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Journal of Bacteriology, May 2004, p. 2735-2744, Vol. 186, No. 9
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.9.2735-2744.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Involvement of Error-Prone DNA Polymerase IV in Stationary-Phase Mutagenesis in Pseudomonas putida

Radi Tegova, Andres Tover, Kairi Tarassova, Mariliis Tark, and Maia Kivisaar*

Department of Genetics, Institute of Molecular and Cell Biology, Tartu University and Estonian Biocentre, 51010 Tartu, Estonia

Received 8 December 2003/ Accepted 23 January 2004


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In this work we studied involvement of DNA polymerase IV (Pol IV) (encoded by the dinB gene) in stationary-phase mutagenesis in Pseudomonas putida. For this purpose we constructed a novel set of assay systems that allowed detection of different types of mutations (e.g., 1-bp deletions and different base substitutions) separately. A significant effect of Pol IV became apparent when the frequency of accumulation of 1-bp deletion mutations was compared in the P. putida wild-type strain and its Pol IV-defective dinB knockout derivative. Pol IV-dependent mutagenesis caused a remarkable increase (approximately 10-fold) in the frequency of accumulation of 1-bp deletion mutations on selective plates in wild-type P. putida populations starved for more than 1 week. No effect of Pol IV on the frequency of accumulation of base substitution mutations in starving P. putida cells was observed. The occurrence of 1-bp deletions in P. putida cells did not require a functional RecA protein. RecA independence of Pol IV-associated mutagenesis was also supported by data showing that transcription from the promoter of the P. putida dinB gene was not significantly influenced by the DNA damage-inducing agent mitomycin C. Therefore, we hypothesize that mechanisms different from the classical RecA-dependent SOS response could elevate Pol IV-dependent mutagenesis in starving P. putida cells.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
During the past several years our understanding of mutation mechanisms has been expanded by the discovery of a new superfamily of DNA polymerases, called the Y family (46). The Y-family polymerases have been identified in prokaryotes, archaea, and eukaryotes. Members of this superfamily are devoid of 3'->5' proofreading exonuclease activity and replicate undamaged DNA with low fidelity and low processivity; many of these enzymes can bypass DNA lesions that block chain elongation by replicative DNA polymerases (21-23). According to the concept of specialized polymerases some of these polymerases are able to copy cognate lesions with high genetic fidelity (22). On the other hand, the specialized DNA polymerases are involved in mutation processes when copying noncognate DNA lesions or normal DNA.

In a growth-restricting environment (e.g., during starvation), mutants arise that are able to take over bacterial populations by a process known as stationary-phase mutation (15). One widely discussed idea is that genetic adaptation of microbial populations under environmental stress might be accelerated by stress-induced activation of error-prone DNA polymerases (see, for example, references 16, 50, and 63). In Escherichia coli, two error-prone DNA polymerases, Pol V (UmuD'C) and Pol IV (DinB), and one high-fidelity DNA polymerase, Pol II, are upregulated during the SOS response (23). SOS induction has also been shown to occur spontaneously in static bacterial populations (62). It has been recently demonstrated that error-prone DNA polymerases Pol IV and Pol V are involved in stationary-phase mutagenesis in E. coli (4, 7, 42). The involvement of SOS-induced polymerases (Pol II, Pol IV, and Pol V) in stationary-phase mutagenesis has also been shown in the appearance of GASP (growth advantage in stationary phase) mutants of E. coli (70). E. coli mutants lacking one or more of these polymerases suffered considerable fitness reduction when competing with a wild-type strain under starvation conditions (70).

Although DinB-like polymerases have been identified in essentially all prokaryotic and eukaryotic organisms studied (46), with only a few exceptions (e.g., see reference 61), involvement of Pol IV homologues in stationary-phase mutagenesis in bacteria other than E. coli has not been investigated. Knowledge about mechanisms of Pol IV-dependent stationary-phase mutagenesis obtained using E. coli-based test systems includes studies of the 1-bp deletions occurring in an F' plasmid (see, for example, references 38 and 42) and in the bacterial chromosome (7). In these cases, another copy of the dinB gene was present in the F' plasmid and Pol IV-dependent stationary-phase mutations required the RecA protein. The genus Pseudomonas represents one of the most diverse and ecologically widely distributed groups of bacteria (58). Here, we studied the role of Pol IV in mutagenesis in Pseudomonas putida. We show that Pol IV is involved in generation of 1-bp deletions in starving cells. The occurrence of other types of stationary-phase mutations (e.g., base substitutions) does not require the presence of Pol IV. Pol IV-dependent mutagenesis in P. putida appears to be a RecA-independent process. Moreover, our results demonstrate that involvement of Pol IV in stationary-phase mutagenesis becomes essential only in long-term-starved populations of P. putida, which indicates that Pol IV-dependent mutagenesis can be induced by a so-far uncharacterized mechanism in cells that have suffered long-term starvation stress.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, plasmids, and media. The bacterial strains and plasmids used in this study are described in Table 1. Complete medium was Luria-Bertani (LB) medium (43), and minimal medium was M9 (1). Phenol minimal plates with 1.5% Difco agar contained 2.5 mM phenol as a sole carbon and energy source. Antibiotics were added at the following final concentrations: for E. coli, ampicillin at 100 µg/ml and tetracycline at 10 µg/ml; for P. putida, carbenicillin at 1,000 to 3,000 µg/ml, chloramphenicol at 1,500 to 3,000 µg/ml, and tetracycline at 80 µg/ml; for both organisms, kanamycin at 50 µg/ml and rifampin at 100 µg/ml. E. coli was incubated at 37°C, and P. putida was incubated at 30°C. E. coli was transformed with plasmid DNA as described by Hanahan (24). P. putida was electrotransformed as described by Sharma and Schimke (54). E. coli strain TG1 or DH5{alpha} was used for the DNA cloning procedures.


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  		TABLE 1. Bacterial strains and plasmids used in this study

 
Construction of test systems to study stationary-phase mutations in P. putida. An assay system to test 1-bp deletions (Fig. 1) was constructed by altering the phenol monooxygenase gene (pheA) coding sequence. Insertion of a single A nucleotide 5' to the ACC codon for Thr-56 was performed by PCR amplification of the segment of the pheA gene from plasmid pPU1930 (45) with primer pheAup (5'-AAGGCGCTCCCGTAAGACA-3'), complementary to the sequence at nucleotides –40 to –22 relative to the coding sequence of the pheA gene, and the mutant primer pheA56+A (5'-GGTGATCATAATGTTGCTAATGCCCTGGGTTCGACAGGAACATTGCTGCG-3'), complementary to the pheA coding sequence at nucleotides 147 to 195. The amplified DNA fragment was subcloned into the pBluescript KS(+) EcoRV site to obtain pKSpheA56+A. In addition to the generation of frameshift in the pheA coding sequence, this nucleotide insertion eliminated the SalI restriction site. The mutation was verified by DNA sequencing. The mutated DNA segment was thereafter inserted as the XbaI- and BclI-generated fragment from pKSpheA56+A (the XbaI site was provided by the pBluescript KS multicloning site) into pPU1930 by replacing the original pheA sequence located between the XbaI (the XbaI site is present upstream of the coding sequence of the pheA gene in pPU1930) and BclI sites to generate pPUpheA56+A. A pPU1930 derivative carrying the mutated sequence was first identified by the absence of the SalI restriction site in the pheA sequence. Finally, we cloned the SacI- and PvuII-generated fragment from pPUpheA56+A containing the constitutively expressed promoter and the mutated pheA gene into the broad-host-range plasmid pKT240 (2) to obtain pKTpheA56+A.



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  		FIG. 1. Assay systems allowing measurement of 1-bp deletions (plasmid pKTpheA56+A) or different base substitutions (plasmids pKTpheA22TAG, pKTpheA22TAA, and pKTpheA22TGA) in P. putida stationary-phase cells. Segments of the original sequences and their mutant variants are shown. The test systems were constructed by altering the phenol monooxygenase gene (pheA) coding sequence. The +1 frameshift was constructed in the phenol monooxygenase gene pheA by inserting the A nucleotide (marked in bold) into the ACC codon for Thr-56 (underlined in the original sequence). Assay systems for the isolation of base substitutions were constructed by replacement of the CTG codon for Leu-22 (underlined in the original sequence) with a TGA, TAA, or TAG stop codon (indicated in boldface). Transcription from the pheA gene is initiated from the constitutively expressed promoter (45), indicated by P.

 
Assay systems allowing detection of base substitutions (Fig. 1) were constructed by using a two-step PCR amplification strategy. Mutant oligonucleotides contained specific base substitutions that replaced the CTG codon for Leu-22 in the pheA gene with a TGA, TAA, or TAG stop codon. In the first step, the PCR with oligonucleotides pheAup and pheA22TGA (5'-CGCTTCACCCATCATCAAAAATGACGCTA-3'), pheA22TAA (5'GCGCTTTACCCATCATCAAAAATGACGCTA-3'), or pheA22TAG (5'-GCGCTCTACCCATCATCAAAAATGACGCTA-3'), complementary to the positions 42 to 71 relative to the coding sequence of the pheA gene, was carried out. ExoI treatment followed, and the PCR products were purified and used in a second PCR with the oligonucleotide pheAts (5'-GTTCATGGGGGACTGCTTC-3'), complementary to pheA nucleotides 295 to 313. The amplified DNA fragments were cloned into the EcoRV site of pBluescript KS (+), and the mutations were verified by DNA sequencing. The next steps in the cloning strategy were identical to those used for the construction of the frameshift assay plasmid pKTpheA56+A. Finally, plasmids pKTpheA22TGA, pKTpheA22TAA, and pKTpheA22TAG were obtained.

Construction of P. putida dinB and recA knockout mutants. The dinB and recA gene sequences of P. putida KT2440 were obtained from The Institute for Genomic Research website (http://www.tigr.org). The dinB gene was amplified by PCR from genomic DNA of P. putida PaW85, which is isogenic to P. putida strain KT2440. Two primers, dinBFw (5'-GGCCTTTTCTTGAATCTGGTTGCG-3'), complementary to the sequence –518 to –496 upstream of the ATG initiator codon, and dinBRev (5'-GCGGATCCAGGCGTGCATTATTAG-3'), complementary to the sequence 24 to 47 nucleotides downstream of the TGA stop codon of the dinB gene, were used for PCR amplification. The amplified DNA fragment containing the dinB gene was subcloned into EcoRV-cleaved pBluescript KS (+), to obtain pKSdinB. The EcoRI- and Van9I-generated DNA fragment containing the Tetr gene from pBR322 was inserted into the Van91-cleaved dinB gene. The resulting dinB::tet sequence from pKSdinB::tet was inserted into plasmid pGP704 L (48) by using Acc65I and XbaI sites. Plasmid pGP704dinB::tet was selected in E. coli strain CC118 {lambda}pir (30). The interrupted dinB gene was inserted into the chromosome of P. putida PaW85 by homologous recombination. Plasmid pGP704dinB::tet, not able to replicate in hosts other than E. coli CC118 {lambda}pir, was conjugatively transferred into P. putida PaW85 by using a helper plasmid, pRK2013 (13). The PaW85 dinB::tet knockout strain was verified by PCR analysis.

For the construction of a recA knockout mutant of P. putida, the genomic DNA of PaW85 was amplified with oligonucleotides RecAFw (5'-AACAAGAAGCGCGCCTTGGTC-3') and RecARev (5'-ATCAGCTTCAGCAGCAGCAGCTT-3'), complementary to the sequences –66 to –45 upstream of the ATG initiator codon and 37 to 60 nucleotides 5' to the stop codon of the recA gene, respectively. The amplified DNA fragment was subcloned into EcoRV-cleaved pBluescript KS (+) to obtain pKSrecA. The EcoRI- and Van9I-cleaved DNA fragment containing the Tetr gene from pBR322 was blunt ended and inserted into the BclI-cleaved recA gene (the BclI ends were blunt ended before the ligation as well), resulting in pKSrecA::tet. Finally, the DNA fragment containing the recA::tet sequence was inserted with XbaI and Ecl136II ends into pGP704 L to obtain pGP704recA::tet. The wild-type recA sequence of P. putida was replaced with the interrupted recA::tet sequence by homologous recombination using a procedure similar to that described above. The knockout mutant was verified by PCR analysis. Additionally, a UV sensitivity test (43) was performed to verify the RecA-defective strain PaW85 recA::tet.

DNA sequence analysis. In Phe+ mutants, an approximately 350-bp DNA region covering the area of the pheA gene containing potential reversion mutations was analyzed by DNA sequencing. The DNA segment containing this region was amplified by PCR using the primers pheAup and pheAts. The nucleotide sequences were determined using the DYEnamic ET Terminator Cycle Sequencing kit (Amersham Pharmacia Biotech, Inc). The oligonucleotides used in sequencing of the mutant DNA region were the same as those used in PCRs. The DNA sequencing reactions were analyzed with an ABI Prism 377 DNA Sequencer (Perkin-Elmer).

Overexpression of the P. putida dinB gene. The dinB gene lacking any regulatory sequences upstream of its ATG initiator codon was obtained by PCR amplification of P. putida PaW85 chromosomal DNA with primers dinBNde (5'-CATATGTCCTTGCGCAAGATCATCCA-3'), complementary to the nucleotides –3 to 23 relative to the dinB ATG initiator codon, and dinBRev. The amplified DNA fragment was cloned into the pBluescript KS (+) EcoRV site. We chose the construct carrying the dinB gene in the pBluescript vector in the opposite direction relative to the transcription direction from the lac promoter and named pKSATGdinB. By using the PstI and Acc65I restriction sites present in the pBluescript multicloning site, the dinB gene was further subcloned into the pBRlacItac plasmid (47) under the control of the Ptac promoter and lacIq repressor to generate plasmid pBRlacItacdinB. In order to overexpress the dinB gene in P. putida, the dinB expression cassette was finally inserted as the NheI- and Acc65I-generated DNA fragment into the XbaI- and Acc65I-cleaved single-copy broad-host-range plasmid pPR9TT (52) to obtain p9TTlacItacdinB. To avoid a leaky transcription of the dinB gene from the Ptac promoter, which is deleterious to P. putida cells, plasmid pKTlacI carrying an additional copy of the lacI gene was constructed by inserting the DNA fragment containing the lacI gene from pREP4 (QIAGEN) with Eco47III- and SmaI-generated ends into the Ecl136II site of the pKT240 vector. The dinB overexpression studies with E. coli were performed with cells carrying plasmid pBRlacItacdinB; P. putida PaW85 carried two plasmids, p9TTlacItacdinB and pKTlacI. The expression level of dinB was artificially increased by addition of 1 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) to the growth medium of 1.5-ml separate cultures of exponentially growing bacteria. After 6 h of cultivation of bacteria, 0.1-ml samples were taken from the cultures and spread onto LB plates containing 100 µg of rifampin/ml. Colonies were counted on plates incubated for 24 h. The control experiments were performed using bacteria carrying the same vector plasmids lacking the dinB gene. The frequency of mutation to Rifr per 109 cells was calculated for at least 35 independent cultures.

Studies of transcription from the dinB promoter. A DNA fragment containing a putative dinB promoter region was amplified by PCR with oligonucleotides dinBBamHI (5'-CCTTGGATCCAAGCTTTTTAACGGGCAAAGAAA-3') and dinBXho (5'-ACGCTGCGATCGAGATGCGCTCGAGAAG-3'), complementary to the nucleotides at positions –328 to –350 and 40 to 58, respectively, relative to the ATG initiator codon of the dinB gene. The amplified DNA fragment was cut with BamHI and XhoI and inserted into BamHI- and XhoI-cleaved pBluescript KS (+) to obtain pKSPdinB. To construct the luxAB reporter plasmid replicating in P. putida, the promoterless luxAB genes were subcloned within the BamHI-cleaved DNA fragment from pGP704 L into pBluescript KS (+) to obtain pKSluxAB. Then, by using the SacI and XhoI restriction sites provided by the pBluescript vector multicloning sites, the luxAB genes were inserted into SacI- and XhoI-cleaved pKT240. The resulting plasmid, named pKTluxAB, was used as a multicopy promoter-probe vector for study of the transcription from the dinB promoter. The putative promoter region of the dinB gene was cloned from plasmid pKSPdinB into pKTluxAB, using restrictases SmaI and Acc65I. The resulting plasmid was named pKTPdinBluxAB. In order to study the regulation of transcription from a single copy of the dinB promoter, the PdinB-luxAB expression cassette was subcloned from pKTPdinBluxAB into the NotI site of pPR9TT to obtain p9TTPdinBluxAB.

P. putida cells carrying pKTPdinBluxAB were grown exponentially in LB medium. In order to study whether DNA damage could induce transcription from the dinB promoter, mitomycin C (at a final concentration of 2 µg/ml) was added to cultures. Samples for luciferase assay were taken from P. putida exponential cultures grown in the presence or absence of mitomycin C. In all cases at least five independent measurements were made. The luciferase assay was performed as follows: 990 µl of phosphate buffer (100 mM Na2HPO4/KH2PO4 [pH 7]) and 10 µl of decanal (5 mM decanal in ethanol) were mixed in a test tube; 10 µl of bacterial culture was then added, and light emission was measured after 5 min of incubation with a luminometer (TD-20/20; Turner Designs).

To study the effect of the growth phase of bacteria on transcription from the dinB promoter, the promoter was subcloned from pKSPdinB into the promoter-probe vector pKTlacZ, using restrictases BamHI and XhoI, to obtain plasmid pKTPdinBlacZ. ß-Galactosidase (ß-Gal) activities in P. putida cells carrying pKTPdinBlacZ were measured by a modification of the standard protocol of Miller (43) as specified previously (64). Bacteria were grown in LB medium.

To map the transcription initiation site of the P. putida dinB gene, a reverse transcriptase reaction was carried out to identify the 5' end of the dinB-specific mRNA. Total RNA was isolated from exponentially growing cells of P. putida using the QIAGEN RNeasy total RNA kit. Ten micrograms of purified RNA was used as a template in primer extension reactions with 10 pmol of [{gamma}-32P]ATP-labeled primer dinBXho. DNA sequencing reactions were performed using a Sequenase version 2.0 kit (U.S. Biochemicals) and the same primer, and the reaction mixtures were loaded onto a sequencing gel as size markers. A dried gel was exposed to a PhosphorImager screen (Molecular Dynamics).

Isolation of Phe+ mutants. P. putida cells carrying different assay systems for the detection of Phe+ revertants were grown overnight in LB medium. Cells sampled from the culture were harvested by centrifugation and washed in M9 solution. Approximately 5 x 108 to 1 x 109 cells were spread onto phenol minimal plates. A few Phe+ colonies appearing on phenol minimal plates on day 2 contained mutations that occurred before the plating in a growing culture, whereas the colonies that emerged on selective plates on day 3 and later contained mutations that occurred after the cells were plated. To control whether the late-arising mutants could form a colony on phenol selective plates with a speed similar to that of those that emerged earlier, we performed reconstruction experiments. Plating of such mutants (approximately 100 cells) in the presence of 1 x 109 nonmutant cells onto phenol minimal plates demonstrated that all revertants tested were able to form visible Phe+ colonies on day 2 after plating of the mixed cultures. No more Phe+ colonies accumulated on these plates during the next days of incubation. This confirmed that the late revertants are truly stationary-phase mutants and that they are formed in P. putida populations after prolonged starvation. The pKT240-based plasmids carrying the test systems for detection of Phe+ mutations in P. putida are derivatives of the broad-host-range plasmid RSF1010 (2). We have previously shown (33) that the copy number of RSF1010-based plasmids is not affected by the growth phase of the bacteria.

Measurement of viability of Pol IV-defective and RecA-defective P. putida on phenol minimal plates. The viability of bacteria was determined on the same plates that were used for the isolation of Phe+ mutants. Using sterile 1-ml pipette tips, small plugs were cut out from the phenol-containing minimal plates, avoiding Phe+ colonies. The plugs were suspended in M9 solution by shaking for 10 min. Approximately 105-fold dilutions were plated onto LB plates, and the number of CFU was determined for at least five independent starving cultures.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
P. putida contains a DNA polymerase Pol IV homologue. Analysis of complete genome sequences of P. putida KT2440 revealed that the putative Pol IV homologue in this strain has 50% identity with the E. coli Pol IV sequence. According to the literature, Pol IV overproduction in E. coli growing cells causes hypermutation, including –1 frameshifts and some base substitutions (35, 67). To study whether overexpression of the P. putida dinB gene would also elevate the frequency of occurrence of spontaneous mutations in growing cells, we amplified the P. putida dinB (dinP) gene, encoding Pol IV, by PCR from chromosomal DNA of P. putida PaW85 (this strain is identical to KT2440) and cloned the gene into plasmid pBRlacItac (47) under the control of the Ptac promoter. We found that overexpression of the P. putida dinB gene in growing cells of E. coli carrying pBRlacItacdinB led to an approximately 24-fold increase in the frequency of appearance of Rif-resistant mutants (Table 2), which indicates that P. putida Pol IV is functional in E. coli.


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  		TABLE 2. Effect of P. putida dinB overexpression on frequency of mutations in growing cellsa

 
To study the effects of overexpression of Pol IV on the frequency of mutations in P. putida, the dinB gene placed under the control of the Ptac promoter and the lacIq repressor gene was inserted into a broad-host-range single-copy plasmid, pPR9TT (52). In contrast to E. coli, P. putida cells did not tolerate this construct, possibly due to a leakiness of the Ptac promoter. The colonies of p9TTlacItacdinB-carrying transformants that appeared on selective plates remained tiny, and cells picked up from these colonies were not further culturable. This indicated that artificial overexpression of the dinB gene in P. putida cells strongly inhibits growth of the bacteria and is probably lethal to the cells. To overcome the deleterious effect of leakiness of transcription of the dinB gene to P. putida cells, additional copies of the lacIq repressor gene, present in the pKT240-derivative pKTlacI, were introduced into P. putida carrying the dinB expression construct p9TTlacItacdinB. In this case, by manipulating the IPTG concentrations, we found that addition of IPTG at a 1 mM concentration (but not below or above that concentration) resulted in an increase in the frequency of appearance of Rif-resistant mutants in P. putida cells (Table 2).

Construction of novel test systems for the study of mechanisms of stationary-phase mutations in P. putida. P. putida PaW85 cells carrying the phenol monooxygenase gene, pheA, are able to grow on minimal medium containing phenol as the only carbon source (45). Selection of mutants able to grow on phenol minimal plates due to the activation of a silent pheA gene has been used by us to study stationary-phase mutagenesis in P. putida (33, 53). The assay system employed in our previous studies allowed isolation of different types of mutations, base substitutions, deletions, and insertions, which all led to the same outcome—generation of a functional promoter. In the current study, in order to study the effects of different genetic backgrounds on the frequency of different types of mutations separately, we constructed a novel set of assay systems. Details of the construction of test systems are presented in Materials and Methods. One of these systems measures reversion of +1 frameshift within the phenol monooxygenase gene, pheA (Fig. 1). Three other test systems, designed for the detection of base substitutions, contain different stop codons, TAG, TAA, or TGA, introduced into the same position (Leu-22) of the pheA coding sequence (Fig. 1).

To control whether the test systems constructed really detect the expected mutations, some of the Phe+ revertants accumulating on phenol-containing selective plates were subjected to DNA sequence analysis. Sequence analysis of Phe+ mutants emerging in P. putida carrying the assay system which should measure a reversion of +1 frameshift in plasmid pKTpheA56+A revealed that out of 15 mutants studied, 90% of frameshifts occurred at the neighboring site, in the CCC repeat flanking the inserted A nucleotide. Only 10% of the mutants contained an A nucleotide deletion. The –1 deletion at the CCC repeat restored the pheA reading frame by replacing the ACC codon (Thr) with the AAC codon (Asn).

Phe+ mutants isolated using the assay systems carrying different stop codons instead of the codon for Leu-22 in the pheA sequence were true revertants. No suppressors were isolated among approximately 100 mutants analyzed per test system. The absence of suppressor mutations was confirmed by the finding that all mutants analyzed contained base substitutions in the pheA gene which eliminated the stop codon by replacing it with codons for different amino acids (Table 3). Also, in the case of every revertant studied, the Phe+ phenotype was conjugatively transferred to the P. putida Phe– recipient strain and the transconjugants carrying different mutant plasmids grew equally well on phenol minimal plates. Similar growth characteristics of different Phe+ transconjugants on phenol minimal plates indicated that the amino acid Leu-22 in the PheA protein is replaceable with many other amino acids without changing the wild-type phenotype of this enzyme. Analysis of the Phe+ revertants which appeared due to base substitutions revealed different spectra of mutations, depending on the sequence of the stop codon used in the particular assay system (Table 2). The spectrum of changes that occurred in the case of the assay system carrying the TAA stop codon (plasmid pKTpheA22TAA) was most homogeneous: among 43 mutants analyzed, 88% contained a T-to-C transition and the rest (12%) had a T-to-G transversion. The T-to-C transition was also dominant in the case of mutants isolated using the assay systems carrying either the TGA or TAG codon (plasmids pKTpheA22TGA and pKTpheA22TAG, respectively), although several other DNA changes were identified as well (Table 3).


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  		TABLE 3. Reversion of different nonsense mutations (TGA, TAA, and TAG) in independent Phe+ mutants accumulating in starving cell populations of P. putida PaW85

 
Pol IV contributes to –1 deletions in starving cells of P. putida. In order to study possible involvement of the error-prone DNA polymerase Pol IV in occurrence of stationary-phase mutations in P. putida, we constructed a dinB knockout mutant of P. putida PaW85 (see Materials and Methods). Based on analysis of the sequence of the P. putida genome, the dinB gene does not belong to an operon of other genes. Therefore, possible negative effects of disruption of the dinB gene on transcription of other genes are excluded. Cells of wild-type P. putida PaW85 and its Pol IV-defective derivative PaW85 dinB::tet carrying different test systems (plasmid pKTpheA56+A, pKTpheA22TAA, pKTpheA22TGA, or pKTpheA22TAG) were plated onto phenol minimal plates containing phenol as the only carbon source. Results presented in Fig. 2A clearly demonstrate that the occurrence of 1-bp deletion mutations depends on the presence of the functional dinB gene in P. putida. Starting from day 9, the frequency of accumulation of Phe+ mutants on selective plates increased remarkably, and on day 11 and later the accumulation frequency of the Phe+ revertants in the P. putida wild-type strain was approximately 10 times higher than in the Pol IV-defective mutant. The lower frequency of accumulation of 1-bp deletion mutants in PaW85 dinB::tet could not be ascribed to a lower viability of dinB-defective cells under starvation conditions: during the 13 days studied, the Pol IV-defective strain survived as well as the wild-type strain (Fig. 2B). Thus, Pol IV-dependent mutagenesis could be elevated under conditions of long-term starvation of P. putida cells.



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  		FIG. 2. (A) Accumulation of Phe+ mutants on phenol minimal plates of P. putida wild-type strain PaW85 (WT) and in its dinB-defective (dinB) and recA-defective (recA) derivatives carrying plasmid pKTpheA56+A. About 5 x 108 P. putida cells were plated from overnight LB-grown cultures onto phenol minimal plates. Data for at least five parallel experiments are presented. Means ± standard deviations (error bars) for 10 plates calculated per 5 x 108 cells are shown. (B) Viability of P. putida wild-type strain PaW85 (WT) and in its dinB-defective (dinB) and recA-defective (recA) derivatives on phenol minimal plates. Means ± standard deviations for at least five cultures are shown. 1.0E + 08, for example, indicates 108 viable cells.

 
No effect of the dinB knockout became apparent if we measured the frequency of appearance of Phe+ mutants in starving cell populations of P. putida carrying test systems which allowed detection of base substitutions (data not shown). This indicated that Pol IV contributes only to a particular type of stationary-phase mutations in P. putida.

RecA is not required for Pol IV-dependent mutagenesis in P. putida. In E. coli, dinB is one of the genes induced in response to DNA damage (SOS response). After RecA-mediated cleavage of the LexA repressor, SOS regulon genes including dinB become upregulated (11, 38). DNA polymerase Pol IV-dependent stationary-phase mutations in the well-studied E. coli FC40 Lac+ reversion system require a functional RecA protein (7, 42). In order to study whether the occurrence of dinB-dependent stationary-phase mutations are affected by RecA functionality in P. putida, we constructed a RecA-defective mutant of P. putida strain PaW85 (see Materials and Methods). Our experiments showed that the rate of accumulation of 1-bp deletion mutants was similar in the wild-type P. putida and its RecA-deficient derivative PaW85 recA::tet (Fig. 2A). This demonstrated that the occurrence of Pol IV-dependent stationary-phase mutations observed in this study does not require the RecA protein.

Transcriptional analysis of the P. putida dinB promoter. In order to study transcription from the dinB promoter, the putative promoter region of the dinB gene was cloned upstream of the luxAB reporter into plasmid pKTluxAB. The resulting plasmid, pKTPdinBluxAB, expressed a high constitutive level of luciferase activity (Fig. 3). Mapping of the transcription initiation site by primer extension analysis localized the 5'end of the dinB-specific mRNA at the G nucleotide which is six nucleotides downstream from the putative promoter sequence, resembling the {sigma}70-type promoter consensus TTGACAN16-18TATAAT (Fig. 4). The sequence GTTTCA, resembling the –35 hexamer, and TACTAT, similar to the –10 hexamer, of this promoter were separated by a 17-nucleotide spacer. The P. putida dinB promoter region contains a sequence exhibiting similarity to the LexA binding consensus CTG-N10-CAG (69) (Fig. 4B). This sequence overlaps the –10 hexameric sequence of the dinB promoter by two nucleotides. Therefore, one may expect repression of this promoter by a LexA protein (the P. putida genome contains two different lexA gene homologues; see, e.g., http://www.tigr.org). However, despite the presence of putative LexA binding sites in the dinB promoter region, addition of the DNA damage-inducing agent mitomycin C to the growth medium of bacteria had only a minor increasing effect (up to twofold) on the level of transcription from the dinB promoter (Fig. 3). The similar up-to-twofold positive effect of addition of mitomycin C became evident when the level of transcription from the dinB promoter was monitored in a single-copy plasmid, p9TTPdinBluxAB (data not shown). The latter fact excludes the possibility that the cellular amount of LexA repressor might be too low to saturate all its binding sites when the dinB promoter is expressed in a medium-copy-number plasmid, pKTPdinBluxAB (approximately 10 copies per cell). Thus, transcription from the dinB promoter is poorly stimulated after DNA damage in P. putida cells.



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  		FIG. 3. Study of effects of DNA damage on transcription from the dinB promoter in P. putida PaW85. The promoter was cloned upstream of the reporter genes luxAB, encoding luciferase, and the expression of the transcriptional fusion was measured on pKT240-derived broad-host-range plasmid pKTPdinBluxAB. Transcription from the dinB promoter was assayed by measuring the luciferase activity (relative luciferase units/optical density unit at 580 nm) in cells grown in LB medium in the presence or absence of DNA-damaging agent mitomycin C (2 µg/ml). We determined that addition of higher concentrations of mitomycin C (up to 20 µg/ml) to the growth medium of the bacteria did not cause greater effects on transcription from the dinB promoter.

 


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  		FIG. 4. (A) Mapping of the transcription initiation site for the P. putida dinB promoter by reverse transcriptase. Lanes G, A, T, and C show DNA-sequencing reactions of the dinB promoter region. Lanes marked by – and + represent primer extension reactions carried out with total RNA isolated from a P. putida PaW85 plasmid-free strain and from the same strain carrying extra copies of the dinB promoter region in medium-copy-number plasmid pKTPdinBluxAB, respectively. The sequence of the dinB promoter region, including the –10 sequence of the promoter (boxed) and the transcription start point (indicated by an asterisk), are shown on the right. (B) Nucleotide sequence of the P. putida dinB promoter region. The putative –35 and –10 hexamers of the promoter are boxed, and the 5' end of the dinB mRNA is indicated by an asterisk. The potential LexA binding site is underlined (LexA binding consensus nucleotides [69] are in boldface). The translation start site of the dinB gene is indicated by an arrow.

 
Recently, Layton and Foster (38) demonstrated that in E. coli, the level of Pol IV is controlled by the stationary-phase sigma factor RpoS. RpoS is shown to also be a transcription regulator in Pseudomonas; however, it is less important in stress survival and has more specific roles related to virulence and colonization (reviewed in reference 66). In order to study whether the level of the transcription from the dinB promoter could be elevated in P. putida stationary-phase cells and to investigate possible effects of RpoS on transcription, we decided to compare expression of the reporter gene under the control of the dinB promoter in the wild type and RpoS-defective bacteria sampled from different time points of the batch culture. Because the luxAB reporter system is sensitive only in exponentially growing cells, we used another reporter, the lacZ gene, and monitored the level of ß-Gal activity in P. putida cells carrying a dinB promoter-lacZ transcriptional fusion in the plasmid pKTPdinBlacZ during 100 h of cultivation of bacteria. Results shown in Fig. 5 demonstrate that transcription from the dinB promoter is not controlled by RpoS. In both strains studied (the wild type and the RpoS mutant), we observed a modest increase (approximately twofold) in the level of ß-Gal expression if the activities from exponentially growing cells and stationary-phase cells sampled at h 12 were compared. During the prolonged incubation of bacteria in stationary phase the ß-Gal activities increased but these changes were also small: an approximately threefold difference became apparent if transcription in exponentially growing and late-stationary-phase cells was compared. This experiment indicated that, although the transcription of the dinB gene in P. putida is affected by the growth conditions of the bacteria, RpoS does not play any role in dinB transcription.



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  		FIG. 5. Effect of growth phase of bacteria on transcription from the dinB promoter. ß-Gal activity was measured in P. putida wild-type strain PaW85 (WT) and its RpoS-defective derivative PKS54 (rpoS) carrying the dinB promoter-lacZ fusion in plasmid pKTPdinBlacZ. Bacteria were grown in LB medium. Growth curves of bacteria are indicated by dashed lines. Results of four independent experiments are presented. The standard deviations are shown. OD580, optical density at 580 nm.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Stationary-phase mutagenesis mechanisms have mostly been studied during short-term experiments, usually lasting less than 1 week. However, a large majority of microorganisms in their natural environments face far longer periods of starvation. Results of our recently published study (53) suggested that mutation processes in cells that have been starving for a short period are not entirely compatible with those of a prolonged starvation. The data presented by us herein demonstrate an increase in the frequency of accumulation of Phe+ mutants in populations of P. putida during prolonged starvation. The test system used for the isolation of these mutants is based on reversion of +1 frameshift in the pheA gene. We found that the error-prone DNA polymerase Pol IV was specifically required for most 1-bp deletions detected by measuring the frequency of appearance of Phe+ revertants in long-term-starved populations of P. putida (Fig. 2A). Meanwhile, this effect of Pol IV became clearly apparent only in cell populations that had already been starved for a carbon source for more than 7 days. This implies that Pol IV-dependent mutagenesis could be induced under conditions of long-term environmental stress.

The idea that some mechanisms have probably evolved to control the mutation rate in a cell is a subject of intense scientific debate (see, for example, references 36, 50, 51, 55, and 63). Evidence supporting the hypothesis that stress-induced mutagenesis is a genetically programmed strategy has been found in different organisms, which indicates that regulation of stress-induced mutagenesis is general to all microorganisms. However, different control mechanisms may exist. For example, mutagenesis in resting organisms in structured environments (ROSE mutagenesis), which occurs in E. coli aging colonies, involves control by the SOS system and the catabolite repression system (62). Studies by Bjedov et al. (5) revealed that the frequency of mutations increased remarkably between 1-day- and 7-day-old colonies of most natural isolates of E. coli, from diverse habitats worldwide, and the mutagenesis in aging colonies, in one natural isolate tested, was genetically controlled by RpoS and carbon-sensing regulators. Sung and Yasbin (60) demonstrated that accumulation of prototrophic revertants among Bacillus subtilis cells required activity of genes that are involved in regulation of bacterial differentiation. In our previous study of P. putida (53), we demonstrated that the spectrum of stationary-phase mutations among early-arising mutants (picked up on days 3 and 4) differs from that of later-arising ones (picked up on days 6 and 7). The occurrence of mutations, the number of which started to increase later (e.g., 2- to 3-bp deletions), was dependent on the stationary-phase sigma factor RpoS (53). At the same time, the transposition of IS1411, which also increased with time of starvation and resulted in emergence of Phe+ mutants, was under the negative control of RpoS (53). Most recently, Layton and Foster (38) provided evidence that the cellular amount of Pol IV in E. coli FC40 is controlled by RpoS, and Pol IV is required for stationary-phase mutation in that system (7, 42). Results presented here demonstrate that the frequency of accumulation of Pol IV-dependent mutants increases in P. putida populations during prolonged starvation. Whether RpoS is involved in Pol IV-dependent stationary-phase mutagenesis in the +1 frameshift reversion examined by us in the current study is difficult to determine because P. putida cells lacking functional RpoS die in cultures that have been starved more than 1 week (32).

Mechanisms of Pol IV-dependent stationary-phase mutations studied by us in P. putida differ in some aspects from those of the well-studied model organism E. coli. RecA is both a signal sensor/transducer molecule for the SOS response and a recombination protein (20). In the case of Pol IV-dependent mutagenesis in E. coli strain FC40, which has become a paradigm of stationary-phase mutation, the Lac+ mutations that arise in starving-cell populations on lactose selective plates require RecA function and a RecBCD double-stranded-break repair system (8, 19, 27, 28). The recombination-dependent stationary-phase mutations are proposed to result from erroneous DNA replication at sites of double-stranded-break repair via homologous recombination (for a review of this model, see, for example, references 15, 17, and 50). The results reported by McKenzie et al. (41) indicate that the SOS activation function of RecA is also required for Lac+ reversion in starving cells. DNA polymerase Pol IV, which is required specifically for 1-bp deletions in the Lac+ reversion system in stationary-phase cells (but not in growing cells), is up-regulated during the SOS response (7, 42). An alternative amplification-mutagenesis model for RecA function in Pol IV-dependent Lac+ reversion phenomena has also been proposed (29, 56). In the current study we showed that Pol IV-dependent mutagenesis in P. putida is RecA independent. Also, in contrast to E. coli, transcription from the dinB promoter in P. putida cells occurs at a high basal level both in the presence and absence of the DNA damage-inducing agent mitomycin C: we observed only a slight, maximum twofold increase after induction of DNA damage (Fig. 3). In comparison with E. coli, the SOS response in P. putida has been only poorly studied. However, the existence of a similar DNA damage-inducible response has been found by analyzing DNA damage-mediated induction of the P. putida lexA gene (9).

E. coli strains which have been used to study the role of Pol IV in stationary-phase mutations carry two copies of dinB, one in the chromosome and the other in the F' plasmid (42). The higher expression of dinB in E. coli starving cells, resulting in stationary-phase mutagenesis, is hypothesized to be connected with sporadic amplification of the F' plasmid DNA region carrying the dinB gene (37, 55). However, the increase in dinB expression in stationary phase shown by Layton and Foster (38) was observed in the absence of selection for dinB amplification. Differently from E. coli, the occurrence of stationary-phase mutations studied by us in P. putida depends on the presence of a single chromosomal copy of the dinB gene. If compared to exponentially growing cells, the level of transcription from the dinB promoter in P. putida was increased threefold in late-stationary-phase cells (Fig. 5). However, unlike expression of dinB in E. coli, the transcription of dinB in P. putida was not dependent on RpoS. This and other differences discussed above indicate that mechanisms distinct from those proposed to control expression of dinB in E. coli may up-regulate Pol IV-dependent mutagenesis under conditions of long-term environmental stress in P. putida starving cells.

There are at least three nonexclusive explanations of how the frequency of accumulation of Pol IV-dependent mutants can increase in long-term-starved cell populations: (i) some posttranscriptional/posttranslational mechanisms may control the activity of Pol IV in stressed P. putida cells; (ii) levels of Pol IV protein might increase in P. putida stationary-phase cells; and (iii) DNA repair can be depressed during prolonged starvation. Studies of stationary-phase mutagenesis in E. coli FC40 have demonstrated that defects in methyl-directed mismatch repair (MMR) resulted in a great increase in the number of Lac+ mutants that arose with time after lactose selection (18). The spectra of Lac+ reversion mutations observed in growing cells of MMR-deficient strains (1-bp deletions in small mononucleotide repeats) were indistinguishable from the spectrum of Lac+ reversion mutations characterized in stationary-phase cells of the wild-type strain (40). MMR has been shown to be down-regulated in stationary phase (12, 65). Moreover, results by Harris et al. (26) imply that the MMR protein MutL becomes limiting during stationary-phase mutation. The role of down-regulation of MMR in accumulation of Lac+ mutants in the FC40 system has stimulated an active dispute (14, 25). The idea that down-regulation of MMR might be involved in stationary-phase mutagenesis is supported by the finding that an MMR-defective mutant did not show any significant elevation of mutagenesis in aging colonies (5). The 1-bp deletions measured in starving cells of FC40 which resulted in Lac+ reversion were mostly produced by Pol IV (7, 42). Wagner and Nohmi (67) have also shown that many Pol IV-induced errors are corrected by MMR. Moreover, that report (67) demonstrates saturation of the MMR system as a result of accumulation of errors made by overproduced Pol IV. Hence, drawing parallels with data obtained in E. coli, we hypothesize that the increase in the frequency of Pol IV-dependent mutations observed by us might be caused by the malfunctioning of MMR in P. putida starving cells. Moreover, one may speculate that MMR might be disabled due to a higher activity of Pol IV in long-term-starved P. putida. Further and more straightforward investigations are needed to test these hypotheses.

Rifampin mutation assay scores only base substitutions, but the most frequently observed mutations associated with Pol IV activity in E. coli are frameshifts (35). Artificial overproduction of Pol IV in P. putida growing cells resulted in an elevated frequency of occurrence of Rif-resistant mutants, which indicates that Pol IV can contribute to base substitutions in this organism as well. However, the occurrence of base substitutions in starving P. putida did not require the activity of Pol IV. Depending upon the DNA lesion and its sequence context, different DNA polymerases are involved in generation of mutations in E. coli (22, 44, 68). Thus, if only a specific type of mutation in a specific sequence context is detectable in a given assay system, some assay systems may be less relevant for the detection of Pol IV-generated mutations than others. Base substitutions in the rpoB gene measured by us (Rif-resistant mutants in growing cell populations) and in the test systems detecting Phe+ reversion mutations also occurred in different sequence contexts. Therefore, the failure to detect Pol IV-dependent base substitutions in starving cell populations in this study can be explained by the use of different assay systems for growing and stationary-phase cells. At the same time, results presented by Wagner and Nohmi (67) suggested a bias for Pol IV-generated mutations occurring in sequences with a guanine base at the 5' position of the mutated base. Our test systems designed for the measurement of base substitutions in the pheA gene detected mostly T-to-C transitions either in the sequence 5'-GGGTAA-3' or 5'-GGGTAG-3'. According to Wagner and Nohmi (67) both these sequences should be favored for base substitutions generated by Pol IV. Therefore, an alternative explanation for the phenomenon of why we did not observe any effect of Pol IV on generation of base substitutions in starving cells would be that the role of Pol IV in mutagenesis may vary between growing and stationary-phase cells.

It was recently shown that stationary-phase E. coli cells lacking one or more SOS-induced DNA polymerases (Pol II, Pol IV, and Pol V) are less fit when grown in the presence of wild-type cells under conditions in which the ability to generate GASP mutations (growth advantage in stationary phase) is under selection (70). Base pair substitution mutations occurring in stationary-phase cells of E. coli have been shown to be dependent on Pol V (4). Among gram-negative bacteria, Pseudomonas species examined so far, like many other nonenteric bacteria, lack chromosomally encoded Pol V. Although genes encoding Pol V homologues are frequently found in plasmids (34, 46, 59), bacteria lacking these plasmids accumulate stationary-phase mutations as well. Results of our current study demonstrate that only a particular type of stationary-phase mutations in P. putida requires Pol IV activity. Hence, the question arises whether these bacteria would express some other error-prone DNA polymerase activities involved in stationary-phase mutagenesis. Experiments to address this question are currently in progress.


   ACKNOWLEDGMENTS
 
We thank T. Alamäe, R. Hõrak, and other coworkers for critically reading the manuscript and J. Truu for help in statistical analysis of the data.

This work was supported by grants 4481 and 4482 from the Estonian Science Foundation and by grant HHMI 55000316 from the Howard Hughes Medical Institute International Research Scholars Program.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Genetics, Institute of Molecular and Cell Biology, Tartu University and Estonian Biocentre, 23 Riia Street, 51010 Tartu, Estonia. Phone: 372-7-375036. Fax: 372-7-420286. E-mail: maiak{at}ebc.ee. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Adams, M. H. 1959. Bacteriophages, p. 445-447. Interscience Publishers Inc., New York, N.Y.
  2. Bagdasarian, M. M., E. Ammann, R. Lurz, B. Ruckert, and M. Bagdasarian. 1983. Activity of the hybrid trp-lac (tac) promoter of Escherichia coli in Pseudomonas putida. Construction of broad-host-range, controlled-expression vectors. Gene 26:273-282.[CrossRef][Medline]
  3. Bayley, S. A., C. J. Duggleby, M. J. Worsey, P. A. Williams, K. G. Hardy, and P. Broda. 1977. Two modes of loss of the TOL function from Pseudomonas putida mt-2. Mol. Gen. Genet. 154:203-204.[CrossRef][Medline]
  4. Bhamre, S., B. B. Gadea, C. A. Koyama, S. J. White, and R. G. Fowler. 2001. An aerobic recA, umuC-dependent pathway of spontaneous base-pair substitution mutagenesis in Escherichia coli. Mutat. Res. 473:229-247.[Medline]
  5. Bjedov, I., O. Tenaillon, B. Gérard, V. Souza, E. Denamur, M. Radman, F. Taddei, and I. Matic. 2003. Stress-induced mutagenesis in bacteria. Science 300:1404-1409.[Abstract/Free Full Text]
  6. Bolivar, F., R. L. Rodriguez, P. J. Greene, M. C. Betlach, H. L. Heyneker, and H. W. Boyer. 1977. Construction and characterization of new cloning vehicles. II. A multipurpose cloning system. Gene 2:95-113.[Medline]
  7. Bull, H. J., M.-J. Lombardo, and S. M. Rosenberg. 2001. Stationary-phase mutation in the bacterial chromosome: recombination protein and DNA polymerase IV dependence. Proc. Natl. Acad. Sci. USA 98:8334-8341.[Abstract/Free Full Text]
  8. Cairns, J., and P. L. Foster. 1991. Adaptive reversion of a frameshift mutation in Escherichia coli. Genetics 128:695-701.[Abstract]
  9. Calero, S., X. Garriga, and J. Barbé. 1993. Analysis of the DNA damage-mediated induction of Pseudomonas putida and Pseudomonas aeruginosa lexA genes. FEMS Microbiol. Lett. 110:65-70.[CrossRef][Medline]
  10. Carter, P., H. Bedouelle, and G. Winter. 1985. Improved oligonucleotide site-directed mutagenesis using M13 vectors. Nucleic Acids Res. 13:4431-4443.[Abstract/Free Full Text]
  11. Courcelle, J., A. Khodursky, B. Peter, P. O. Brown, and P. C. Hanawalt. 2001. Comparative gene expression profiles following UV exposure in wild-type and SOS-deficient Escherichia coli. Genetics 158:41-64.[Abstract/Free Full Text]
  12. Feng, G., H.-C. T. Tsui, and M. E. Winkler. 1996. Depletion of the cellular amounts of the MutS and MutH methyl-directed mismatch repair proteins in stationary-phase Escherichia coli K-12 cells. J. Bacteriol. 178:2388-2396.[Abstract/Free Full Text]
  13. Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76:1648-1652.[Abstract/Free Full Text]
  14. Foster, P. L. 1999. Are adaptive mutations due to decline in mismatch repair? The evidence is lacking. Mutat. Res. 436:179-184.[CrossRef][Medline]
  15. Foster, P. L. 1999. Mechanisms of stationary phase mutation: a decade of adaptive mutation. Annu. Rev. Genet. 33:57-88.[CrossRef][Medline]
  16. Foster, P. L. 2000. Adaptive mutation in Escherichia coli. Cold Spring Harbor Symp. Quant. Biol. 65:21-29.
  17. Foster, P. L. 2001. Adaptive mutation: implications for evolution. BioEssays. 22:1067-1074.
  18. Foster, P. L., and J. Cairns. 1992. Mechanisms of directed mutation. Genetics 131:783-789.[Abstract]
  19. Foster, P. L., J. M. Trimarchi, and R. A. Maurer. 1996. Two enzymes, both of which process recombination intermediates, have opposite effects on adaptive mutation in Escherichia coli. Genetics 142:25-37.[Abstract]
  20. Friedberg, E. C., G. C. Walker, and W. Siede. 1995. DNA repair and mutagenesis. ASM Press, Washington, D.C.
  21. Friedberg, E. C., P. L. Fischhaber, and C. Kisker. 2001. Error-prone DNA polymerases: novel structures and the benefits of infidelity. Cell 107:9-12.[CrossRef][Medline]
  22. Friedberg, E. C., R. Wagner, and M. Radman. 2002. Specialized DNA polymerases, cellular survival, and the genesis of mutations. Science 296:1627-1630.[Abstract/Free Full Text]
  23. Goodman, M. F. 2002. Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 71:17-50.[CrossRef][Medline]
  24. Hanahan, D. 1983. Studies on the transformation of E. coli with plasmids. J. Mol. Biol. 166:577-580.
  25. Harris, R. S., G. Feng, K. J. Ross, R. Sidhu, C. Thulin, S. Longerich, S. K. Szigety, P. J. Hastings, M. E. Winkler, and S. M. Rosenberg. 1999. Mismatch repair is diminished during stationary-phase mutation. Mutat. Res. 437:51-60.[CrossRef][Medline]
  26. Harris, R. S., G. Feng, K. J. Ross, R. Sidhu, C. Thulin, S. Longerich, S. K. Szigety, M. E. Winkler, and S. M. Rosenberg. 1997. Mismatch repair protein MutL becomes limiting during stationary-phase mutation. Genes Dev. 11:2426-2437.[Abstract/Free Full Text]
  27. Harris, R. S., S. Longerich, and S. M. Rosenberg. 1994. Recombination in adaptive mutation. Science 264:258-260.[Abstract/Free Full Text]
  28. Harris, R. S., K. J. Ross, and S. M. Rosenberg. 1996. Opposing roles of the Holliday junction processing systems of Escherichia coli in recombination-dependent adaptive mutation. Genetics 142:681-691.[Abstract]
  29. Hendrickson, H., E. S. Slechta, U. Bergthorsson, D. I. Andersson, and J. R. Roth. 2002. Amplification-mutagenesis: evidence that "directed" adaptive mutation and general hypermutability result from growth with a selected gene amplification. Proc. Natl. Acad. Sci. USA 99:2164-2169.[Abstract/Free Full Text]
  30. Herrero, M., V. de Lorenzo, and K. N. Timmis. 1990. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosomal insertion of foreign genes in gram-negative bacteria. J. Bacteriol. 172:6557-6567.[Abstract/Free Full Text]
  31. Hõrak, R., and M. Kivisaar. 1998. Expression of the transposase gene tnpA of Tn4652 is positively affected by integration host factor. J. Bacteriol. 180:2822-2829.[Abstract/Free Full Text]
  32. Ilves, H., R. Hõrak, and M. Kivisaar. 2001. Involvement of {sigma}S in starvation-induced transposition of Pseudomonas putida transposon Tn4652. J. Bacteriol. 183:5445-5448.[Abstract/Free Full Text]
  33. Kasak, L., R. Hõrak, and M. Kivisaar. 1997. Promoter-creating mutations in Pseudomonas putida: a model system for the study of mutation in starving bacteria. Proc. Natl. Acad. Sci. USA 94:3134-3139.[Abstract/Free Full Text]
  34. Kim, J. J., and G. W. Sundin. 2000. Regulation of the rulAB mutagenic DNA repair operon of Pseudomonas syringae by UV-B (290 to 320 nanometers) radiation and analysis of rulAB-mediated mutability in vitro in planta. J. Bacteriol. 182:6137-6144.[Abstract/Free Full Text]
  35. Kim, S.-R., G. Maenhaut-Michel, M. Yamada, Y. Yamamoto, K. Matsui, T. Sofuni, T. Nohmi, and H. Ohmori. 1997. Multiple pathways for SOS-induced mutagenesis in Eschericha coli: an overexpression of dinB/dinP results in strongly enhancing mutagenesis in the absence of any exogeneous treatment to damage DNA. Proc. Natl. Acad. Sci. USA 94:13792-13797.[Abstract/Free Full Text]
  36. Kivisaar, M. 2003. Stationary phase mutagenesis: mechanisms that accelerate adaptation of microbial populations under environmental stress. Environ. Microbiol. 5:814-827.[CrossRef][Medline]
  37. Kofoid, E., U. Bergthorsson, E. S. Slechta, and J. R. Roth. 2003. Formation of an F' plasmid by recombination between imperfectly repeated chromosomal Rep sequences: a closer look at an old friend (F'128 pro lac). J. Bacteriol. 185:660-663.[Abstract/Free Full Text]
  38. Layton, J. C., and P. L. Foster. 2003. Error-prone DNA polymerase IV is controlled by the stress-response sigma factor, RpoS, in Escherichia coli. Mol. Microbiol. 50:549-561.[CrossRef][Medline]
  39. Lea, D. E., and C. A. Coulson. 1949. The distribution of the numbers of mutants in bacterial populations. J. Genet. 49:264-265.
  40. Longerich, S., A. M. Galloway, R. S. Harris, C. Wong, and S. M. Rosenberg. 1995. Adaptive mutation sequences reproduced by mismatch repair deficiency. Proc. Natl. Acad. Sci. USA 92:12017-12020.[Abstract/Free Full Text]
  41. McKenzie, G. J., R. S. Harris, P. L. Lee, and S. M. Rosenberg. 2000. The SOS response regulates adaptive mutation. Proc. Natl. Acad. Sci. USA 97:6646-6651.[Abstract/Free Full Text]
  42. McKenzie, G. J., P. L. Lee, M.-J. Lombardo, P. J. Hastings, and S. M. Rosenberg. 2001. SOS mutator DNA polymerase IV functions in adaptive mutation and not adaptive amplification. Mol. Cell 7:571-579.[CrossRef][Medline]
  43. Miller, J. H. 1992. A short course in bacterial genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
  44. Napolitano, R., R. Janel-Bintz, J. Wagner, and R. P. P. Fuchs. 2000. All three SOS-inducible DNA polymerases (Pol II, Pol IV and Pol V) are involved in induced mutagenesis. EMBO J. 19:6259-6265.[CrossRef][Medline]
  45. Nurk, A., L. Kasak, and M. Kivisaar. 1991. Sequence of the gene (pheA) encoding phenol monooxygenase from Pseudomonas sp. EST1001: expression in Escherichia coli and Pseudomonas putida. Gene 102:13-18.[CrossRef][Medline]
  46. Ohmori, H., E. C. Friedberg, R. P. P. Fuchs, M. F. Goodman, F. Hanaoka, D. Hinkle, T. A. Kunkel, C. W. Lawrence, Z. Livneh, T. Nohmi, L. Prakash, S. Prakash, T. Todo, G. C. Walker, Z. Wang, and R. Woodgate. 2001. The Y-family of DNA polymerases. Mol. Cell 8:7-8.[CrossRef][Medline]
  47. Ojangu, E.-L., A. Tover, R. Teras, and M. Kivisaar. 2000. Effects of combination of different –10 hexamers and downstream sequences on stationary-phase-specific sigma factor {sigma}S-dependent transcription in Pseudomonas putida. J. Bacteriol. 182:6707-6713.[Abstract/Free Full Text]
  48. Pavel, H., M. Forsman, and V. Shingler. 1994. An aromatic effector specificity mutant of the transcriptional regulator DmpR overcomes the growth constraints of Pseudomonas sp. strain CF600 on para-substituted methylphenols. J. Bacteriol. 176:7550-7557.[Abstract/Free Full Text]
  49. Rosche, W. A., and P. L. Foster. 2000. Determining mutation rates in bacterial populations. Methods 20:4-17.[CrossRef][Medline]
  50. Rosenberg, S. M. 2001. Evolving responsively: adaptive mutation. Nat. Rev. Genet. 2:504-515.[Medline]
  51. Roth, J. R., E. Kofoid, F. P. Roth, O. G. Berg, J. Seger, and D. I. Andersson. 2003. Regulating general mutation rates: examination of the hypermutable state model for cairnsian adaptive mutation. Genetics 163:1483-1496.[Abstract/Free Full Text]
  52. Santos, P. M., I. Di Bartolo, J. M. Blatny, E. Zennaro, and S. Valla. 2001. New broad-host-range promoter probe vectors based on the plasmid RK2 replicon. FEMS Microbiol. Lett. 195:91-96.[CrossRef][Medline]
  53. Saumaa, S., A. Tover, L. Kasak, and M. Kivisaar. 2002. Different spectra of stationary-phase mutations in early-arising versus late-arising mutants of Pseudomonas putida: involvement of the DNA repair enzyme MutY and stationary-phase sigma factor RpoS. J. Bacteriol. 184:6957-6965.[Abstract/Free Full Text]
  54. Sharma, R. C., and R. T. Schimke. 1996. Preparation of electro-competent E. coli using salt-free growth medium. BioTechniques 20:42-44.[Medline]
  55. Slechta, E. S., K. L. Bunny, E. Kugelberg, E. Kofoid, D. I. Andersson, and J. R. Roth. 2003. Adaptive mutation: general mutagenesis is not a programmed response to stress but results from rare coamplification of dinB with lac. Proc. Natl. Acad. Sci. USA 100:12847-12852.[Abstract/Free Full Text]
  56. Slechta, E. S., J. Liu, D. I. Andersson, and J. R. Roth. 2002. Evidence that selected amplification of a bacterial lac frameshift allele stimulates Lac+ reversion (adaptive mutation) with or without general hypermutability. Genetics 161:945-956.[Abstract/Free Full Text]
  57. Sokal, R. R., and F. J. Rohlf. 1981. Biometry. W. H. Freeman, San Francisco, Calif.
  58. Spiers, A. J., A. Buckling, and P. B. Rainey. 2000. The causes of Pseudomonas diversity. Microbiology 146:2345-2350.[Free Full Text]
  59. Sundin, G. W., S. P. Kidambi, M. Ullrich, and C. L. Bender. 1996. Resistance to ultraviolet light in Pseudomonas syringae: sequence and functional analysis of the plasmid-encoded rulAB genes. Gene 177:77-81.[CrossRef][Medline]
  60. Sung, H.-M., and R. E. Yasbin. 2002. Adaptive, or stationary-phase, mutagenesis, a component of bacterial differentiation in Bacillus subtilis. J. Bacteriol. 184:5641-5653.[Abstract/Free Full Text]
  61. Sung, H.-M., G. Yeamans, C. A. Ross, and R. E. Yasbin. 2003. Roles of YgjH and YgjW, homologs of the Escherichia coli UmuC/DinB or Y superfamily of DNA polymerases, in stationary-phase mutagenesis and UV-induced mutagenesis of Bacillus subtilis. J. Bacteriol. 185:2153-2160.[Abstract/Free Full Text]
  62. Taddei, F., I. Matic, and M. Radman. 1995. cAMP-dependent SOS induction and mutagenesis in resting bacterial populations. Proc. Natl. Acad. Sci. USA 92:11736-11740.[Abstract/Free Full Text]
  63. Tenaillon, O., F. Taddei, M. Radman, and I. Matic. 2001. Second-order selection in bacterial evolution: selection acting on mutation and recombination rates in the course of adaptation. Res. Microbiol. 152:11-16.[Medline]
  64. Tover, A., E.-L. Ojangu, and M. Kivisaar. 2001. Growth medium composition-determined regulatory mechanisms are superimposed on CatR-mediated transcription from the pheBA and catBCA promoters in Pseudomonas putida. Microbiology 147:2149-2156.[Abstract/Free Full Text]
  65. Tsui, H.-C. T., G. Feng, and M. E. Winkler. 1997. Negative regulation of mutS and mutH repair gene expression by the Hfq and RpoS global regulators of Escherichia coli K-12. J. Bacteriol. 179:7476-7487.[Abstract/Free Full Text]
  66. Venturi, V. 2003. Control of rpoS transcription in Escherichia coli and Pseudomonas: why so different? Mol. Microbiol. 49:1-9.[CrossRef][Medline]
  67. Wagner, J., and T. Nohmi. 2000. Escherichia coli DNA polymerase IV mutator activity: genetic requirements and mutational specificity. J. Bacteriol. 182:4587-4595.[Abstract/Free Full Text]
  68. Wagner, J., H. Etienne, R. Janel-Bintz, and R. P. Fuchs. 2002. Genetics of mutagenesis in E. coli: various combinations of translesion polymerases (Pol II, IV and V) deal with lesion/sequence context diversity. DNA Repair 1:159-167.[Medline]
  69. Walker, G. C. 1984. Mutagenesis and inducible responses to deoxyribonucleic acid damage in Escherichia coli. Microbiol. Rev. 48:60-93.[Free Full Text]
  70. Yeiser, B., E. D. Pepper, M. F. Goodman, and S. E. Finkel. 2002. SOS-induced DNA polymerases enhance long-term survival and evolutionary fitness. Proc. Natl. Acad. Sci. USA 99:8737-8741.[Abstract/Free Full Text]

Journal of Bacteriology, May 2004, p. 2735-2744, Vol. 186, No. 9
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.9.2735-2744.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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