Sulfate Ester Detergent Degradation in Pseudomonas aeruginosa Is Subject to both Positive and Negative Regulation
ABSTRACT
Bacteria using toxic chemicals, such as detergents, as growth substrates face the challenge of exposing themselves to cell-damaging effects that require protection mechanisms, which demand energy delivered from catabolism of the toxic compound. Thus, adaptations are necessary for ensuring the rapid onset of substrate degradation and the integrity of the cells. Pseudomonas aeruginosa strain PAO1 can use the toxic detergent sodium dodecyl sulfate (SDS) as a growth substrate and employs, among others, cell aggregation as a protection mechanism. The degradation itself is also a protection mechanism and has to be rapidly induced upon contact to SDS. In this study, gene regulation of the enzymes initiating SDS degradation in strain PAO1 was studied. The gene and an atypical DNA-binding site of the LysR-type regulator SdsB1 were identified and shown to activate expression of the alkylsulfatase SdsA1 initiating SDS degradation. Further degradation of the resulting 1-dodecanol is catalyzed by enzymes encoded by laoCBA, which were shown to form an operon. Expression of this operon is regulated by the TetR-type repressor LaoR. Studies with purified LaoR identified its DNA-binding site and 1-dodecanoyl coenzyme A as the ligand causing detachment of LaoR from the DNA. Transcriptional studies revealed that the sulfate ester detergent sodium lauryl ether sulfate (SLES) induced expression of sdsA1 and the lao operon. Growth experiments revealed an essential involvement of the alkylsulfatase SdsA1 for SLES degradation. This study revealed that the genes for the enzymes initiating the degradation of toxic sulfate-ester detergents are induced stepwise by a positive and a negative regulator in P. aeruginosa strain PAO1.
IMPORTANCE Bacterial degradation of toxic compounds is important not only for bioremediation but also for the colonization of hostile anthropogenic environments in which biocides are being used. This study with Pseudomonas aeruginosa expands our knowledge of gene regulation of the enzymes initiating degradation of sulfate ester detergents, which occurs in many hygiene and household products and, consequently, also in wastewater. As an opportunistic pathogen, P. aeruginosa causes severe hygienic problems because of its pronounced biocide resistance and its metabolic versatility, often combined with its pronounced biofilm formation. Knowledge about the regulation of detergent degradation, especially regarding the ligands of DNA-binding regulators, may lead to the rational development of specific inhibitors for restricting growth and biofilm formation of P. aeruginosa in hygienic settings. In addition, it may also contribute to optimizing bioremediation strategies not only for detergents but also for alkanes, which when degraded merge with sulfate ester degradation at the level of long-chain alcohols.
INTRODUCTION
Bioremediation processes such as wastewater treatment and decontamination of soils rely on the ability of bacteria to use toxic organic compounds, such as solvents or detergents, as growth substrates. The presence of such toxic compounds is generally a challenge for bacteria and requires energy-consuming protection mechanisms such as efflux pumps (1) and chaperones (2). If the toxic compound is the only source of energy and carbon, this challenge is increased because the bacterial cells have to take up and expose their cell membrane and cytosol to these chemicals. Since the energy needed for protection against this chemical stress has to originate from the catabolism of the toxic substrate, this process requires elaborated adaptations for providing a well-balanced regulation of catabolism and protection mechanisms to avoid detrimental cell damage.
Degradation of the anionic detergent sodium dodecyl sulfate (SDS) by Pseudomonas aeruginosa is a suitable paradigm for studying how bacteria cope to grow with a toxic compound as a carbon and energy source. In the past we have shown that a sufficient energy status of the cells is crucial for survival, e.g., for operating proton-motif-force-dependent efflux-pumps such as MexEF-OprM (3–5). In addition, we were able to show that cell aggregation serves as a preadaptive survival strategy of P. aeruginosa for growth with SDS (6). Cell aggregation is induced via a signal transduction system encoded by siaABCD and increases the survival rate upon exposure to SDS-related stress several hundredfold compared to suspended single cells (4, 7). Since the degradation of the toxic compounds also contributes to minimizing the toxic effects, the rapid induction of synthesis of the respective enzymes is crucial. This calls attention to the genetic regulation of the enzymes that initiate SDS degradation in P. aeruginosa. While the ability of SDS degradation in pseudomonads has been known for many years (8–12), the regulation of the involved enzymes and stimulus perception is poorly understood.
The initial step in SDS degradation by P. aeruginosa strain PAO1 starts with the hydrolysis of the sulfate ester group catalyzed by the well-studied alkylsulfatase SdsA1 (PA0740) (13) (Fig. 1, left panel). We recently identified the Lao system to be involved in the following oxidation steps of the first intermediate, the long-chain primary alcohol 1-dodecanol, and the emerging aldehyde 1-dodecanal (14). In particular, LaoA and LaoB (PA0364-65) were found to be responsible but not essential for the 1-dodecanol oxidation. The respective genes are located together with laoC (PA0366), encoding an aldehyde dehydrogenase, in one gene cluster. In addition, the Lao system has been shown to participate in the oxidation of long-chain alcohols originating from alkane degradation (14). Thus, the Lao system is apparently disconnected from the initial SDS degradation step not only by its genomic localization but also by its substrate specificity. Accordingly, different regulatory pathways are required to enable the expression of the Lao system in the absence of SDS. Thus far, the LysR-type regulator SdsB for the alkylsulfatase SdsA in Pseudomonas sp. strain ATCC 19151 has been specified and analyzed. This regulator gene has 54% identity to PA0739 (15, 16), which is located directly adjacent to sdsA1 in an opposite direction on the P. aeruginosa PAO1 genome (Fig. 1, right panel). In addition, we have already shown that the TetR-type regulator LaoR (formerly PA0367), the gene of which is located directly adjacent to laoCBA in the opposite direction (Fig. 1, right panel), regulates the genes encoding the Lao enzymes (14).
FIG 1
The goal of our study was to study the question how P. aeruginosa copes to grow with SDS as a carbon and energy source from the point of genetic regulation. In particular, we sought to identify the regulator for sdsA1, as well as inducers for expression of the initiating enzymes in the SDS degradation.
RESULTS
Identification of the regulator SdsB1.
For analyzing whether the gene PA0739 is involved in the SDS metabolism of P. aeruginosa strain PAO1, a chromosomal gene deletion strain was constructed. Growth of the wild-type strain PAO1 was compared to that of the PAO1 ΔsdsB1 deletion mutant strain and the complemented mutant. During growth with succinate, no differences between strain PAO1 and the PAO1 ΔsdsB1 deletion mutant strain were observable (data not shown). With SDS as the sole carbon and energy source (Fig. 2), the PAO1 ΔsdsB1 deletion mutant strain exhibited almost no growth, as well as no SDS degradation (Fig. 2A). Growth could be restored to wild-type level by providing an intact copy of PA0739 on the plasmid pUCP18 (Fig. 2B). These results demonstrated that PA0739 constituted the respective regulator for sdsA1, which encodes the essential alkylsulfatase for growth with SDS (13, 16). Hence, PA0739 was named sdsB1.
FIG 2
Analysis of the DNA-binding site of SsdB1.
To identify the DNA-binding site of SdsB1, we purified His-tagged SdsB1 and performed electrophoretic mobility shift assays (EMSAs) with different DNA fragments of the intergenic region between sdsB1 and sdsA1 (Fig. 3). These experiments revealed a binding within the first 40 bp of the intergenic region (Fig. 3B). This DNA fragment exhibits two inverted repeats (IRs), which partly overlap. The IRs were either solely or both exchanged by the nucleotide exchanges depicted in Fig. 3C. Changes in one of these IRs revealed a reduced SdsB1 DNA binding, while the change of both IRs completely abolished SdsB1 DNA binding (Fig. 3D). This result revealed binding of SdsB1 to an atypical LysR-type regulatory binding site (RBS) with a binding motif which is different from the classical and conserved T-N11-A binding motif (17, 18).
FIG 3
Reverse transcriptase PCR analysis of the gene cluster laoCBA.
The genes laoCBA are located in a gene cluster on the minus strand separated by only two 53- to 54-bp-long intergenic regions, suggesting an operon organization (Fig. 1). Reverse transcriptase PCR was used to identify such putative polycistronic mRNA in cells grown with 1-dodecanol compared to cells grown with succinate (Fig. 4). Succinate-grown cells have already been demonstrated to not exhibit 1-dodecanol oxidation activity by the Lao system (14). As shown in Fig. 4, polycistronic mRNA could be detected in cells grown with 1-dodecanol but not in succinate-grown cells. As a result, the laoCBA genes constitute an operon, which is induced during growth with 1-dodecanol.
FIG 4
Analysis of the DNA-binding site of LaoR.
In the next step, we examined whether the TetR-type repressor LaoR binds to the intergenic region between laoC and laoR. For this purpose, His-tagged LaoR was purified and EMSAs with different DNA fragments of the 288-bp intergenic region were performed (Fig. 5). LaoR was found to bind within the central 88-bp fragment. Two inverted repeats (within fragment C1 or C2) of this central 88-bp fragment were assumed to constitute the possible binding sites (Fig. 5A and C). Both IRs, as well as exchanges of nucleotides within fragment C1, were analyzed by EMSAs (Fig. 5B and D). The IR of fragment C1 was shown to be the actual binding site for LaoR.
FIG 5
Analysis of SdsB1 and LaoR binding sites by thermal shift assays.
Thermal shift assays are used as a biochemical method for the detection of protein stability by quantification of their melting temperature. Because DNA binding should have a stabilizing effect on regulator proteins, this method should be applicable to analyze DNA-binding sites but has, to our knowledge, not been reported thus far. The DNA-regulator binding activities for the regulators SdsB1 (Fig. 6A) and LaoR (Fig. 6B) showed an increased protein melting temperature in the presence of the specific DNA-binding sites identified by the EMSA analysis. Thus, the thermal shift assays could constitute an alternative and rapid method for analyzing DNA-binding sites of regulators.
FIG 6
Ligand analysis of LaoR.
The classical TetR-type regulators are released from the DNA by ligand binding, which leads to the expression of their controlled genes (19). To identify the corresponding ligand for LaoR, EMSAs with SDS and intermediates of SDS degradation were performed (Fig. 7). Increasing SDS concentrations exhibited an increasing influence on the LaoR-DNA complex resulting in a detachment of LaoR from its DNA-binding site (Fig. 7A). This effect can probably be explained by the protein denaturing property of the toxic detergent SDS (20). The degradation intermediate 1-dodecanal also exhibited an influence on the LaoR-DNA complex, with a slight detachment of the regulator from the DNA binding site, which also might be due to denaturing effects on the protein since this is well known for reactive aldehydes (21). The SDS degradation intermediates 1-dodecanol and 1-dodecanoic acid, as well as coenzyme A (CoA) or a combination of 1-dodecanoic acid and CoA, exhibited no detachment. Strikingly, the degradation intermediate 1-dodecanoyl-CoA influenced the LaoR-DNA complex, resulting in a detachment detected by removal of the shifted band (Fig. 7B). Decanoyl-CoA caused detachment only at higher concentrations, while hexanoyl-CoA did not cause detachment even at the highest concentration. Accordingly, this experiment reveals specificity for long-chain acyl-CoA esters as ligands for LaoR.
FIG 7
Transcriptional analysis of sdsA1 and laoCBA.
In the next step, we analyzed whether the SDS substrate analogues sodium lauryl ether sulfate (SLES) and dodecane sulfate would also induce the expression of sdsA1 and the lao operon. For this purpose, transcriptional fusions of the intergenic regions between genes sdsB1 and sdsA1 and between genes laoC and laoR with the lacZ gene were constructed and analyzed by supplying suspensions of noninduced succinate-grown cells with the putative inducers (Fig. 8A). These experiments showed that gene expression of both systems is induced in the presence of the sulfate esters SDS and SLES. In contrast, dodecane sulfonate, in which the sulfur atom is directly attached to the alkyl chain by a C-S bond, did not induce gene expression. The expression of the lao operon was additionally induced in the presence of the LaoAB substrate 1-dodecanol. In the ΔsdsB1 mutant, expression of sdsA1 could not be induced (Fig. 8A), while in the ΔlaoR mutant, expression of the lao operon was constitutively and strongly induced, reaching Miller unit values of about 2,000 even for succinate-grown cells (data not shown). Furthermore, we analyzed the mRNA levels of sdsA1 and laoA in SDS-, SLES-, and 1-dodecanol-grown cells in comparison to succinate-grown cells by quantitative real-time PCR (qRT-PCR) (Fig. 8B). An induced expression of sdsA1 in the presence of SDS and SLES could be confirmed (Fig. 8BI). The reason for the large quantitative difference of induction between the lacZ reporter gene assay (∼3.5-fold) and the quantitative real-time PCR measurement (∼140-fold) is unknown. It might be due to the different experimental setups (induction assays with cell suspensions and growth experiments, respectively) or to an influence of SDS on mRNA stability. The lao operon was shown to be induced with 1-dodecanol and SDS, while an induction by SLES was hardly detectable (mRNA fold change, 2.4 ± 0.9) (Fig. 8BII).
FIG 8
We also analyzed the mRNA levels of the regulators SdsB1 and LaoR themselves. While the mRNA level of sdsB1 was not increased in SDS- and SLES-grown cells compared to succinate-grown cells, the mRNA level of laoR was increased in SDS- and 1-dodecanol-grown cells (Fig. 8BIII and IV), indicating an autoinduction of laoR expression.
Growth of P. aeruginosa with SLES.
The inducing effects of SLES raised the question whether this detergent is also a growth substrate for strain PAO1 and, if so, whether SdsA1 and LaoABC are also involved in its degradation. In the respective growth experiments, the PAO1 ΔsdsB1 mutant strain exhibited almost no growth in comparison to the wild-type and complemented strains (Fig. 9A), and analysis of the SLES concentration revealed no SLES degradation for the PAO1 ΔsdsB1 mutant strain (Fig. 9BI). Thus, the alkylsulfatase SdsA1 was apparently essential for SLES degradation in P. aeruginosa PAO1. The growth rate of the PAO1 ΔlaoA mutant strain (μ = 0.06 h−1) was about 50% lower than that of the wild type (μ = 0.13 h−1) during growth with SLES. In contrast, SLES degradation was similar to that of the wild type (Fig. 8BII). Cultures of strain PAO1 growing with SLES revealed a very similar formation of macroscopic cell aggregates, as has been described during growth with SDS (5).
FIG 9
DISCUSSION
The goal of this study was to elucidate how the expression of the enzymes catalyzing the initial reactions of SDS degradation, namely, of the alkylsulfatase SdsA1 and the long-chain alcohol-oxidizing enzymes encoded by laoCBA, in P. aeruginosa strain PAO1 is regulated. We could show that these reactions are induced in a two-step manner by a consecutive positive and negative regulation. In the first step, the transcription of sdsA1 encoding the sulfatase converting SDS into sulfate and 1-dodecanol (13) is activated by the LysR-type regulator SdsB1 (PA0739), which we identified in this study. This activation was caused by SDS, as well as by SLES, another sulfate ester detergent and commonly used alternative for SDS. In the second step, transcription of the laoCBA operon is induced by a detachment of the TetR-type repressor LaoR from its DNA binding site. Our analysis revealed that long-chain acyl-CoA esters can act as ligands, with dodecanoyl-CoA exhibiting the highest efficiency of LaoR detachment from its DNA-binding site. Thus, derepression of the laoCBA operon would require basal expression of the encoded enzymes and an as-yet-unknown acyl-CoA-ligase for the formation of this ligand. This two-step induction of SDS degradation could be the outcome of a strategy that ensures rapid inactivation of a toxic sulfate ester detergent, while the degradation of the resulting long-chain alcohol is only induced when it can sufficiently be further converted into a metabolite that signals the availability of an energy-rich substrate for β-oxidation. This derepression of the laoCBA operon would result in the increased formation of the inductor 1-dodecanoyl-CoA and thus in a positive regulatory feedback loop. Further degradation of the activated fatty acid by β-oxidation then enables both growth and the operation of protection mechanisms against SDS. The latter is particularly important for survival since we have previously concluded that dividing cells are more vulnerable to the damaging effects of SDS (5). The observation that the laoR gene is subject to positive autoregulation suggests that the lao operon can be rapidly repressed again when the inducer is depleted, thereby preventing undue expression. In this respect it is unlikely that SDS, although it caused detachment of LaoR in the EMSAs, is a specific inducer of the lao operon, which is also required for the degradation of alkanes that are much more abundant in nature than SDS (16).
EMSA analyses revealed that SdsB1 possesses an atypical DNA-binding site within the first 40-bp DNA fragment upstream of sdsB1, which is located in the intergenic region between sdsB1 and sdsA1 (Fig. 3B). LysR-type regulators often bind to an RBS and a distinct activation binding site (18). The RBS generally has a recognized LTTR (LysR-type transcriptional regulator) box with the conserved palindromic sequence T-N11-A (18, 22) that was not identified within the fragment bound by SdsB1. Instead, we identified two IRs, gcat-N16-atgc and aatg-N13-catt, which partly overlap within the bound DNA fragment (Fig. 3). An in silico protein domain analysis (Interpro) (23) of SdsB1 exhibited a conserved CysB-like substrate binding domain (IPR 000847). Interestingly, complex DNA-binding with multiple overlapping DNA binding sites is also described for CysB from Salmonella enterica serovar Typhimurium (24, 25). Moreover, the binding site of the master regulator CysB from P. aeruginosa (26–28) contains two inverted repeats, which partly overlap (28). However, an alignment of SdsB1 and CysB from P. aeruginosa, as well as S. Typhimurium, revealed only 24% identity in each case (BLASTP) (29). Nevertheless, both CysB regulators share their involvement in metabolism of organic sulfur compounds with SdsB1 (30, 31). In addition, a recent study identified sulfate ions captured within the cleft of the crystallized C-terminal regulatory domain (substrate binding domain) of CysB from P. aeruginosa, pointing toward sulfate ions as potential ligands influencing the function of this regulator (27). Since sulfate was the sulfur source for P. aeruginosa in our experiments, it is unlikely to be the inducer for SdsB1. The expression of sdsA1 via SdsB1 was induced only by substrates of SdsA1, namely, SDS and SLES, but not by the structural analogue dodecyl sulfonate, which cannot be cleaved by SdsA1. Thus, a sulfate group attached to a long-chain alcohol is apparently a decisive structural element for a ligand of SdsB1, thereby supporting the similarities between SdsB1 and CysB.
A typical DNA-binding site was identified to be occupied by the TetR-type regulator LaoR. Ligands of TetR-type regulators comprise a remarkable variety of compounds, including antibiotics, bile acids, cell-cell signaling molecules, proteins, metal ions, or fatty acids (19, 32). Also, acyl-CoA esters such as oleoyl-CoA and palmitoyl-CoA for the regulator DesT, which is involved in lipid metabolism of P. aeruginosa, are known as ligands (33). One example very similar to our proposed model for the positive-feedback regulation for LaoR is the TetR regulator AlkX from Dietzia sp. strain DQ12-45-1b (34). This regulator controls the expression of genes involved in the degradation of alkanes and binds long-chain fatty acids derived from alkane degradation as ligands. The relatively high concentration of dodecanoyl-CoA required for the detachment of LaoR from its DNA-binding site might indicate that a threshold concentration is required for inducing the postulated positive-feedback regulation. However, since acyl-CoA esters are already unstable in aqueous solution at neutral pH values, the actual concentration of this ligand might have been overestimated in the respective experiments.
The finding that SLES serves as an inducer for the SdsB1-mediated sdsA1 transcription prompted us to test whether this sulfate ester detergent could also serve as a carbon and energy source. To the best of our knowledge, the growth of P. aeruginosa strain PAO1 with SLES has not yet been described. This milder detergent often replaces SDS in cosmetic and hygienic products due to its reduced skin irritation properties (35–37). Since SLES also induced cell aggregation in P. aeruginosa, this sulfate ester detergent apparently also activates the Sia system, indicating that it presumably has toxic effects similar to those of SDS. Generally, SLES degradation has been observed in other pseudomonads, but the degradation pathway is not completely understood. Metabolite analysis in Pseudomonas sp. strain Des1 (38, 39), Pseudomonas sp. strain SC25A (40), and P. nitroreducens (41) revealed that degradation starts with cleavage of the ether bonds, but cleavage of the sulfate ester bond has also been shown to occur concomitantly to ether cleavage in some of these strains (41, 42). However, in strain PAO1, the sdsB1 deletion mutant, which does not express the alkylsulfatase SdsA1, could not no longer grow with SLES, indicating that SLES degradation is compulsively initiated by sulfate ester hydrolysis. This conclusion is supported by the fact that the stains-all assay (see Materials and Methods) used to quantify sulfate ester detergents did not indicate a decrease in SLES concentration. While SdsA1 apparently had an essential role in SLES metabolism, the Lao system is clearly involved but not essential. The reduced growth rate of the PAO1 ΔlaoA strain with SLES could plausibly be explained by the presumptive formation of 1-dodecanol after cleavage of the ethoxy units from the SLES molecule. There are many different ether-cleaving reactions known among bacteria (43), but they are unknown for P. aeruginosa and are therefore under investigation in our laboratory.
In conclusion, our study is a further step toward the understanding of how P. aeruginosa copes to grow with toxic detergent SDS as a carbon and energy source. The next steps for pursuing this research question further involve analysis of how expression of the metabolic pathway and the protection mechanisms, especially cell aggregation and efflux pumps, are coordinated.
MATERIALS AND METHODS
Bacterial strains and growth experiments.
Bacterial strains and plasmids used in this study are described in Table 1. Cells were grown in lysogeny broth (LB) medium or in modified M9 mineral medium (5, 44) with variable carbon and energy sources, which were supplied at the following concentrations (the percent purity is indicated in parentheses): 10 mM succinate (≥99%), 3.5 mM SDS (≥99.5%), and 3.5 mM 1-dodecanol (≥98.0%) (Sigma-Aldrich, St. Louis, MO). A 27% SLES aqueous solution was purchased from Caelo (Hilden, Germany) with an average Mw of 382 g mol−1, an average of two degrees of ethoxylation (n = 2), and an average alkyl chain length of 12 carbons (information given by the manufacturer). Escherichia coli strains harboring plasmids were maintained and selected on LB agar plates (1.5% [wt/vol]) with 100 μg ml−1 carbenicillin (Carl Roth GmbH, Karlsruhe, Germany). For E. coli ST18, media were supplemented with 50 μg ml−1 5-aminolevulinic acid (Sigma-Aldrich). Plasmid-harboring P. aeruginosa strains were maintained and selected on LB agar plates (1.5% [wt/vol]) with 200 μg ml−1 carbenicillin. Growth was monitored as the optical density at 600 nm (OD600) with the UV-mini 1240 photometer (Shimadzu, Kyoto, Japan). Overnight precultures in 10-ml test tubes with 3 ml of LB were inoculated from LB agar plates and incubated at 30°C and 200 rpm in a rotary shaker (INFORS HT; Minitron, GmbH, Bottmingen, Switzerland) for 10 to 16 h. Cells for experiments were centrifuged for 1.5 min at 10,000 × g from precultures and resuspended in M9 medium without a C source. For growth experiments with SDS or SLES, C-source volumes of 75 ml of M9 medium in 500 ml in Erlenmeyer flasks without baffles were used. Main cultures were inoculated with cells from precultures to an OD600 of 0.01 and cultivated at 30°C and 200 rpm. Immediately after inoculation and at several time points thereafter, samples from the cultures, avoiding the macroscopic cell aggregates as described previously (5), were taken to measure the OD600 and quantify the substrate concentrations. Samples for substrate quantification were centrifuged at 18,500 × g for 10 min at room temperature. Supernatants were transferred to new plastic tubes and stored at –20°C.
TABLE 1
| Strain or plasmid | Relevant characteristicsa | Source or reference |
|---|---|---|
| Strains | ||
| Pseudomonas aeruginosa | ||
| PAO1 | PAO1 Nottingham wild-type strain | Holloway collection |
| PAO1 ΔsdsB1 | sdsB1 deletion mutant | This study |
| PAO1 ΔsdsB1/pUCP18[sdsB1] | sdsB1 deletion mutant complementation of sdsB1 | This study |
| PAO1 ΔsdsB1/pUCP18 | sdsB1 deletion mutant pUCP18 (empty vector control) | This study |
| Escherichia coli | ||
| DH5α | recA1 endA1 hsdR17 thi-1 supE44 gyrA96 relA1 deoR | 55 |
| Δ(lacZYA-argF)U196 (ϕ80lacZΔM15) | ||
| ST18 | pro thi hsdR+ Tpr Smr; chromosome::RP4-2 Tc::Mu-Kan::Tn7/λpir ΔhemA | 56 |
| C41(DE3) | F– ompT gal dcm hsdSB[rB– mB–] | 57 |
| C41(DE3)/pET23a[his-sdsB1] | Overexpression of his-sdsB1 | This study |
| C41/pET23a[laoR] | Overexpression of laoR | This study |
| Plasmids | ||
| pEX18Ap | Gene replacement vector; Apr, sacB | 58 |
| pUCP18 | Escherichia-Pseudomonas shuttle vector; Apr | 59 |
| pBBR1MCS2Y[lacZ] | pBBR1MCS2 vector backbone modified as reporter gene vector with lacZ (without promotor); Apr | M. Czieborowski et al., unpublished data |
| pET23a | Overexpression vector; Apr, C-terminal His tag | Merck Millipore |
| pET23a[his-sdsB1] | Vector with his-sdsB1 for purification | This study |
| pET23a[laoR] | Vector with laoR for purification | This study |
| pUCP18[sdsB1] | Vector for sdsB1 complementation | This study |
| pBBR1MCS2[PsdsA1-lacZ] | Gene fusion of intergenic sdsB1/sdsA1 region with lacZ | This study |
| pBBR1MCS2[PlaoCBA-lacZ] | Gene fusion of intergenic laoR/laoCBA region with lacZ | This study |
a
Apr, ampicillin resistance; Smr, streptomycin resistance; Tpr, trimethoprim resistance.
Construction of unmarked P. aeruginosa deletion mutant and complementation/overexpression/reporter gene plasmids.
The PAO1 ΔsdsB1 deletion strain was generated with primers 1 to 4 (Table 2) as already described (14). Selected mutants were checked for correct deletion by colony PCR amplifying the region of interest with primers 5 and 6 (Table 2). For complementation, the gene of interest was amplified from the genomic DNA of strain PAO1 (primers 5 and 6, Table 2) and cloned into the vector pUCP18 (Table 1) into the XbaI and HindIII restriction sites. For overexpression of sdsB1 and laoR, the genes were amplified with primers 7 to 10 (Table 2) from the genomic DNA of strain PAO1 and cloned into the NdeI and HindIII restriction sites into vector pET23a (Table 1). For reporter gene assay of the putative promoter regions of sdsA1 and laoCBA, the respective intergenic regions were amplified with primers 11 to 14 (Table 2) from genomic DNA of strain PAO1 and cloned into the BamHI and XhoI restriction sites into vector pBBR1MCS2::lacZ (Table 1). Plasmids for complementation, overexpression, or reporter genes (Table 1) were transferred into the respective strains by transformation, and the strains were checked by colony PCR amplifying the multiple cloning site of the vector (primers 15 to 20, Table 2). All plasmids were verified by DNA sequencing prior to transformation (Microsynth Seqlab GmbH, Göttingen, Germany).
TABLE 2
| No. | Oligonucleotide | Sequencea |
|---|---|---|
| 1 | sdsB1_Up_fw | tatatctagaCCGCTGTCGCCGCGGATGAAG |
| 2 | sdsB1_Up_rev | GCGGACTCCGCATCGAATG |
| 3 | sdsB1_Dn_fw | CATTCGATGCGGAGTCCGCGGCTCAGCGCCAGCGAGCG |
| 4 | sdsB1_Dn_rev | tataaagcttGTCAACCTGGCCTACCCGC |
| 5 | sdsB1_fw | tatatctagaATGAGCGATCTACGCCAGTTC |
| 6 | sdsB1_rev | tataaagcttTCAGGTGGCGGCGAAGC |
| 7 | his-sdsB1_fw | tatacatatgcaccaccaccaccaccacAGCGATCTACGCCAGTTC |
| 8 | his-sdsB1_rev | tataaagcttTCAGGTGGCGGCGAAG |
| 9 | laoR-his_fw | tatacatatgGCACCACGCATCAAGACCC |
| 10 | laoR-his_rev | tataaagcttGAGCGGCACGTAGAGCCGC |
| 11 | PsdsA1_fw | tataggatccGCGGACTCCGCATCGAATGTG |
| 12 | PsdsA1_rev | tatactcgagCGGATACTCCGTTTTATTGTTGTGGG |
| 13 | PlaoCBA_fw | tataggatccGGGGACGACGGGCTCGTTCG |
| 14 | PlaoCBA_rev | tatactcgagGGGCTGGCTCCTCGGCGAAG |
| 15 | pUCP18_fw | GAGCGGATAACAATTTCACACAGG |
| 16 | pUCP18_rev | AGGGTTTTCCCAGTCACGACGTT |
| 17 | pET23a_fw | GGAGACCACAACGGTTTCCC |
| 18 | pET23a_rev | TTGTTAGCAGCCGGATCTCA |
| 19 | pBBR1_fw | TCACACTGCTTCCGGTAGTC |
| 20 | lacZ_rev | TCCGTGGGAACAAACGGCGG |
| 21 | SdsB1_A_fw | GCGGACTCCGCATCGAATG |
| 22 | SdsB1_A_rev | CGGATACTCCGTTTTATTG |
| 23 | SdsB1_B_fw | GCGGACTCCGCATCGAATGTGGACCTGGAATGCATTGGAA |
| 24 | SdsB1_B_rev | TTCCAATGCATTCCAGGTCCACATTCGATGCGGAGTCCGC |
| 25 | SdsB1_C_fw | TTATTGCCGTGCCGCTCGGCATAGTGCCAGCCGAAGCCCA |
| 26 | SdsB1_C_rev | TGGGCTTCGGCTGGCACTATGCCGAGCGGCACGGCAATAA |
| 27 | SdsB1_D_fw | CAACAATAAAACGGAGTATCCG |
| 28 | SdsB1_D_rev | CGGATACTCCGTTTTATTGTTG |
| 29 | SdsB1_B*_fw | GCGGACTCCATGCCGAATGTGGACCTGGAATGCATTGGAA |
| 30 | SdsB1_B*_rev | TTCCAATGCATTCCAGGTCCACATTCGGCATGGAGTCCGC |
| 31 | SdsB1_B**_fw | GCGGACTCCGCATCGGGCATGGACCTGGAATGCATTGGAA |
| 32 | SdsB1_B**_rev | TTCCAATGCATTCCAGGTCCATGCCCGATGCGGAGTCCGC |
| 33 | SdsB1_B***_fw | GCGGACTCCATGCCGGGCATGGACCTGGAATGCATTGGAA |
| 34 | SdsB1_B***_rev | TTCCAATGCATTCCAGGTCCATGCCCGGCATGGAGTCCGC |
| 35 | LaoR_A_fw | tataGGGCTGGCTCCTCGGCGAAG |
| 36 | LaoR_A_rev | tataGGGCTCGTTCGGGTAACAGT |
| 37 | LaoR_B_fw | tataGGGCTGGCTCCTCGGCGAAG |
| 38 | LaoR_B_rev | tataAATCCAGCGACGCAGCGAAA |
| 39 | LaoR_C_fw | tataTCTAGAGTAAATACTCTAAAG |
| 40 | LaoR_C_rev | tataGACCGGCCCGCCTTTCGTTG |
| 41 | LaoR_C1_fw | TTTCGCTGCGTCGCTGGATTTCTAGAGTAAATACTCTAAAGTGTCAAACA |
| 42 | LaoR_C1_rev | TGTTTGACACTTTAGAGTATTTACTCTAGAAATCCAGCGACGCAGCGAAA |
| 43 | LaoR_C1*_fw | TTTCGCTGCGTCGCTGGATTTCTAGAGTAAACGTCTCGAAGTGTCAAACA |
| 44 | LaoR_C1*_rev | TGTTTGACACTTCGAGACGTTTACTCTAGAAATCCAGCGACGCAGCGAAA |
| 45 | LaoR_C1**_fw | TTTCGCTGCGTCGCTGGATTTCCGAGACGAACGTCTCGAAGTGTCAAACA |
| 46 | LaoR_C1**_rev | TGTTTGACACTTCGAGACGTTCGTCTCGGAAATCCAGCGACGCAGCGAAA |
| 47 | LaoR_C2_fw | GCATGCCTGCCAGCCCGGCGCTTGCGTACCCAACGAAAGGCGGGCCGGTC |
| 48 | LaoR_C2_rev | GACCGGCCCGCCTTTCGTTGGGTACGCAAGCGCCGGGCTGGCAGGCATGC |
| 49 | LaoR_C_fw_JOE | JOE-TTTCGCTGCGTCGCTGGATTTCTAGAGTAAATACTCTAAAGTGTCAAACA |
| 50 | Lao_operon_fw | GGTCGTCGTCCAGGCGCGAG |
| 51 | Lao_operon_rev | GGCCAAGGGCGTGTTCAGCAAG |
| 52 | qRT_rpoS_fw | CTCCCCGGGCAACTCCAAAAG |
| 53 | qRT_rpoS_rev | CGATCATCCGCTTCCGACCAG |
| 54 | qRT_sdsB1_fw | CACGCGTGCGCGTCGTACTG |
| 55 | qRT_sdsB1_rev | GGATGTCCGGGGCGGCAGAA |
| 56 | qRT_sdsA1_fw | CAATCCCAGCCTGCAACGCCA |
| 57 | qRT_sdsA1_rev | GTACGGATCGGCCTCTCGCCC |
| 58 | qRT_laoR_fw | CCTGCTGGAAAGCGACCCGGA |
| 59 | qRT_laoR_rev | GGAAGCGCACCCAGGAGGTGA |
| 60 | qRT_laoA_fw | GCGGCACCACCGCGGAAATC |
| 61 | qRT_laoA_rev | GGTGATGGCGCCGTCCTTGC |
a
Restriction sites or nucleotides complementary to a vector are underlined. Lowercase letters indicate bases that are not complementary to the respective genome or vector sequence. Nucleotide exchanges are indicated in boldface.
SDS and SLES quantification.
Quantifications of the anionic detergents SDS and SLES were performed by a modified “stains-all” assay (14) as previously described. Briefly, stains-all solution was supplied to culture supernatants, and quantification of the absorption at an 438-nm wavelength was measured using a UV-VIS spectrophotometer (UV-2600; Shimadzu, Kyoto, Japan). Absorption spectra recorded with a double-beam spectrophotometer revealed that SLES and SDS reacted similarly with the stains-all solution (45), and calibration curves with SLES revealed linearity within the range of 0 to 20 μM.
Purification of regulatory proteins.
E. coli C41(DE3) was transformed with either pET23a[laoR] or pET23a[his-sdsB1]. For the expression of laoR and sdsB1, the cells were incubated at 30°C in liquid LB containing 100 μg ml−1 ampicillin. At an OD600 of 0.4, the expression was induced by addition of 0.4 mM IPTG (isopropyl-β-d-thiogalactopyranoside), and the temperature was reduced to 20°C. After overnight incubation, the cells were harvested and disrupted using a French press (American Instrument Company, Silver Spring, MD). The soluble fraction was used for further purification steps. Affinity chromatography with a 1-ml HisTrap Fast Flow column (GE Healthcare, Chicago, IL) was performed to purify LaoR and SdsB1 containing C-terminal and N-terminal His6 tags, respectively. The column was equilibrated with binding buffer (100 mM Tris, 500 mM NaCl, 40 mM imidazole [pH 7.4]), and the soluble fraction was loaded into the column and washed with binding buffer. Elution was performed with buffer containing 500 mM imidazole. Protein concentrations were quantified by a bicinchoninic acid assay (Pierce/Thermo Scientific, Rockford, IL) with bovine serum albumin as the standard. Purification results were verified by SDS-PAGE as described previously (14).
Electrophoretic mobility shift assay.
Binding of SdsB1 and LaoR on the intergenic DNA sequences was analyzed using EMSAs. Putative binding regions were amplified by PCR with primers 21 and 22 and primers 35 to 40 (Table 2). DNA at 0.2 μM was incubated with 0.5 μM protein in a 10-μl total volume in binding buffer (1 mM EDTA, 10 mM Tris/HCl [pH 7.0], 80 mM NaCl, 10 mM β-mercaptoethanol, and 5% [wt/vol] glycerol) (46). For fine localization of the binding sequences, oligonucleotides 23 to 34 and 41 to 48 (Table 2) were annealed within a C1000 Touch thermal cycler (Bio-Rad, Hercules, CA) according to the following modified program: 2 min at 94°C, a rapid ramp down to 70°C at maximal cycler speed (3°C/s), followed by slow (0.1°C/min) cooling to 12°C (47). The obtained DNA fragments were tested at concentrations of 0.5 and 0.1 μM with 2.5 μM SdsB1 and 0.4 μM LaoR, respectively. For ligand analysis, 5′ JOE (6-carboxy-4′,5′-dichloro-2′,7′-dimenthoxyfluoresceine)-labeled oligonucleotides (Eurofins Genomics Germany GmbH, Ebersberg, Germany) (oligonucleotides 48 and 42 in Table 2) were used to achieve a more sensitive detection, and samples were prepared as described previously with the addition of the putative ligand. In general, after 30 min of incubation at room temperature, the samples were mixed with 2 μl of loading dye (25% [wt/vol] sucrose and 0.05% [wt/vol] bromophenol blue in 20 mM Tris/HCl [pH 8.0]) and separated in a native 8% (wt/vol) polyacrylamide gel containing TBE buffer (89 mM Tris, 89 mM boric acid, 2 mM Na2-EDTA) (46). Gels were run at 10 V cm−1 on ice in TBE buffer, and DNA was detected either with ethidium bromide or, in the case of labeled DNA visualization, with “Alexa488” since an excitation/emission method at a ChemiDoc MP imaging system (Bio-Rad) was used.
Thermal shift assay.
To investigate the thermal stability of the regulator SdsB1 or LaoR alone and in the presence of DNA fragments containing the putative binding site or not, thermal shift assays using a Sypro Orange dye (50×; Sigma-Aldrich, St. Louis, MO) were performed (48, 49). Briefly, 20-μl reaction mixtures containing 2 μM regulator protein, 1× Sypro Orange dye, 1 μM DNA fragment, and 1× binding buffer from EMSAs (as described above) were heated from 4 to 99°C in 0.5°C steps using a CFX96 Touch real-time PCR detection system (Bio-Rad). The fluorescence intensity was measured at excitation/emission wavelengths of 490/575 nm. The midpoint temperature of transition between folded and unfolded states, also known as the melting temperature (Tm), was calculated by using CFX Manager software (Bio-Rad).
β-Galactosidase assay.
To analyze β-galactosidase expression from the putative sdsA1 and the lao gene promoter, precultures of strain PAO1 harboring the respective reporter gene lacZ fusion plasmids (Table 1) were incubated in 3 ml of M9 medium with 10 mM succinate containing 200 μg ml−1 carbenicillin for 6 to 7 h at 30°C and 200 rpm. Main cultures containing 50 ml of fresh M9 medium with 10 mM succinate and 200 μg ml−1 carbenicillin were inoculated to an OD600 of 0.001, followed by incubation for 10 h under the same conditions. The cells were harvested in their exponential phase and washed with M9 medium without a carbon source, and cell suspensions at an OD600 of 0.3 to 0.4 were prepared with M9 medium without a carbon source. A 1-ml portion of cell suspension in a 1.5-ml reaction vessel was then supplied with 1 mM putative inducer substrate and incubated for 3 h at 30°C and 200 rpm. The OD600 values of the culture samples were measured, and the cells were then treated as previously described (50). Samples with a volume of 200 μl were resuspended in 800 μl of buffer Z (50). After the addition of 50 μl of SDS (0.1% [wt/vol]) and 25 μl of chloroform, the samples were vortexed and incubated on ice for 20 min. After incubation at 30°C for 5 min, 200 μl of o-nitrophenyl-β-d-galactopyranoside (ONPG; 4 mg ml−1, freshly prepared) was added. After 5 to 30 min of incubation, the reaction was stopped by the addition of 500 μl of 1 M Na2CO. The samples were then centrifuged at maximum speed for 10 min before the absorption at 405 nm was measured. The β-galactosidase activities were calculated as described previously (50).
Reverse transcriptase PCR.
For reverse transcriptase PCR assays, RNA was extracted from 3 ml of exponentially growing P. aeruginosa strain PAO1 cultures with 3.5 mM 1-dodecanol or 10 mM succinate as the substrate. Cells were mixed with 1 ml of RNAlater stabilization solution (Thermo Scientific, Waltham, MA), and RNA was extracted using a PureLink RNA minikit (Thermo Scientific) according to the manufacturer’s protocol. RNA was then treated with a Turbo DNA-Free kit (Invitrogen, Waltham, MA), and a PCR analysis with the primer pair 44/45 (Table 2) was performed with wild-type DNA as a positive control to check for contaminating chromosomal DNA. The RNA amounts were then adjusted and utilized for cDNA synthesis using a RevertAid H Minus first-strand cDNA synthesis kit (Thermo Scientific) with random hexamer primers according to the manufacturer’s instructions. Reverse transcriptase PCR was then performed with 5 μl of cDNA and Q5 polymerase and the primer pair 50/51 (Table 2) for a transcript spanning laoABC (see Fig. 5A). As a positive control, 20 ng of chromosomal DNA was utilized. After 30 s of denaturation at 98°C, the following reaction regimen was used for 30 cycles: 98°C for 10 s, 72°C for 15 s, and 72°C for 50 s. The PCR products were detected on a 1% (wt/vol) agarose gel and stained with ethidium bromide. Experiments were performed in biological triplicates.
Quantitative real-time PCR.
For quantitative real-time PCR (qRT-PCR) analysis, 3- ml portions of exponentially growing P. aeruginosa strain PAO1 cultures with 10 mM succinate or 3.5 mM 1-dodecanol as the substrate were extracted. As cells grown with 3.5 mM SDS or SLES formed macroscopic cell aggregates (6), 4-ml portions of nonaggregating, freely suspended cells from exponentially growing strain PAO1 cultures were extracted when these sulfate ester detergents were used as the substrates. RNA extraction was performed as described for reverse transcriptase PCR. RNA (500 ng) was treated with DNase I (using a Turbo DNA-Free kit) according to the manufacturer’s instructions, and DNA contamination was checked by quantitative real-time PCR using 40 ng of RNA and the reference gene primer pair, as described above. Synthesis of cDNA was performed as described for reverse transcriptase PCR. Gene-specific primers were designed using Primer-BLAST software (NCBI Primer-BLAST) (51) to avoid nonspecific amplification of P. aeruginosa DNA. The well-established reference gene primer pair for rpoS was used (52, 53) (Table 2). For every sample, an adjusted amount of cDNA (50 ng) was amplified with gene-specific primers 54 to 61 (Table 2) and the iTaq Universal SYBR green Supermix (Bio-Rad) within the CFX96 Touch real-time PCR detection system (Bio-Rad). After 3 min of denaturation at 95°C, the following reaction cycle was used for 30 cycles: 95°C for 10 s, 60°C for 5 s, and 72°C for 10 s with data acquisition. Melting-curve analysis was performed in the range from 55 to 95°C at 0.5°C intervals to confirm specific amplification of single PCR products. Cycle threshold (CT) values were generated by using CFX Manager software (Bio-Rad) and used to quantify the relative levels of gene expression by the 2–ΔΔCT method with the calculated reaction efficiencies of the respective primer pair and the rpoS gene as an internal control (54).
ACKNOWLEDGMENTS
We thank Karin Niermann for experimental support and Michael Czieborowski for providing the reporter gene vector as well as for support in the preparation of some of the figures.
This study was funded by grants from the German Federal Ministry for Economic Affairs and Energy (BMWi) through its Central Innovation Program for SMEs (ZIM) and by the German Research Foundation (DFG) to B.P. (grants 16KNO13527 PakuNaS and PH71/3-2, respectively).
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Information & Contributors
Information
Published In
Applied and Environmental Microbiology
Volume 85 • Number 23 • 1 December 2019
eLocator: e01352-19
Editor: M. Julia Pettinari, University of Buenos Aires
PubMed: 31540990
Copyright
© 2019 American Society for Microbiology. All Rights Reserved.
History
Received: 14 June 2019
Accepted: 14 September 2019
Published online: 14 November 2019
Keywords
Contributors
Editor
M. Julia Pettinari
Editor
University of Buenos Aires
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Sulfate Ester Detergent Degradation in Pseudomonas aeruginosa Is Subject to both Positive and Negative Regulation. Appl Environ Microbiol
85:e01352-19.
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