Photoreactivation of Escherichia coli is impaired at high growth temperatures

Journal of Photochemistry and Photobiology B: Biology 147 (2015) 37–46

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Photoreactivation of Escherichia coli is impaired at high growth temperatures Lei Xu a,b, Changqing Tian a, Xiaohua Lu b, Liefeng Ling b, Jun Lv b, Mingcai Wu a,b, Guoping Zhu a,⇑ a b

Institute of Molecular Biology and Biotechnology, Anhui Normal University, Wuhu, Anhui, China Anhui Province Key Laboratory of Active Biological Macro-molecules, Wannan Medical College, Wuhu, Anhui, China

a r t i c l e

i n f o

Article history: Received 5 October 2014 Received in revised form 26 February 2015 Accepted 2 March 2015 Available online 23 March 2015

a b s t r a c t Photolyase repairs UV-induced lesions in DNA using light energy, which is the principle of photoreactivation. Active photolyase contains the two-electron-reduced flavin cofactor. We observed that photoreactivation of Escherichia coli was impaired at growth temperatures P37 °C, and growth in this temperature range also resulted in decreased photolyase protein levels in the cells. However, the levels of phr transcripts (encoding photolyase) were almost unchanged at the various growth temperatures. A lacZ-reporter under transcriptional control of the phr promoter showed no temperature-dependent expression. However, a translational reporter consisting of the photolyase N-terminal a/b domain–LacZ fusion protein exhibited lower b-galactosidase activity at high growth temperatures (37–42 °C). These results indicated that the change in photolyase levels at different growth temperatures is post-transcriptional in nature. Limited proteolysis identified several susceptible cleavage sites in E. coli photolyase. In vitro differential scanning calorimetry and activity assays revealed that denaturation of active photolyase occurs at temperatures P37 °C, while apo-photolyase unfolds at temperatures P25 °C. Evidence from temperature-shift experiments also implies that active photolyase is protected from thermal unfolding and proteolysis in vivo, even at 42 °C. These results suggest that thermal unfolding and proteolysis of newly synthesized apo-photolyase, but not active photolyase, is responsible for the impaired photoreactivation at high growth temperatures (37–42 °C). Ó 2015 Elsevier B.V. All rights reserved.

1. Introduction UV radiation in the UV-B (280–315 nm) and UV-C (100– 280 nm) spectral regions stimulates adjacent pyrimidine bases in DNA to form dimer lesions. The two major dimer lesions, the cyclobutane pyrimidine dimer (CPD) and the (6-4) pyrimidinepyrimidone photoproduct [(6-4) photoproduct], constitute 70–90% and 10–30% of the total lesions, respectively [1,2]. The UV-induced lesions inhibit replication and transcription in cells [2], which enables UV-C to inactivate many microorganisms such as bacteria, viruses and protozoa [3]. Currently, UV disinfection technology is widely used in hospitals, the food industry and wastewater treatment plants. There are many advantages of UV disinfection over other technologies, including convenience, safety, a reduced space requirement, and the absence of a chemical smell or taste. Furthermore, the UV disinfection process is insensitive to temperature variations [4,5]. However, a significant disadvantage ⇑ Corresponding author at: Institute of Molecular Biology and Biotechnology, Anhui Normal University, No. 1 Beijing East Road, Wuhu 241000, Anhui, China. Tel./fax: +86 553 3883592. E-mail address: [email protected] (G. Zhu). http://dx.doi.org/10.1016/j.jphotobiol.2015.03.012 1011-1344/Ó 2015 Elsevier B.V. All rights reserved.

of UV disinfection is that many microorganisms can reverse UV-induced lesions by photoreactivation, which is an enzymatic reaction that may be affected by the pre- and post-UV irradiation conditions [3,6,7]. Photoreactivation was discovered on the basis of the observation that the lethal effect of UV radiation on some organisms could be reversed following visible light illumination [8]. It was later demonstrated that photolyase (EC 4.1.99.3) is the enzyme responsible for photoreactivation [9]. Photoreactivation by photolyase is considered to be a more direct, energy-saving and less error-prone pathway to restore UV-induced lesions compared with the other repair pathways [2]. In Escherichia coli, photolyase can efficiently repair CPD in DNA utilizing blue or UV-A light (350–450 nm) as the energy source. The enzyme is a monomeric protein of 50 kDa and has two non-covalently bound cofactors. Flavin adenine dinucleotide in the two-electron-reduced state (FADH) is essential for catalysis [1]. The second cofactor 5,10methenyltetrahydrofolate (MTHF) is not necessary for catalysis but functions as an antenna to increase the catalytic efficiency of the enzyme under limiting light conditions [1]. E. coli photolyase is encoded by the phr gene, which is preceded by a function-unknown gene ybgA (formerly named orf169) in the E. coli genome

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to form a transcription unit [10,11]. Two promoters (P1 and P2) control the transcription of the phr gene [11] which are not induced by the SOS response [1,12]. Additionally, some organisms possess 6-4 photolyases that can repair (6-4) photoproducts in DNA using light energy [13]. However, 6-4 photolyase is not found in E. coli. The reaction of photoreactivation follows a two-step scheme [1,2,14]: a light-independent step to form a complex between photolyase and a dimer lesion, followed by a light-dependent step to photorepair the lesion. The enzyme-substrate complex formation step is dependent on temperature, pH, and ionic strength, making the entire photoreactivation process to be influenced by these post-UV irradiation conditions [6,15–17]. However, the effects of pre-UV irradiation conditions such as culture time and culture temperature on photoreactivation are largely unknown. It has been reported that seasonal differences significantly affect photoreactivation that summer coliform populations in wastewater have a lower level of photoreactivation than winter populations, but the reason for this remains obscure [6]. Further explorations into the effects of the pre-UV conditions on photoreactivation will not only provide new insights into photoreactivation, but also help enhance the efficiency of UV disinfection and reduce the risk of microorganism pollution. As a model organism, E. coli has many similarities with other coliforms. Furthermore, photolyases of coliforms have at least 70% sequence identities with E. coliphotolyase (Table 1). In this study, we investigated the photoreactivation of E. coli cells cultured at various temperatures. The total protein levels of photolyase and the steady-state transcript levels of the phr gene encoding photolyase were analyzed by Western blotting and real-time quantitative reverse transcription–PCR (RT–PCR). Plasmids containing transcriptional and translational lacZ reporters under the control

Table 1 Comparing the protein sequences of photolyases of coliforms and other species of Enterobacteriaceae with E. coli photolyase using the Basic Local Alignment Search Tool (BLAST, http://blast.ncbi.nlm.nih.gov). Lengtha

Identitiesb

Positivesc

Escherichia coli

472

Citrobacter freundii

472

Citrobacter amalonaticus Enterobacter aerogenes Enterobacter cloacae

471

Klebsiella oxytoca

472

Klebsiella pneumoniae

471

472/ 472(100%) 382/ 472(81%) 385/ 471(82%) 335/ 469(71%) 351/ 470(75%) 334/ 472(71%) 329/ 471(70%)

472/ 472(100%) 431/ 472(91%) 421/ 471(89%) 394/ 469(84%) 402/ 470(85%) 391/ 472(82%) 380/ 471(80%)

Shigella dysenteriae

471

Shigella sonnei

472

Salmonella enterica

472

Pantoea agglomerans

473

Yersinia enterocolitica Hafnia alvei

476

459/ 471(97%) 466/ 472(99%) 372/ 472(79%) 294/ 473(62%) 292/ 476(61%) 284/ 472(60%)

466/ 471(98%) 467/ 472(98%) 416/ 472(88%) 356/ 473(75%) 361/ 476(75%) 350/ 472(74%)

Species Coliforms

Other species

a

469 470

472

The number of total protein residues. The number of identical residues/the number of total residues; the ratio is provided in parentheses. c The number of conservative substitutions/the number of total residues; the ratio is provided in parentheses. b

of the phr promoters were constructed to further unravel the temperature-dependent expression of the phr gene. Furthermore, the proteolytic susceptibility and the thermal stability of E. coli photolyase were studied in vitro using limited proteolysis, activity assay and differential scanning calorimetry. And the in vivo behavior of E. coli photolyase was verified by temperature-shift experiment.

2. Material and methods 2.1. Photoreactivation experiments under continuous illumination or flash light Photoreactivation experiments under continuous illumination were performed as described previously [18] with some modifications. To study the kinetics of photoreactivation with less interference by dark repair, E. coli strain AB2463 (recA13) was used. In this strain, recombination repair [19] and the SOS response [20] are deficient, but photoreactivation is proficient. Bacterial cultures were grown in Luria broth (LB) medium at various temperatures (15–42 °C) with vigorous shaking. Stationary phase or exponential phase cells at certain cell densities were harvested, washed and resuspended in buffered saline (pH 7.0) to a proper scale. For UV irradiation, the cell suspension (10 ml) was placed in a Petri dish (90 mm diameter) and irradiated with 253.7 nm UV-C light from a low-pressure mercury lamp whose dose had been measured by the iodide–iodate chemical actinometer [21] or a calibrated UV-C radiometer (ZDZ-1, Shanghai). For photoreactivation, a high power UV-A light emitting diode (LED, kmax = 385 nm, light intensity is 100 W m2 at a distance of 10 cm) was used as the light source, and a glass cover was used to filter out any wavelengths below 300 nm. Control experiments demonstrated that this light source did not change the survival of undamaged E. coli cells. Aliquots of illuminated samples were withdrawn at intervals and plated onto LB plates in triplicate, and colonies were counted after incubation at 37 °C overnight. The culturable survival was defined as the ratio of the colony-forming units (CFU) of UV irradiated or photoreactivated samples to a control sample kept in the dark. At least three independent experiments were performed for the stationary phase cells at each growth temperature. All experiments were performed under a red LED lamp with an ambient temperature of 23 °C. In the flash photoreactivation experiments, the extremely UVsensitive strain AB2480 (recA13, uvrA6) was used. In this strain, excision repair [28], recombination repair and the SOS response are deficient, but photoreactivation is proficient. The experimental procedure was adapted from an earlier report [22]. A home-made flash apparatus containing two oppositely mounted flash units was used. The flash windows were attached with three pieces of glass slides to filter out wavelengths below 300 nm. Suspension of UV-irradiated cells was pipetted into a clear plastic tube (Axygen) and incubated in the dark for at least 15 min prior to the flash. The tube containing the cell suspension was placed in the middle of the flash apparatus, with a distance to each flash window of approximately 1 cm. The two flash units were discharged simultaneously to provide a 1-ms duration flash, which was sufficiently intense to repair essentially all enzyme-substrate complexes present at the time. After applying the flash, aliquots of the sample were spread onto plates and immediately incubated at 37 °C. The colonies were counted after overnight incubation. The maximum extent of photoreactivation was determined by illuminating UV-C irradiated cell suspensions with a 30 W daylight fluorescent lamp at a distance of 10 cm for 1 h as a reference. Three independent experiments were performed for the cells at each growth temperature. All experiments were performed under a red LED lamp with an ambient temperature of 23 °C.

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2.2. Antibody preparation and Western blotting His-tagged E. coli photolyase was purified as previously reported [18,23]. The protein was overexpressed from plasmid phr(N + X), which harboring the E. coli phr gene followed by a six-histidine-tag coding sequence, in E. coli strain BL21(DE3). The cells containing overexpressed photolyase were collected and disrupted. Fresh photolyase in the radical form (E-FADH) was purified from the lysate supernatant via Ni2+-activated chelating Sepharose and DNA cellulose columns. A polyclonal primary antibody against E. coli photolyase was raised by injecting the purified His-tagged protein with Freund’s incomplete adjuvant into rabbits. E. coli strains AB1157 (‘‘wild type’’), AB2463 (recA13) and AB2480 (recA13, uvrA6) were used to analyze the total photolyase levels in the cells cultured at various temperatures and to verify whether the recA and uvrA genes affected the photolyase levels. The CFUs of the samples were determined by dilution and plating. Each sample containing 109 cells was lysed with 80 ll lysis solution containing 50 mM Tris-HCl (pH 7.0), 100 mM NaCl, 25 mM MgCl2, 200 lg ml1 lysozyme, 60 U ml1 DNase I and 1 mM phenylmethanesulfonyl fluoride (PMSF). The lysate was then mixed with 20 ll 5  SDS PAGE loading buffer, boiled for 5 min and centrifuged. Ten-microliter aliquots of supernatant (from 108 cells) were loaded and electrophoresed on SDS–polyacrylamide gels. The proteins were transferred to nitrocellulose membranes using a Trans-Blot SD semi-dry electrophoretic transfer cell (Bio-Rad). Photolyase was detected by incubating the membranes with the rabbit polyclonal primary antibody and the horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (Sigma) followed by the enhanced chemiluminescence substrate (Applygen). Each experiment was repeated at least three times.

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pTrcHisA (Invitrogen) between the HindIII and PstI restriction sites to obtain the pTrcLac plasmid. pLLac was constructed by inserting a ybgA-phr fragment (738,031–740,153 of the E. coli MG1655 genome, containing the ybgA-phr transcription unit with the phr gene and the preceding ybgA gene as well as the two phr promoters, but without the terminator) into pTrcLac between SphI and PstI restriction sites, resulting in a lacZ gene under the transcriptional control of the phr promoters (P1 and P2) [11]. A shorter fragment (738,031–739,135 of the E. coli MG1655 genome) was inserted into pTrcLac instead to obtain pSLac, which encodes a photolyase N-terminal a/b domain (residue 1–134)-LacZ fusion protein under the control of the phr promoters. DNA sequence analyses confirmed that the plasmid constructions were successful. To inspect the growth temperature-dependent expression of the lacZ reporters under the control of the phr promoters, pLLac and pSLac were transformed into DH5a cells [D(argF-lac)169, u80dlacZ58(M15), recA1]. The transformants were inoculated into LB containing 100 lg/ml ampicillin and cultured at various temperatures. At intervals, the cultures at different cell densities were withdrawn for CFU counting by dilution and plating and for b-galactosidase activity assay. A culture of 200 ll was mixed with 800 ll assay buffer containing 100 mM sodium phosphate, 40 mM 2-mercaptoethanol, 10 mM KCl, 1 mM MgSO4 and 0.005% SDS (pH 7.0). The cells were permeabilized by the addition of 100 ll chloroform and vortexed. Then, a 200 ll solution of o-nitrophenyl-b-Dgalactoside (ONPG) at 4 mg ml1 was added and mixed to start the reaction at 28 °C. The reaction was stopped by adding 500 ll of 1 M Na2CO3 at time t, the cell debris and chloroform was removed by centrifugation, and the absorbance at 420 nm (A420) was determined. The b-galactosidase activity was defined as 1010  A420/(CFU  t) (min1), where the CFU value was the total colony-forming units of the culture of 200 ll.

2.3. RNA isolation and real-time quantitative RT–PCR 2.5. Limited proteolysis Genomic-DNA-free total RNA samples of E. coli cells cultured at different temperatures in the exponential or stationary phases were isolated using a modified SDS-phenol method [24]. Reverse transcription was performed with 1 lg of total RNA per sample, 5 lM random hexamer primers, 200 lM of each dNTP and 20 U AMV reverse transcriptase (Sangon) in a 20 ll final reaction volume. The reactions were performed at 42 °C for 45 min. Then, 2 ll aliquots of cDNA were employed as the templates in 25 ll real-time PCR reactions with 0.2 lM of each primer. Specific primers phrL (50 -ACCCATCTGGTCTGGTTTC-30 ) and phrR (50 -CCCGCTCATTCACTTCAT-30 ) were used to determine the expression of the target gene phr under different conditions. Specific primers pgiL (50 -GCCGTTACTCTTTGTGGTC-30 ) and pgiR (50 -GCGGGTTATGGGTGATAG-30 ) were used to amplify the pgi gene, a housekeeping gene encoding the phosphoglucose isomerase whose expression in E. coli is almost constant at various growth temperatures [25], to serve as the reference gene. The PCR conditions were: 3 min at 95 °C, followed by 40 cycles of 15 s at 95 °C, 20 s at 60 °C, 20 s at 72 °C and 20 s at 83 °C for data acquisitions. Each reaction was performed in triplicate wells. The assays were carried out using an iQ5 real-time PCR detection system (Bio-Rad). The amplification data were analyzed by the iQ5 optical system software (Bio-Rad) with the normalized expression (DDCT) analysis method. 2.4. Construction of plasmids containing lacZ reporters and assay of bgalactosidase activity Two plasmids (pLLac and pSLac) containing transcriptional and translational lacZ reporters under the control of the phr promoters were constructed. A lacZ fragment (362,435–365,541 of the E. coli MG1655 genome, containing the lacZ coding sequence and its ribosome binding site) was amplified by PCR and inserted into

C-terminal His-tagged photolyase with two-electron-reduced FAD (E-FADH) was obtained by reducing the freshly purified enzyme with 3 mM dithionite. Apo-photolyase was prepared by incubating the freshly purified enzyme with 0.5 M imidazole (pH 10.0) as previously described [23]. The photolyase samples in these two forms were digested using trypsin, chymotrypsin and proteinase K (all from Sigma) at 23 °C for 15 min with enzyme:substrate ratios of 1:100, 1:100 and 1:200, respectively. The reactions were terminated by the addition of SDS loading buffer and boiled for 5 min. The mixtures were resolved by 10% SDS– PAGE and electrotransferred to nitrocellulose. The fragments containing the intact C-terminal His-tag were detected using an anti-6  His rabbit polyclonal antibody (BBI, AB10002). 2.6. Photolyase activity assays in vitro E. coli photolyase in the E-FADH form was incubated at various temperatures and withdrawn at intervals. The activity of the enzyme retained after incubation was evaluated by the method described previously [23] with some modifications. The assay volume was 500 ll and contained 0.1 lM enzyme with 5 lM UV-C-irradiated oligothymidylates [UV-oligo (dT)16] (containing 4 dimers per molecule). The high power UV-A LED (kmax = 385 nm, light intensity is 100 W m2 at a distance of 10 cm) was used as the source of illumination. The activity was monitored as the increase of absorbance at 265 nm at 23 °C. 2.7. Differential scanning calorimetry The thermal stability of E. coli photolyase in the E-FADH form, the oxidized form (E-FADox) or the apo form was studied by

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differential scanning calorimetry (DSC). E-FADox was gained by reconstituting apo-photolyase with excess oxidized FAD [23]. Micro DSC analyses were performed on a VP-DSC instrument (MicroCal) as described previously [18]. The volume of the sample cell was 0.509 ml. The scans were performed at the rate of 1 °C min1 from 10–75 °C. Data were analyzed by Origin 7.0 (Microcal) with the VP-DSC add-on. 2.8. Temperature-shift experiment E. coli strain AB2463 (recA13) cells were initially grown at 25 °C or 42 °C to the stationary phase or exponential phase, transferred to 42 °C or 25 °C and cultured for another 24 h. At intervals, the photoreactivation kinetics of the cells were determined under continuous illumination as described in Section 2.1, and the photolyase protein levels in the cells were analyzed by Western blotting as described in Section 2.2. 3. Results 3.1. Photoreactivation of E. coli cells under continuous illumination All E. strains containing intact phr gene are photoreactivation proficient, because the gene product photolyase can remove UV-induced CPD lesions in DNA using light energy [1,2]. To detect photoreactivation kinetics with less interference by dark repair, we used a recA strain AB2463 which is deficient in recombination repair [19] and the SOS response [20]. AB2463 has a higher UV sensitivity than normal E. coli strains. Stationary phase AB2463 cells can be reduced to sufficiently low culturable survival (103) by a low UV-C dose (10 J m2); in contrast, the ‘‘wild type’’ strain AB1157 requires 100 J m2 to obtain comparable culturable survival (Fig. S1A). Using a lower dose minimizes the side effects of UV-C to cells (i.e. free-radical formation). Moreover, 10 J m2 of UV-C can produce sufficient CPD lesions per genome, and thereby serve as the substrate of photolyase to determine the photoreactivation kinetics. It is widely accepted that photoreactivation is an enzymatic process catalyzed by photolyase that follows the conventional Michaelis–Menten scheme [1,2,14]: k1

k3 ¼kp I

k2

hv

ƒ! E þ S ƒƒƒƒ ƒ ES ƒƒƒƒƒƒ! E þ P

ð1Þ

where the rate constant k3 is dependent on the light intensity I and photolytic constant kp. If the light intensity is sufficiently large, any enzyme-substrate complex (ES) formed can be immediately converted to product (P) and enzyme (E). Then the photoreactivation kinetics will only depend on the kinetics of the formation of ES, and the free enzyme concentration [E] will remain constant. Then we can obtain

d½P d½S   k1 ½E½S dt dt

ð2Þ

or

ln

½S  k1 ½Et ½S0

ð3Þ

where the indices 0 and t specify the substrate concentrations [S] at t = 0 and t. For this reason, we used a high power UV-A LED as the light source for continuous illumination; this light source emits 100 W m2 385 nm light on the illumination surface. Using this high intensity light source, the above assumption can be roughly met. Thus, we can obtain a good estimation for active photolyase content in the cells.

Fig. 1. The kinetics of photoreactivation of AB2463 (recA13) cells. The cells were grown at various temperatures (, 15 °C; h, 20 °C; j, 25 °C; s, 30 °C; d, 37 °C; D, 42 °C) to stationary phase, harvested and diluted, given 10 J m2 of UV-C, and illuminated under a high power UV-A LED (kmax = 385 nm, light intensity is 100 W m2 at a distance of 10 cm). The culturable survival was plotted vs. the illuminating time. Each point represents 3 data measurements, and the bars represent the standard deviations.

The photoreactivation kinetics were determined with stationary phase AB2463 cells that had been cultured at various temperatures between 15 and 42 °C. The cell suspension of each sample was given a dose of 10 J m2 of UV-C. Then, the cells were immediately illuminated under continuous 385 nm light with an ambient temperature of 23 °C for photoreactivation. The culturable survival values of the illuminated samples were recorded at intervals (Fig. 1). No significant difference was observed in the photoreactivation kinetics from the cells cultured between 15 and 30 °C, although a slight decrease was observed at 30 °C. In contract, there was a significant decrease in photoreactivation efficiency in the cells cultured at 37 °C and a drastic decline in those cultured at 42 °C (Fig. 1 and Table S1). The enzyme concentrations at various growth temperatures were calculated after converting the culturable survival values from the photoreactivation kinetics data into the substrate concentrations with the aid of the UV survival rates and the assumption that k1 does not change with the growth temperature (Fig. S1 and Table S1, details of the derivation are provided in the Supplementary material). The numbers of active photolyase per cell were roughly estimated to be 224 ± 52, 99 ± 11 and 11 ± 6 in cells grown at 25 °C, 37 °C and 42 °C, respectively. Similarly, we measured the photoreactivation kinetics of exponential phase AB2463 cells under continuous illumination. The UV sensitivity of AB2463 cells increased dramatically during the exponential phase, such that only 2 J m2 of UV-C reduced the culturable survival to <103. Suspensions of the exponential phase cells at certain optical densities were irradiated with 2 J m2 of UV-C and then immediately illuminated under the LED light at 385 nm for photoreactivation. The kinetics results are listed in Table S2. The observed behavior was similar to that observed in the stationary phase cells, in which the free enzyme concentration was significantly higher in the cells cultured at 25 °C compared to those cultured at 37 °C or 42 °C. These results suggest that there are reduced amounts of active photolyase in E. coli cells cultured at higher temperatures, and this phenomenon may be responsible for the decline in photoreactivation efficiency. However, a less likely possible explanation that the enzyme in the cells cultured at high temperatures had lower association rates for the substrate (k1) could not be ruled out.

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3.2. Photoreactivation of E. coli cells under flash light A conventional method to determine the number of active photolyase per cell is the flash repair technique [22,26,27]. The principle of the method is that a sufficiently intense and short flash light can repair essentially all enzyme-substrate complexes that are present at the time, without new complex formation or dissociation. A prerequisite for the application of the method is that the strain used must be sufficiently UV-sensitive that the relatively low number of lesions repaired by one flash would result in a significant increase in culturable survival. E. coli strain AB2480 (recA13, uvrA6) is deficient in excision repair [28] as well as recombination repair and the SOS response; thus, this strain is extremely sensitive to UV-C irradiation and should be suitable for the use of the flash photoreactivation method to determine the active photolyase content in the cells. The number of enzyme-substrate complexes present per cell is proportional to the dose decrement (DD) (i.e. the amount of UV dose that in effect was annulled by the flash illumination), which can be determined with the aid of the UV survival curves before and after the flash [22,26,27]. Fig. 2 shows the results of the flash repair experiments with stationary phase AB2480 cells cultured at 25 °C, 37 °C and 42 °C. The maximum extents of photoreactivation (achieved by 1 h whitelight illumination after UV-C irradiation) in AB2480 cells cultured at various temperatures were similar. However, there was a clear tendency for reduced DD at higher growth temperatures. This result also indicates that the active photolyase contents were reduced in the cells cultured at higher temperatures. From the data of the flash repair experiments, AB2480 cells cultured at 25 °C, 37 °C and 42 °C were estimated to contain approximately 250 ± 23, 126 ± 11 and 21 ± 9 molecules of active photolyase per cell, respectively (Fig. S2, details of the derivation are provided in the Supplementary material). These values are closely similar to those estimated based on the photoreactivation kinetics data (Section 3.1). The possibility of the lower k1 in the cells cultured at higher temperatures can now be excluded, because during the flash repair experiments the binding of the enzyme and substrate is in equilibrium.

3.3. Photolyase protein levels but not steady-state phr transcript levels are decreased at higher growth temperatures To verify the effect of growth temperatures on the total photolyase protein levels, Western blotting analyses were performed with E. coli cells cultured at various temperatures. The results obtained with stationary phase cells of several E. coli strains (AB1157, AB2463 and AB2480) are shown in Fig. 3. The standard lanes containing purified photolyase produced a single band at 50 kDa. In all strains, the photolyase protein levels were decreased with increasing growth temperatures. These results are consistent with the results from the photoreactivation kinetics (Section 3.1) and the flash repair experiments (Section 3.2). The similar behavior observed in the ‘‘wild type’’ strain AB1157 rules out the possibility that the mutated genes (recA and uvrA) affected the photolyase protein levels. To investigate whether the transcription of the phr gene of E. coli was temperature-dependent, total RNA samples were isolated from AB2463 cells cultured at various temperatures at different cell densities; these RNA samples served as templates for real-time quantitative RT–PCR analyses. The results revealed that the steady-state transcript levels of the phr gene were not significantly affected by cell density and growth temperature (two-way ANOVA, both P > 0.05, Fig. S3), indicating that the change in photolyase content in cells cultured at different temperatures is post-transcriptional in nature.

Fig. 2. Flash repair experiments of AB2480 (recA13, uvrA6) cells. The cells were cultured at 25 °C (A), 37 °C (B) and 42 °C (C) to the stationary phase, harvested and resuspended. Then, the cell suspensions were UV irradiated and flash-repaired or photoreactivated under white-light (WL). The culturable survival values of the cells after given various UV-C doses (j, UV), and flash-repair (D, UV + Flash) or maximal photoreactivation (s, UV + WL 1 h) were determined and plotted. Dose decrements (DD) are indicated by double arrows. Each plot consists of combined data from three independent experiments.

3.4. Growth temperature-dependent expression of the lacZ reporters under phr promoters Further information on photolyase expression was gained from the lacZ reporters under control of the phr promoters. In plasmid pLLac, a lacZ fragment with an intact Shine–Dalgarno sequence

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Fig. 3. Western blotting analyses of purified photolyase and the photolyase levels in AB2463 (recA13), AB1157 (‘‘wild type’’) and AB2480 (recA13, uvrA6) cells cultured at different temperatures. For purified photolyase, the amounts of protein loaded (ng) were indicated. For the cell samples, the lysate from 108 cells was loaded in each lane. Loading controls were not included because CFU calibrations were performed. The experiments were repeated at least three times, and representative results are shown.

but no promoter (362,435–365,541 of the E. coli MG1655 genome) was placed downstream of a full length ybgA-phr fragment (738,031–740,153 of the E. coli MG1655 genome), which contained the phr promoters [11] and the intact coding regions of ybgA and phr. This resulted in a phr-lacZ fusion protein in which the transcription of lacZ is under control of the phr promoters in tandem with ybgA and phr. The pLLac plasmid was transformed into DH5a cells [D(argF-lac)169, u80dlacZ58(M15), recA1], and the activity of b-galactosidase was monitored with the cells cultured at various temperatures at different cell densities. The activity unit was defined as 1010 times of the increment of the absorbance at 420 nm (the maximum absorbance wavelength of the product o-nitrophenol) per minute per CFU. The results in Fig. 4A reveal that there was no significant difference in b-galactosidase expression under control of the phr promoters at various growth temperatures, which is in agreement with the quantitative RT–PCR analyses (Section 3.3). Plasmid pSLac is similar to pLLac, but contains a shorter ybgAphr fragment (738,031–739,135 of the E. coli MG1655 genome). This fragment contains an incomplete phr gene that encodes only the N-terminal a/b domain of photolyase (residue 1–134) [29]. The downstream lacZ gene was placed in the reading frame of the incomplete phr gene, resulting in a phr-lacZ fusion protein that encodes a photolyase N-terminal a/b domain-LacZ fusion protein under control of the phr promoters. In contrast to the results obtained with pLLac, much lower b-galactosidase activity was detected in the cells with pSLac cultured at 37–42 °C in both the exponential and stationary phases (Fig. 4B). The b-galactosidase activity units of stationary cultures with pSLac were 25, 18 and 8 min1 at growth temperatures of 25 °C, 37 °C and 42 °C, respectively. Considering that the molar absorption coefficient of o-nitrophenol at 420 nm is 4500 M1 cm1, the turnover number of bgalactosidase is 0.4 mol min1 g1 [30], the molecular weight of the b-galactosidase monomer is 116 kDa, the total reaction volume is 1.7 ml, and there are 30 plasmids per cell, the numbers of the photolyase-LacZ fusion protein expressed by one plasmid at stationary phase were estimated to be approximately 408, 294 and 131 at growth temperatures of 25 °C, 37 °C and 42 °C, respectively. These values are comparable to the photolyase numbers per cell determined by the kinetic assay and flash repair technique. These results further corroborate that the photolyase content in E. coli

Fig. 4. The b-galactosidase activity of DH5a cells [D(argF-lac)169, u80dlacZ58(M15), recA1] with (A) plasmid pLLac, containing a transcriptional phr-lacZ fusion; or with (B) plasmid pSLac, expressing a photolyase N-terminal a/b domain-LacZ fusion protein under control of the phr promoters. The cells were grown at various temperatures (j, 25 °C; d, 37 °C; D, 42 °C) to different cell densities (CFU/ml) before activity determination. The basal b-galactosidase activity of DH5a was not detectable.

cells is reduced at higher growth temperatures in a manner that should be due to post-transcriptional regulation. The N-terminal a/b domain of photolyase is at least partially responsible for the post-transcriptional regulation. Furthermore, the regulation is unlikely to be due to decreased translational initiation efficiency of the phr gene at high growth temperatures because this may cause a polar effect on the translation of downstream gene(s) [31] and this effect was not observed in the experiments with plasmid pLLac (Fig. 4A). Thus, the regulation is most likely post-translational, and may be the result of factors such as thermal unfolding and/or proteolytic degradation at high growth temperatures. 3.5. The proteolytic susceptibility of E. coli photolyase E. coli photolyase might be degraded by specific proteolysis at high temperatures, thereby resulting in the lower photoreactivation efficiency. Indeed, several protease active sites were found in Anacystis nidulans photolyase, some of which were within the N-terminal a/b domain [32]. To investigate the proteolytic susceptibility of E. coli photolyase, C-terminal His-tagged photolyase in the two-electron-reduced form (E-FADH) or the apo form was digested using trypsin, chymotrypsin and proteinase K. The

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Fig. 5. The proteolytic susceptibility of E. coli photolyase. (A) C-terminal His-tagged photolyase in the two-electron-reduced form (E-FADH, re) or the apo form (apo) was digested using trypsin (T), chymotrypsin (C) and proteinase K (K) at 23 °C for 15 min. The fragments containing intact C-terminal His-tags were detected by Western blotting using an anti-6  His primary antibody. The bands produced by trypsin and chymotrypsin are indicated by numbers and letters, respectively. The control lane of undigested full-length photolyase is indicated by ‘‘PL’’. (B) The possible cleavage sites (indicated by arrows) of trypsin (T) and chymotrypsin (C) to produce the resulting fragments. For trypsin cleavage sites, the boxes indicate the fragments formed both in E-FADH and the apo form, the underlines indicate the fragments dominantly formed in the apo form and the overline indicates the fragment only formed in E-FADH. (C) The possible cleavage sites of trypsin are shown in the crystal structure of E. coli photolyase (Protein Data Bank ID code 1DNP) in stick representations (brown sites, cleaved both in E-FADH and the apo form; pink sites, dominantly cleaved in the apo form; yellow site, dominantly cleaved in the E-FADH form). The FAD cofactor is shown in green stick representation.

fragments with intact C-terminal His-tags were detected by Western blotting (Fig. 5A). Reaction of E-FADH with trypsin generated four dominant proteolytic fragments with intact C-terminal His-tags (bands 1, 2, 3 and 4) that migrated at 41, 33, 28 and 21 kDa, respectively. However, the digestion of apo-photolyase with trypsin yielded seven dominant fragments with intact Cterminal His-tags (bands 1⁄, 1, 2⁄, 2, 3⁄, 3 and 4⁄) that migrated at 45, 41, 38, 33, 31, 28 and 26 kDa, respectively. The digestion pattern of photolyase in the E-FADH or the apo form with chymotrypsin was also significantly different. The former generated five dominant fragments with intact C-terminal His-tags (bands a, b, c, d and e) that migrated at 48, 45, 38, 31 and 26 kDa, respectively; while the later generated six dominant fragments with intact C-terminal His-tags (bands b, a⁄, c, d, b⁄ and e) that migrated at 45, 41, 38, 31, 28 and 26 kDa, respectively. The possible cleavage sites to produce these fragments are shown in Fig. 5B in the sequence and Fig. 5C in the crystal structure of E. coli photolyase. Reaction of E-FADH or apo-photolyase with proteinase K yielded similar fragments, which might be due to the fact that highly reactive proteinase K could digest the two photolyase forms irrespective of their structural differences. The above results reveal that there are some sites with high proteolytic susceptibility in E. coli photolyase and that the apo form photolyase seems to have more cleavage sites, especially in the N-terminal a/b domain.

3.6. The thermal stability of E. coli photolyase in vitro Activity assays were performed in vitro using the E-FADH form photolyase incubated at various temperatures for various time intervals to repair the UV-C irradiated oligothymidylates [UV-oligo (dT)16]. The enzyme retained 80% activity after 10 min of incubation at 25 °C. However, only 30% and <5% activity remained after 1 min of incubation at 37 °C and 42 °C, respectively. The result indicates that the E-FADH form photolyase is thermally labile in vitro at temperatures P37 °C. Thermal stability of photolyase in the E-FADH form, the oxidized form (E-FADox) and the apo form were analyzed by differential scanning calorimetry. The thermograms of these photolyase forms are shown in Fig. 6. Apo-photolyase has one symmetric endothermic peak with a melting temperature (Tm) of 33 °C and the transition beginning at 25 °C. E-FADH has one endothermic peak with an apparent Tm of 48 °C and the transition beginning at 37 °C. E-FADox has two endothermic peaks. The higher peaks at 48 °C is attributed to holo-photolyase, while the lower peaks at 33 °C is attributed to apo-photolyase present in the solution. This indicated that E. coli photolyase binds FADox relatively weakly, but associates with FADH tightly; this finding is consistent with our previous observations [18,23]. All photolyase forms are thermal unstable P37 °C, and the apo form and the E-FADox form are

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Fig. 6. Differential scanning calorimetric thermograms of E. coli photolyase in different forms. The dotted trace, the apo form (apo); the solid trace, the twoelectron-reduced form (E-FADH, re); the dashed trace, the oxidized form (E-FADox, ox). The concentrations of samples were approximately 20 lM. The scan rates were 1 °C/min. The transition beginning temperatures of the apo form and the E-FADH form are indicated (25 °C and 37 °C, respectively).

even more unstable than the E-FADH form. Thus, the thermal unfolding of photolyase may be responsible for the impaired photoreactivation of E. coli cells grown at temperatures P37 °C. 3.7. Temperature-shift experiments Temperature-shift experiments were performed to verify whether thermal unfolding and/or proteolytic degradation of photolyase were responsible for the impaired photoreactivation of E. coli cells grown at higher temperatures. When stationary phase AB2463 cells grown at 25 °C were transferred to 42 °C, the photolyase activity was gradually lost during culturing (Fig. 7A). The progress was very slow, with approximately half of the enzyme inactivated after 4 h; after 16–24 h, the photoreactivation efficiency dropped to a level consistent with the stationary phase cells continuously grown at 42 °C. This results is in sharp contrast to the in vitro experiments, in which >95% photolyase activity was lost after only 1 min incubation at 42 °C (Section 3.6). Furthermore, in contrast to the stationary phase cells cultured at 42 °C (Fig. 3), the photolyase protein levels were only slightly decreased in the stationary cells transferred from 25 °C to 42 °C for 16–24 h, although most of the enzyme had been already inactivated (Fig. 7B). These results imply that the photolyase protein is relatively stable in vivo once it has gained its activity (i.e. associating with the two-electron-reduced flavin cofactor). Active photolyase may be protected from thermal unfolding and proteolysis by unknown factors. Indeed, a large-scale identification of protein– protein interactions in E. coli revealed that photolyase interacted with the chaperonin GroEL, which can help proteins properly fold [33]. The photoreactivation efficiency and the photolyase protein levels are reduced in stationary phase cells grown at 42 °C (Figs. 1, 3 and 7). This reduction may be due to the fact that newly synthesized apo-photolyase is more easily unfolded and more susceptible to proteolysis than active photolyase when cells are continuously grown at higher temperatures. It was intriguing that after transferring stationary AB2463 cells from 42 °C to 25 °C for up to 24 h, neither the photoreactivation efficiency nor the photolyase protein levels were increased (Fig. 7). Moreover, the steady-state phr transcript levels were not significantly changed after temperature shifting (data not shown). One possible explanation is that the translation of phr mRNA is shut down after several

Fig. 7. Temperature-shift experiments of AB2463 (recA13) cells. The cells were initially grown at 25 °C or 42 °C to stationary phase, then transferred to 42 °C or 25 °C and cultured for another 24 h. (A) The photolyase activity (shown in k1[E]) in the cells after the temperature-shift was determined by the photoreactivation kinetic assay under continuous illumination (j, 25 °C to 42 °C; h, 42 °C to 25 °C). Each point is represented by 3 data points, and the bars represent the standard deviations. (B) The photolyase levels in the cells after temperature-shift were analyzed by Western blotting. The lysate from 108 cells was loaded into each lane. Loading controls were not included because CFU calibrations were performed. The experiment was repeated three times, and a representative result is shown.

rounds of expression and cannot be reestablished even after the translated enzyme is inactivated or degraded. Nevertheless, when the exponential cells cultured at 42 °C were transferred to 25 °C and grown to stationary phase, the photoreactivation efficiency was almost the same as that of the stationary phase cells continuously grown at 25 °C. This result indicates that the newly divided cells regained the ability for photolyase expression.

4. Discussion In this study, we found that the photoreactivation efficiency of E. coli is significantly impaired at growth temperatures P37 °C (and a slight but not significant decrease tendency was observed at 30 °C). This result is reminiscent of the observation that the photoreactivation extent of summer coliform populations in wastewater is significantly lower than that of winter populations [6]. The phenomenon may be partially due to the different growth temperatures of coliforms in different seasons. Although summer temperatures are not always higher than 37 °C, photoreactivation of coliforms might be impaired at slightly lower growth temperatures in natural environments that are not ideal for the growth of coliforms. Nevertheless, other factors may contribute to the seasonal difference in photoreactivation, such as a higher fraction of exponential phase cells (which are more UV-sensitive) in summer coliform populations and higher natural exposure to UV light during the summer months. The lower photoreactivation efficiency in the cells cultured at temperatures P37 °C is due to the presence of fewer molecules

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of active photolyase. Western blotting experiments also revealed that total protein levels of photolyase are lower in cells cultured at temperatures P37 °C. However, we found that growth temperature does not affect the steady-state transcript level of the phr gene. Furthermore, the b-galactosidase activity of a lacZ-reporter under transcriptional control of the phr promoters is not affected by growth temperature, but lower expression levels of the photolyase N-terminal a/b domain-LacZ fusion protein were observed at high growth temperatures (37–42 °C). All of these results indicate that the regulation of photolyase levels at different growth temperatures is post-transcriptional, and most likely to be posttranslational in nature. This finding is reminiscent of the photolyase amplification in purA mutants during adenine deprivation, which was also due to a post-translational change [27]. Both results are consistent in that no specific transcriptional regulatory mechanism has been identified for the E. coli phr gene to date [1]. Thus, thermal unfolding and/or specific proteolysis of photolyase at high growth temperatures might represent the post-translational regulation mechanism. The calorimetry and activity assay revealed that the active form photolyase (E-FADH) unfolds at temperatures P37 °C in vitro. In contrast, the active photolyase resisted thermal unfolding and proteolysis in E. coli cells even after being transferred to 42 °C for a long period of time. These results suggest that some factors such as chaperons might protect active photolyase in vivo. Thus, only warming samples containing coliforms for a relatively short period of time (<4 h) before UV treatment may not efficiently reduce photoreactivation. Furthermore, most wastewater treatment plants operate at temperatures less than or equal to 35 °C, and it would require far too much energy to heat the wastewater to higher temperatures. Therefore, raising the growth temperature of coliforms by heating the wastewater to suppress photoreactivation may not be a good choice for wastewater treatment plants. Apo-photolyase is more thermolabile and unfolds at temperatures P25 °C in vitro. More susceptible cleavage sites were detected in apo-photolyase, particularly in the N-terminal domain. Thus, the lower protein levels in the cells cultured at temperatures P37 °C might be due to the thermal unfolding and proteolytic degradation of newly synthesized apo-photolyase, but not active photolyase, which associates with the reduced FAD cofactor. The calorimetric data also revealed that the oxidized form of photolyase (E-FADox) contained some apo-photolyase in solution due to its weaker binding affinity for the FAD cofactor. Thus, the E-FADox form photolyase is more unstable than the E-FADH form. If photolyase is oxidized in vivo by some reagents, photoreactivation will be more readily suppressed. This inference is in line with the report that combined peracetic acid and UV processes can reduce photoreactivation [34]. Although some factors may protect E. coli photolyase from thermal unfolding and proteolysis in vivo, the enzyme is not very stable at the optimal growth temperature of E. coli (37 °C). This prompts us to question why E. coli has not evolved a more stable photolyase. It may be due to the fact that the natural habitat of E. coli is the mammalian intestines, where both the optimum growth temperature and protection from UV irradiation are provided [35]. Moreover, the non-specific DNA binding ability of photolyase may have negative effects on cells with high metabolic activity. When E. coli cells grow vigorously in the intestines, decreasing the stability of photolyase can avoid the potential interference with other DNA-binding proteins and has little influence on DNA repair because there is no light. Nevertheless, when E. coli cells are outside the mammalian body, where the ambient temperature is generally lower and the levels of both UV irradiation and visible light are high, the enzyme is relatively stable. This allows the bacteria to repair UV lesions using photolyase with external light energy [35]. Even when the cells are in the conditions where the

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ambient temperature and the level of UV irradiation are both high, the high growth rates of the cells will provide enough survival for selection. Under these conditions, mutations can actually be advantageous for promoting the evolution of the bacteria. Hence, a more stable photolyase is not required for E. coli. The thermal instability of E. coli photolyase may be an ‘‘intrinsic factor’’ that regulates its level in vivo under different conditions. Although this regulation seems to be disadvantageous for wasting energy on gene transcription and protein synthesis, it would be more economical than evolving a more complex regulation system considering the small numbers of the enzyme per cell. Interestingly, some plant photolyases were also found to be thermally unstable [36,37]. Cryptochromes are homologs of photolyases, but do not possess repair activity. Instead, they are circadian genes that maintain the circadian rhythm of organisms [1,38,39]. Some cryptochromes are not stable under light, such as Drosophila cryptochrome and Arabidopsis cryptochrome 2, which can be degraded in a lightdependent manner [40,41]. The thermal instability of photolyases may provide a prototype for the evolution of a light labile cryptochromes. Indeed, our preliminary study has shown that replacement of one or several residues of E. coli photolyase can confer the property of light-dependent unfolding on the mutants. Stationary phase cells grown at 42 °C rarely express active photolyase again even after being transferred to 25 °C for a long period of time, implying that there are mechanisms to down-regulate the translation of phr mRNA. The phr gene can be co-transcribed with an upstream gene ybgA, whose function remains unclear, to form a polycistronic mRNA [11]. There is a possibility that the YbgA protein is translated together with photolyase, and this protein may interact with the mRNA as a translational inhibitor to close its expression. Thus, the expression of the mRNA will not be opened when photolyase is inactivated but YbgA is not. When the cell divides and a new cell cycle begins, new message will be transcribed and new photolyase molecules will be synthesized. This hypothesis needs to be verified by future research.

Acknowledgements The research was supported by the National High Technology Research and Development Program (‘‘863’’ Program: 2012AA02A708) and the National Natural Science Foundation of China. (30900243; 31170005). We are grateful to Yanwei Ding at University of Science and Technology of China for his assistance in the calorimetric experiments. We also thank Ronald Gehr at McGill University, Hsin-Hou Chang at Tzu-Chi University, Martin Byrdin at CEA Saclay and Joseph L. Alcorn at University of Texas Health Science Center at Houston for helpful advice and discussion.

Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jphotobiol.2015. 03.012.

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