Escherichia coli photolyase uses blue light to repair cyclobutane pyrimidine dimers which are formed upon irradiation of DNA with ultraviolet (UV) light. E. coli photolyase is a flavoenzyme which contains a flavin adenine dinucleolide (FAD) in its active site and a 5,10-methenyltetrahydrofolate (MTHF) as a light-harvesting pigment. In the isolated enzyme, the FAD cofactor is present as a stable neutral radical semiquinone (FADH(.)). In this paper, we investigate the interaction between photolyase and UV-damaged DNA by using resonance Raman and UV-vis spectroscopy. Substrate binding results in intensity changes and frequency shifts of the FADH(.) vibrations and also induces electrochromic shifts of the FADH(.) electronic transitions because of the substrate electric dipole moment. The intensity changes in the resonance Raman spectra can be largely explained by changes in the Raman excitation profiles because of the electrochromic shift. The size of the electrochromic shift suggests that the substrate binding geometry is similar to that of oxidized FAD in reconstituted photolyase. The frequency changes are partially a manifestation of the vibrational Stark effect induced by the substrate electric dipole moment but also because of small perturbations of the hydrogen-bonding environment of FADH(.) upon substrate binding. Furthermore, differences in the resonance Raman spectra of MTHF-containing photolyase and of an MTHF-less mutant suggests that MTHF may play a structural role in stabilizing the active site of photolyase while comparison to other flavoproteins indicates that the FAD cofactor has a strong hydrogen-bonding protein environment. Finally, we show that the electrochromic shift can be used as a direct method to measure photolyase-substrate binding kinetics.