The nature of the binding site of the quinone acceptor A1 in Photosystem I (PSI) is studied by modeling the protein and cofactor on the basis of structural data derived from the intermediate resolution 4 Å X-ray diffraction electron density map, the position and orientation of A1 as evaluated from EPR data, and the histidine ligation of P700 as deduced from mutation experiments. Several models are constructed within the degrees of freedom allowed by the experimental constraints. In all cases a close interaction between the A1 headgroup and the side chain of PsaA-Trp697 (PsaB-Trp677) is found. The model is compared to the known binding site of QA in bacterial reaction centers (bRC) in which a similar quinone−tryptophan arrangement has been established. The results are also compared for consistency with published magnetic resonance data. The influences of the protein environment on the semiquinone g-tensor and hyperfine couplings are considerably different in PSI and bRC. It is argued that this is mainly a result of differences in the hydrogen bonding to the protein, in the strength of the π−π interactions with the tryptophan, and in the protein induced asymmetry in the spin density of the respective quinone radical anion.
A clear picture: In situ EPR and FTIR spectroscopic studies on the soluble, NAD+-reducing [NiFe]-hydrogenase of Ralstonia eutropha reveal that the catalytic site resides predominantly in the intermediate Nia-C state within whole cells. This state, can either be reversibly oxidized to a "Nir-B"-like state or further reduced to various Nia-SR species. The data suggest that the iron center in the active site contains a standard set (one CO and two CN−) of inorganic ligands.
An entry from the Cambridge Structural Database, the world’s repository for small molecule crystal structures. The entry contains experimental data from a crystal diffraction study. The deposited dataset for this entry is freely available from the CCDC and typically includes 3D coordinates, cell parameters, space group, experimental conditions and quality measures.
Covalent dimers, particularly pentacenes, are the dominant platform for developing a mechanistic understanding of intramolecular singlet fission (iSF). Numerous studies have demonstrated that a photoexcited singlet state in these structures can rapidly and efficiently undergo exciton multiplication to form a correlated pair of triplets within a single molecule, with potential applications from photovoltaics to quantum information science. One of the most significant barriers limiting such dimers is the fast recombination of the triplet pair, which prevents spatial separation and the formation of long-lived triplet states. There is an ever-growing need to develop general synthetic strategies to control the evolution of triplets following iSF and enhance their lifetime. Here, we rationally tune the dihedral angle and interchromophore separation between pairs of pentacenes in a systematic series of bridging units to facilitate triplet separation. Through a combination of transient optical and spin-resonance techniques, we demonstrate that torsion within the linker provides a simple synthetic handle to tune the fine balance between through-bond and through-space interchromophore couplings that steer iSF. We show that the full iSF pathway from femtosecond to microsecond timescales is tuned through the static coupling set by molecular design and structural fluctuations that can be biased through steric control. Our approach highlights a straightforward design principle to generate paramagnetic spin pair states with higher yields.
Pulsed ENDOR and HYSCORE measurements were carried out to characterize the active site of the oxygen-tolerant NAD+-reducing hydrogenase of Ralstonia eutropha. The catalytically active Nia-C state exhibits a bridging hydride between iron and nickel in the active site, which is photodissociated upon illumination. Its hyperfine coupling is comparable to that of standard hydrogenases. In addition, a histidine residue could be identified, which shows hyperfine and nuclear quadrupole parameters in significant variance from comparable histidine residues that are conserved in standard [NiFe] hydrogenases, and might be related to the O2 tolerance of the enzyme.
Significance Pairs of spins in molecular materials have attracted significant interest as intermediates in photovoltaic devices and light-emitting diodes. However, isolating the local spin and electronic environments of such intermediates has proved challenging due to the complex structures in which they reside. Here we show how exchange coupling can be used to select and characterize multiple coexisting pairs, enabling joint measurement of their exchange interactions and optical profiles. We apply this to spin-1 pairs formed by photon absorption whose coupling gives rise to total-spin S=0,1 and 2-pair configurations with drastically different properties. This presents a way of identifying the molecular conformations involved in spin-pair processes and generating design rules for more effective use of interacting spins.
Cryptochromes are blue light-sensing photoreceptors found in plants, animals, and humans. They are known to play key roles in the regulation of the circadian clock and in development. However, despite striking structural similarities to photolyase DNA repair enzymes, cryptochromes do not repair double-stranded DNA, and their mechanism of action is unknown. Recently, a blue light-dependent intramolecular electron transfer to the excited state flavin was characterized and proposed as the primary mechanism of light activation. The resulting formation of a stable neutral flavin semiquinone intermediate enables the photoreceptor to absorb green/yellow light (500–630 nm) in addition to blue light in vitro. Here, we demonstrate that Arabidopsis cryptochrome activation by blue light can be inhibited by green light in vivo consistent with a change of the cofactor redox state. We further characterize light-dependent changes in the cryptochrome1 (cry1) protein in living cells, which match photoreduction of the purified cry1 in vitro. These experiments were performed using fluorescence absorption/emission and EPR on whole cells and thereby represent one of the few examples of the active state of a known photoreceptor being monitored in vivo. These results indicate that cry1 activation via blue light initiates formation of a flavosemiquinone signaling state that can be converted by green light to an inactive form. In summary, cryptochrome activation via flavin photoreduction is a reversible mechanism novel to blue light photoreceptors. This photocycle may have adaptive significance for sensing the quality of the light environment in multiple organisms. Cryptochromes are blue light-sensing photoreceptors found in plants, animals, and humans. They are known to play key roles in the regulation of the circadian clock and in development. However, despite striking structural similarities to photolyase DNA repair enzymes, cryptochromes do not repair double-stranded DNA, and their mechanism of action is unknown. Recently, a blue light-dependent intramolecular electron transfer to the excited state flavin was characterized and proposed as the primary mechanism of light activation. The resulting formation of a stable neutral flavin semiquinone intermediate enables the photoreceptor to absorb green/yellow light (500–630 nm) in addition to blue light in vitro. Here, we demonstrate that Arabidopsis cryptochrome activation by blue light can be inhibited by green light in vivo consistent with a change of the cofactor redox state. We further characterize light-dependent changes in the cryptochrome1 (cry1) protein in living cells, which match photoreduction of the purified cry1 in vitro. These experiments were performed using fluorescence absorption/emission and EPR on whole cells and thereby represent one of the few examples of the active state of a known photoreceptor being monitored in vivo. These results indicate that cry1 activation via blue light initiates formation of a flavosemiquinone signaling state that can be converted by green light to an inactive form. In summary, cryptochrome activation via flavin photoreduction is a reversible mechanism novel to blue light photoreceptors. This photocycle may have adaptive significance for sensing the quality of the light environment in multiple organisms. Cryptochromes are flavin-type blue light photoreceptors found in many organisms including bacteria, plants, animals, and humans (1Banerjee R. Batschauer A. Planta. 2005; 220: 498-502Crossref PubMed Scopus (78) Google Scholar, 2Lin C. Todo T. Genome Biol. 2005; 6: 220-229Crossref PubMed Scopus (274) Google Scholar, 3Briggs W.R. Schäfer E. Nagy F. 3rd Ed. Photomorphogenesis in Plants and Bacteria. Springer, Dordrecht, Netherlands2006: 171-197Google Scholar). They mediate a variety of blue light-dependent responses including photomorphogenesis and growth responses in plants and entrainment of the circadian clock in animals such as Drosophila and mouse (4Li Q.-H. Yang H.-Q. Photochem. Photobiol. 2006; (in press)Google Scholar, 5Van Gelder R.N. J. Biol. Rhythms. 2002; 17: 110-120Crossref PubMed Scopus (32) Google Scholar, 6Sancar A. J. Biol. Chem. 2004; 279: 34079-34082Abstract Full Text Full Text PDF PubMed Scopus (112) Google Scholar). The common feature of cryptochrome photoreceptors is their marked homology to photolyase DNA repair enzymes, such that the primary amino acid sequence homology of cryptochromes to photolyases is in many instances as high as the homology between photolyases from different species to each other (7Sancar A. Chem. Rev. 2003; 103: 2203-2237Crossref PubMed Scopus (1042) Google Scholar, 8Todo T. Mutat. Res. 1999; 434: 89-97Crossref PubMed Scopus (153) Google Scholar). Like photolyases, cryptochromes bind flavin chromophores, and x-ray crystallographic analysis indicates marked structural similarity to photolyases, particularly in the flavin-binding pocket (9Brudler R. Hitomi K. Daiyasu H. Toh H. Kucho K. Ishiura M. Kanehisa M. Roberts V.A. Todo T. Trainer A. Getzoff E.D. Mol. Cell. 2003; 11: 59-67Abstract Full Text Full Text PDF PubMed Scopus (271) Google Scholar, 10Brautigam C.A. Smith B.S. Ma Z. Palnitkar M. Tomchick D.R. Machius M. Deisenhofer J. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 12142-12147Crossref PubMed Scopus (247) Google Scholar). Cryptochromes are divided into three subclasses: animal, plant, and DASH 3The abbreviations used are: DASH, Drosophila, Arabidopsis, Synechocystis, human; CPD, cyclobutane-pyrimidine dimer; FAD, flavin adenine dinucleotide. cryptochromes (2Lin C. Todo T. Genome Biol. 2005; 6: 220-229Crossref PubMed Scopus (274) Google Scholar). However, cryptochromes differ from photolyases in that they generally show no DNA repair activity. Only single-stranded DNA repair activity in vitro was shown recently for DASH cryptochromes (11Selby C.P. Sancar A. Proc. Natl. Acad. Sci. U. S. A. 2006; 103: 17696-17700Crossref PubMed Scopus (247) Google Scholar). Furthermore, they generally contain C-terminal extensions of variable sizes, which may be important for protein-protein interactions with signaling partners. In Arabidopsis, signaling partners include the regulatory protein COP1, an E3 ubiquitin-protein ligase that plays a key role in photomorphogenesis and development. This protein is known to interact to the C-terminal extensions of both cry1 and cry2 (12Wang H. Ma L.-G. Li J.-M. Zhao H.-Y. Deng X.W. Science. 2001; 294: 154-158Crossref PubMed Scopus (399) Google Scholar, 13Yang H-Q. Tang R.-H. Cashmore A.R. Plant Cell. 2001; 13: 2573-2587Crossref PubMed Scopus (304) Google Scholar). In animals such as Drosophila and mouse, cryptochromes interact with components of the circadian clock (14Green C.B. Curr. Biol. 2004; 14: R847-R849Abstract Full Text Full Text PDF PubMed Scopus (19) Google Scholar). Experiments probing the function of particular domains of cryptochrome receptors suggest that light initiates a conformational change in the protein permitting interaction with downstream signaling partners (15Yang H.-Q. Wu Y.-J. Tang R.-H. Liu D. Liu Y. Cashmore A.R. Cell. 2000; 103: 815-827Abstract Full Text Full Text PDF PubMed Scopus (332) Google Scholar). Direct evidence for light-induced conformational change has been recently obtained with purified Arabidopsis cry1 both by partial proteolysis (16Partch C.L. Clarkson M.W. Ozgur S. Lee A.L. Sancar A. Biochemistry. 2005; 44: 3795-3805Crossref PubMed Scopus (156) Google Scholar) and Fourier transform infrared techniques (17Kottke T. Batschauer A. Ahmad M. Heberle J. Biochemistry. 2006; 45: 2472-2479Crossref PubMed Scopus (99) Google Scholar). However, the primary photoreaction of cryptochromes, as well as the means whereby the light signal is transduced into a signaling cascade by the photoreceptor, has remained a matter of debate (18Partch C. Sancar A. Photochem. Photobiol. 2005; 81: 1291-1304Crossref PubMed Scopus (105) Google Scholar). Given the marked similarity of cryptochromes to photolyases, it is reasonable to assume that their primary photoreactions may be related. The generally accepted mechanism of DNA repair in photolyases (19Carell T. Burgdorf L.T. Kundu L.M. Cichon M. Curr. Opin. Chem. Biol. 2001; 5: 491-498Crossref PubMed Scopus (147) Google Scholar, 20Weber S. Biochim. Biophys. Acta. 2005; 1707: 1-23Crossref PubMed Scopus (293) Google Scholar) involves light-dependent electron transfer from fully reduced flavin to either cyclobutane-pyrimidine dimers (CPD, in the case of CPD photolyases) or to 6-4 photoproducts (in the case of 6-4 photolyases) in UV-damaged double-stranded DNA. The resulting reaction is catalytic, in that the DNA lesion is repaired and the electron subsequently returns to the flavin cofactor, restoring fully reduced flavin. Because this photoreaction does not occur to any significant extent in most cryptochromes (1Banerjee R. Batschauer A. Planta. 2005; 220: 498-502Crossref PubMed Scopus (78) Google Scholar, 2Lin C. Todo T. Genome Biol. 2005; 6: 220-229Crossref PubMed Scopus (274) Google Scholar, 3Briggs W.R. Schäfer E. Nagy F. 3rd Ed. Photomorphogenesis in Plants and Bacteria. Springer, Dordrecht, Netherlands2006: 171-197Google Scholar), it cannot be involved in blue light signaling. However, in addition to DNA repair, photolyases are capable of undergoing another photoreaction known as "photoactivation," which has been demonstrated in purified preparations of photolyases in which the flavin cofactor is not in its catalytically active, fully reduced, redox state (20Weber S. Biochim. Biophys. Acta. 2005; 1707: 1-23Crossref PubMed Scopus (293) Google Scholar, 21Byrdin M. Sartora V. Eker A.P. Vos M.H. Aubert C. Brettel K. Mathis P. Biochim. Biophys. Acta. 2004; 1655: 64-70Crossref PubMed Scopus (75) Google Scholar). In this reaction, light induces electron transfer to the excited flavin through a chain of amino acids (tryptophan or tyrosine residues) from the surface of the protein, thereby reducing the flavin. This reaction occurs both in classical type I CPD photolyases (to which plant cryptochromes are most closely related) and in 6-4 photolyases (to which animal type cryptochromes are more closely related) (21Byrdin M. Sartora V. Eker A.P. Vos M.H. Aubert C. Brettel K. Mathis P. Biochim. Biophys. Acta. 2004; 1655: 64-70Crossref PubMed Scopus (75) Google Scholar). Because photolyases contain mostly fully reduced flavin in living cells (22Kavakli I.H. Sancar A. Biochemistry. 2004; 43: 15103-15110Crossref PubMed Scopus (73) Google Scholar), this photoreaction is not important for DNA repair. However, because the Trp residues forming the intramolecular electron transfer are conserved in most cryptochrome sequences that are known currently, the question has arisen as to whether this photoreaction may be important for cryptochrome signaling. Evidence favoring this possibility has come from a number of experimental approaches; action spectroscopy for cry1-dependent plant hypocotyl growth inhibition has revealed a wavelength sensitivity (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar) consistent with oxidized flavin (peak absorption at 450 nm) as the active photopigment in vivo, rather than reduced flavin, as is the case for photolyases. Experiments with purified cry1 protein showed that photoreduction of oxidized flavin occurs in vitro (24Lin C. Robertson D.E. Ahmad M. Raibekas A.A. Schuman Jornes M. Dutton P.L. Cashmore A.R. Science. 1995; 269: 968-970Crossref PubMed Scopus (377) Google Scholar), and the relative stability of the flavosemiquinone intermediate was noted. Recently, a chain of intramolecular electron transfer events to the excited state flavin subsequent to light activation of purified cry1 protein (25Giovani B. Byrdin M. Ahmad M. Brettel K. Nat. Struct. Biol. 2003; 10: 489-490Crossref PubMed Scopus (242) Google Scholar) has been documented. Because such reactions involving light-driven electron transfer chains are rare in biological systems, these data provided support for a functional role of this pathway. Further evidence of a biological role for photoreduction in the activation of cryptochromes was provided by the observation that single amino acid substitution of tryptophan residues necessary for this reaction in vitro results in virtual loss of biological function in vivo (26Zeugner A. Byrdin M. Bouly J.P. Bakrim N. Giovani B. Brettel K. Ahmad M. J. Biol. Chem. 2005; 280: 19437-19440Abstract Full Text Full Text PDF PubMed Scopus (138) Google Scholar). Finally, magnetic effects on cryptochrome-dependent growth responses have been identified in Arabidopsis, which could only result from a mechanism of photoreceptor activation involving radical pair formation (27Ahmad M. Galland P. Ritz T. Wiltschko R. Wiltschko W. Planta. 2006; 224: 995-1003Crossref PubMed Scopus (93) Google Scholar), as occurs in the course of light-driven electron transfer. Therefore, light-mediated electron transfer leading to flavin photoreduction has been proposed as the primary signaling reaction for cryptochrome photoreceptors in vivo. In the present work, we have provided conclusive evidence for this mechanism by demonstrating that flavin photoreduction follows light activation of Arabidopsis cry1 in vivo using a combination of physiological, spectroscopic, and biophysical techniques. We show that cryptochrome activation by blue light in Arabidopsis is inhibited by co-irradiation with green light in a manner consistent with the formation of a long-lived neutral flavin radical (green-light absorbing) intermediate. Light-dependent changes in the cry1 protein are then characterized directly in living insect cells expressing high levels of recombinant cry1. Whole cell fluorescence techniques are used to measure the decrease of oxidized flavin associated with cry1 following light activation in these cells. The resulting accumulation of a metastable semiquinone intermediate is demonstrated by EPR using this same whole cell expression assay. Finally, we demonstrate interconversion between redox forms of purified cry1 and show that the effect of green light in vitro corresponds to the redox changes observed in vivo. The present work also represents one of the few studies, other than of the classic plant phytochromes (28Smith H. Nature. 2000; 407: 589Google Scholar) or in rhodopsins (29Spudich J.L. Bogomolni R.A. Nature. 1984; 312: 509-513Crossref PubMed Scopus (251) Google Scholar), where photoreversible changes in a known photoreceptor have been determined within living cells. Plant Material, Light Treatments, and Growth Conditions—Seeds of Arabidopsis thaliana of the indicated genotypes (see legends for Figs. 1 and 2) were sterilized and sown on 0.5× MS salts as described (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar). After 48 h at 277 K for inbibition, plates were transferred to white light for a further 48 h until radicle emergence to ensure synchronous germination (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar). Plates were subsequently transferred to the test conditions, and hypocotyl length was determined from an average of at least 20 seedlings/test condition. Anthocyanin levels were determined as described (30Ahmad M. Lin C. Cashmore A.R. Plant J. 1995; 8: 653-658Crossref PubMed Scopus (179) Google Scholar) for 30 seedlings at a time according to the formula A530–0.25 × A657 of the extracted pigments. Interference filters generating monochromatic light in all experiments were from Schott Industries or Corion Co.FIGURE 2Antagonistic effect of green light on rapid cryptochrome responses. Seedlings for all experiments were germinated as described (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar) and returned to the dark for an additional 48 h prior to light treatments (etiolated seedlings). Hypocotyl lengths prior to irradiations were between 0.6 and 0.8 mm. a, time course of cry2 degradation in 4 μmol m–2s–1 blue (450 ± 10 nm); bichromatic blue + green (blue 20 μmol m–2s–1, green (582 ± 10 nm)); or blue + red (blue 25 μmol m–2s–1, red (667 ± 10 nm)). b, inhibition of cry2 degradation by multiple wavelengths of green light. Seedlings were irradiated for 30 min either with blue light (2 μmol m–2s–1) alone or simultaneously with the indicated wavelength (±10 nm) at 20 μmol m–2s–1. c, fluence dependence of inhibition of cry2 degradation. Irradiation was performed for 30 min with blue light (4 μmol m–2s–1) and green (582 ± 10 nm) light at 3, 10, and 60 μmol m–2s–1. d, pulse experiment. Etiolated seedlings were exposed to eight cycles of 3-min blue light (10 μmol m–2s–1) pulses followed by 3-min dark (blue pulse) or 3-min green (559 nm, 30 μmol m–2s–1) given monochromatically in between blue light pulses (blue + green pulse). cblue is continuous blue light (48 min). a.u., arbitrary units; D, dark; B, blue light; G, green light; R, red light.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Western Blot and Protein Analysis—Protein extraction from seedlings and Western blot analysis with anti-cry2 antibody was performed as described (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar, 31Ahmad M. Jarillo J. Cashmore A.R. Plant Cell. 1998; 10: 197-208Crossref PubMed Scopus (142) Google Scholar). Seedlings were harvested and immediately ground and boiled in SDS sample loading buffer. Prior to running protein gels, protein concentration was quantified by Bio-Rad protein assay to ensure equal loading of wells. The load was subsequently verified after running of the gel and transfer to membrane, either by staining the blots (Fig. 2, a–c) or by reprobing the blots with anti-DnaK antibody (Fig. 2d) (32Neumann D. Emmermann M. Thierfelder J.M. zur Nieden U. Clericus M. Braun H.P. Nover L. Schmitz U.K. Planta. 1993; 190: 32-43Crossref PubMed Scopus (42) Google Scholar). Bands were quantified from scanned photographic images using Quantity One imaging software from Bio-Rad. All experiments were repeated in a minimum of three independent trials with qualitatively similar results. Whole Cell Fluorescence Emission Experiments—Living Sf21 insect cells expressing cry1 protein or uninfected controls were centrifuged from culture medium, resuspended in phosphate-buffered saline (pH 7.4), and placed directly into cuvettes at 283 K for measurement of fluorescence spectra. Fluorescence emission at 525 nm was monitored in a Varian fluorescence spectrophotometer over a range of excitation wavelengths or at a single designated wavelength as indicated (see Fig. 3 legend). Excitation and/or emission spectra were always determined in parallel both for infected (cry1-expressing) and uninfected cell cultures at identical cell density. For light treatments, samples were removed from the spectrophotometer and placed on ice. Illumination was carried on for the indicated times using the designated interference filters placed before a slide projector to provide sufficient light intensity. Samples were then returned to the fluorescence spectrophotometer to monitor differences in excitation and emission spectra. All experiments were repeated for a minimum of five independent trials with qualitatively similar results. UV-visible Spectroscopy—In vivo absorption difference spectra of cry1 were determined directly from living insect cells in an Uvikon 930 spectrophotometer. For these experiments absorption spectra of whole cells expressing cry1 protein were taken at 283 K and set as blank. Cells from the identical culture were subsequently transferred to ice and irradiated for 10 min with blue light at the indicated wavelength and light intensity. Samples were returned to the spectrophotometer for measurement and absorption spectra were taken with reference to the initial time point (t0). Difference spectra plotted were the average of 11 such measurements, obtained independently on the identical cell cultures. For spectra obtained from purified protein, A. thaliana cry1 or A. thaliana CPD photolyase were purified by established techniques. A. thaliana cry1 and CPD photolyase were then transferred into buffer (0.3 m NaCl, 0.05 m sodium phosphate (pH 8.0), 20% (v/v) glycerol, and 2 mm dithiothreitol) at identical concentrations estimated from FAD absorbance at 450 nm (ϵ450 = 1.12 × 104 m–1cm–1). Optical spectroscopy was carried out at 290 K at the indicated times and light treatments. EPR Spectroscopy—X-band cw-EPR spectra were recorded using a pulsed EPR spectrometer (Bruker Elexsys E580) with a cavity resonator (Bruker SHQE-4122-W1) and helium cryostat (Oxford CF-910). X-band pulsed ENDOR spectra were recorded on the same spectrometer using an ENDOR accessory (Bruker E560-DP), an rf amplifier (Amplifier Research 250A250A), and a dielectric-ring ENDOR resonator (Bruker EN4118X-MD-4W1) immersed in a helium gas flow cryostat (Oxford CF-935). The temperature was regulated to ±0.1 K by a temperature controller (Oxford ITC-503S). The cw-EPR spectra were recorded at 120 K with a microwave power of 3.0 microwatts, at 9.38 GHz microwave frequency with field modulation amplitude of 0.3 millitesla (at 100 kHz modulation frequency). For Davies-type ENDOR spectroscopy, a microwave pulse sequence π -T-π /2-τ-π with 64 and 128 ns π /2 and π pulses, respectively, and a RF pulse of 10-μs duration starting 1 μs after the first microwave pulse were used. The pulse separations T and τ between the microwave pulses were selected to be 13 μs and 500 ns, respectively. To avoid saturation effects due to long relaxation times, the entire pulse pattern was repeated with a low repetition frequency of 200 Hz. Spectra were taken at a magnetic field of 345.7 millitorrs and a microwave frequency of 9.71 GHz. Sf21 insect cells expressing N-terminal His-tagged cry1 and control Sf21 cells were resuspended in phosphate-buffered saline supplemented with 50% (v/v) glycerol in the dark. Aliquots were transferred into EPR quartz tubes (3 mm inner diameter) and illuminated for different times at 290 K with blue light (Halolux 30HL, Streppel, Wermelskirchen-Tente, Germany) using a 420–470-nm band filter (Schott, Mainz, Germany). Samples were then frozen rapidly under illumination in liquid nitrogen and stored therein. Cryptochrome responds primarily to UVA/blue light (peak near 450 nm) and only weakly above 500 nm, consistent with oxidized flavin as a primary photosensor (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar, 33Lin C. Ahmad M. Gordon D. Cashmore A.R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 8423-8427Crossref PubMed Scopus (163) Google Scholar). If the primary photoreaction of cryptochrome involves photoreduction of the flavin to the radical state (24Lin C. Robertson D.E. Ahmad M. Raibekas A.A. Schuman Jornes M. Dutton P.L. Cashmore A.R. Science. 1995; 269: 968-970Crossref PubMed Scopus (377) Google Scholar, 25Giovani B. Byrdin M. Ahmad M. Brettel K. Nat. Struct. Biol. 2003; 10: 489-490Crossref PubMed Scopus (242) Google Scholar, 26Zeugner A. Byrdin M. Bouly J.P. Bakrim N. Giovani B. Brettel K. Ahmad M. J. Biol. Chem. 2005; 280: 19437-19440Abstract Full Text Full Text PDF PubMed Scopus (138) Google Scholar), it follows that activation of the (oxidized flavin-containing) photoreceptor by blue light should lead to the formation of a semiquinone intermediate that can efficiently absorb green (500–600 nm) light. It further follows that bichromatic irradiations (green light given simultaneously with blue light) should decrease the responsiveness of cry1 because the added green light would decrease (by formation of fully reduced flavin, which absorbs only weak in the visible range between 400–500 nm) the levels of active semiquinone intermediate. To test this mechanism, blue light-dependent hypocotyl growth inhibition, which is largely cry1-dependent in Arabidopsis (23Ahmad M. Grancher N. Heil M. Black R.C. Giovani B. Galland P. Lardemer D. Plant Physiol. 2002; 129: 774-785Crossref PubMed Scopus (172) Google Scholar), was investigated under conditions of both monochromatic (blue light) and bichromatic (blue plus green light) irradiation. Consistent with the proposed mechanism, hypocotyl growth inhibition was significantly more pronounced in blue light (472 nm) than in seedlings co-irradiated with green light (564 nm) under the identical blue light intensity (Fig. 1a). Green light irradiation by itself under these conditions resulted in little difference in hypocotyl growth as compared with dark grown seedlings, in contrast to short-term illumination conditions (34Folta K.M. Plant Physiol. 2004; 135: 1407-1416Crossref PubMed Scopus (156) Google Scholar) or broad bandwidth green light (28Smith H. Nature. 2000; 407: 589Google Scholar). In phytochrome-deficient phyAphyB mutants, green light also acts antagonistically to blue light but not in cryptochrome-deficient cry1cry2 double mutant seedlings (Fig. 1a). Therefore the antagonistic effect of green light on hypocotyl growth inhibition requires cry1 and/or cry2. Qualitatively similar results (antagonistic effect of green light) were obtained for anthocyanin accumulation, which is also largely under the control of cry1 in blue light (Fig. 1b). In nature, plants under a canopy are exposed to light enriched in green wavelengths (35Franklin K.A. Whitelam G.C. Ann. Bot. (Lond.). 2005; 96: 169-175Crossref PubMed Scopus (392) Google Scholar, 36Vandenbussche F. Pierik R. Millenaar F.F. Voesenek L.A. van der Straeten D. Curr. Opin. Plant Biol. 2005; 8: 462-468Crossref PubMed Scopus (198) Google Scholar). Therefore, antagonistic effects of green and blue light might have adaptive significance, leading to increased elongation growth (decreased hypocotyl growth inhibition) as a response to shading. We show that for hypocotyl growth inhibition (Fig. 1c), green light also acts antagonistically to white light and that this response is cryptochrome-mediated, thereby suggesting adaptive significance in analogy to the classic phytochrome-dependent shade avoidance response mediated by phyB-D-E (35Franklin K.A. Whitelam G.C. Ann. Bot. (Lond.). 2005; 96: 169-175Crossref PubMed Scopus (392) Google Scholar, 36Vandenbussche F. Pierik R. Millenaar F.F. Voesenek L.A. van der Straeten D. Curr. Opin. Plant Biol. 2005; 8: 462-468Crossref PubMed Scopus (198) Google Scholar). Blue light-mediated degradation of cry2 protein (31Ahmad M. Jarillo J. Cashmore A.R. Plant Cell. 1998; 10: 197-208Crossref PubMed Scopus (142) Google Scholar, 37Lin C. Yang H. Guo H. Mockler T. Chen J. Cashmore A.R. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 2686-2690Crossref PubMed Scopus (395) Google Scholar) is a rapid (within minutes) and direct assay for photoreceptor activation. Etiolated seedlings accumulate high levels of cry2 protein in the dark, which decline rapidly subsequent to blue light (B) irradiation (Fig. 2a). Bichromatic irradiation with blue light and 582 nm green light (Fig. 2a, B+G) caused significant reduction in the rate of cry2 degradation as compared with blue light control seedlings. Red light, which is not absorbed by the flavosemiquinone, was ineffective in retarding cry2 degradation (Fig. 2a, compare B+R with B+G). Bichromatic irradiation was then performed at wavelengths at which the flavosemiquinone can efficiently absorb light (531, 540, 567, and 591 nm). At these wavelengths of green light, responsiveness of cry1 to blue light was significantly reduced (Fig. 2b). Inhibition by 582 nm green light occurred most efficiently at 10 and 60 μmol m–2s–1, well above the blue light intensity used in these experiments (Fig. 2c). These data are consistent with a mechanism involving alteration of levels of an existing semiquinone pool. Finally, pulse experiments showed that the effects of blue and green light are separable in time, consistent with a long-lived semiqinone intermediate compatible with a role in signaling (Fig. 2d). Direct determination of the redox state of cry1 in living plants is difficult for technical reasons such as low levels of cry1 protein and high pigment background. We consequently adopted a whole cell approach (38Galland P. Toelle N. Planta. 2003; 217: 971-982Crossref PubMed Scopus (17) Google Scholar) using baculovirus-infected insect cells expressing cry1 protein to high levels. Oxidized flavin can be detected in these living cells by directly monitoring fluorescence emission at 525 nm subsequent to excitation at different wavelengths in a fluorimeter. The excitation spectrum is essentially similar to oxidized flavin (Fig. 3a), although light scattering and masking by other pigments in these living cells presumably causes the loss of fine structure as compared with purified cryptochrome in solution. When this assay was performed on insect cells expressing cry1 protein, emission at 525 nm was severalfold higher than in uninfected cells; this increased signal was generated by the large quantities of oxidized flavin bound to cry1protein in vivo (24Lin C. Robertson D.E. Ahmad M. Raibekas A.A. Schuman Jornes M. Dutton P.L. Cashmore A.R. Science. 1995; 269: 968-970Crossref PubMed Scopus (377) Google Scholar, 25Giovani B. Byrdin M. Ahmad M. Brettel K. Nat. Struct. Biol. 2003; 10: 489-490Crossref PubMed Scopus (242) Google Scholar) (Fig. 3a). The fact that cry1 in the
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