JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M609842200 on January 19, 2007

J. Biol. Chem., Vol. 282, Issue 13, 9383-9391, March 30, 2007
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
282/13/9383    most recent
M609842200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bouly, J.-P.
Right arrow Articles by Ahmad, M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bouly, J.-P.
Right arrow Articles by Ahmad, M.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Cryptochrome Blue Light Photoreceptors Are Activated through Interconversion of Flavin Redox States*Formula

Jean-Pierre Bouly{ddagger}, Erik Schleicher§, Maribel Dionisio-Sese1, Filip Vandenbussche{ddagger}||, Dominique Van Der Straeten**, Nadia Bakrim{ddagger}, Stefan Meier, Alfred Batschauer, Paul Galland, Robert Bittl§, and Margaret Ahmad{ddagger}**2

From the {ddagger}Université Paris VI, FRE-CNRS 2846, Casier 156, 4 Place Jussieu, 75005 Paris, France, §Freie Universität Berlin, Fachbereich Physik, Arnimallee 14, 14195 Berlin, Germany, FB Biologie-Pflanzenphysiologie, Philipps-Universität, Karl-von-Frisch-Strasse 8, 35032 Marburg, Germany, ||Unit Plant Hormone Signaling and Bio-imaging, Department of Molecular Genetics, Ghent University, Ledeganckstraat 35, B-9000 Ghent, Belgium, and **Pennsylvania State University, Media, Pennsylvania 19063

Received for publication, October 19, 2006 , and in revised form, January 19, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
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.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cryptochromes are flavin-type blue light photoreceptors found in many organisms including bacteria, plants, animals, and humans (1–3). 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 (4–6). 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 (7, 8). Like photolyases, cryptochromes bind flavin chromophores, and x-ray crystallographic analysis indicates marked structural similarity to photolyases, particularly in the flavin-binding pocket (9, 10). Cryptochromes are divided into three subclasses: animal, plant, and DASH3 cryptochromes (2). 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 (11). 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 (12, 13). In animals such as Drosophila and mouse, cryptochromes interact with components of the circadian clock (14). 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 (15). Direct evidence for light-induced conformational change has been recently obtained with purified Arabidopsis cry1 both by partial proteolysis (16) and Fourier transform infrared techniques (17). 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 (18).

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 (19, 20) 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 (1–3), 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 (20, 21). 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) (21). Because photolyases contain mostly fully reduced flavin in living cells (22), 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 (23) 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 (24), 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 (25) 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 (26). 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 (27), 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 (28) or in rhodopsins (29), where photoreversible changes in a known photoreceptor have been determined within living cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
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 0.5x MS salts as described (23). 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 (23). 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 (30) for 30 seedlings at a time according to the formula A530–0.25 x A657 of the extracted pigments. Interference filters generating monochromatic light in all experiments were from Schott Industries or Corion Co.

Western Blot and Protein Analysis—Protein extraction from seedlings and Western blot analysis with anti-cry2 antibody was performed as described (23, 31). 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) (32). 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 ({epsilon}450 = 1.12 x 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 {pi} -T-{pi} /2-{tau}-{pi} with 64 and 128 ns {pi} /2 and {pi} 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 {tau} 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.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
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 (23, 33). If the primary photoreaction of cryptochrome involves photoreduction of the flavin to the radical state (24–26), 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 (23), 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 (34) or broad bandwidth green light (28). 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 (35, 36). 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 (35, 36).

Blue light-mediated degradation of cry2 protein (31, 37) 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).


Figure 1
View larger version (47K):
[in this window]
[in a new window]

  		FIGURE 1.
Antagonistic effect of green light on cry1 activity. a, Arabidopsis seedlings (Ler background) of the indicated genotypes were sown and germinated as described (21) prior to transfer to either blue (472 ± 10 nm, 20 µmol m–2s–1) or blue supplemented with green (563 ± 10 nm, 10 µmol m–2s–1) bandwidth light. Hypocotyl lengths were measured after 3 days of light treatment. Error bars represent S.E. b, anthocyanin accumulation was determined essentially as described (31). Seedlings were germinated and placed for 48 h in blue light (445 ± 10 nm, 10 µmol m–2s–1) or supplemented with green (582 ± 10 nm, 20 µmol m–2s–1) light. Error bars represent the S.D. of three trials. c, seedlings of the indicated genotypes were germinated and placed under 25 µmol m–2s–1 white light (W; Philips "cool white" fluorescent tubes). Green light (G; 550 ± 25 nm) was at a fluence rate of 50 µmol m–2s–1. Error bars indicate S.D. Wt, wild type; Ws, Arabidopsis Wassilewskija.

 
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 (38) 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 (24, 25) (Fig. 3a). The fact that cry1 in these living cells is found with flavin in the oxidized (rather than reduced) state is consistent with data from action spectroscopy showing that oxidized flavin is bound to the photoreceptor in living plants (23).

To determine whether flavin associated with cry1 undergoes photoreduction in vivo, cry1-expressing cell cultures were irradiated with blue light and the excitation spectrum taken for fluorescence emission at 525 nm. The difference spectrum generated from these data matches light-dependent photoreduction of oxidized flavin as assessed by a decreased absorption at 450 nm over time (Fig. 3b). To identify this photoreaction more directly, absorbance changes were monitored. Living cells were placed in a spectrophotometer, and wavelength scans were taken before and after short periods of irradiation by light. The difference between scans before and after light treatment were plotted (difference absorption spectrum) (Fig. 3b, inset). A measurable cry-dependent light induced absorbance change was observed matching reduction of oxidized flavin. Taken together, these data represent the first in vivo difference spectrum generated for cryptochrome activation and match that of the purified protein in vitro.

Levels of semiquinone intermediate could not be determined directly by fluorescence techniques alone, because this method allows only the detection of the fully oxidized flavin. However, bichromatic irradiations with green light resulted in significant reduction in oxidized flavin content as compared with the effect of blue light alone in living cells (Fig. 3c). This effect of green light was not observed in uninfected control cells. These data are consistent with a shift of the flavin photoequilibrium in vivo by green light toward the fully reduced form of flavin, thereby reducing overall levels of both oxidized and semi-reduced forms of cry1. Such a shift could only occur if there was prior accumulation of the semi-reduced (green light-absorbing) form of the photoreceptor.


Figure 2
View larger version (39K):
[in this window]
[in a new window]

  		FIGURE 2.
Antagonistic effect of green light on rapid cryptochrome responses. Seedlings for all experiments were germinated as described (23) 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.

 
To directly demonstrate the accumulation of semireduced flavin in living cells, EPR was used for radical detection. In these experiments, intact cells expressing plant cry1 and uninfected control cells were irradiated in parallel with the identical intensity of blue light and rapidly frozen for EPR analysis. A paramagnetic species that does not accumulate in control cells was induced by blue light irradiation in cry1-expressing cells (Fig. 4, upper panel, B–D). This species was further characterized by X-band pulsed (Davies) ENDOR spectroscopy (Fig. 4, lower panel); the spectrum obtained with insect cells (H) was identical to that obtained for purified photoreduced cry1 protein (I). The spectra are similar to photolyase bound FADH•, thus confirming the photoinduced species in the whole cells as a neutral flavin radical. It is noteworthy that the spectra show distinct changes of the characteristic 8{alpha} hyperfine coupling compared with photolyase, reflecting subtle differences in the respective binding pockets. These data are comparable with similar experiments performed with insect cells overexpressing Arabidopsis cry2 protein.4

In past experiments with purified cry1 protein, only the effects of blue or white light on flavin photochemistry had been evaluated (24–26). Because these are absorbed by the oxidized form of cryptochrome in addition to the radical, we examined the effect of green light, which is specifically absorbed by the radical, in purified preparations of cry1 protein. Blue light irradiation of purified cry1 protein under anaerobic and reducing conditions resulted in the formation of the flavin radical, as described previously (24–26) (see Fig. 3d and supplemental Fig. 1A). Subsequent transfer to darkness resulted in slow reoxidation, presumably because of remaining oxygen in the cuvette. When green light was given subsequent to blue light, the rate of reoxidation of cry1 was significantly decreased as compared with dark controls (Fig. 3d), consistent with formation of fully reduced flavin and a concomitant shift in flavin state equilibrium. Irrespective of the likelihood that flavin reoxidation occurs via different mechanisms in vivo and in vitro, the effect of green light in vitro is a clear indication of reduced flavin (FADH–) formation in cry1. Because this fully reduced intermediate does not accumulate to high levels except under strong white light and is not detectable at all under aerobic conditions (not shown), it is either unstable and/or rapidly reoxidized in comparison with the other redox intermediates of cry1 (25, 26) (see also supplemental Fig. 1A). These data show that green light shifts the flavin redox state in a manner consistent with its effect on biological activity in vivo, supporting the mechanism for cry1 photoactivation presented in the model (Fig. 5a). Furthermore, fully reduced flavin is a short-lived intermediate that does not contribute significantly to the flavin photoequilibrium in either dark or light, in marked contrast to photolyases, where the fully reduced form of the enzyme predominates under these conditions (Fig. 5b) and is catalytically active in vivo (19, 20).


Figure 3
View larger version (39K):
[in this window]
[in a new window]

  		FIGURE 3.
Redox state of cry1 protein in vivo. Living Sf21 insect cells 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. a, fluorescence emission at 525 nm was monitored over a range of excitation wavelengths. Excitation spectra are presented for infected (cry1-expressing) and uninfected cell cultures at identical cell density. Increase in peak intensity at 450 nm is due to oxidized flavin associated with cry1 in infected cells. b, cry1-overexpressing insect cells were irradiated with blue light (445 ± 10 nm, 25 µmol m–2s–1), and excitation spectra were taken at the indicated intervals by monitoring emission at 450 nm. An in vivo difference spectrum was plotted by subtracting subsequent excitation spectra from the (dark) base-line spectrum. Peak excitation efficiency at 450 nm decreased as a function of time, consistent with photoreduction of oxidized flavin in vivo. Inset, absorption difference spectrum determined directly from living insect cells subjected to a 10-min blue light pulse. c, flavin photoreduction in living insect cells under conditions of bichromatic irradiation. Peak excitation efficiency at 450 nm was monitored in duplicate samples over the indicated time course. Samples were treated with either blue light at 25µmol m–2s–1 or bichromatically with blue + green (550 ± 25 nm, 150 µmol m–2s–1) light over the same interval. Deduced 450 nm peak intensity was plotted over time. d, green light effect on purified cry1. cry1 protein was photoreduced under anaerobic conditions. Once radical formation was established, absorbance was measured at 446 nm (oxidized flavin) and 552 nm (radical flavin) at 2-min intervals in either darkness (reoxidation of the protein, open symbols) or green light (552 nm, solid symbols). a.u., arbitrary units.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
We have provided evidence that the Arabidopsis cry1 photocycle consists of light-dependent transition between the three interconvertible redox forms of FAD, of which the flavosemiquinone form (FADH•) is the signaling state of the receptor. These data are consistent with prior studies in vitro showing an unusual stability of the flavosemiquinone form of cry1 (24, 25) and with studies implicating the electron transfer chain required for photoreduction as essential for biological activity (26). In this mechanism, inactive cry1, containing oxidized flavin, accumulates in the dark. Blue light triggers flavin photoreduction such that the resulting photoequilibrium strongly favors the radical signaling state. Green light reduces cryptochrome activity by reducing the concentration of the radical in favor of the inactive fully reduced form, which is subsequently reoxidized. In keeping with this proposed mechanism, the radical form of cry1 is far more stable than that of photolyases, for which we provide evidence that FAD is rapidly reduced to the fully reduced (catalytic) form under identical illumination conditions (Fig. 5c; see also supplemental Fig. 1). Thus, the cry1 photocycle differs fundamentally from that of its photolyase progenitors.


Figure 4
View larger version (27K):
[in this window]
[in a new window]

  		FIGURE 4.
X-band cw-EPR frozen-solution spectra of intact Sf21 cells expressing cry1 and Sf21 control cells. A–D, Sf21 insect cells expressing cry1 after different blue light illumination times. A, 0 min; B, 6 min; C, 12 min; D, 18 min; E–G, same amount of control Sf21 cells after different blue light illumination times. E, 0 min; F, 12 min; G, 18 min. All spectra were recorded at 120 K with a microwave power of 3.0 microwatts and a magnetic field modulation amplitude of 0.3 millitesla (modulation frequency, 100 kHz). X-band pulsed (Davies) ENDOR spectrum of sample C is shown in lane H. Lane I represents the X-band pulsed (Davies) ENDOR spectrum of purified cry1 protein. Both spectra were recorded at 80 K. a.u., arbitrary units.

 
In prior studies with cryptochrome-overexpressing plants, it was observed that they were somewhat hypersensitive to broad bandwidth (500–600 nm) green light, indicating a weak but measurable positive effect on cryptochrome activation (24, 28). These data appear to be in contradiction to the role of green light documented in the present study, where under bichromatic (blue plus green) light the effect of green light is to oppose cryptochrome activation by blue light. The reason for this seeming discrepancy became apparent when growth inhibition studies were performed under a series of narrow bandwidth green light filters (supplemental Fig. 2) instead of under broad bandwidth (500–600 nm) green light as in the prior studies. It was observed that only the wavelengths closest to 500 nm (518 nm) were effective in mediating growth inhibition. Longer wavelengths were progressively less effective (supplemental Fig. 2). The small positive effect of broad bandwidth green light on cryptochrome activation can therefore be fully explained by the weak residual absorption of oxidized flavin above 500 nm, in agreement with the present study, and could not result from activation by green light of the radical form of cryptochrome as suggested previously (24, 33).


Figure 5
View larger version (21K):
[in this window]
[in a new window]

  		FIGURE 5.
a, model of cry1 photocycle. cry1 exists in three interconvertible redox forms, FAD, FADH•, and FADH–. The FAD form is inactive and accumulates to high levels in the dark. Blue light triggers photoreduction of FAD to establish a photoequilibrium that favors FADH• over FAD or FADH–. The flavosemiquinone state is the signaling state of the receptor. Green light is absorbed by the radical and shifts the photoequilibrium to the fully reduced form (FADH–), which is inactive. Reversion to oxidized flavin occurs in the dark, involving electron acceptors that are as yet uncharacterized. Dark reversion occurs either from semiquinone flavin directly to oxidized flavin or from fully reduced flavin to the oxidized FAD. b, comparison between radical formation in photolyases and cry1. Purified A. thaliana CPD photolyase (47) and cry1 at the identical protein concentration (48) were subject to photoreduction under identical conditions (20 µmol m–2s–1 blue (400–500 nm) light, 2 mM dithiothreitol) and absorption spectra taken at the indicated intervals (see supplemental Fig. 1 for complete spectra). Building and decay kinetics of the FADH• absorption are presented at 550 nm for cry1 (open squares) and at 595 nm for photolyase (open circles). a.u., arbitrary units.

 
Although plants are poorly responsive to green light, there is recent evidence of a distinct green light photoreceptor that promotes hypocotyl growth under conditions of short-term irradiation (34). However, there was no effect of the novel green light photoreceptor on promoting hypocotyl growth under the long-term irradiation conditions used in our study (34, 39, 40) (see also supplemental Fig. 2) or on cry2 degradation (Fig. 2). The possibility of unrelated green light photoreceptors contributing to the green light inhibition of cryptochrome responses therefore appears remote.

Given its signaling role, it would be expected that the flavin radical form of cryptochrome would have a reasonable lifetime to interact with downstream signaling partners. Further experiments varying the time between successive blue and green light pulses should allow an estimate of this lifetime, in analogy to the "escape" time for photoreversibility of phytochrome responses by far red light (28). How the formation of such a long-lived radical initiates cryptochrome signaling has yet to be determined, but it likely involves changes in conformation necessary for interaction with signaling partners (12, 13, 15). Fourier transform infrared studies of cryptochrome photoreceptors upon light activation have revealed small but potentially significant structural changes that could lead to changes in surface properties of the receptor (17), and studies with partial proteolysis of purified protein also reveal conformational changes subsequent to light activation (16).

It is not clear whether all known cryptochromes function with the currently derived photocycle. In fact, the cryptochrome family is highly diverse, and its sole defining characteristic appears to be homology to photolyases coupled with signaling activity. Cryptochromes apparently developed multiple times, independently from different photolyase ancestors, during the course of evolution (7, 8). The classic plant cryptochromes (comprising Arabidopsis cry1 and cry2) for example originated from a type I CPD photolyase ancestor, whereas the classical animal-type cryptochromes such as Drosophila and mammalian cryptochrome evolved from a distinct 6-4 type photolyase ancestor. An additional category of cryptochromes known as the cryDASH family, found in bacteria and microbes as well as plants (Arabidopsis cry3) (9), apparently evolved independently from yet another distinct photolyase ancestor (2). In the case of those animal-type cryptochromes that are light-activated, studies of wavelength sensitivity of Drosophila cryptochrome showed little activity above 500 nm (41) and the classic peaks and shoulders,5 consistent with oxidized flavin as the primary photosensor. Furthermore, the closely related type 6-4 photolyases also undergo photoreduction in vitro by an electron transfer pathway similarly to plant cryptochromes (20). It is therefore possible that a similar mechanism of light activation, involving photoreduction of oxidized flavin, may have evolved for both plant and animal-type photoreceptors. By contrast, the cryDASH type cryptochromes seem not to accumulate the flavin radical upon light activation in vitro and instead undergo rapid photoconversion to the fully reduced flavin, similar to photolyases (42, 43).

In summary, the plant cryptochrome photocycle represents a fundamental departure in mechanism from photolyases. Flavin is found in the oxidized state in vivo and undergoes photoreduction to accumulate the neutral radical form as its signaling state. Because of this property, cryptochrome responses can be partially reversed under conditions of bichromatic irradiation with saturating green light. In the case of plants, cryptochrome photoreversion may contribute to shade avoidance responses under a canopy and thereby be of adaptive significance. It will be intriguing to determine whether other instances of antagonistic blue-green interactions in plants and other organisms (38, 44–46) may also somehow be linked to cryptochromes.


   FOOTNOTES
 
* This work was supported by Grant 0343737 from the National Science Foundation and by CNRS, Action Concertée Incitative/Biologie Cellulaire, Moleculaire et Structurale (ACI/BCMS), Deutsche Forschungsgemeinschaft (FOR526 BA985/10-2, BI464/8-2), and the Research Foundation-Flanders (postdoctoral fellowship to F. V.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1 and 2. Back

1 Permanent address: Inst. of Biological Sciences, College of Arts and Sciences, University of the Philippines Los Banos College, Laguna 4031, Philippines. Back

2 To whom correspondence should be addressed: Université Paris VI, FRE-CNRS 2846, Casier 156, 4 Place Jussieu, 75005 Paris. Tel.: 33-1-44272916; Fax: 33-1-44272916; E-mail: ahmad{at}ccr.jussieu.fr.

3 The abbreviations used are: DASH, Drosophila, Arabidopsis, Synechocystis, human; CPD, cyclobutane-pyrimidine dimer; FAD, flavin adenine dinucleotide. Back

4 R. Banerjee, E. Schleicher, S. Meier, R. M. Viana, R. Pokorny, M. Ahmad, R. Bittl, and A. Batschauer, unpublished results. Back

5 M. Ahmad, unpublished results. Back


   ACKNOWLEDGMENTS
 
We are indebted to Winslow Briggs (Stanford University) for critical reading of the manuscript, Hans Matthijs (Vrije Universiteit) for help with in vivo absorption studies, Alexandre Degrave (Université Paris VI) for assay development, and Eberhard Schlodder (Technical University Berlin) for use of a UV-visible spectrometer.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Banerjee, R., and Batschauer, A. (2005) Planta 220, 498–502[CrossRef][Medline] [Order article via Infotrieve]
  2. Lin, C., and Todo, T. (2005) Genome Biol. 6, 220–229[CrossRef][Medline] [Order article via Infotrieve]
  3. Briggs, W. R. (2006) in Photomorphogenesis in Plants and Bacteria (Schäfer, E., and Nagy, F., eds) 3rd Ed., pp. 171–197, Springer, Dordrecht, Netherlands
  4. Li, Q.-H., and Yang, H.-Q. (2006) Photochem. Photobiol., in press
  5. Van Gelder, R. N. (2002) J. Biol. Rhythms 17, 110–120[Abstract]
  6. Sancar, A. (2004) J. Biol. Chem. 279, 34079–34082[Free Full Text]
  7. Sancar, A. (2003) Chem. Rev. 103, 2203–2237[CrossRef][Medline] [Order article via Infotrieve]
  8. Todo, T. (1999) Mutat. Res. 434, 89–97[Medline] [Order article via Infotrieve]
  9. Brudler, R., Hitomi, K., Daiyasu, H., Toh, H., Kucho, K., Ishiura, M., Kanehisa, M., Roberts, V. A., Todo, T., Trainer, A., and Getzoff, E. D. (2003) Mol. Cell 11, 59–67[CrossRef][Medline] [Order article via Infotrieve]
  10. Brautigam, C. A., Smith, B. S., Ma, Z., Palnitkar, M., Tomchick, D. R., Machius, M., and Deisenhofer, J. (2004) Proc. Natl. Acad. Sci. U. S. A. 101, 12142–12147[Abstract/Free Full Text]
  11. Selby, C. P., and Sancar, A. (2006) Proc. Natl. Acad. Sci. U. S. A. 103, 17696–17700[Abstract/Free Full Text]
  12. Wang, H., Ma, L.-G., Li, J.-M., Zhao, H.-Y., and Deng, X. W. (2001) Science 294, 154–158[Abstract/Free Full Text]
  13. Yang, H.-Q., Tang, R.-H., and Cashmore, A. R. (2001) Plant Cell 13, 2573–2587[Abstract/Free Full Text]
  14. Green, C. B. (2004) Curr. Biol. 14, R847–R849[CrossRef][Medline] [Order article via Infotrieve]
  15. Yang, H.-Q., Wu, Y.-J., Tang, R.-H., Liu, D., Liu, Y., and Cashmore, A. R. (2000) Cell 103, 815–827[CrossRef][Medline] [Order article via Infotrieve]
  16. Partch, C. L., Clarkson, M. W., Ozgur, S., Lee, A. L., and Sancar, A. (2005) Biochemistry 44, 3795–3805[CrossRef][Medline] [Order article via Infotrieve]
  17. Kottke, T., Batschauer, A., Ahmad, M., and Heberle, J. (2006) Biochemistry 45, 2472–2479[CrossRef][Medline] [Order article via Infotrieve]
  18. Partch, C., and Sancar, A. (2005) Photochem. Photobiol. 81, 1291–1304[CrossRef][Medline] [Order article via Infotrieve]
  19. Carell, T., Burgdorf, L. T., Kundu, L. M., and Cichon, M. (2001) Curr. Opin. Chem. Biol. 5, 491–498[CrossRef][Medline] [Order article via Infotrieve]
  20. Weber, S. (2005) Biochim. Biophys. Acta 1707, 1–23[Medline] [Order article via Infotrieve]
  21. Byrdin, M., Sartora, V., Eker, A. P., Vos, M. H., Aubert, C., Brettel, K., and Mathis, P. (2004) Biochim. Biophys. Acta 1655, 64–70[Medline] [Order article via Infotrieve]
  22. Kavakli, I. H., and Sancar, A. (2004) Biochemistry 43, 15103–15110[CrossRef][Medline] [Order article via Infotrieve]
  23. Ahmad, M., Grancher, N., Heil, M., Black, R. C., Giovani, B., Galland, P., and Lardemer, D. (2002) Plant Physiol. 129, 774–785[Abstract/Free Full Text]
  24. Lin, C., Robertson, D. E., Ahmad, M., Raibekas, A. A., Schuman Jornes, M., Dutton, P. L., and Cashmore, A. R. (1995) Science 269, 968–970[Abstract/Free Full Text]
  25. Giovani, B., Byrdin, M., Ahmad, M., and Brettel, K. (2003) Nat. Struct. Biol. 10, 489–490[CrossRef][Medline] [Order article via Infotrieve]
  26. Zeugner, A., Byrdin, M., Bouly, J. P., Bakrim, N., Giovani, B., Brettel, K., and Ahmad, M. (2005) J. Biol. Chem. 280, 19437–19440[Abstract/Free Full Text]
  27. Ahmad, M., Galland, P., Ritz, T., Wiltschko, R., and Wiltschko, W. (2006) Planta 224, 995–1003[CrossRef][Medline] [Order article via Infotrieve]
  28. Smith, H. (2000) Nature 407, 589
  29. Spudich, J. L., and Bogomolni, R. A. (1984) Nature 312, 509–513[CrossRef][Medline] [Order article via Infotrieve]
  30. Ahmad, M., Lin, C., and Cashmore, A. R. (1995) Plant J. 8, 653–658[CrossRef][Medline] [Order article via Infotrieve]
  31. Ahmad, M., Jarillo, J., and Cashmore, A. R. (1998) Plant Cell 10, 197–208[Abstract/Free Full Text]
  32. Neumann, D., Emmermann, M., Thierfelder, J. M., zur Nieden, U., Clericus, M., Braun, H. P., Nover, L., and Schmitz, U. K. (1993) Planta 190, 32–43[Medline] [Order article via Infotrieve]
  33. Lin, C., Ahmad, M., Gordon, D., and Cashmore, A. R. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8423–8427[Abstract/Free Full Text]
  34. Folta, K. M. (2004) Plant Physiol. 135, 1407–1416[Abstract/Free Full Text]
  35. Franklin, K. A., and Whitelam, G. C. (2005) Ann. Bot. (Lond.) 96, 169–175[Abstract/Free Full Text]
  36. Vandenbussche, F., Pierik, R., Millenaar, F. F., Voesenek, L. A., and van der Straeten, D. (2005) Curr. Opin. Plant Biol. 8, 462–468[CrossRef][Medline] [Order article via Infotrieve]
  37. Lin, C., Yang, H., Guo, H., Mockler, T., Chen, J., and Cashmore, A. R. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 2686–2690[Abstract/Free Full Text]
  38. Galland, P., and Toelle, N. (2003) Planta 217, 971–982[CrossRef][Medline] [Order article via Infotrieve]
  39. Young, J., Liscum, E., and Hangarter, R. J. (1992) Planta 188, 106–114
  40. Goto, N., Yamamoto, K., and Watanabe, M. (1993) Photochem. Photobiol. 57, 867–871
  41. Busza, A., Emery-Le, M., Rosbash, M., and Emery, P. (2004) Science 304, 1503–1506[Abstract/Free Full Text]
  42. Saxena, C., Wang, H. Y., Kavakli, H., Sancar, A., and Zhong, D. P. (2005) J. Am. Chem. Soc. 127, 7984–7985[CrossRef][Medline] [Order article via Infotrieve]
  43. Song, S. H., Dick, B., Penzkofer, A., Pokorny, R., Batschauer, A., and Essen, L.-O. (2006) J. Photochem. Photobiol. 85, 1–6
  44. Schmidt, W., and Galland, P. (1999) Planta 208, 274–282[CrossRef][Medline] [Order article via Infotrieve]
  45. Frechilla, S., Talbott, L. D., Bogomolni, R. A., and Zeiger, E. (2000) Plant. Cell Physiol. 41, 171–176[Medline] [Order article via Infotrieve]
  46. Mao, J., Zhang, Y. C., Sang, Y., Li, Q. H., and Yang, H. Q. (2005) Proc. Natl. Acad. Sci. U. S. A. 102, 12270–12275[Abstract/Free Full Text]
  47. Kleiner, O., Butenandt, J., Carell, T., and Batschauer, A. (1999) Eur. J. Biochem. 264, 161–167[Medline] [Order article via Infotrieve]
  48. Wang, B. Y., and Jorns, M. S. (1989) Biochemistry 28, 1148–1152[CrossRef][Medline] [Order article via Infotrieve]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 Biophys. JHome page
K. C. Toh, I. H. M. van Stokkum, J. Hendriks, M. T. A. Alexandre, J. C. Arents, M. A. Perez, R. van Grondelle, K. J. Hellingwerf, and J. T. M. Kennis
On the Signaling Mechanism and the Absence of Photoreversibility in the AppA BLUF Domain
Biophys. J., July 1, 2008; 95(1): 312 - 321.
[Abstract] [Full Text] [PDF]

Home page
 Biophys. JHome page
T. Domratcheva, B. L. Grigorenko, I. Schlichting, and A. V. Nemukhin
Molecular Models Predict Light-Induced Glutamine Tautomerization in BLUF Photoreceptors
Biophys. J., May 15, 2008; 94(10): 3872 - 3879.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
N. Ozturk, S.-H. Song, C. P. Selby, and A. Sancar
Animal Type 1 Cryptochromes: ANALYSIS OF THE REDOX STATE OF THE FLAVIN COFACTOR BY SITE-DIRECTED MUTAGENESIS
J. Biol. Chem., February 8, 2008; 283(6): 3256 - 3263.
[Abstract] [Full Text] [PDF]

Home page
 Mol PlantHome page
N. Hoang, J.-P. Bouly, and M. Ahmad
Evidence of a Light-Sensing Role for Folate in Arabidopsis Cryptochrome Blue-Light Receptors
Mol Plant, January 1, 2008; 1(1): 68 - 74.
[Abstract] [Full Text] [PDF]

Home page
 Mol PlantHome page
Y.-J. Yang, Z.-C. Zuo, X.-Y. Zhao, X. Li, J. Klejnot, Y. Li, P. Chen, S.-P. Liang, X.-H. Yu, X.-M. Liu, et al.
Blue-Light-Independent Activity of Arabidopsis Cryptochromes in the Regulation of Steady-State Levels of Protein and mRNA Expression
Mol Plant, January 1, 2008; 1(1): 167 - 177.
[Abstract] [Full Text] [PDF]

Home page
 Plant CellHome page
X. Yu, J. Klejnot, X. Zhao, D. Shalitin, M. Maymon, H. Yang, J. Lee, X. Liu, J. Lopez, and C. Lin
Arabidopsis Cryptochrome 2 Completes Its Posttranslational Life Cycle in the Nucleus
PLANT CELL, October 1, 2007; 19(10): 3146 - 3156.
[Abstract] [Full Text] [PDF]

Home page
 J. Exp. Biol.Home page
S. Johnsen, E. Mattern, and T. Ritz
Light-dependent magnetoreception: quantum catches and opponency mechanisms of possible photosensitive molecules
J. Exp. Biol., September 15, 2007; 210(18): 3171 - 3178.
[Abstract] [Full Text] [PDF]

Home page
 J Exp BotHome page
P. F. Devlin, J. M. Christie, and M. J. Terry
Many hands make light work
J. Exp. Bot., September 1, 2007; 58(12): 3071 - 3077.
[Full Text] [PDF]