadecane as sole carbon source. glu, glucose; n-C16, n-hexadecane; WT, Dietzia sp. DQ12-45-1b (wild-type strain); K, DQ4502 strain (alkX mutant); X, DQ4504 strain (alkX mutant complemented with plasmid pXL1817 containing alkX gene); C, DQ4503 strain (alkX mutant complemented with control plasmid pNV18Sm without alkX gene). The bands marked with a solid arrow refer to AlkW1 protein, which has a molecular mass of 57 kDa.


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Fig. 2. Promoter analysis of alkW1. A. The alkW1 promoter. The −10 and −35 regions are shown in red. The transcriptional start site is shown in bold red and indicated with an arrow. The AlkW1 start codon is also indicated with an arrow. B. Determination of the alkW1 promoter region with deletion analysis. The PCR fragments were cloned into EGFP reporter vector pXL1803 for RFU measurement with EcoRI and BamHI restriction sites. The upper left panel shows the construction of the 5′ promoter deletions, whereas the lower left panel shows the construction of the 3′ promoter deletions. The right panel shows the RFU values of cells with 5′ and 3′ deletions measured after 5 h of incubation. The RFU values are shown as the means ± standard deviations (n = 3). The x-axis labels refer to E. coli DH5α transformed with the plasmids (Table S1). **P < 0.01 compared with pXL1803 control as determined by Student’s t-test. C. Dietzia sp. DQ12-45-1b alkW1 promoter activity induced by n-alkanes of various chain lengths. Values represent the means ± standard deviations (n = 3). *P < 0.05; **P < 0.01 compared with glucose control as determined by Student’s t-test.

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(Table S1), the expression pattern of AlkW1 in the alkX mutant was similar to that observed in the wild-type strain. These results suggested that AlkX serves as a repressor for alkW1 expression and that n-alkane releases its repressive effect. As alkX was cotranscribed with alkW1, it negatively autoregulated its own expression at the same time. Negative autoregulation is common in many transcription families but has been less explored in TFRs (Cuthbertson and Nodwell, 2013).

Characterization of the alkW1 promoter and its inducibility by a broad range of n-alkanes

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The results of 5′ rapid amplification of complementary DNA ends (5′ RACE) analysis revealed that the transcriptional start site (TSS) of alkW1 is localized at the di-deoxy nucleotide A, which is 48 bp upstream of the alkW1 start codon (Fig. 2A). The putative −10 (TAACCT) and −35 (TAGACA) regions, with a spacing of 17 bp, were pro© 2015 John Wiley & Sons Ltd, Molecular Microbiology

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posed as the core promoter sequences recognized by σ70-RNA polymerase (RNAP). For localization of the alkW1 promoter, various deleted fragments of the 255 bp intergenic region upstream of alkW1 were incorporated into the reporter plasmid pXL1803, forming 5′ set and 3′ set plasmids (Table S1 and Fig. 2B), and transformed into Escherichia coli DH5α. Compared with the pXL1803 control, the E. coli recombinants containing the 50 bp to 150 bp deletions from the 5′ terminus and 10 bp to 40 bp deletions from the 3′ terminus produced 17.3- to 41.8-fold and 8.1- to 28.9-fold higher fluorescence respectively (P < 0.01). However, when the 200 bp or 60 bp fragment was deleted from the 5′ terminus or 3′ terminus, respectively, no significant fluorescence was detected (Fig. 2B). These results along with those of 5′ RACE analysis clearly revealed that the alkW1 promoter is located between −57 and +8 of alkW1 TSS. To investigate alkW1 promoter activity further, we constructed the PalkW1-lacZ fusion vector containing the 255 bp fragment upstream of alkW1 and a 117 bp fragment in alkW1 and transformed into Dietzia sp. DQ1245-1b to measure alkW1 promoter activities in response to various n-alkanes. The β-galactosidase activity of the cells induced by C8 to C36 n-alkanes were 2.3- to 12.4fold higher (P < 0.01 or 0.05) than that of cells grown in glucose (Fig. 2C). These results agreed with measurements of in vivo alkW1 expression (Nie et al., 2011), further demonstrating that promoter activity is induced by a broad range of n-alkanes. Because the highest inducing capacity was observed with n-hexadecane (2,267 nmol mg of protein−1 min−1; Fig. 2C), the experiments described below were carried out with n-hexadecane as the inducer.

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Direct binding of AlkX to the inverted repeat upstream of alkW1

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The recombinant AlkX protein with a 6 × His tag at the N terminus was expressed in E. coli and purified for promoter binding analysis (Fig. S2). To investigate whether AlkX interacts directly with alkW1 promoter, we carried out in vitro DNase I footprinting assays. The results showed that AlkX bound directly to the −62 to −6 region according to alkW1 TSS, which covered most of alkW1 promoter (between −57 and +8 of alkW1 TSS) (Fig. 3A). DNA sequence analysis revealed that the protein-protecting region included an unusually long 48 bp operator (5′TGGACAAA-N11-TGTCTAGACA-N11-TTTGTCTA-3′), which covered both −10 and −35 regions. Within the operator, there were four special sequences, which were designated as OP1 (5′-TGGACAAA-3′), OP2 (5′TTTGTCTA-3′), OP3 (5′-TGTCT-3′) and OP4 (5′-AGACA3′), with OP1 and OP2 being palindrome while OP3 and OP4 were also palindrome (Fig. 3A). In addition, OP1 and

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OP3 were presented in half of the operator, i.e., (5′TGGACAAA-N11-TGTCT-3′), whereas OP2 and OP4 were presented in another half operator, i.e. (5′-AGACAN11-TTTGTCTA-3′). Electrophoretic mobility shift assay (EMSA) results also confirmed that AlkX binds to the 70 bp DNA fragments containing the 48 bp operator (Fig. 3B, Table S2). When the protein concentration was low (0.1 μM), there was only one faster shift band; when protein concentration was up to 0.5 μM, there were two shift bands; with an even higher protein concentration (5 μM), there was almost only one slower shift band (Fig. 3B). However, control experiments showed that AlkX did not bind to the randomly selected DNA sequences, which indicated AlkX bound to the 70 bp DNA fragments in a specific manner (Fig. S3). Taken together, the results indicated the existence of two different AlkX-DNA complexes, in which either half of the operator, or the whole operator could be occupied by AlkX.

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A pair of AlkX dimers binds to the operator

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First, sedimentation velocity experiment was carried out with AlkX to evaluate protein homogeneity and selfassociation. The protein concentration (in the 4–35 μM range) had no significant impact on the sedimentation behavior (Fig. 4A and Fig. S4), which indicates that AlkX formed stable dimers in solution under the experimental conditions we used. Taking the case of 14 μM AlkX as an example, a major species (> 92.3% of total peak area in the sedimentation coefficient distribution) was found sedimented with a standard s-value of 3.13 (± 0.20) S, indicating the molecular weight of protein was 50 982 (± 5,922) Da. This is compatible with the molecular weight of the AlkX dimer being 52 470 Da, which is calculated based on the sequence-derived molecular weight of the AlkX monomer being 26 235 Da. Second, velocity sedimentation experiments were carried out with a mixture containing 1–4 μM AlkX and 0.4–1 μM 52 bp operator (Table S2) with the protein/DNA ratios varied from 1:1 to 10:1. The results showed the existence of two different nucleoprotein complexes, i.e., one dimer per operator and two dimers per operator (Fig. 4 and Fig. S4). For example, the velocity sedimentation experiments of a mixture containing 4 μM AlkX and 0.4 μM DNA (10:1) showed a biphasic behavior with two separate peaks in the sedimentation coefficient distribution (Fig. 4B). The slower sedimentation species, characterized by a standard s-value of 3.23 (± 0.44) S, corresponded to unbound AlkX dimers. The faster species sedimented with a standard s-value of 6.96 (± 0.57) S and represented the nucleoprotein complex. From the differences between the integrated peak areas of the free protein obtained in the absence and presence of DNA

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Fig. 3. DNA-binding assays for AlkX. A. Sequence of the protected region of alkW1 identified with DNase I footprinting assays. The 255 bp DNA fragment containing the alkW1 promoter sequence was labeled with 3′ HEX, incubated with AlkX or BSA, and subjected to DNase I digestion and fragment length analysis. The fluorescence signal of the labeled DNA fragments is plotted against the sequence of the fragment. The blue line shows the BSA-protected trace, and the red line shows the AlkX-protected trace from the top strands. The palindromic motifs are shown in red. The figure below summarizes the DNase I footprinting analysis data. The protected regions are boxed. The OP1 and OP2 sequences are shown with red arrows, whereas OP3 and OP4 sequences are shown with black arrows. The −10 and −35 regions are shown in red, and the alkW1 transcriptional start site is indicated by an arrow. B. AlkX binds to the 70 bp DNA fragments including the palindromic sequences upstream alkW1 gene. The two AlkX-DNA shift bands are indicated with solid and dotted arrows. Free DNA is indicated with a dashed arrow.



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Fig. 4. Analytical ultracentrifugation analysis of AlkX and AlkX-DNA complex. A. Sedimentation coefficient distributions c(s) from the analysis of the sedimentation profiles of 14 μM AlkX. The major species is characterized by s-value of 3.13 (± 0.20) S (indicated with a dotted and solid arrow). B. Sedimentation coefficient distributions c(s) from the analysis of the sedimentation profiles of the mixture of 4 μM AlkX and 0.4 μM DNA fragment. The solid curve shows the sample of free AlkX with s-value of 3.22 (± 0.19) S (indicated with a dotted and solid arrow) and AlkX-DNA mixture with s-values of 6.96 (± 0.57) S (indicated with a dash arrow) respectively. C. Sedimentation coefficient distributions c(s) from the analysis of the sedimentation profiles of the mixture of 2 μM AlkX and 1 μM DNA fragment. The solid curve shows the sample of free DNA fragments with s-value of 3.69 (± 0.36) S (indicated with a solid dashed arrow) and AlkX-DNA mixture with s-values of 5.92 (± 0.43) S (indicated with a solid arrow) respectively. D. Sedimentation coefficient distributions c(s) from the analysis of the sedimentation profiles of 1 μM DNA. The solid curve shows the sample of free DNA fragments with s-value of 3.60 (± 0.27) S (indicated with a solid dashed arrow).

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fragments, the amount of DNA-bound protein was determined as two AlkX dimers per operator. Meanwhile, the velocity sedimentation experiments of a mixture containing 2 μM AlkX and 1 μM DNA (2:1) showed a different biphasic behavior (Fig. 4C). The slower species, characterized by a standard s-value of 3.60 (± 0.27) S, corresponded to free 52 bp operator (Fig. 4D). However, the faster species sedimented with a standard s-value of 5.92 (± 0.43) S and represented another nucleoprotein complex, which indicated one AlkX dimer bound to per operator. These results were compatible with those obtained by EMSA (Fig. 3B). The faster species in the © 2015 John Wiley & Sons Ltd, Molecular Microbiology

AUC corresponded to the slower shift band in EMSA, which suggested two dimers bound to one operator, whereas the faster shift band showed in EMSA suggested the binding condition that one dimer bound to one operator.

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Mutation analysis to determine the roles of palindromic motifs

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Based on the AUC results as well as the operator sequence analysis, we proposed that AlkX might specifically bind to the OP1–OP4 sequences. To further test our

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8 J.-L. Liang et al. ■ 1 2 3

Table 1. Thermodynamic parameters for the binding of AlkX to the operator.

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1.64 ± 0.03

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Notes: The Hill coefficient, nH, was calculated using: nH = 2/[1 + (k/k′)] (Krell et al., 2007).

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hypothesis, we designed a series of mutated DNA fragments for EMSA analysis (Fig. 5A). When AlkX was incubated with the wild-type DNA fragment, only the slower shift band was observed (Fig. 5A, lane W) without a free DNA band, indicating that the operator was fully occupied with AlkX. When we first completely mutated both OP1 and OP3, only the faster shift band was observed (Fig. 5A, lane P13), suggesting that only one dimer bound to OP2 and OP4. When only OP3 was mutated, both the faster and slower AlkX-DNA shift bands were observed (Fig. 5A, lane P3), which suggested that OP3 mutation did not hinder AlkX dimer binding to OP1, OP2 and OP4, but with slightly disrupted binding affinity (because free DNA band was observed). When OP1 was completely mutated, only the faster shift band was observed (Fig. 5A, lane P1) as in the case of P13, which suggested that the dimer only bound to OP2 and OP4 and the binding to OP1 and OP3 was disrupted without OP1. Therefore, compared with OP3, OP1 seemed to have a more important role in AlkX-DNA binding. Meanwhile, the mutation of another half of the operator (OP2, OP4, and both OP2 and OP4) exhibited the similar results as OP1, OP3 and both OP1 and OP3 respectively (Fig. 5A, lane P4, P2 and P24). Our results indicated that OP1–OP4 could be the possible binding sites. Moreover, when OP1, OP3 and OP4 were simultaneously mutated, only the faster band was observed as in case of lane P13 (Fig. 5A, lane P134) but with a greater amount of free DNA. The result indicated that OP1 and OP3 might be bound by one dimer, and OP4 might be bound by another dimer (Fig. 5B). When OP1, OP2 and OP3 were mutated, no protein-binding band was observed (Fig. 5A, lane P123), which indicated that OP2 seemed to be more important than OP4. All these results indicated that OP1 and OP2 might be essential for specific binding and that binding affinity was severely disrupted in their absence, whereas OP3 and OP4 might provide appropriate spacing for AlkX binding. Furthermore, we performed the AUC with different mutated DNA incubated with AlkX. Our results showed that only the initial (one dimer per operator) nucleoprotein complex existed when OP1 or OP2 was mutated and both OP1 and OP3, or both OP2 and OP4 were mutated (Fig. 5C– F). The results were in accordance with those in EMSA (Fig. 5A). Taken together, the results may support our

assumption that one dimer bound to OP1/OP3, whereas another dimer bound to OP2/OP4 (Fig. 5B).

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Cooperative binding of AlkX to its operator

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Because OP1 and OP2 seemed to equally have a more critical role in AlkX binding, we investigated the role of OP1 in alkW1 regulation in vivo. Plasmid pXL1901 (Table S1) was cotransformed into E. coli DH5α with plasmid pXL1815 (with OP1-OP4 included) or pXL1816 (with OP1 mutated; Table S1). Compared with that of the control E. coli DH5α (pXL1815, pUC19), the expression levels of green fluorescence in E. coli DH5α (pXL1815, pXL1901) were 20.9-fold decreased (Fig. 6A), again confirming that AlkX repressed alkW1 transcriptional expression. However, the expression level of E. coli DH5α (pXL1816, pXL1901) was only 1.7-fold lower than that of E. coli DH5α (pXL1816, pUC19), which was considered relatively weak repression. In addition, the expression level in E. coli DH5α (pXL1815, pXL1901) was 11.7-fold lower than that in E. coli DH5α (pXL1816, pXL1901), indicating that the mutated OP1 diminished the DNA binding affinity of AlkX. The results agreed with the EMSA results and confirmed the importance of OP1. Moreover, the results also indicated that two dimers bound to the operator in a cooperative way. To further confirm our assumption, we used isothermal titration calorimetry (ITC) to characterize the AlkX–DNA interaction by titrations of the same 52 bp DNA fragment as used in AUC with increasing concentration of AlkX. Binding of the operator by AlkX was characterized by exothermic heat changes (Fig. 6B, upper panel), and the integrated peak areas for the binding reaction showed biphasic behavior (Fig. 6B, lower panel). Analysis of these data with the models based on the binding of a ligand to one or two binding sites in Origin software resulted in an unacceptable curve fit. However, we analyzed the data with the model for cooperative binding of two dimers to one operator (Krell et al., 2007) gave a satisfactory curve fit. It was thus assumed that binding of AlkX to its operator occurred in a cooperative fashion, with the thermodynamic parameters listed in Table 1. The initial dissociation constant for binding of the first AlkX dimer to 52 bp DNA fragment was calculated as 1.06 (± 0.16) μM, whereas dissociation constant of the second dimer was 0.05

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Fig. 5. The palindromic sequences in the operator had different roles of AlkX-DNA binding. A. Mutation analysis of different parts in the operator. DNA fragments of both wild type (W) and mutated (P3-P24) designed for EMSA are shown and the OP1-OP4 sequences are indicated with red or black arrows. The mutated sequences are shown in bold red. Also shown are the EMSA results for the DNA binding activity of AlkX on various DNA fragments. The two AlkX-DNA shift bands are indicated with solid and dotted arrows. Free DNA is indicated with a dashed arrow. B. Schematic illustration of AlkX-DNA binding fashion. Two AlkX dimers bind to one 48 bp operator. One dimer binds to OP1 (indicated as a red arrow) and OP3 (indicated as a black arrow) with one monomer binding to OP1 and the other binding to OP3, whereas the other dimer binds to OP2 (indicated as a red arrow) and OP4 (indicated as a black arrow) with one monomer binding to OP2 and the other binding to OP4. (C)-(F) Sedimentation coefficient distributions c(s) from the analysis of the sedimentation profiles of the mixture of 4 μM AlkX and 1 μM mutated operator. The solid arrow indicated the initial nucleoprotein complex (one dimer per operator).



Fig. 6. Cooperative binding of AlkX to the operator. A. Green fluorescence expression detection from the cotransformed cells. Reporter vectors with the alkW1 promoter and AlkX expression vectors were cotransformed into E. coli DH5α to measure green fluorescence expression. 1: pXL1815 + pUC19; 1t: pXL1815 + pXL1901; 2: pXL1816 + pUC19; 2t: pXL1816 + pXL1901. Values represent the means ± standard deviations (n = 3). **P < 0.01 compared with relative control as determined by Student’s t-test. B. Cooperative binding of AlkX to 52 bp DNA fragment containing the 48 bp operator monitored by ITC. Upper panel: injection of a single 0.5 μl aliquot and a series of 2 μl aliquots of 87.7 μM AlkX (dimer) into the 52 bp DNA fragment (6.0 μM). Lower panel: integrated and corrected peak areas of titration of DNA fragment with AlkX in the upper panel. The black crosses represent the original data and the red circles represent the corresponding data generated by the fitting curve. Derived thermodynamic parameters are listed in Table 1.

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(± 0.01) μM. The Hill coefficient was commonly used as a measurement of cooperativity, and the value of more than 1 is indicative of positive cooperative system (Krell et al., 2007). The Hill coefficient was 1.64 (± 0.03) in our results, which proved the positive cooperative binding of AlkX to its operator.

Long-chain fatty acids (LCFAs) specifically modulate the DNA binding capability of AlkX

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Because alkW1 promoter activity was significantly induced by n-alkanes in Dietzia sp. DQ12-45-1b, the regulatory effect of n-alkanes on AlkX-DNA binding activity was determined with EMSA. However, n-tetradecane and n-hexadecane did not inhibit AlkX-DNA binding (Fig. S5A). Because n-alkanes are reportedly oxidized through n-alcohols to fatty acids and, finally, undergo β-oxidation (Fig. S5B) (Feng et al., 2007), the regulatory effects of 1-alcohols and fatty acids on AlkX-DNA binding activity were also evaluated with EMSA. Interestingly, when the myristic acid or palmitic acid were incubated with AlkX and DNA fragment, protein–DNA interaction was disrupted, resulting in the release of free DNA fragment, whereas tetradecanol or hexadecanol did not interfere with AlkXDNA binding activity (Fig. S5A). Effect of titration of palmitic acids on AlkX-DNA binding affinity showed that 15 μM palmitic acid could partially dissociate the AlkX-DNA complex, and the concentration of palmitic acid able to completely dissociate the AlkX-DNA complex was 30 μM (30-fold higher than protein concentration) (Fig. 7A). Furthermore, the same and even higher concentration of palmitic acid (15–150 μM) did not interfere with the binding affinity of RNAP to alkW1 promoter (Fig. S6), which suggested that palmitic acid might be the true effector of the AlkX. Moreover, we used other LCFAs (with carbon atoms being C10, C12, C14, C18, C20 and C24) to determine the ligand profile of AlkX. The results showed that all these LCFAs could modulate the DNA-binding affinity, and the palmitic acid and myristic acid had the strongest impact (Fig. 7B). In addition, we also test the effects of other different small molecule acid compounds like acetic acid, propanoic acid, lactic acid and pyruvic acid, and our results showed that these acids could not modulate the AlkX-DNA binding affinity (Fig. S7). To test whether LCFAs could also de-repress alkW1 expression in Dietzia sp. DQ12-45-1b, we tested the β-

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Fig. 7. LCFAs modulate the AlkX-DNA interaction. A. Effects on AlkX-DNA binding affinity by titrations of palmitic acid. The concentrations of palmitic acid varied from 0.75 μM to 150 μM. The two AlkX-DNA shift bands are indicated with solid and dotted arrows. Free DNA is indicated with a dashed arrow. B. Effects on AlkX-DNA binding affinity by different LCFAs. The two AlkX-DNA shift bands are indicated with solid and dotted arrows. Free DNA is indicated with a dashed arrow. C. Dietzia sp. DQ12-45-1b alkW1 promoter activity induced by 1-hexadecanol and palmitic acid. Values represent the means ± standard deviations (n = 3). **P < 0.01 compared with glucose control, as determined by Student’s t-test.

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galactosidase activities of strains grown on 1-hexadecanol and palmitic acid. Interestingly, alkW1 promoter activities induced by 1-hexadecanol and palmitic acid were about 2.6-fold higher than that caused by glucose (P < 0.01; Fig. 7C). These results indicated that, like n-hexadecane, metabolic products derived from n-hexadecane degradation could be inducers of alkW1. Furthermore, palmitic acid was also detected in cell lysate of Dietzia sp. DQ12-45-1b by gas chromatography-mass spectrometry (GC-MS) when cells were cultivated in minimal medium with n-hexadecane as sole carbon source (Table S4). To further test these results, we constructed a lacZ-fusion reporter vector with alkW1 promoter and alkX gene and transformed into the alkB-deficient mutant Pseudomonas fluorescens KOB2Δ1 and its wild-type strain P. fluorescens CHA0. The β-galactosidase results showed that n-alkanes (C12–C16) could induce the alkW1 promoter activity in the CHA0 strain but could not induce that in the alkB-deficient mutant (Fig. S8). Taken together, the results indicated that it was not n-alkanes as commonly believed, but LCFAs that © 2015 John Wiley & Sons Ltd, Molecular Microbiology

might bind to AlkX, release it from DNA, and consequently initiate alkW1 expression, which is indicative of a product positive feedback mechanism.

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Highly conserved sequences of alkW1-alkX operons in Dietzia strains and some Actinobacteria

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The corresponding alkB-tetR regions in Dietzia sp. UCDTHP, Dietzia sp. E1, D. alimentaria 72 and D. cinnamea P4 have the same alkW1-alkX gene arrangement as that in Dietzia sp. DQ12-45-1b. The AlkB and TetR proteins of these five Dietzia strains share high amino acid sequence similarities, and the palindromic motifs are also highly conserved, suggesting a potentially similar alkB regulation mechanism among the strains. Furthermore, we collected and analyzed 22 additional Dietzia strains originally isolated from geographically distant places worldwide, including type strains of Dietzia species (Table S3). All the 27 strains share the same gene arrangement and have the same inverted repeat (5′-TGGACAAA-N11-

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TGTCTAGACA-N11-TTTGTCTA-3′) in the putative alkB promoter region (Fig. S9). Moreover, phylogeny based on AlkB and TetR amino acid sequences showed that the proteins in Dietzia clustered together (Fig. S10 and S11) with the sequence similarities of 65–99% and 82–99% respectively. The results indicated that the transcriptional regulation mechanism of alkB expression was potentially common in Dietzia strains. Besides, phylogenetic analysis also showed that AlkW1 and AlkX are closely related to the proteins from other Gram-positive bacteria such as Rhodococcus and Mycobacterium (Fig. S10 and S11). In addition, the same gene arrangement of alkW1-alkX in Dietzia sp. DQ12-45-1b was shared by the Gram-positive bacteria such as Prauserella, Nocardioides, Rhodococcus and Mycobacterium (Fig. S12), also suggesting the cotranscription of alkB and tetR. The conserved inverted repeats (5′-TTTACAAA-N9-TTTGTCTA-3′) were also found in Mycobacterium, Rhodococcus, Nocardia and Gordonia (Fig. S13), which was similar to the OP1 and OP2 motifs in Dietzia species. Moreover, these conserved sequences may be the binding sites for TFRs because the HTH domains of these TFRs share high similarities. Taken together, the similarities in alkW1 transcriptional regulation mechanisms might be common not only in Dietzia strains but also in other Gram-positive bacteria such as Actinobacteria.

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Discussion

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Alkane hydroxylase and the regulation of its expression are central to alkane degradation in natural biogeochemical processes and industrial applications. As destroying alkanes can have negative impacts on both plant and human hosts, alkB gene regulation may be also important to the pathogenesis as well as to clinical cures for infections caused by Dietzia pathogens. As members of the third most common transcriptional regulator family in bacteria, TFRs have broad roles in prokaryotic physiology (Ramos et al., 2005; Cuthbertson and Nodwell, 2013), but the regulation of TFRs in n-alkane metabolism has never been reported before. The present study is also the first to demonstrate alkB regulation in Gram-positive bacteria experimentally, which differs completely from those observed in Gram-negative bacteria. First, the reported regulators in Gram-negative bacteria mostly belong to AraC (Ratajczak et al., 1998; Tani et al., 2001; Liu et al., 2011) or LuxR family (van Beilen et al., 2001; 2004), which serves as activators instead of TetR family repressors. Second, some of the regulators in Gramnegative bacteria can directly respond to n-alkanes and induce alkB gene expression (Marin et al., 2001; Tani et al., 2001), whereas AlkX interacts with the intermediate metabolite palmitic acid and de-represses alkW1 expres-

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sion. Third, those regulators are divergently transcribed with alkB (Whyte et al., 2002) while AlkX is cotranscribed with alkW1.

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Cooperative binding of AlkX to an unusually long 48 bp operator upstream alkW1

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There are two DNA-binding fashions in TFRs. The TFRs in the first class bind to the operators as a dimer and those in the second class bind as a pair of dimers (Bhukya et al., 2014). Our results showed that AlkX cooperatively bound to the unusually long 48 bp operator as a pair of dimers (Figs 3, 4 and 6). One of the striking difference of the two binding fashions is that the TFRs belonging to the second class in general [QacR, 28 bp (Schumacher et al., 2002), CgmR, 32 bp (Itou et al., 2010), MS6564, 31 bp (Yang et al., 2013), SlmA, 20 bp (Tonthat et al., 2013) and CprB, 22 bp (Bhukya et al., 2014)] recognize longer DNA sequences than those in the first class [TetR, 15 bp (Orth et al., 2000), SimR, 17 bp (Le et al., 2011) and DesT, 17 bp (Miller et al., 2010)]. In fact, this kind of cooperative effect is often found in the binding of relatively small proteins, which are often in the form of multimers, to different sites of the relatively long operators, not only in the TFRs, but also in other protein families like MetJ (Somers and Phillips, 1992), TrpR (Lawson and Carey, 1993) and DtxR (White et al., 1998). The cooperative binding may contribute to energetic DNA binding and saturation of the operator at low protein concentrations in cells (Smith and Sauer, 1995). Although further structural studies are still needed to address our proposed model, our EMSA results are indicative of a different binding mode (Fig. 5A) for AlkX compared with other TFRs that bind as a pair of dimers like QacR (Schumacher et al., 2002), CgmR (Itou et al., 2010) and CprB (Bhukya et al., 2014) or in other protein family like DxtR (Chen et al., 2000). First of all, our results indicated that the homodimer of AlkX may bind to both OP1 and OP3 at the same time (Fig. 5A); meanwhile, the homodimers of other TFRs like QacR, CgmR and CprB bind to equivalently OP1 and OP4 instead. The indicative different binding fashion of AlkX might allow longer DNA binding to the extent of 48 bp. On the other hand, the EMSA results of AlkX-DNA complex showed that two bands were observed in the proper protein concentration; however, the QacRDNA complex in the EMSA exhibited only one band regardless of the protein concentration (Grkovic et al., 1998). The QacR-DNA structure also indicated that QacR could bind to the operator only when two pairs of dimers functioned together. On the contrary, AlkX could bind to the operator both as a dimer and as a pair of dimers (Figs 3 and 5), in which case, the binding of the first dimer to the operator allowed the second dimer to bind more easily (Fig. 6). These results indicated that AlkX could bind to the operator and thus the RNAP open complex formation was pre-

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AlkX regulates alkane hydroxylase gene expression 13 1 2

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vented even at low protein concentration. This cooperative binding of AlkX could be important for Dietzia strains, which are generally detected in harsh environments – for example, with high pH, high osmotic pressure or petroleum related (Mayilraj et al., 2006; Khodaiyan et al., 2007; Li et al., 2009; Kim et al., 2011; Sun et al., 2014), where nutrients such as available nitrogen and phosphorus are less abundant. Therefore, these bacteria usually maintain low protein concentrations for better adaptation to these environments even though n-alkanes are ubiquitous. However, further study is needed to clarify the mechanism.

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Novel product positive feedback regulation of alkW1 by AlkX

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For the first time, we found that LCFAs might be the direct inducers of the alkW1-alkX promoter (Fig. 7). This indicative product positive feedback mechanism in the regulation of n-alkane degradation differs from those in Gram-negative bacteria. In Acinetobacter sp. M-1 and Pseudomonas aeruginosa, n-alkanes instead of fatty acids induced alkB gene expression (Ratajczak et al., 1998; Marin et al., 2003), whereas in P. putida GPo1 and Burkholderia cepacia RR10, the inducers were alkanes and alkanols (Grund et al., 1975; Marin et al., 2001). Product positive feedback is important but less reported. The limited examples included the positive feedback controlled by extracellular end-products in bacteria, which is central to cell–cell communication and the regulation of antibiotic biosynthesis (Bassler, 1999; Haas et al., 2000). Another example is the positive allosteric feedback regulation by a direct product of enzyme RelA in E.coli (Shyp et al., 2012). This indicative product positive feedback regulation could be beneficial for Dietzia sp. DQ12-45-1b to degrade n-alkanes, especially the long-chain n-alkanes. AlkX, as a regulator, is a cytoplasmic protein and interacts with the DNA in the cytoplasm. It is therefore convenient and energy-saving to interact with ligands in the cytoplasm. Generally speaking, alkanes with more than eight carbon atoms have poor water solubility and tend to accumulate in the cytoplasmic membrane (Rojo, 2010). However, the long-chain n-alkanes can be hydroxylated to n-alcohol by AlkB in the cytoplasmic membrane and, finally, oxidized to fatty acids in the cytoplasm (van Beilen et al., 2003). Compared with the long-chain n-alkanes, the LCFAs in the cytoplasm could interact much more easily with cytoplasmic AlkX. In addition, the basal level of AlkW1 in the membrane is enough to facilitate a rapid response for the hydroxylation of n-alkanes and downstream products could initiate AlkW1 and AlkX expression. In this way, negative autoregulation speeds the transcription response (Rosenfeld et al., 2002) and allows a fast switch-off when alkanes are consumed.

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Although direct evidence is lacking, the accumulation of downstream products is speculated to be a signal to repress alkB promoter expression when the β-oxidation pathway is overloaded (Marin et al., 2001). The expression level of alkW1 is low when cells are grown with glucose (Fig. 2) or sucrose as sole carbon source (Nie et al., 2011), suggesting the catabolites produced in tricarboxylic acid cycle might cause repression as reported in other alkane-degrading bacteria (Marin et al., 2001). In this scenario, the appropriate concentrations of AlkW1 and AlkX are maintained. Although product feedback regulation is energy saving and efficient, it is unknown why the Gram-negative n-alkane degrading bacteria do not utilize this mechanism. In Alcanivorax borkumensis, the AlkB expression regulator AlkS appeared associated to the membrane fraction, rather than to the cytoplasmic fraction in a proteomic study (Sabirova et al., 2006), which might be convenient to interact with n-alkane. In Alcanivorax dieselolei, there are three outer membrane transporter proteins that can selectively transport n-alkanes into the cytoplasm (Wang and Shao, 2014). Further study of n-alkane regulation is still needed.

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Model of AlkX regulating alkW1 expression might be universal among Actinobacteria

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Based on the results, a model for the regulation of alkW1 expression by AlkX was proposed (Fig. 8). In the absence of n-alkane, AlkX was slightly expressed and autoregulated the transcription of alkW1-alkX operon to maintain the basal optimal protein concentrations of AlkW1 and AlkX. When n-alkanes were present, they (such as n-hexadecane) were hydroxylated by the basal low concentration of AlkW1 and further oxidized to LCFAs (such as palmitic acid) in the cytoplasm. LCFAs, in turn, might bind to AlkX at its C-terminal ligand-binding domain and release it from the alkW1-alkX promoter by changing AlkX conformation. Finally, the transcriptional expression of alkW1-alkX operon was initiated. Similar gene arrangements of alkB-tetR gene clusters and partially similar inverted repeats upstream of alkB in some Actinobacteria such as Rhodococcus and Mycobacterium (Figs S12 and S13) suggested that TetR repression of AlkB expression might be a common regulation mechanism in Actinobacteria. Indeed, Actinobacteria are frequently reported to be capable of utilizing long-chain n-alkane, which is barely soluble in water and therefore extremely difficult to enter the cells. In this case, LCFAs are effective inducers of a quick gene response initiation. The high similarity of alkB-tetR gene clusters in different microorganisms might indicate their similar ancestor origins, although the exact mechanism is unknown (Nie et al., 2014). ‘Everything is everywhere, but, the environment

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Colour


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Fig. 8. Model of alkW1 gene regulation via a fatty acid positive feedback mechanism. In the absence of n-alkane, AlkX was slightly expressed and autoregulated the transcription of alkW1-alkX operon to maintain the basal optimal protein concentrations of AlkW1 and AlkX. When n-alkanes were present, they (such as n-hexadecane) were hydroxylated by the basal low concentration of AlkW1, and further oxidized to LCFAs (such as palmitic acid) in the cytoplasm. Palmitic acid, in turn, bound to AlkX at its C-terminal ligand-binding domain and released it from the alkW1-alkX promoter by changing AlkX conformation. Finally, the transcriptional expression of alkW1-alkX operon was initiated.

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selects’ (Baas-Becking, 1934). These similarities are likely resulted from gene evolution in response to long-time adaptation to long-chain-alkane-rich terrestrial environments (van Beilen et al., 2003) where higher abundance of Actinobacteria is detected (Venter et al., 2004; Giovannoni and Stingl, 2005). Therefore, TetR repression of AlkB expression and the product positive feedback mechanism might be a potentially universal mechanism in Actinobacteria. However, as the inverted repeats in the relative promoter regions of the alkB-tetR operons are shorter than that of alkW1-alkX, the corresponding TFRs may not use the cooperative binding mechanism as AlkX does. In conclusion, this study thoroughly characterized the mechanism of the transcriptional regulation of alkW1alkX operon in Dietzia sp. DQ12-45-1b. The TFR AlkX repressed alkW1 expression by cooperatively binding to the inverted repeat in alkW1 promoter. The LCFAs, downstream products of n-alkane oxidation, might be the effectors that modulated the DNA-binding affinity of AlkX and de-repressed alkW1 expression. This mechanism might be common in Dietzia strains and even in Actinobacteria. Alkane hydroxylase gene regulation is not only central to alkane degradation in both natural and industrial processes but also potentially critical in the

pathogenesis and clinical cures of infections caused by pathogens.

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Experimental procedures

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Growth media and culture conditions

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The bacterial strains and plasmids used in this study are listed in Table S1. E. coli DH5α, BL21 (DE3) and their recombinants were grown in Lysogeny Broth (LB) medium at 37°C on a shaker incubator at 150 r.p.m. P. fluorescens CHA0 and its alkB deletion mutant KOB2Δ1 as well as their recombinants were grown in LB medium at 30°C. Dietzia sp. DQ1245-1b and its alkX mutant were grown in GPY medium (10 g l−1 glucose, 10 g l−1 tryptone, 5 g l−1 yeast extract) at 30°C with shaking at 150 r.p.m. For RNA extraction and 5′ RACE, Dietzia sp. DQ12-45-1b was grown in minimal medium (Bihari et al., 2011) containing 1 ml l−1 n-hexadecane as sole carbon source. For western blot analysis, Dietzia sp. DQ12-45-1b and its alkX mutant were grown in minimal medium with 10 g l−1 glucose or 1 ml l−1 n-hexadecane. In the β-galactosidase assay, the Dietzia sp. DQ12-45-1b recombinant was grown in minimal medium supplemented with 10 g l−1 glucose, 1 ml l−1 liquid n-alkanes, 1 g l−1 solid n-alkanes, 1 g l−1 hexadecanol or 1 g l−1 palmitic acid as sole carbon source. The recombinants of P. fluorescens CHA0 and its alkB deletion mutant KOB2Δ1 were grown in E2 © 2015 John Wiley & Sons Ltd, Molecular Microbiology

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AlkX regulates alkane hydroxylase gene expression 15 1 2 3 4 5 6 7

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medium supplied with 1 ml l−1 MT microelements (Lageveen et al., 1988) and 5 ml l−1 liquid alkanes (C12–C16). Recombinants of E. coli, P. fluorescens or Dietzia sp. DQ12-45-1b strains were grown with appropriate antibiotics: ampicillin, 100 μg ml−1; streptomycin, 30 μg ml−1; kanamycin, 50 μg ml−1; gentamicin, 10 μg ml−1 for the E. coli recombinants and 100 μg ml−1 for the P. fluorescens recombinants respectively).

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Nucleic acid extraction and manipulation

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Plasmid and chromosomal DNA extraction, purification, PCR, enzymatic digestion, DNA ligation and transformation into E. coli were performed using standard molecular techniques (Sambrook and Russell, 2001). Total RNA was extracted using TRIzol reagent (Invitrogen, Waltham, MA, USA) according to the manufacturer’s instructions, treated with DNase I, and purified using TRIzol reagent. The primer sequences used for PCR are listed in Table S2. All PCRs without specific definition were performed using PrimeSTAR DNA Polymerase (TaKaRa, Beijing, China) according to the user manual: 3 min at 94°C; 30 cycles of 10 s at 98°C, 5 s at 60°C and 1–5 min at 72°C (depending on sequence lengths of PCR products); and finally maintaining the PCR products at 10°C.

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Reverse transcription of alkB-tetR mRNA and PCR amplification

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Total RNA was extracted from Dietzia sp. DQ12-45-1b cells harvested in the logarithmic phase in minimal medium supplemented with n-hexadecane, as previously described (Nie et al., 2011). Reverse transcription was performed using 0.5 μg of total RNA with primer alkXRT (Table S2) and a ReverTra Ace reverse transcription kit (TOYOBO, Shanghai, China). Control reactions to assess the level of DNA contamination in the RNA samples were carried out by omitting only the ReverTra Ace reverse transcriptase. Primers alkwrtF and alkwrtR (Table S2) were used for the PCR amplification. PCR using rTaq polymerase (TaKaRa) was performed as follows: 3 min at 94°C; followed by 30 cycles of 30 s at 94°C, 30 s at 55°C and 1 min at 72°C; 10 min at 72°C; and finally maintaining the PCR products at 10°C.

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Construction and complementation of the alkX disruption mutant

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The alkX mutant was constructed using a mono-homologous recombination method described elsewhere (Fig. 1A) (Lu et al. unpublished). Briefly, a selective cassette (EGFP-Kan) was amplified from vector pXL1819 (Lu et al. unpublished) using the primers P1 and P2 (Table S2), which had a 24 bp overlapping end with the alkX 400 bp fragment (see below) 5′ terminus and an ‘A’ in primer P2 for frameshift mutation. A 400 bp fragment of alkX was amplified from the chromosomal DNA of Dietzia sp. DQ12-45-1b using the primers P3 and P4 (Table S2). The selective cassette fragment and alkX 400 bp fragment were then fused into a single molecule with all sequences in the desired order using primers P1 and P4. The generated linear DNA fragment was introduced © 2015 John Wiley & Sons Ltd, Molecular Microbiology

into Dietzia sp. DQ12-45-1b cells via electroporation (2,000 V mm−1, 12 ms) as described previously (Lu et al., 2014), and the kanamycin-resistance clones were identified by amplifying and sequencing the alkW1 and alkX fragments using the primers alkw1F and alkXR (Table S2). The correct chromosomal integration of the fused DNA fragments resulted in a kanamycin-resistance disruption mutant designated DQ4502. The expression vectors for complementation studies were constructed as follows. The alkX fragment was amplified from chromosomal DNA from Dietzia sp. DQ1245-1b with the primers alkXF and alkXR (Table S2). The obtained 687 bp DNA fragment was ligated into vector pXL1804 (Table S1; for details, see below), which contained the promoter for both alkW1 and alkX, using BamHI and PstI restriction sites to construct a new plasmid, pXL1817 (Table S1). Vector pXL1817 was introduced into alkX mutant via electroporation as described elsewhere (Lu et al., 2014). Control transformation with Dietzia-E. coli shuttle vector pNV18Sm (Szvetnik et al., 2010) was also included.

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To explore the physiological function of AlkX on AlkW1 expression in Dietzia sp. DQ12-45-1b, we detected AlkW1 protein in the strain DQ12-45-1b and its alkX mutant with western blot. Cells grown in minimal medium amended with glucose or n-hexadecane were harvested in the logarithmic phase and broken with glass beading. The lysate was centrifuged at 9,279 × g (15 min, 4°C), and the supernatants were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis on 8% polyacrylamide gel, then transferred to a polyvinylidene fluoride membrane at 100 mA for approximately 30 min. Immunoblotting was performed using antiAlkW1 antibody (the mouse antisera against AlkW1 was provided by Guang Hu) with 1:1,000-fold dilution, horseradish peroxidase-conjugated secondary antibody with 1:1,000-fold dilution and 0.05% 3, 3-diaminobenzidine tetrahydrochloride reagent (Amresco, Solon, OH, USA).

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The 5′ terminal sequence of alkW1 was cloned using a 5′-Full RACE kit (TaKaRa) following the manufacturer’s instructions. A sample of the total RNA of Dietzia sp. DQ12-45-1b (approximately 2 μg) was dephosphorylated using alkaline phosphatase and pyrophosphorylated using tobacco acid pyrophosphatase followed by ligation of the 5′ RACE adaptor and reverse transcription using specific transcriptional primer alkW1rt (Table S2). The transcribed product (5 μl of the reaction mixture) was then used as a template for the first round of PCR with the GSP1 primer and the 5′ RACE outer primer supplied with the kit. Subsequently, the product (1 μl of the reaction mixture) was used as a template for the second round of PCR with the GSP2 primer and the 5′ RACE inner primer supplied with the kit. PCR reaction conditions for these two rounds of PCR using LA Taq (TaKaRa) were set at 3 min at 94°C; 20 or 30 cycles of 30 s at 94°C, 30 s at 60°C and 90 s at 72°C; 10 min at 72°C; and maintenance at 10°C. The target DNA fragment was then cloned into the pGEM-T easy vector (Promega, Fitchburg, WI, USA) and sequenced.

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Deletion analysis

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A large fragment of vector pNV18Sm was first amplified with primer pNV18EF and pNV18ER (Table S2). The resulting 5,424 bp linear fragment lacking the lacZ promoter was then digested with EcoRI and self-ligated, resulting in promoterless Dietzia-E. coli shuttle plasmid pXL1801. The DNA fragment of EGFP-Kan was obtained from vector pXL1819 (Table S1) via enzyme digestion by BamHI and HindIII and was then ligated into vector pXL1801, resulting in plasmid pXL1803. PCR was used to generate fragments with deletions of different lengths in the 255 bp fragment upstream of alkW1. Both the 5′ and the 3′ (Fig. 2B) deletion sets were cloned into pXL1803 with EcoRI and BamHI restriction sites and transformed into E. coli DH5α (Table S1). The generated DH5α recombinants were cultured in LB medium for 5 h, and then 1 ml of cells was collected via centrifugation at 2,320 × g (1 min) at room temperature (RT) and suspended in phosphate-buffered saline containing 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 2 mM KH2PO4. The optical density of these cells was measured at 600 nm and diluted to 1.0 using phosphate-buffered saline. The relative fluorescence unit (RFU) values were measured on a SpectraMax M5 microplate reader (excitation/emission: 485 nm/538 nm; Molecular Devices, Sunnyvale, CA, USA). Each experiment was performed at least three times.

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We amplified lacZ from the E. coli MG1655 genome with PCR using primers LacZF and LacZRP containing the PstI restriction site (Table S2). The DNA fragment that included the 255 bp alkW1 upstream region and the 117 bp alkW1 coding region from ATG was amplified with PCR using primers alkwFB and alkX-117 containing the BamHI restriction site. The two DNA fragments were fused using PCR and cloned into pXL1801. The resulting vector was designated pXL1802 and was introduced into Dietzia sp. DQ12-45-1b via electroporation as described previously (Lu et al., 2014). The Dietzia sp. DQ12-45-1b recombinant containing pXL1802 was grown in 50 ml GPY medium with streptomycin. After phosphate buffer washing, the cells were suspended in 50 ml minimal medium amended with various carbon sources and streptomycin and incubated at 30°C on a rotary shaker at 150 r.p.m. After 48 or 72 h of incubation, 50 ml aliquots were collected via centrifugation at 2,320 × g (5 min, 4°C) and suspended in 100 μl breaking buffer containing 100 mM TrisHCl (pH 7.5) and 20% glycerol. Buffer Z (400 μl) containing 40 mM Na2HPO4·7H2O, 60 mM NaH2PO4 ·H2O, 10 mM KCl, 1 mM MgSO4·7H2O and 50 mM 2-mercaptoethanol (pH 7.0) was added to each aliquot, which was then bead-beaten for 30 s three times at 2,500 r.p.m. The supernatant was collected after centrifugation at 9,279 × g (10 min, 4°C), and β-galactosidase assays were then performed using a method described previously (Rose and Botstein, 1983). The reactions were initiated by the addition of 90 μl ortho2-nitrophenyl-β-D-galactopyranoside solution (4 mg ml−1 in buffer Z) to 10 μl supernatant and stopped by the addition 150 μl of 1 M Na2CO3. A blank sample without biomass was prepared each time when buffer Z and ortho-2-nitrophenyl-βD-galactopyranoside were added to a final volume of 250 μl.

Proteins in the supernatant were later quantified using the Bradford Protein Assay Kit (QIANGEN, Beijing, China) with bovine serum albumin (BSA) as a standard. Each experiment was performed at least three times. The unit of β-galactosidase activity was defined as 1 nmol of o-nitrophenol released per mg of protein per min (nmol mg of protein−1 min−1).

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Preparation and purification of recombinant AlkX protein

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The alkX gene was cloned using primers alkXFH and alkXRE (Table S2) and ligated into plasmid pET-28a with HindIII and EcoRI restriction sites, thus yielding plasmid pXL2801. The overexpression of AlkX was performed in E. coli BL21 (DE3) with pXL2801, and the recombinant protein was purified with His tag. In brief, cells transformed with pXL2801 were grown in LB medium with kanamycin at 37°C on a rotary shaker at 150 r.p.m. One milliliter of the overnight culture was used to inoculate 100 ml fresh medium. After the bacteria grew to an optical density of 0.4–0.6 at 600 nm, target protein expression was induced by the addition of isopropyl-β-Dthiogalactoside to a final concentration of 1 mM. After induction for 4 h at 37°C, the cells were harvested via centrifugation at 2,320 × g (5 min, 4°C). After the pellet was resuspended in 10 ml lysis buffer (50 mM NaH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0), the cells were lysed via sonication followed by centrifugation at 20,400 × g (60 min, 4°C). The supernatant was incubated in Ni Sepharose™ High Performance Media (GE Healthcare, Little Chalfont, UK) for 20 min. After three washes with washing buffer (50 mM NaH2PO4, 300 mM NaCl, 40 mM imidazole, pH8.0), AlkX was eluted from the medium by applying 100 μl elution buffer four times (50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, pH8.0). TGN buffer (50 mM Tris-HCl, 50 mM NaCl, 5% glycerol, pH 8.0) was added to purified AlkX to maintain protein activity. Soluble AlkX was assessed with sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The concentration of purified protein was determined using the Bradford method as described for promoter activity analysis.

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The DNase I footprinting experiment was performed as described previously (Zianni et al., 2006). Briefly, 255 bp DNA fragments upstream of alkW1 were generated with PCR using plasmid pXL1802 as the template and primers 255F and 255R labeled with 6-carboxy-2′,4,4′,5′,7,7′-hexachlorofluorescein (HEX). The 3′ HEX-labeled PCR fragments were purified with a DNA purification kit (BioTeke Corporation, Beijing, China) and quantified using the UV1700 UV-VIS Spectrophotometer (SHIMADZU, Kyoto, Japan). The labeled DNA fragment (170 ng) was diluted in 50 μl binding buffer consisting of 10 mM Tris-HCl (pH 7.6), 30 mM KCl, 1 mM ethylenediaminetetraacetic acid (EDTA), 10 mM (NH4)2SO4, 1 mM dithiothreitol and 0.2% (w/v) Tween-20 and then incubated with 30 μg AlkX or 30 μg BSA as the control. After a 20 min incubation at RT, 0.07 units of DNase I (TaKaRa, Beijing, China) was added to the reaction mixture and incubated for an additional 12 min at RT. The reaction was stopped by heating at 80°C for 10 min. The DNA fragments were purified with phenol-chloroform and © 2015 John Wiley & Sons Ltd, Molecular Microbiology

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AlkX regulates alkane hydroxylase gene expression 17 1 2 3 4 5 6 7

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precipitated with ethanol. The digested DNA was analyzed on a 3130 DNA Analyzer (Applied Biosystems, Waltham, MA, USA) with Genescan-500 LIZ (Applied Biosystems) as the size standard. The results were then analyzed with Genemapper 4.0 software (Applied Biosystems) to determine the protected sequences on the DNase I digestion map (Yu et al., 2010a).

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For EMSA assays, all of the oligonucleotides of both strands (Table S2) were synthesized directly (Sangon Biotech Corporation, Shanghai, China) and were labeled with digoxigenin at the 5′ terminus of the top strand. All the test sequences had 70 bp oligonucleotides, containing a 48 bp wild-type operator/ mutated operator sequence and extra 10 and 12 bps in the two flanking regions. For example, the 70 bp wild-type sequence contained the 48 bp operator sequence, and the mutated sequences were still 70 bp, with the 48 bp sequence containing mutated OP1-OP4 respectively (Fig. 5B). Annealing was carried out by mixing equimolar concentrations (10 μM) of each complementary oligonucleotide in annealing buffer containing 10 mM Tris-HCl (pH 8.0), 50 mM CH3COOK and 1 mM EDTA. The mixture was incubated at 95°C for 2 min and then cooled slowly to RT. Then, 0.1 μM labeled DNA fragment and 0.1–5 μM AlkX were mixed and incubated in binding buffer as described for the DNase I footprinting assay. To determine the possible binding substrate of AlkX, we added 0.2 millimole of the substrates (tetradecane, hexadecane, tetradecanol, hexadecanol, myristic acid and palmitic acid) into 10 ml of 1% Tween-80 solution respectively and kept them at RT for 72 h for emulsification. After excluding the undissolved parts in the emulsified mixture by centrifugation, 1 μl of the dissolved solution was co-incubated with AlkXDNA complex. To investigate the impacts of LCFAs, we performed EMSA by using seven fatty acids, i.e., those with carbon atoms being C10, C12, C14, C16, C18, C20 and C24. Briefly, we added 1 millimole of these LCFAs into 10 ml of 1% Tween-80 solution respectively and kept them at RT for 72 h. The dissolved LCFAs were then determined by the fatty acid methyl esters (FAMEs) method (Liu et al., 2010) as mentioned below. To test the effects of other different small molecule acid compounds, we used 1 mM acetic acid, propanoic acid, lactic acid and pyruvic acid dissolved in the deionized water. These stock solutions were diluted to working solutions with the concentrations as follows: palmitic acids, 0.75– 150 μM; other LCFAs, 30 μM; acetic acid, propanoic acid, lactic acid and pyruvic acid, 100 μM. Then 1 μl of the working solution was added to the AlkX-DNA mixture (1 or 2 μM AlkX and 0.1 μM DNA) respectively. After incubation at RT for 30 min, the mixture samples were separated on nondenaturing 5% polyacrylamide gels in 0.05 M Tris borateEDTA buffer (pH 8.3) for 2 h at 10 V cm−1 and transferred to positively charged nylon membranes for 30 min at 300 mA. Labeled DNA was visualized using a digoxigenin detection kit (Roche, Mannheim, Germany).

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sampled, freeze-dried and incubated with a mixture of 1 ml 100% toluene and 2 ml 1% sulphuric acid in methanol (v/v) overnight at 50°C to form FAMEs. The resulting FAMEs were then extracted twice with hexane (1:1, v/v) and analysis with GC-MS using an Agilent 7890C GC and a 5975C MS (Agilent, Santa Clara, CA, USA) equipped with an Agilent HP-5AS capillary column (30 m by 0.25 mm). Helium was the carrier gas at a flow rate of 3.0 ml min−1. One microliter of sample was injected via a split injection port at a split ratio of 1:19. The injection port and detector temperatures were constant at 250°C and 280°C respectively. The column temperature was kept initially at 150°C, then increased at 10°C min−1 to 200°C, and then the temperature was increased to 250°C at 15°C min−1 and held for 3 min. FAMEs were identified by chromatographic comparison with authentic standards (National Institute of Standards and Technology). The quantities of LCFAs were estimated from the peak areas on the chromatogram using heptadecanoic acid as the internal standard. The experiment was performed at least three times. The fatty acids were also analyzed with the lysate of cells grown with n-hexadecane as sole carbon source. Briefly, Dietzia sp. DQ12-45-1b cells grown in minimal medium amended with n-hexadecane as sole carbon source were harvested in the stationary phase. The cell pellet treatments for obtaining the organic compounds in the cell lysates are described elsewhere (Kim et al., 2002). Lysate components were quantified with GC-MS in the similar method described below. The injection port and detector temperatures were constant at 250°C and 300°C respectively. The column temperature was kept initially at 100°C for 1 min, then increased at 20°C min−1 to 260°C and held for 5 min. The temperature was then increased to 280°C at 20°C min−1 and held for 5 min. The mass of components accumulated in cells was calculated using calibration curves and normalized to the internal standard.

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The AUC analysis of AlkX (4–35 μM), AlkX-wild-type DNA mixtures (1–4 μM protein and 0.4–1 μM DNA) and AlkXmutated DNA mixtures (4 μM protein and 1 μM mutated DNA) was performed according to standard protocol (Lebowitz et al., 2002). The protein/wild-type DNA ratios were chosen as 1:1 (1 μM AlkX and 1 μM DNA), 2:1 (2 μM AlkX and 1 μM DNA), 4:1 (4 μM AlkX and 1 μM DNA), 8:1 (4 μM AlkX and 0.5 μM DNA) and 10:1 (4 μM AlkX and 0.4 μM DNA). The 52 bp DNA fragment contained 48 bp operator with or without mutations and extra 2 bps in each flanking of the operator (Table S2). All samples were prepared in 20 mM Tris-HCl (pH 8.0) and 300 mM NaCl. The sedimentation velocity experiments were carried out at 40,000 r.p.m. and 20°C in an XL-A analytical ultracentrifuge (Beckman Coulter, Fullerton, CA, USA) with a UV-VIS optics detection system, using an An60Ti rotor and 12 mm double-sector centerpieces. Sedimentation profiles were recorded every 5 min at 280 nm. The sedimentation coefficient distributions were calculated by leastsquares boundary modeling of sedimentation velocity data using the c(s) method in the SEDFIT program (Schuck, 2000) from http://www.analyticalultracentrifugation.com/. The 3 s-values were corrected to standard conditions (water, 20°C, and infinite dilution) by SEDNTERP program to obtain

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After the above LCFAs were emulsified in 1% Tween-80 at RT for 72 h respectively, the dissolved solution (1 ml) was © 2015 John Wiley & Sons Ltd, Molecular Microbiology

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18 J.-L. Liang et al. ■ 1 2 3 4

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the corresponding standard s-values (s20, w) (Schuck, 2003; Brown and Schuck, 2006). Each experiment was performed at least three times. The s-values were from the fit of a single experiment.

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Cotransformation experiments and RFU detection

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Vectors pXL1815 (with OP1–OP4 included) or pXL1816 (with OP1 mutated; Table S1) were constructed with the method described in deletion analysis. The DNA fragments were amplified with primers 107FE or MFE and alkwBR and then ligated into plasmid pXL1803 with EcoRI and BamHI restriction sites. The alkX fragment was cloned with primers alkXFH and alkXRE and ligated into plasmid pUC19 using HindIII and EcoRI restriction sites, resulting in plasmid pXL1901 (Table S1). Plasmids pXL1901 and pXL1815 or pXL1816 were cotransformed into E. coli DH5α. Plasmid pUC19 was also cotransformed with pXL1815 or pXL1816 as the control. The transformed cells were incubated overnight in LB medium with streptomycin and ampicillin at 37°C on a rotary shaker at 150 r.p.m. The cells were then collected and the RFU values measured as mentioned above. Each experiment was performed at least three times.

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then ligated into pXL8001. The ligated product was transformed into E.coli DH5α and the successful cloning was verified by sequencing. The plasmid was designated as pXL8002 (Table S1). Next, we amplified the alkW1 promoter with primers alkwEF and alkwurR, and the alkX gene with primers alkXF and alkXRB. Then we fused the two fragments together with primers alkwEF and alkXRB (Table S2). The fused PCR product was digested with restriction enzymes EcoRI and BglII and then ligated into the pXL8002. The ligated product was transformed into E.coli DH5α and the successful cloning was verified by sequencing. The plasmid was designated as pXL8003 (Table S1). The plasmid pXL8003 was then transformed into the P. fluorescens CHA0 and KOB2Δ1 cells respectively by electroporation (Højberg et al., 1999). All of the recombinants were grown with 100 μg ml−1 gentamicin. P. fluorescens CHA0 and KOB2Δ1 recombinants were first grown in LB medium. The cells from the LB medium were washed three times with E2 medium and inoculated into 50 ml of E2 medium (with the final concentration expressed as an optical density at 600 nm of 0.5) with different n-alkanes as sole carbon sources. The cells were then cultured at 30°C and shaken at 150 r.p.m. for 24 h. The cells were then harvested and measurements of the β-galactosidase activity were performed as described above. All the experiments described above were repeated in triplicate.

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ITC experiments were performed at 30°C using the iTC200 (Microcal, Northampton, MA, USA). Protein and DNA fragments were prepared in the same buffer containing 20 mM Tris-HCl (pH 8.0), 300 mM NaCl and 5% glycerol. Forward titrations (AlkX into DNA) consisted of the injection of 87.7 μM AlkX dimer into 6.0–8.0 μM DNA. All data were corrected using the heat changes arising from injection of the ligand in the syringe into the buffer. Data were analyzed with a model for the cooperative binding of two ligands to a macromolecule (Krell et al., 2007), and also the ‘one set of sites’ and ‘two set of sites’ models implemented in the MicroCal version of ORIGIN (Wiseman et al., 1989). The fitting curve was generated with MATLAB R2014a software (The MathWorks, Natick, MA, USA). Each experiment was performed at least three times. The errors corresponded to that of curve fitting of a single experiment.

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Sequence analysis of alkB-tetR gene clusters inadditional 22 Dietzia strains

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The Dietzia strains used in this study are listed in Table S3. These strains were purchased from other laboratories or isolated in our own laboratory from oil field samples. Primers F and R (Table S2) were used to amplify the alkB-tetR gene clusters via PCR with a DNA fragment length of approximately 3,000 bp. The PCR products were purified and ligated into the pGEM-T easy vector (Promega, Fitchburg, WI, USA), transformed into E. coli DH5α and sequenced. These sequences were submitted to GenBank under accession numbers KM019144-KM019165 (http://www.ncbi.nlm.nih.gov/).

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To further test the hypothesis that LCFAs worked as inducers in vivo, we constructed the lacZ-fusion reporter vector with alkW1 promoter and AlkX. Briefly, we first amplified the E.coliPseudomonas shuttle vector pCom8 (Nie et al., 2011) with primers p8EF and p8ER (Table S2). Then we digested the PCR product with restriction enzyme EcoRI and self-ligated the DNA fragment. The ligated product was transformed into E.coli DH5α and the successful cloning was verified by sequencing. The plasmid was designated as pXL8001 (Table S1). Later, we amplified the lacZ from the E. coli MG1655 genome with primers LacZFB and LacZRP (Table S2) containing the BamHI and PstI restriction sites. The PCR product was digested with BamHI and PstI restriction enzymes and

Primer design, sequence analysis and assembly were performed with Vector NTI Advanced 11 (Invitrogen). The nucleotide and amino acid sequences of the AlkB and TetR family proteins in this research were compared with those in the GenBank database using the blastn and blastp programs (http://blast.ncbi.nlm.nih.gov/). Multiple sequence alignments were generated with ClustalW2 (Larkin et al., 2007). The phylogenetic trees for the AlkB and TetR family proteins were determined using the neighbor-joining algorithm and the Jukes-Cantor correction factor and generated with MEGA 5.0 (Tamura et al., 2011).

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Calculations and statistical analyses were performed using Origin 8.0 software (OriginLab, Northampton, MA, USA). The © 2015 John Wiley & Sons Ltd, Molecular Microbiology

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AlkX regulates alkane hydroxylase gene expression 19 1 2 3 4

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values are presented as means ± standard deviations. Data were analyzed statistically using a one-tailed Student’s t-test, and P values of 0.05 or less were considered statistically significant.

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Acknowledgements

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We thank Dong-Ling Ma for gas chromatograph-mass spectrometry performance and relative data analysis. We thank Xiao-Xia Yu (Institute of Biophysics, Chinese Academy of Sciences) for technical assistance in analytical ultracentrifugation experiments. We thank Theo Smits for kindly providing P. fluorescens CHA0 and KOB2Δ1 and the pCom8 plasmid. This study was supported by National Natural Science Foundation of China (31225001 and 31300108) and National High Technology Research and Development Program of China (2012AA02A703). All authors declare no conflict of interest.

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Alonso, A., Rojo, F., and Martínez, J.L. (1999) Environmental and clinical isolates of Pseudomonas aeruginosa show pathogenic and biodegradative properties irrespective of their origin. Environ Microbiol 1: 421–430. Baas-Becking, L.G.M. (1934) Geobiologie of inleiding tot de Milieukunde. The Hague, the Netherlands: W.P. Van Stockum & Zoon. Bassler, B.L. (1999) How bacteria talk to each other: regulation of gene expression by quorum sensing. Curr Opin Microbiol 2: 582–587. van Beilen, J.B., Wubbolts, M.G., and Witholt, B. (1994) Genetics of alkane oxidation by Pseudomonas oleovorans. Biodegradation 5: 161–174. van Beilen, J.B., Panke, S., Lucchini, S., Franchini, A.G., Röthlisberger, M., and Witholt, B. (2001) Analysis of Pseudomonas putida alkane-degradation gene clusters and flanking insertion sequences: evolution and regulation of the alk genes. Microbiology 147: 1621–1630. van Beilen, J.B., Smits, T.H., Whyte, L.G., Schorcht, S., Röthlisberger, M., Plaggemeier, T., et al. (2002) Alkane hydroxylase homologues in Gram-positive strains. Environ Microbiol 4: 676–682. van Beilen, J.B., Li, Z., Duetz, W.A., Smits, T.H.M., and Witholt, B. (2003) Diversity of alkane hydroxylase systems in the environment. Oil Gas Sci Technol 58: 427–440. van Beilen, J.B., Marín, M.M., Smits, T.H., Röthlisberger, M., Franchini, A.G., Witholt, B., and Rojo, F. (2004) Characterization of two alkane hydroxylase genes from the marine hydrocarbonoclastic bacterium Alcanivorax borkumensis. Environ Microbiol 6: 264–273. Bertrand, J.C., Almallah, M., Acquaviva, M., and Mille, G. (1990) Biodegradation of hydrocarbons by an extremely halophilic archaebacterium. Lett Appl Microbiol 11: 260– 263. Bhukya, H., Bhujbalrao, R., Bitra, A., and Anand, R. (2014) Structural and functional basis of transcriptional regulation by TetR family protein CprB from S. coelicolor A3(2). Nucleic Acids Res 42: 10122–10133. Bihari, Z., Szvetnik, A., Szabó, Z., Blastyák, A., Zombori, Z., Balázs, M., and Kiss, I. (2011) Functional analysis of long-

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AUTHOR QUERY FORM Dear Author, During the preparation of your manuscript for publication, the questions listed below have arisen. Please attend to these matters and return this form with your proof. Many thanks for your assistance.

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