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Applied and Environmental Microbiology, January 2008, p. 327-328, Vol. 74, No. 1
0099-2240/08/$08.00+0     doi:10.1128/AEM.00013-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Effects of UV Radiation on Photolyase and Implications with Regards to Photoreactivation following Low- and Medium-Pressure UV Disinfection{triangledown}

Jiangyong Hu* and Puay Hoon Quek

Centre for Water Research, Division of Environmental Science and Engineering, National University of Singapore, 10 Kent Ridge Crescent, Singapore 119260

Received 3 January 2007/ Accepted 24 October 2007


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 REFERENCES
 
Photolyase activity following exposure to low-pressure (LP) and medium-pressure (MP) UV lamps was evaluated. MP UV irradiation resulted in a greater reduction in photolyase activity than LP UV radiation. The results suggest that oxidation of the flavin adenine dinucleotide in photolyase may have caused the decrease in activity.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
REFERENCES
 
UV disinfection inactivates microorganisms through the formation of cyclobutane pyrimidine dimers in their DNA (4). However, these cyclobutane pyrimidine dimers may be removed via photoreactivation, which allows inactivated microorganisms to recontaminate the water. Studies have found that photoreactivation of Escherichia coli following medium-pressure (MP) UV disinfection is lower than that following low-pressure (LP) UV disinfection (9, 14). A hypothesized explanation for this is damage to photolyase, because it contains a tryptophan-rich flavin adenine dinucleotide (FAD) cofactor that has a peak absorbance at 280 nm (4), a wavelength emitted by MP and not LP UV lamps (8). Nevertheless, there is currently no evidence to test the hypothesis, and research on this is limited. This study therefore aims to investigate the effects of LP and MP UV radiation on photolyase activity and to propose the mechanism for photoreactivation suppression by MP UV radiation reported previously.

Photolyase was extracted and purified from E. coli containing plasmid with the phr gene in accordance with the methods described by Sancar and Sancar (12). One hundred microliters of photolyase (1 x 10–7 to 5 x 10–7 M) was dispensed onto a microcentrifuge tube cap and placed under a collimated beam apparatus (Calgon Carbon Corporation) with interchangeable LP and MP UV lamps. The UV intensity of the lamps was measured with a radiometer (IL1400A; International Light, Inc.) with a SED240 sensor. UV doses were calculated by the method of Bolton and Linden (1) and applied by varying the exposure times. Photolyase activity was determined using a spectrophotometric assay modified from that developed by Jorns et al. (5). Dimer substrate was mixed with irradiated photolyase, incubated in the dark for 3 to 5 min, and then exposed to 365 nm light (9 W) at a distance of 5 cm. Absorption spectra (250 nm to 320 nm) were taken at 2-min intervals. In other experiments, 5 mM dithiothreitol (DTT) was added to the mixture to investigate whether oxidation of the FAD caused the decrease in photolyase activity, and absorption spectra were taken at 1-min intervals. The increase in absorbance at 260 nm was plotted against the time, and the gradient of the straight-line portion was used to calculate the rate of dimer repair by applying the Beer-Lambert law, with {varepsilon}260 of thymine monomers taken to be 8.3 x 103 M–1 cm–1 (5).

The calculated rates of dimer repair by LP and MP UV-irradiated photolyase based on absorbance changes in the substrate are summarized in Fig. 1. For LP UV-irradiated photolyase, the dimer repair rate was unaffected up to a dose of 10 mJ cm–2 (rates varied 3% between 0.467 and 0.480 M dimer M–1 photolyase min–1) and then decreased significantly such that the dimer repair rate at 40 mJ cm–2 was only 72% of that of unirradiated photolyase. This suggests that a minimum radiation threshold is required before the dimer repair ability of photolyase is affected; this is analogous to the multihit inactivation model (4). In contrast, the dimer repair rates of MP UV-irradiated photolyase decreased by 20% at a UV dose as low as 2 mJ cm–2 and remained relatively constant above 10 mJ cm–2. This may be due to the high absorption of photolyase at wavelengths such as 280 nm and 384 nm that are produced by MP UV lamps, which could have caused photolyase damage either structurally (irreversible) or via oxidation (reversible), thereby affecting its dimer repair ability.


Figure 1
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  		FIG. 1. Repair rates of photolyase exposed to various doses of LP and MP UV radiation. Error bars represent the standard deviations of three to five experiments for each experimental condition.

 
To investigate whether oxidative damage to photolyase resulted in the decrease in its dimer repair rates, experiments with DTT were conducted, and the results are presented in Fig. 2. DTT is a reducing agent used to stabilize enzymes and has been shown to restore activity to enzymes whose activities have been lost due to the oxidation of certain functional groups (3). With DTT addition, it was clear that dimer repair rates of photolyase were unaffected by UV radiation exposure up to 40 mJ cm–2. There was also no difference in the dimer repair rates of LP and MP UV-irradiated photolyase with addition of 5 mM DTT at 10 mJ cm–2 (1.815 ± 0.203 [117%] and 1.758 ± 0.151 [113%], respectively [values in brackets denote the rate of dimer repair as a percentage of that of unirradiated photolyase under the same experimental conditions]). In comparison to the repair rates without DTT addition at 10 mJ cm–2 (0.667 ± 0.089 [97%] and 0.446 ± 0.030 [65%] for LP and MP UV-irradiated photolyases, respectively), the addition of a reducing agent eradicated differences between the dimer repair rates of LP and MP UV-irradiated photolyases. This suggests that reversible oxidative damage to photolyase was the cause for decreased dimer repair rates. DTT addition also increased dimer repair rates by more than two times the rates without DTT addition, because it fully reduced the neutral flavin to the catalytically active state (11).


Figure 2
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  		FIG. 2. Repair rates of photolyase exposed to various doses of LP and MP UV radiation and chemically reduced by the addition of 5 mM DTT. Error bars represent the standard deviations of three to five experiments for each experimental condition.

 
The spectrophotometric assay used to measure the activity of photolyase in vitro is based on the principle that thymine monomers absorb strongly at 260 nm while dimers do not (5, 12). Therefore, control experiments conducted with photolyase or the substrate alone showed no increase in absorbance (data not shown), suggesting that dimer repair, instead of conformational changes in the substrate, was responsible for the increase in absorbance observed during the experiments.

This study shows that the dimer repair rates of photolyase decreased when exposed to both LP and MP UV radiation in vitro, an observation that differs from results of Oguma et al. (10), who did not detect any change in the activity of MP UV-exposed photolyase. This could be because of the higher dimer concentration in the substrate used in this study and the lack of interferences from other photoproducts, which allowed for more sensitive detection of photolyase activity.

Photolyase in vitro contains a neutral FAD while photolyase in vivo has a reduced FAD (2, 6, 13). As such, the results in the current study may not represent the actual situation during UV irradiation and photoreactivation inside the E. coli cell, which is unknown since in vivo studies on photolyase and photoreactivation are ongoing (7). Nevertheless, the results provide evidence in favor of the hypothesis that photolyase damage results in a reduced photoreactivation rate when both LP and MP UV radiation are applied and also suggest the mechanism by which the damage occurs in the in vitro system.


   ACKNOWLEDGMENTS
 
We thank Aziz Sancar (University of North Carolina, Chapel Hill) for his generous gift of the photolyase-overproducing plasmid pMS969S, and we thank his graduate student Sezgin Özgür for providing helpful comments and suggestions for the purification of photolyase. We also thank Seeram Ramakrishna (National University of Singapore) for the use of the protein purification system.


   FOOTNOTES
 
* Corresponding author. Mailing address: Division of Environmental Science and Engineering, Faculty of Engineering, National University of Singapore, 9 Engineering Drive 1, EA 07-23, Singapore 117576. Phone: (65)-65164540. Fax: (65)-67744202. E-mail: esehujy{at}nus.edu.sg Back

{triangledown} Published ahead of print on 2 November 2007. Back


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 REFERENCES
 

  1. Bolton, J. R., and K. G. Linden. 2003. Standardization of methods for fluence (UV dose) determination in bench-scale UV experiments. J. Environ. Eng. 129:209-215.[CrossRef]
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  5. Jorns, M. S., G. B. Sancar, and A. Sancar. 1985. Identification of oligothymidylates as new simple substrates for Escherichia coli DNA photolyase and their use in a rapid spectrophotometric enzyme assay. Biochemistry 24:1856-1861.[CrossRef][Medline]
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  8. Masschelein, W. J. 2002. Available lamp (or burner) technologies, p. 9-58. In R. G. Rice (ed.), Ultraviolet light in water and wastewater sanitation. Lewis Publishers, Boca Raton, FL.
  9. Oguma, K., H. Katayama, H. Mitani, T. Hirata, and S. Ohgaki. 2001. Determination of pyrimidine dimers in Escherichia coli and Cryptosporidium parvum in UV light inactivation, photoreactivation, and dark repair. Appl. Environ. Microbiol. 67:4630-4637.[Abstract/Free Full Text]
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  11. Payne, G., P. F. Heelis, B. R. Rohrs, and A. Sancar. 1987. The active form of Escherichia coli DNA photolyase contains a fully reduced flavin and not a flavin radical, both in vivo and in vitro. Biochemistry 26:7121-7127.[CrossRef][Medline]
  12. Sancar, G. B., and A. Sancar. 1988. DNA photolyase: purification and use, p. 461-478. In E. C. Friedberg and P. C. Hanawalt (ed.), DNA repair: a laboratory manual of research procedures, vol. 3. Marcel Dekker, Inc., New York, NY.
  13. Weber, S. 2005. Light-driven enzymatic catalysis of DNA repair: a review of recent biophysical studies on photolyase. Biochim. Biophys. Acta 1707:1-23.[Medline]
  14. Zimmer, J. L., and R. M. Slawson. 2002. Potential repair of Escherichia coli DNA following exposure to UV radiation from both medium- and low-pressure UV sources used in drinking water treatment. Appl. Environ. Microbiol. 68:3293-3299.[Abstract/Free Full Text]

Applied and Environmental Microbiology, January 2008, p. 327-328, Vol. 74, No. 1
0099-2240/08/$08.00+0     doi:10.1128/AEM.00013-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.




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