GacS-Dependent Regulation of Polyhydroxyalkanoate Synthesis in Pseudomonas putida CA-3

William J Ryan; Niall D O'Leary; Mark O'Mahony; Alan D W Dobson

doi:10.1128/AEM.02962-12

. 2013 Mar;79(6):1795–1802. doi: 10.1128/AEM.02962-12

GacS-Dependent Regulation of Polyhydroxyalkanoate Synthesis in Pseudomonas putida CA-3

William J Ryan ^a,^b, Niall D O'Leary ^a,^b,^✉, Mark O'Mahony ^a, Alan D W Dobson ^a,^b

PMCID: PMC3592236 PMID: 23291549

Abstract

To date, limited reports are available on the regulatory systems exerting control over bacterial synthesis of the biodegradable polyester group known as polyhydroxyalkanoates (PHAs). In this study, we performed random mini-Tn5 mutagenesis of the Pseudomonas putida CA-3 genome and screened transconjugants on nitrogen-limited medium for reduced PHA accumulation phenotypes. Disruption of a GacS sensor kinase in one such mutant was found to eliminate medium-chain-length PHA production in Pseudomonas putida CA-3. Recombinant expression of wild-type gacS from a pBBRgacS vector fully restored PHA accumulation capacity in the mutant strain. PCR-based screening of the P. putida CA-3 genome identified gene homologues of the GacS/GacA-rsm small RNA (sRNA) regulatory cascade with 96% similarity to published P. putida genomes. However, reverse transcription-PCR (RT-PCR) analyses revealed active transcription of the rsmY and rsmZ sRNAs in gacS-disrupted P. putida CA-3, which is atypical of the commonly reported Gac/Rsm regulatory cascade. Quantitative real-time RT-PCR analyses of the phaC1 synthase responsible for polymer formation in P. putida CA-3 indicated no statistically significant difference in transcript levels between the wild-type and gacS-disrupted strains. Subsequently, SDS-PAGE protein analyses of these strains identified posttranscriptional control of phaC1 synthase as a key aspect in the regulation of PHA synthesis by P. putida CA-3.

INTRODUCTION

Polyhydroxyalkanoates (PHAs) are biodegradable polyesters produced by numerous bacterial species typically synthesized when an essential nutrient such as nitrogen, sulfur, or phosphorus becomes limiting in the presence of an excess of carbon (1–4). Several comprehensive recent reviews are available in the field of microbial PHA synthesis (5–7). In summary, PHAs are condensation polyesters of various hydroxyalkanoic acids, with 2 major classes identified based on the respective chain lengths of the monomer units that make up the polymers. Short-chain-length PHAs (scl-PHAs) consist of 3- to 5-carbon-atom monomers, and medium-chain-length PHAs (mcl-PHAs) consist of 6- to 14-carbon-atom monomers (8–10). Generation of the various scl and mcl monomers can occur by several routes. In the case of scl-PHAs, 3-hydroxybutyrate monomers are derived via condensation of acetyl coenzyme A (acetyl-CoA) moieties from the tricarboxylic acid (TCA) cycle (e.g., Ralstonia eutropha) (11). Alternatively, fatty acid metabolism can generate various hydroxyalkanoate monomers for PHA biosynthesis via substrate-related conversions of keto-acyl-CoAs to corresponding 3-hydroxyacyl-CoAs during β-oxidation (e.g., Pseudomonas aeruginosa) (12). PHA monomers can also be derived from unrelated carbon sources via acetyl-CoA diversion to de novo fatty acid synthesis, yielding various 3-hydroxyacyl–acyl carrier protein (ACP) intermediates. Conversion to the required 3-hydroxyacyl-CoA monomers is achieved by 3-hydroxy-ACP-CoA transacylases (e.g., Pseudomonas putida) (13). The latter pathway is of specific biotechnological interest as it presents the potential for PHA production from organic wastes (5).

Metabolic networks and carbon flux play a central role in PHA production, and a number of core cellular metabolic enzymes have been manipulated in attempts to improve PHA yields in various bacterial species (7, 14, 15). Signal transduction systems associated with the detection of extracellular conditions and the coordination of central metabolic pathways involved have also been implicated in PHA production control. The arc system in Escherichia coli, which regulates the expression of several operons relative to the redox state of the environment, has previously been manipulated to improve E. coli as a host strain for poly-β-hydroxybutyrate (PHB) biosynthesis (16). In addition, arc gene mutants enable the expression of TCA cycle components under microaerobic conditions, elevating levels of reducing equivalents, which can be channeled into PHB production. The CreBC two-component system controls several genes involved in carbon flux in E. coli (17). Nikel et al. (18) achieved increased PHB yields in an arcA creC double mutant of E. coli, owing to the induction of a reduced redox state coupled with increased carbon utilization, respectively.

However, despite an increased understanding of the role of metabolic intermediates, carbon flux, and redox potentials affecting PHA synthesis, relatively little remains known in relation to global cellular regulatory systems directly controlling PHA synthesis processes. Such understanding would likely be of considerable value in future recombinant strategies for the optimization of PHA production from pure cultures on diverse substrates. Our group has previously utilized a solid medium screen incorporating nitrogen-limited conditions in conjunction with random Tn5 mutagenesis to identify functional PHA biosynthetic genes in P. putida CA-3 (19). In a separate, more recent investigation of aromatic hydrocarbon degradation by P. putida CA-3, random Tn5 mutagenesis successfully revealed the importance of RpoN-dependent regulation of a permease essential for phenylacetic acid uptake (20). In the current study, we therefore sought to employ a similar approach to determine whether functional links could be established between global regulatory/signaling mechanisms and the PHA synthetic pathway within the well-characterized mcl-PHA producer P. putida CA-3. Random mini-Tn5 mutagenesis of the P. putida CA-3 genome was combined with a solid medium screen to identify deficient/altered PHA accumulation phenotypes. Approximately 13,500 mutants were generated by this process, of which approximately 44 demonstrated altered PHA accumulation profiles. Here, we present our findings on the characterization of one of these mutants, PHA45A, which hosted a gacS gene disruption and established a direct regulatory influence at the posttranscriptional level on PHA synthesis in P. putida CA-3.

MATERIALS AND METHODS

Bacterial strains, plasmids, culture media, and growth conditions.

Pseudomonas putida CA-3, a styrene-degrading bioreactor isolate, has been described previously (21). E. coli CC118λpir hosted the mini-Tn5 derivative pUT-Km1 (R6K origin of replication), which encodes resistance to kanamycin and ampicillin (22). Plasmid pRK600 (Cm^r) was used as a helper in triparental mating experiments and encodes the tra functions facilitating pUT-Km1 mobilization. pCR2.1-TOPO vector (Invitrogen, CA) was used in the cloning of PCR amplification products. The expression vector pBBR1MCS-5 was used for expression of cloned genes in complementation studies as described below (23). Pseudomonas putida CA-3 was routinely grown on E2 minimal medium (24) containing 23 mM citrate as the sole carbon source. PHA accumulation was achieved by reducing the nitrogen content of the E2 medium in the form of NH₄SO₄ from 8 mM to 1.5 mM and allowing cultures to grow for ∼8 h. All E. coli vector hosts were maintained on standard LB agar plates containing the appropriate antibiotic(s) and were inoculated into 10-ml LB overnight broths prior to desired applications.

Cultures were routinely tested for PHA accumulation by microscopic fluorescent visualization of Nile Red-stained samples exposed to UV excitation. One-milliliter samples of respective cultures actively growing under nitrogen-limiting (1.5 mM) conditions were centrifuged (16,400 × g), the supernatants were removed, and pellets were washed in phosphate-buffered saline (PBS) before being stained with 10 μl of 10 mg/ml Nile Red (excitation, 485 nm; emission, 525 nm) in dimethyl sulfoxide (DMSO). The stained pellets were incubated at room temperature for 5 min and protected from ambient light exposure before being made up to 1 ml with PBS. Cells were visualized under fluorescence (filter I3; excitation, 450 nm/490 nm) at an ×1,000 magnification for the presence of PHA granules by using a Leica DM3000 microscope fitted with a Leica DFC490 camera and analyzed with Leica Application suite v3.1.0.

Random mini-Tn5 mutagenesis.

A triparental mating approach was used to introduce pUT-Km1 into P. putida CA-3. The mating mixture (1 ml), containing mid-log-phase LB-grown recipient, donor, and helper strains at a ratio of 7:2:1, was centrifuged for 2 min at a relative centrifugal force (RCF) of 16,400, resuspended in 50 μl fresh LB medium, spotted onto an LB agar plate, and incubated at 28°C for 24 h. P. putida CA-3 transconjugants expressing kanamycin resistance were subsequently isolated by plating serial dilutions of the mating mixture onto E2-citrate medium containing kanamycin (50 μg/ml). Approximately 13,500 isolated colonies were individually transferred to 96-well microtiter plates containing 200 μl of E2-citrate broth in each well. These master plates were used to identify transposition events resulting in a qualitative loss/reduction of the parent strain's ability to accumulate PHAs from the unrelated carbon source citrate. PHA-negative phenotypes were identified by first growing colonies in E2-citrate broth containing 1.5 mM nitrogen (28°C for 24 h), followed by transfer to E2-citrate agar plates lacking any nitrogen, incubation for 48 h at 30°C, and monitoring thereafter for an absence of colony opacity associated with PHA-accumulating microorganisms (19).

Mapping of transposon insertion sites.

Arbitrarily primed PCR was employed to map the gene disruption sites by utilizing previously published oligonucleotide primers and appropriate thermal cycling parameters (25). Products were visualized on 1% agarose gels, purified using a Qiagen QIAquick gel extraction kit, and sequenced using the mini-Tn5 internal primer, TNINT (see Table S1 in the supplemental material).

Mutant complementation.

Genomic DNA was isolated from P. putida CA-3 according to the method of Ausubel et al. (26). The primer pair FgacS-FW and FgacS-RV (see Table S1 in the supplemental material) was designed based on available Pseudomonas putida gacS gene sequences in GenBank. PCR amplification of the full-length gacS gene (GenBank accession number JX826483) from genomic DNA involved 35 cycles of PCR amplification with Pfu proofreading polymerase followed by the addition of 2.5 U Taq polymerase and incubation at 72°C for 10 min to add a poly(A) tail to facilitate cloning into the pCR2.1 cloning vector (Invitrogen). The cloned fragments were then transformed into Top10 E. coli cells (Invitrogen) per the manufacturer's instructions. Transformants were plated on LB agar plates containing 50 μg/ml kanamycin which were overlaid with 40 μl of 40 mg/ml X-Gal (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside) to facilitate blue/white screening of colonies. Several of the white colonies generated were inoculated into liquid LB medium containing kanamycin and subjected to plasmid minipreps (Fermentas) per the manufacturer's instructions. The resultant plasmid was cut with XbaI and HindIII enzymes in a double digest at 37°C for 90 min with Tango buffer (Fermentas). The pBBR1MCS-5 expression vector was also digested in the same manner. The gacS fragment obtained from the double digest was ligated into the pBBR1MCS-5 vector using T4 DNA ligase (Fermentas) per the manufacturer's instructions to yield pBBRgacS. Transformation of pBBRgacS into chemically competent Top10 E. coli cells (Invitrogen) was achieved as previously described, and the resultant transformants were screened on LB agar plates containing 20 μg/ml gentamicin. Transconjugation of pBBRgacS expression vector into the P. putida CA-3 gacS::Tn5 PHA45A mutant was performed via triparental mating as before. Successful transconjugants were isolated on solid E2 minimal medium containing 50 μg/ml kanamycin and 20 μg/ml gentamicin. Successful mating reactions yielded the complemented gacS::Tn5 mutant named PHA45AgacS+. Complementation of the full-length phaC1 gene in the phaC1::Tn5 mutant B11 to yield B11phaC1+ was achieved in the same manner as detailed above for gacS complementation. PCR amplification of the full-length phaC1 gene from P. putida CA-3 utilized the primers FphaC1-FW and FphaC1-RV (see Table S1), with thermal cycling conditions as described above for amplification of full-length gacS.

RT-PCR analyses.

Total RNA was isolated from P. putida CA-3 wild type (WT) and Tn5 insertion mutants using a Fermentas GeneJet RNA purification kit, per the manufacturer's instructions. The purified RNA was treated with Turbo DNA-free DNase (Ambion) to ensure complete removal of DNA. Reverse transcription (RT) was performed with 1 μg of total RNA using random hexamer priming, 1 mM deoxynucleoside triphosphates (dNTPs), and 10 U BioScript reverse transcriptase with 1× reaction buffer (Bioline) in a 20-μl reaction volume. Reaction mixtures were incubated at 25°C for 10 min, followed by 30 min at 42°C and finally 10 min at 70°C. One microliter of the respective RT reactions was employed as the template in subsequent PCRs to confirm generation of cDNA. Amplification of the 16S rRNA gene from CA-3 genomic DNA acted as a positive control for RT analyses and was performed with the universal primers 27f and 1429r (see Table S1 in the supplemental material) at an annealing temperature of 55°C and under standard thermocycling conditions. 16S rRNA gene reverse transcription-PCR (RT-PCR) amplification was used for optimization of RT-PCR product equivalence to ensure that template cDNA concentrations were comparable between samples. Equivalence was achieved using the SynGene GeneTools v3.07 band comparison software to compare band intensities between samples. PCR cycle numbers were also optimized in assessing Gac/Rsm homologue expression, to ensure that amplifications did not reach a plateau phase.

Oligonucleotide primers (see Table S1 in the supplemental material) were also designed for transcriptional profiling of pathway-related genes: gacS, encoding the GacS sensor kinase, and gacA, encoding the response regulator of the GacS/GacA two-component system; rsmA, encoding the secondary metabolism regulator RsmA; and both rsmY and rsmZ noncoding small RNAs (sRNAs).

Bioinformatic analysis of the published rsmZ and rsmY gene sequences identified poor sequence conservation among various Pseudomonas species (data not shown). However, the genome positions of rsmZ and rsmY demonstrated high degrees of conservation within the genes flanking the respective sRNA sites. A flanking PCR and sequencing approach was therefore employed to obtain the sequence of both rsmY and rsmZ in P. putida CA-3. PCR utilizing an rpoS gene-based forward primer (rpoS-817) coupled with an fes gene-specific reverse primer (feS-45) (see Table S1 in the supplemental material) was employed to amplify the intergenic region harboring the rsmZ gene. The PCR product was sequenced and found to contain a full-length rsmZ sRNA (GenBank accession number JX826485), which was subsequently used to design P. putida CA-3-specific rsmZ primers. A similar flanking PCR approach utilizing primers ACDH-1520 and lysR-524 was used to amplify the intergenic region harboring the rsmY gene located between the lysR and ACDH (acyl-CoA dehydrogenase) genes. The amplicon generated included the full-length rsmY gene sequence (GenBank accession number JX826484), which was used to design the rsmY-specific primers rsmY-FW and rsmY-RV. Transcriptional profiling of the genes required for PHA synthesis, i.e., phaC1 (encoding the mcl-PHA synthase) and phaG (encoding an ACP-CoA transferase), was also performed using previously published primer pairs C1-F/C1-R and G-F/G-R, respectively (19).

qRT-PCR analyses of phaC1 transcripts.

Total RNA was isolated from wild-type and gacS::Tn5 P. putida CA-3 cultures grown in triplicate under PHA-accumulating conditions using a GeneJet RNA purification kit (Fermentas). Samples were subsequently DNase treated using Turbo DNA Free (Ambion) according to the manufacturer's instructions. 16S rRNA gene PCR analyses of samples were conducted to ensure the absence of contaminating DNA. cDNA synthesis was performed with random hexamers and a BioScript Moloney murine leukemia virus (MMLV)-based enzyme and buffer system (Bioline) as described previously. cDNA concentrations were calculated using a NanoDrop 2000 spectrophotometer (Thermo Scientific), and all samples were diluted to achieve final concentrations of 50 ng/μl. Oligonucleotide primer pairs suitable for quantitative reverse transcription-PCR (qRT-PCR) were designed to amplify a 177-bp fragment of the 16S rRNA gene (control) and a 158-bp fragment of the phaC1 gene target of interest. The primer sequences are listed in Table S1 in the supplemental material as 16S-915F/16S-1092R and PhaC-1289F/PhaC-1447R, respectively. PCR amplifications of internal 16S rRNA and phaC1 targets were optimized at 60°C with genomic DNA templates. Amplicon identities were confirmed by sequencing (GATC, Germany), and the purified products were used to generate serial dilutions (10⁹ to 10⁰) for standard melting curves. qRT-PCR was performed using a SYBR green QuantiTect kit (Qiagen) with a Rotor-Gene 3000, per the manufacturer's instructions.

SDS-PAGE protein analysis.

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis was performed on soluble protein fractions from wild-type P. putida CA-3, mutant PHA45A (gacS::Tn5), PHA45AgacS+ (gacS complement), B11 (phaC1::Tn5), and B11 pBBRphaC1 (phaC1 complement) under PHA accumulation conditions. Samples were incubated at 28°C with shaking at 180 rpm under nitrogen-limiting conditions as described above. After approximately 6 h, cultures were sampled and analyzed for PHA accumulation as previously described. Upon confirmation of PHA accumulation, 20 ml of actively growing culture was removed and centrifuged for 10 min at 5,000 × g. Total protein was extracted from the resulting pellet using the B-PER Plus Halt protease inhibitor cocktail kit (Thermo Scientific). Pellets were resuspended in 4 ml of buffer containing 2 μl of 20 mg/ml lysozyme (Sigma) and 2 μl DNase I (Fermentas) and incubated for 15 min at room temperature. Separation of soluble and insoluble protein fractions was achieved by centrifugation of samples at 15,000 × g for 5 min. Resultant soluble protein fractions were resuspended in sample buffer and heated for 10 min at 100°C. The protein concentration of the crude extract was determined by the Bradford assay against a bovine serine albumin standard curve (Sigma-Aldrich). Five micrograms of each protein sample was used for SDS-PAGE using a Bio-Rad Protean II Xi system (Bio-Rad) according to the manufacturer's instructions. Gels were run for 2 h at 30 mA through a 4% acrylamide stacking gel and a 12% separating gel. Prestained protein marker weight standards (New England BioLabs) were used for molecular mass estimation. Gels were stained with PageBlue stain (Fermentas) overnight per the manufacturer's instruction and destained with distilled water (dH₂O) for 10 min.

Nucleotide sequence accession numbers.

Newly determined accession numbers obtained as a result of this study have been deposited in GenBank under accession numbers JX826483, JX826484, and JX826485.

RESULTS

Cloning and characterization of PHA-negative mutants.

Random mini-Tn5 mutagenesis of the P. putida CA-3 genome produced 13,500 kanamycin-resistant transconjugants. Solid medium screening on E2 nitrogen-limited medium identified 44 transconjugants exhibiting reduced opacity, indicative of reduced PHA accumulation capacity (Fig. 1A). Fluorescence microscopic analyses further confirmed PHA accumulation deficiencies within these isolates (Fig. 1B). The mini-Tn5 insertion site was mapped in each isolate using two consecutive rounds of arbitrary PCR as previously described (25), and the resulting amplicons were sequenced. Comparative analyses of the sequence data against the GenBank nonredundant nucleotide database were performed using the NCBI BLASTn algorithm (http://blast.ncbi.nlm.nih.gov/). Our analyses revealed a wide array of gene disruption events in putative genes from diverse metabolic pathways, which could not be readily associated with PHA production. These included regions encoding putative ABC and MFS1 efflux transporters, translation initiation factors, LysR family proteins, DNA chaperones, amino acid biosynthesis proteins, and phosphoesterase/phosphoglucerate mutase proteins. However, two of the isolates (PHA11C and PHA22C) were found to harbor disruptions within the phaC1 synthase gene, which has previously been shown to be crucial for PHA accumulation in P. putida CA-3 (19). This finding confirmed the validity/efficacy of this screening strategy for identifying functionally significant genes/enzymes involved in PHA accumulation. It should be noted that the presence of a phaC2 gene has been previously reported in the pha operon of P. putida CA-3. However, it is not expressed under nitrogen-limiting conditions and does not appear to play a role in PHA synthesis in our strain (19). The regions downstream of the mini-Tn5 insertion in two of the other transconjugants, PHA39B and PHA45A, were found to share over 96% sequence similarity with gacS gene sequences from other P. putida strains. In order to rule out any possible polar effects caused by the insertion of mini-Tn5, the gacS gene disruption in PHA45A was complemented. Expression of gacS from the pBBR1MCS-5-derived gacS expression vector pBBRgacS in PHA45AgacS+ was found to completely restore the strain's ability to accumulate intracellular PHA (Fig. 2B).

Fig 1 — (A) Solid medium screen of Tn5 mutants on nitrogen-limiting E2-minimal medium supplemented with 60 mM citrate. The arrow indicates a mutant exhibiting the reduced-opacity phenotype. (B) Fluorescence microscopy of Tn5 mutant PHA45A and wild-type *P. putida* CA-3 grown on nitrogen-limiting E2-minimal medium supplemented with 60 mM citrate and stained with Nile Red at an ×1,000 magnification, using filter I3.

Fig 2 — Fluorescence microscopy of Tn5 mutants PHA45A (A) and PHA45AgacS+ (B) and *P. putida* CA-3 wild type (C) grown on nitrogen-limiting E2-minimal medium supplemented with 60 mM citrate and stained with Nile Red at an ×1,000 magnification, using filter I3.

Identification and transcriptional analyses of Gac/Rsm cascade gene homologues in P. putida CA-3.

The GacS/GacA two-component system, which positively controls the expression of regulatory sRNAs, has been reported in a variety of different bacterial species (reviewed in reference 27). The proposed mechanistic model of the cascade is presented in Fig. 3. A PCR approach was employed to screen the P. putida CA-3 genome for homologues of genes potentially involved in the Gac/Rsm regulatory cascade in the strain, namely, gacS, gacA, rsmA, rsmY, and rsmZ. Amplicons corresponding to the appropriate sizes of gacS and gacA genes were obtained in each case from P. putida CA-3 and, following sequence analyses, were found to possess high levels of nucleotide identity (85 to 99%) with the respective genes from previously published P. putida genomes (Table 1). PCR primers specific to conserved regions of previously published rsmA sequences were also employed to identify an rsmA homologue in CA-3; the resultant amplicon, when sequenced, was found to possess a nucleotide identity of 88 to 100% to other known PHA-accumulating Pseudomonas species. Due to poor sequence conservation in published genomes for the rsmY and rsmZ sRNA genes, flanking PCR approaches were employed to facilitate the design of CA-3-specific primers as previously described. Subsequent identification of rsmY and rsmZ gene homologues in P. putida CA-3 revealed 89 to 99% and 91 to 97% identities, respectively, compared to putative gene sequences obtained from published P. putida genomes (Table 1). An rsmX homologue could not be identified in P. putida CA-3, possibly due to the low conservation in the genes flanking rsmX in species in which this sRNA has previously been reported.

Fig 3 — Current understanding of the GacS/GacA cascade. Under repressive conditions, mRNA-binding RsmA sequesters target mRNA, preventing RNA polymerase binding and protein synthesis. Under nonrepressive conditions, RsmA is bound by *rsmY*, *rsmZ*, and *rsmX* sRNAs, allowing translation of mRNA to form protein.

Table 1.

BLASTn percent nucleotide identity^a

P. putida CA-3 gene(s)	% identity with strain:
P. putida CA-3 gene(s)	P. putida F1	P. putida GB1	P. putida KT2440	P. putida BIRD1	P. putida W619	P. fluorescens Pf-5
phaC1/phaA	98.1	95	98.5	97.6	89.3	83.6
phaG	99	93.7	98.8	98.3	86.8	75.8
gacS	98.3	92.6	98.6	98.4	83.3	77.7
gacA	99.3	91.5	98	98.5	84.5	79.3
rsmA/csrA	100	98.4	100	87.8	92.9	87.4
rsmY	98.4	98.4	98.4	99.2	88.9	83.1
rsmZ	97.3	94.5	97.3	95.5	90.8	73

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Comparison of the pha and Gac/Rsm cascade genes cloned from P. putida CA-3 to those in other PHA-accumulating pseudomonads.

In Pseudomonas species, the GacS/GacR two-component system regulates the expression of sRNAs that in turn sequester the RsmA posttranscriptional regulator (Fig. 3). RT-PCR analysis was employed to assess expression of the sRNA genes in both P. putida CA-3 wild type and the gacS::Tn5 PHA45A mutant. As described in Materials and Methods, the SynGene GeneTools band analysis software was employed to analyze the relative yields of control 16S PCR amplicons, with a <10% difference in band intensity between CA-3 wild-type (WT) and PHA45A cDNA samples being observed. The results of these analyses were used to ensure that equivalent cDNA quantities were employed in the subsequent PCR analysis of Gac/Rsm gene homologue expression. It was observed that gacA, rsmA, rsmY, and rsmZ are all expressed in both the CA-3 wild-type and PHA45A mutant strains under PHA accumulation conditions. The expression of gacS was also observed in wild-type CA-3 but could not be detected in the PHA45A mutant, suggesting that a single functional/transcribed copy exists in the P. putida CA-3 genome (Fig. 4).

Fig 4 — RT-PCR analysis of elements involved in the GacS signaling cascade. Lanes: 1, 16S rRNA control; 2, *gacA* response regulator; 3, *rsmA* mRNA binding protein; 4, *rsmY* sRNA molecule; 5, *rsmZ* sRNA molecule; 6, *gacS* sensor kinase.

Expression of PHA-biosynthetic genes in wild-type P. putida CA-3 and gacS mutant.

Our group has previously reported transcriptional regulation of the key pha genes (phaC1 polymerase and phaG-encoded ACP-CoA transacylase) in P. putida CA-3 under nitrogen-limiting and nonlimiting conditions (19). Here, pha gene expression profiles were examined in both the gacS::Tn5 mutant PHA45A and P. putida CA-3 wild type, with similar levels of transcription for both the synthase and transacylase being observed in the two strains (Fig. 5). qRT-PCR analysis of phaC1 gene transcript levels, relative to 16S rRNA gene expression, in triplicate wild-type and gacS::Tn5 cultures revealed relative expression levels of 1.21E−04 ± 1.7E−05 versus 7.58E−05 ± 2.8E−05, respectively. The overall fold difference in phaC1 transcripts between the wild type and the gacS mutant was determined to be 6.2E−01, with a t test score of sample variance value of 7.6E−02. Therefore, no statistically significant difference could be attributed to phaC1 transcription levels between the WT and gacS mutant strains. We therefore sought to determine whether GacS-induced control of PHA biosynthesis in P. putida CA-3 was mediated at the level of translation of the biosynthetic apparatus.

Fig 5 — RT-PCR analysis of expression of genes required for PHA accumulation. Lanes: 1, *phaC1* class II poly-3-hydroxyalkanoate synthase; 2, *phaG* 3-hydroxyacyl-CoA-ACP acyltransferase.

SDS-PAGE analyses were performed on the soluble protein fractions from wild-type P. putida CA-3, the gacS::Tn5 mutant PHA45A, and gacS-complemented mutant PHA45AgacS+ (Fig. 6). Samples were obtained from cultures grown under PHA accumulation conditions previously described (19, 28). A protein of ∼60 kDa in size which is present in wild-type CA-3 (Fig. 6, lane 1) was found to be absent in the gacS::Tn5 negative mutant PHA45A (Fig. 6, lane 2) and restored in the complemented mutant PHA45gacS+ (Fig. 6, lane 4). Previously, our group reported that the P. putida PHA synthase (phaC1::Tn5)-disrupted mutant B11 was incapable of PHA accumulation (19). Bioinformatic analysis of the P. putida CA-3 phaC1 gene sequence detailed in that study predicted a putative size for the PhaC1 protein of 62.2 kDa. Samples of the soluble protein fraction obtained from the B11 mutant grown under the same conditions were analyzed using SDS-PAGE and compared with the protein profile of gacS::Tn5 negative mutant PHA45A. The comparison of mutant B11 (Fig. 6, lane 3) with PHA45A (Fig. 6, lane 2) identified the absence of a similar ∼60-kDa protein which was present in both wild-type CA-3 (Fig. 6, lane 1) and PHA45AgacS+ (Fig. 6, lane 4). Complementation of phaC1::Tn5 mutant B11 with the full-length phaC1 gene to form B11phaC1+ was found to restore production of this ∼60-kDa protein (Fig. 6, lane 5) as well as to restore PHA granule production (see Fig. S1b in the supplemental material).

Fig 6 — SDS-PAGE of crude soluble protein extracts. Lanes: 1, *P. putida* CA-3 WT; 2, PHA45A; 3, B11; 4, PHA45AgacS+; 5, B11phaC1+; 6, molecular mass standards. The PhaC1 protein band is denoted with an arrow.

DISCUSSION

The two-component GacS/GacA system forms the sensory apparatus of the Gac/Rsm cascade, which involves interactions with small RNA regulatory molecules affecting a range of microbial metabolic pathways (27). A mechanistic model of the cascade is presented in Fig. 3 and demonstrates the sensing of appropriate extracellular signals via the GacS sensor kinase, resulting in autophosphorylation. A phosphor-transfer event is then believed to activate the associated response regulator GacA, which is found distally encoded in P. putida genomes published to date. Phosphorylated GacA acts as a transcriptional activator of several nonoperonic sRNA genes, rsmX, -Y, and -Z. These sRNAs act to sequester a negative regulator of mRNA translation, RsmA, enabling affected protein synthesis to proceed.

While GacS-dependent regulation of mcl-PHA production in Pseudomonas species has not been previously reported, an early study by Castañeda and coworkers observed the loss of scl-PHA production in a gacS-negative mutant of Azotobacter vinelandii (29). A recent follow-up study by Hernandez-Eligio et al. (30) investigated GacA regulation of PHB accumulation in the same strain. The authors reported that regulation of PHB biosynthesis proceeds in Azotobacter vinelandii via posttranscriptional regulation of PhbR, via mRNA binding by RsmA. PhbR has previously been shown to be required for transcriptional activation of the phbBAC biosynthetic operon for PHB. Regulation of phbR mRNA transcripts thus reduces expression from the phbBAC operon, resulting in reduced PHB accumulation by the cell. The correlation between the PHA-deficient phenotypes in the gacS mutants of A. vinelandii and P. putida CA-3, despite the dissimilarities between scl- and mcl-PHA biosynthetic routes, raises the possibility of a common or convergent regulatory mechanism controlling PHA production across many species of bacteria. However, the mechanism of Gac/Rsm-mediated regulation of PHB production proposed by Hernandez-Eligio et al. is unlikely to affect PHA accumulation in P. putida CA-3, as PHA regulation in this strain has not been demonstrated to be dependent on a dedicated transcriptional regulator such as, for example, PhbB in A. vinelandii. Prieto et al. have indicated that PhaF, a PHA granule-associated protein, can exert carbon source-dependent (i.e., octanoate versus citrate) regulatory influences on the expression of phaC1 in Pseudomonas oleovorans GPo1 (31). It would not appear that such an effect is relevant in this study, however, as we report that transcription of the phaC1 and phaG genes, central to PHA biosynthesis, is unaffected by gacS disruption in citrate-grown P. putida CA-3. Thus, the GacS/GacA system does not appear to be involved in controlling expression of any intermediate transcriptional regulator affecting PHA operon gene expression in this strain.

The presence and expression of typical Gac/Rsm cascade genes (gacA, rsmA, rsmY, and rsmZ) were demonstrated in both wild-type and PHA45A mutant strains of P. putida CA-3; however, an rsmX homologue could not be identified. It should be noted that while analyses of sequenced genomes of P. aeruginosa and Pseudomonas entomophila have identified rsmY and rsmZ homologues, rsmX equivalents could not be identified in either strain (reviewed in reference 27). It is possible, therefore, that the inability to clone an rsmX homologue in P. putida CA-3 in this study reflects a similar absence in our strain. Pursuant to the generation of a genome sequence for this strain, it is not possible to be definitive on this matter. Bioinformatic analyses of rsmY and rsmZ sRNA sequences revealed archetypal RsmA binding sites in both. In Pseudomonas fluorescens, RsmA has previously been shown to bind the sequence 5′-^A/_UCANGGANG^U/_A-3′ (32, 33). In our study, a sequence (5′-TCAAGGATGA-3′) matching this binding site was identified in rsmY at the +35 site with respect to the putative transcriptional start site. A similar sequence with a single-base-pair deletion was also identified at the +20 site in the rsmZ sequence, 5′-TCA_GGATGA-3′. However, the observed expression of both rsmY and rsmZ in the PHA45A mutant is not consistent with the widely held model of transcriptional regulation within the cascade, for which GacS is believed to be essential. Therefore, despite the presence in P. putida CA-3 of the key Gac/Rsm regulatory elements (exhibiting conserved structural characteristics, e.g., putative RsmA binding sites within the sRNAs), our findings suggest that GacS may not be the sole factor regulating rsmY and rsmZ sRNAs under PHA-accumulating conditions. Similarly, a recent report by Lalaouna and coworkers reported expression of both rsmY and rsmZ sRNAs in a GacA-negative mutant of Pseudomonas brassicacearum (34). The authors suggested that the expression of the rsmY and rsmZ sRNAs is not completely controlled by the GacS-GacA system and proposed that other regulators may be involved in regulating transcription of these sRNAs. Our identification of rsmY and rsmZ sRNA expression in P. putida CA-3 also suggests that an alternative mediation of sRNA expression by additional, unknown elements may be possible. Despite the expression of these sRNAs in the GacS mutant, gacS disruption in P. putida CA-3 definitively inhibits PHA accumulation. The authors therefore sought to establish whether the loss of GacS in mutant PHA45A affected posttranscriptional expression of the PHA synthetic apparatus in P. putida CA-3.

SDS-PAGE analyses of N-limited cultures indicated an ∼60-kDa protein absent in PHA45A compared with wild-type protein profiles. The size as indicated on the gel is consistent with the predicted size of PhaC1 in P. putida CA-3, ∼62 kDa. A comparative analysis of the protein profile from a previously generated phaC1-disrupted mutant, B11, revealed a similar protein absence from the gel. Complementation of gacS and phaC1 in these mutants, respectively, restored this protein and the PHA accumulation phenotype in both strains (Fig. 6). The restoration of the same band, therefore, in both PHA45AgacS+ and B11phaC1+ complemented mutants (Fig. 6, lanes 4 and 5, respectively) compared with the Tn5 mutants PHA45A and B11 (Fig. 6, lanes 2 and 3, respectively) indicates that the affected band corresponds to the PhaC1 PHA synthase. Thus, it appears that a key interaction between the GacS regulatory cascade and PHA synthesis in P. putida CA-3 occurs via the posttranscriptional regulation of the phaC1 product, a protein which we have previously shown to be required for PHA synthesis in this strain (19). It should be noted that efforts undertaken to identify a consensus RsmA binding site (5′-^A/_UCANGGANG^U/_A-3′) within the promoter region of published phaC1 sequences did not reveal any significant sequence similarity within the promoter sequences. The latter contributes further to the atypical nature of GacS-dependent regulation of PHA synthesis observed in this strain; however, it does not rule out the potential involvement of an as-yet-unidentified RsmA recognition motif associated with phaC1 mRNA.

Regulation of PHA accumulation in bacteria is known to be dependent on many physiological and regulatory factors, including transcriptional regulation, redox potential, nutrient limitation, and availability of metabolic intermediates. The physiological status of the cell is therefore vital to the accumulation of these biopolymers. The findings reported here further develop this understanding as we demonstrate, for the first time, the regulation of mcl-PHA production via GacS. The GacS/GacA system has previously been described in many bacterial genera as regulating a variety of cellular elements, including biocontrol characteristics, stress resistance, motility, virulence, central cellular metabolism, and PHB biosynthesis (reviewed in reference 27). The observance of GacS-mediated regulation of PHA production is perhaps not surprising, given its role in dealing with stress resistance and central metabolism, which often coincide with known PHA accumulation conditions (e.g., N, S, and P limitation). It is possible that the GacS/GacA sensor kinase acts in concert with other environmental sensors involved in carbon availability and redox sensing to coordinate the cell's response, e.g., by switching the metabolic focus from carbon utilization and growth to storage. Such processes must clearly involve global/common regulatory mechanisms, affecting various cellular processes, including the production of PHA in different species. Indeed, this may potentially explain why both scl- and mcl-PHA syntheses are regulated via mechanisms involving GacS, despite being distinct both genetically and biochemically. Future work in this area is required to further explore the role of GacS and the specific operation of the associated sRNA regulatory cascade in mcl-PHA production and to determine whether any biotechnological advances in PHA synthesis can be achieved via recombinant manipulations of the regulatory cascade in PHA-producing strains.

ACKNOWLEDGMENT

This work was funded by the Science, Technology, Research and Innovation for the Environment 2007-2013 (STRIVE) fellowship program of the Irish Environmental Protection Agency (grant no. 2007-PhD-ET-8).

Footnotes

Published ahead of print 4 January 2013

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.02962-12.

REFERENCES

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22. de Lorenzo V, Herrero M, Jakubzik U, Timmis KN. 1990. Mini-Tn5 transposon derivatives for insertion mutagenesis, promoter probing, and chromosomal insertion of cloned DNA in gram-negative eubacteria. J. Bacteriol. 172:6568–6572 [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Kovach ME, Elzer PH, Hill DS, Robertson GT, Farris MA, Roop RM, Peterson KM. 1995. Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene 166:175–176 [DOI] [PubMed] [Google Scholar]
24. Vogel HJ, Bonner DM. 1956. Acetylornithinase of E. coli: partial purification and some properties. J. Biol. Chem. 218:97–106 [PubMed] [Google Scholar]
25. Espinosa-Urgel M, Salido A, Ramos JL. 2000. Genetic analysis of functions involved in adhesion of Pseudomonas putida to seeds. J. Bacteriol. 182:2363–2369 [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K. 1987. Current protocols in molecular biology. Greene Publishing Associates-Wiley Interscience, New York, NY [Google Scholar]
27. Lapouge K, Schubert M, Allain FHT, Haas D. 2008. Gac/Rsm signal transduction pathway of γ-proteobacteria: from RNA recognition to regulation of social behaviour. Mol. Microbiol. 67:241–253 [DOI] [PubMed] [Google Scholar]
28. Ward PG, de Roo G, O'Connor KE. 2005. Accumulation of polyhydroxyalkanoate from styrene and phenylacetic acid by Pseudomonas putida CA-3. Appl. Environ. Microbiol. 74:2046–2052 [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Castañeda M, Guzmán J, Moreno S, Espín G. 2000. The GacS sensor kinase regulates alginate and poly-β-hydroxybutyrate production in Azotobacter vinelandii. J. Bacteriol. 182:2624–2628 [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Hernandez-Eligio A, Moreno S, Castellanos M, Castañeda M, Nuñez C, Muriel-Millan LF, Espín G. 2012. RsmA post-transcriptionally controls PhbR expression and polyhydroxybutyrate biosynthesis in Azotobacter vinelandii. 158:1953–1963 [DOI] [PubMed] [Google Scholar]
31. Prieto MA, Bühler B, Jung K, Witholt B, Kessler B. 1999. PhaF, a polyhydroxyalkanoate-granule-associated protein of Pseudomonas oleovorans GPo1 involved in the regulatory expression system for pha genes. J. Bacteriol. 181:858–868 [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Lapouge K, Sineva E, Lindell M, Starke S, Baker CS, Babitzke P, Haas D. 2007. Mechanism of hcnA mRNA recognition in the Gac/Rsm signal transduction pathway of Pseudomonas fluorescens. Mol. Microbiol. 66:341–356 [DOI] [PubMed] [Google Scholar]
33. Schubert M, Lapouge K, Duss O, Oberstrass FC, Jelesarov I, Haas D, Allain FHT. 2007. Molecular basis of messenger RNA recognition by the specific bacterial repressing clamp RsmA/CsrA. Nat. Struct. Mol. Biol. 14:807–813 [DOI] [PubMed] [Google Scholar]
34. Lalaouna D, Fochesato S, Sanchez L, Schmitt-Kopplin P, Haas D, Heulin T, Achouak W. 2012. Phenotypic switching involves GacS/GacA-dependent Rsm small RNAs in Pseudomonas brassicacearum. Appl. Environ. Microbiol. 78:1658–1665 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B1] 1. Lageveen RG, Huisman GW, Preusting H, Ketelaar P, Eggink G, Witholt B. 1988. Formation of polyesters by Pseudomonas oleovorans: effect of substrates on formation and composition of poly-(R)-3-hydroxyalkanoates and poly-(R)-3-hydroxyalkenoates. Appl. Environ. Microbiol. 54:2924–2932 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B2] 2. Ramsay BA, Saracovan I, Ramsay JA, Marchessault RH. 1992. Effect of nitrogen limitation on long-side-chain poly-beta-hydroxyalkanoate synthesis by Pseudomonas resinovorans. Appl. Environ. Microbiol. 58:744–746 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B3] 3. Huisman GW, Wonink E, Meima R, Kazemier B, Terpstra P, Witholt B. 1991. Metabolism of poly-(3-hydroxyalkanoates) (PHAs) by Pseudomonas oleovorans. J. Biol. Chem. 266:2191–2198 [PubMed] [Google Scholar]

[B4] 4. García B, Olivera ER, Miñambres B, Fernández-Valverde M, Cañedo LM, Prieto MA, García JL, Martínez M, Luengo JM. 1999. Novel biodegradable aromatic plastics from a bacterial source. Genetic and biochemical studies on a route of the phenylacetyl-CoA catabolon. J. Biol. Chem. 274:29228–29241 [DOI] [PubMed] [Google Scholar]

[B5] 5. Philip S, Keshavarz T, Roy I. 2007. Polyhydroxyalkanoates: biodegradable polymers with a range of applications. J. Chem. Technol. Biotechnol. 82:233–247 [Google Scholar]

[B6] 6. Grage K, Jahns AC, Parlane N, Palanisamy R, Rasiah IA, Atwood JA, Rehm BHA. 2009. Bacterial polyhydroxyalkanoate granules: biogenesis, structure, and potential use as nano-/micro-beads in biotechnological and biomedical applications. Biomacromolecules 10:660–669 [DOI] [PubMed] [Google Scholar]

[B7] 7. Gomez JGC, Méndez BS, Nikel PI, Pettinari MJ, Prieto MA, Silva LF. 2012. Making green polymers even greener: towards sustainable production of polyhydroxyalkanoates from agroindustrial by-products, p 41–63 In Petre M. (ed), Advances in applied biotechnology. InTech, New York, NY [Google Scholar]

[B8] 8. van der Walle GA, Buisman GJH, Weusthuis RA, Eggink G. 1999. Development of environmentally friendly coatings and paints using medium-chain-length poly(3-hydroxyalkanoates) as the polymer binder. Int. J. Biol. Macromol. 25:123–128 [DOI] [PubMed] [Google Scholar]

[B9] 9. van der Walle GA, de Koning GJ, Weusthuis RA, Eggink G. 2001. Properties, modifications and applications of biopolyesters. Adv. Biochem. Eng. Biotechnol. 71:263–291 [DOI] [PubMed] [Google Scholar]

[B10] 10. Chen GQ, Wu Q. 2005. The application of polyhydroxyalkanoates as tissue engineering materials. Biomaterials 26:6565–6578 [DOI] [PubMed] [Google Scholar]

[B11] 11. Senior PJ, Dawes EA. 1971. Poly-β-hydroxybutyrate biosynthesis and the regulation of glucose metabolism in Azotobacter beijerinckii. Biochem. J. 125:55–66 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B12] 12. Tsuge T, Fukui T, Matsusaki H, Taguchi S, Kobayashi G, Ishizaki A, Doi Y. 2000. Molecular cloning of two (R)-specific enoyl-CoA hydratase genes from Pseudomonas aeruginosa and their use for polyhydroxyalkanoate synthesis. FEMS Microbiol. Lett. 184:193–198 [DOI] [PubMed] [Google Scholar]

[B13] 13. Rehm BHA, Steinbüchel A. 1999. Biochemical and genetic analysis of PHA synthases and other proteins required for PHA synthesis. Int. J. Biol. Macromol. 25:3–19 [DOI] [PubMed] [Google Scholar]

[B14] 14. Lim SJ, Jung YM, Shin HD, Lee YH. 2002. Amplification of the NADPH-related genes zwf and gnd for the oddball biosynthesis of PHB in an E. coli transformant harboring a cloned phbCAB operon. J. Biosci. Bioeng. 93:543–549 [DOI] [PubMed] [Google Scholar]

[B15] 15. Hong SH, Park SJ, Moon SY, Park JP, Lee SY. 2003. In silico prediction and validation of the importance of the Entner-Doudoroff pathway in poly(3-hydroxybutyrate) production by metabolically engineered Escherichia coli. Biotechnol. Bioeng. 83:854–863 [DOI] [PubMed] [Google Scholar]

[B16] 16. Nikel PI, Pettinari MJ, Galvagno MA, Méndez BS. 2006. Poly(3-hydroxybutyrate) synthesis by recombinant Escherichia coli arcA mutants in microaerobiosis. Appl. Environ. Microbiol. 72:2614–2620 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B17] 17. Avison MB, Horton RE, Walsh TR, Bennett PM. 2001. Escherichia coli CreBC is a global regulator of gene expression that responds to growth in minimal media. J. Biol. Chem. 276:26955–26961 [DOI] [PubMed] [Google Scholar]

[B18] 18. Nikel PI, de Almeida A, Pettinari MJ, Méndez BS. 2008. The legacy of HfrH: mutations in the two-component system CreBC are responsible for the unusual phenotype of an Escherichia coli arcA mutant. J. Bacteriol. 190:3404–3407 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B19] 19. O'Leary ND, O'Connor KE, Ward P, Goff M, Dobson ADW. 2005. Genetic characterization of accumulation of polyhydroxyalkanoate from styrene in Pseudomonas putida CA-3. Appl. Environ. Microbiol. 71:4380–4387 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B20] 20. O'Leary ND, O'Mahony M, Dobson ADW. 2011. Regulation of phenylacetic acid uptake is σ54 dependent in Pseudomonas putida CA-3. BMC Microbiol. 11:229 doi:10.1186/1471-2180-11-229 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21. O'Connor KE, Buckley CM, Hartmans S, Dobson ADW. 1995. Possible regulatory role for nonaromatic carbon sources in styrene degradation by Pseudomonas putida CA-3. Appl. Environ. Microbiol. 61:544–548 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B22] 22. de Lorenzo V, Herrero M, Jakubzik U, Timmis KN. 1990. Mini-Tn5 transposon derivatives for insertion mutagenesis, promoter probing, and chromosomal insertion of cloned DNA in gram-negative eubacteria. J. Bacteriol. 172:6568–6572 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B23] 23. Kovach ME, Elzer PH, Hill DS, Robertson GT, Farris MA, Roop RM, Peterson KM. 1995. Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene 166:175–176 [DOI] [PubMed] [Google Scholar]

[B24] 24. Vogel HJ, Bonner DM. 1956. Acetylornithinase of E. coli: partial purification and some properties. J. Biol. Chem. 218:97–106 [PubMed] [Google Scholar]

[B25] 25. Espinosa-Urgel M, Salido A, Ramos JL. 2000. Genetic analysis of functions involved in adhesion of Pseudomonas putida to seeds. J. Bacteriol. 182:2363–2369 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26. Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K. 1987. Current protocols in molecular biology. Greene Publishing Associates-Wiley Interscience, New York, NY [Google Scholar]

[B27] 27. Lapouge K, Schubert M, Allain FHT, Haas D. 2008. Gac/Rsm signal transduction pathway of γ-proteobacteria: from RNA recognition to regulation of social behaviour. Mol. Microbiol. 67:241–253 [DOI] [PubMed] [Google Scholar]

[B28] 28. Ward PG, de Roo G, O'Connor KE. 2005. Accumulation of polyhydroxyalkanoate from styrene and phenylacetic acid by Pseudomonas putida CA-3. Appl. Environ. Microbiol. 74:2046–2052 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B29] 29. Castañeda M, Guzmán J, Moreno S, Espín G. 2000. The GacS sensor kinase regulates alginate and poly-β-hydroxybutyrate production in Azotobacter vinelandii. J. Bacteriol. 182:2624–2628 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B30] 30. Hernandez-Eligio A, Moreno S, Castellanos M, Castañeda M, Nuñez C, Muriel-Millan LF, Espín G. 2012. RsmA post-transcriptionally controls PhbR expression and polyhydroxybutyrate biosynthesis in Azotobacter vinelandii. 158:1953–1963 [DOI] [PubMed] [Google Scholar]

[B31] 31. Prieto MA, Bühler B, Jung K, Witholt B, Kessler B. 1999. PhaF, a polyhydroxyalkanoate-granule-associated protein of Pseudomonas oleovorans GPo1 involved in the regulatory expression system for pha genes. J. Bacteriol. 181:858–868 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B32] 32. Lapouge K, Sineva E, Lindell M, Starke S, Baker CS, Babitzke P, Haas D. 2007. Mechanism of hcnA mRNA recognition in the Gac/Rsm signal transduction pathway of Pseudomonas fluorescens. Mol. Microbiol. 66:341–356 [DOI] [PubMed] [Google Scholar]

[B33] 33. Schubert M, Lapouge K, Duss O, Oberstrass FC, Jelesarov I, Haas D, Allain FHT. 2007. Molecular basis of messenger RNA recognition by the specific bacterial repressing clamp RsmA/CsrA. Nat. Struct. Mol. Biol. 14:807–813 [DOI] [PubMed] [Google Scholar]

[B34] 34. Lalaouna D, Fochesato S, Sanchez L, Schmitt-Kopplin P, Haas D, Heulin T, Achouak W. 2012. Phenotypic switching involves GacS/GacA-dependent Rsm small RNAs in Pseudomonas brassicacearum. Appl. Environ. Microbiol. 78:1658–1665 [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

GacS-Dependent Regulation of Polyhydroxyalkanoate Synthesis in Pseudomonas putida CA-3

William J Ryan

Niall D O'Leary

Mark O'Mahony

Alan D W Dobson

Abstract

INTRODUCTION

MATERIALS AND METHODS