Handbook of Photosensory Receptors (BRIGGS:PHOTORECEPTORS O-BK) || LOV-Domain Photochemistry

14LOV-domain Photochemistry

Trevor E. Swartz and Roberto A. Bogomolni

14.1Introduction

The LOV domains are the light-sensing modules of the phototropins and relatedchromoproteins. Previous chapters have introduced the physiology mediated by thephototropins in plants and have discussed in general terms the LOV domain-medi-ated signal transduction process that results in activation of their kinase domains(Christie and Briggs, Chapter 13). Brief ly, in this process, light absorbed by the f lavinchromophore results in the transient formation of a covalent f lavoprotein adduct anda localized protein structural perturbation (Briggs et al. 2002). Intramolecular propa-gation of this conformational change results in activation of the kinase moiety andhierarchical autophosphorylation at several sites located in the protein region be-tween the two LOV domains and near the N terminus. Thus, adduct formation isviewed as a light-driven molecular switch that activates the subsequent molecularevents.

Since the discovery of the LOV domains in planta, a variety of LOV domain-con-taining proteins have been identified in prokaryotes and eukaryotes (Crosson et al.2003; Crosson, Chapter 15). The LOV domains are versatile blue-light sensors and arefound coupled to a diverse list of signaling elements. In all cases, it appears that theLOV domain is the primary sensory module that conveys a signal to protein domainswith functions as diverse as regulation of gene expression, regulation of protein ca-tabolism, or serine/threonine kinase activation in eukaryotes and histidine kinasesin prokaryotes. The LOV domains utilize photochemistry unlike that in any otherknown light sensors, such as the rhodopsins, phytochromes, xanthopsins, cryp-tochromes and BLUF proteins (Van der Horst et al. 2004). In addition, the LOV do-mains are versatile light sensors in that they are soluble proteins, absorb maximallyin the blue and UV-A, do not require synthesis of a special chromophore but utilizea ubiquitous cellular component, and are able to adjust their photocycle kinetics andquantum yield depending presumably on the physiology they are mediating.

The initial work on the phototropin LOV domains demonstrated that they bind anFMN chromophore and exhibit reversible photochemistry (Salomon et al. 2000). The

Handbook of Photosensory Receptors. Edited by W. R. Briggs, J. L. SpudichCopyright © 2005 WILEY-VCH Verlag GmbH & Co. KGaA, WeinheimISBN 3-527-31019-3

306

spectral similarities between the light-induced absorbance changes observed in iso-lated LOV domains and those associated to redox reactions in mercuric ion reductaseupon formation of a cysteinyl-f lavin adduct (Miller et al. 1990) strongly suggestedsuch reaction occurring in the LOV domains as a result of photoactivation, with tran-sient formation of a protein-FMN bond (cysteinyl adduct). As expected, these spectralchanges are absent in a mutant in which the reactive cysteine is replaced by alanine(C39A) (Salomon et al. 2000). Time-resolved spectroscopy showed that the LOV do-mains undergo a photocycle, which is characterized by a series of transient interme-diates (Salomon et al. 2000; Swartz et al. 2001; Kottke et al. 2002) with a spontaneousreturn to the ground-state of the protein in the dark. In addition, these studiesshowed that the C39A mutant has in fact a truncated photoreaction in which the sys-tem returns thermally from an excited state to its initial ground state (Swartz et al.2001). We present in this chapter these photochemical processes and the accompa-nying molecular structural perturbations. We first discuss the protein ground-statestructure and spectroscopy and then the photocycle kinetics, structure, and spectralproperties of the intermediates and the mechanism of transitioning between photo-intermediates.

14.2The Chromoprotein Ground State Structure and Spectroscopy

14.2.1Structure of the Chromoprotein and its Chromophore Environment

The first crystal structure of a closely related phototropin LOV domain (Adiantumcapillus-veneris phy3 LOV2) was solved to 2.7 Å (Crosson et al. 2001) and describedelsewhere (Crosson, Chapter 15). Here, we mention some of the key points relevantto the photochemistry discussion. The LOV domain structure clearly demonstratedthe very specific and tight binding of FMN within a protein pocket (Crosson et al.2001; Fedorov et al. 2003). The FMN is held in place non-covalently through interac-tion with a hydrogen-bonding network on the pyrimidime side of the isoalloxazinering and with hydrophobic residues on the dimethylbenzene moiety (Crosson et al.2001). Specifically, the FMN N3, O2, and O4 all hydrogen-bond to protein side chains;in addition, the phosphate group on the FMN ribityl chain interacts with the guani-dinium groups of two arginine residues to form salt bridges. The hydroxl groups ofthe FMN ribityl side chain form hydrogen bonds to the LOV-domain protein. Thereare two water molecules in close proximity to the chromophore. Both are within hy-drogen-bonding distance of the hydroxl groups of the FMN ribityl chain.

14 LOV-domain Photochemistry

307

14.2.2FMN Electrostatic Environment within the Protein

14.2.2.1 Chromophore AbsorptionThe proximity of the FMN molecule to the protein is apparent in the spectroscopicproperties (absorption, f luorescence, and circular dichroism) of the LOV domains ascompared to that of free FMN. Figure 14.1 shows both the absorption spectrum ofAvena sativa (oat) phot1-LOV2 and free FMN in H2O. Typical of the LOV domains(Figure 14.1), the ground form has major absorption bands around 370 nm and450 nm, with vibronic structure at around 360 nm and 380 nm, and 425 nm and475 nm, respectively (Salomon et al. 2000; Swartz et al. 2001). This vibronic structure,which is absent in the absorption of free FMN in H2O at room temperature (Figure14.1), is the consequence of a reduction of inhomogeneous broadening of the f lavinchromphore within the protein binding pocket as compared to that of free FMN(Schuttrigkeit et al. 2003) and is a ref lection of the tight binding of FMN held in placeby both hydrogen bonding and hydrophobic interactions. The reported extinction co-efficients for the two domains of oat phot1 are for LOV1, ε449 = 12 200 mol–1 cm–1

and ε370 = 10 000 mol–1 cm–1; for LOV2, ε447 = 13 800 mol–1 cm–1 andε378 = 8700 mol–1 cm–1 (Salomon et al. 2000). The main absorption band around450 nm is due to the first singlet state transition So→S1, whereas the 360 nm band isassociated with the So→S2 transition (Müller 1991). Stark Spectroscopy provides in-formation on changes in charge distribution in molecules upon optical excitation bymeasuring the changes in permanent dipole moment (permanent polarity) and inmolecular polarizability (inducible polarity) associated with a transition. Work onmodel f lavins (Stanley 2001) has shown that both transitions, S1 and S2, are accom-panied by small changes in the permanent excited state dipole moment relative to theground state. This slight increase in polarity in the excited states (less than 10%) im-plies a small change in the electrostatic interaction of the FMN with the immediateenvironment upon excitation.

In contrast to the minor alterations in permanent dipole moment, the change inpolarizability associated with the So→S2 transition (370 nm band) is much largerthan that of the So→S1 transition (450 nm band), making its absorption spectrumsensitive to the dielectric environment around the f lavin-binding sites in proteins(Stanley 2001). The vibronic components of the absorption spectrum of the LOV do-mains in this 370 nm region differs among different domains, showing changes inboth the wavelengths of maximal absorption and the relative intensities. In oat phot1-LOV2, the 360 nm peak is considerably lower than the 380 nm peak (dome shape),whereas in oat phot1-LOV1 domain, the peaks are slightly shifted and of nearly equalheight (f lat top shape) (Salomon et al. 2000). These features are very similar to thoseobserved between N3 methyl f lavins (Stanley 2001) (f lat top) and free FMN (dometop) at low temperatures, where all f lavins show these vibronic bands.

Within a given LOV domain, the vibronic structure and peak frequencies of the370 nm band are particularly sensitive to replacement of the cys39 with an alanine.This sensitivity strongly suggests the presence of electrostatic interactions betweenthe sulfur of cys39 and the FMN chromophore (Swartz et al. 2001). Comparison of

14.2 The Chromoprotein Ground State Structure and Spectroscopy

308

the absorption spectra of various LOV domains (At-phot1-LOV1, LOV2 and At-phot2-LOV1, LOV2) shows that the major difference in absorption is the change in shape ofthe 360 nm peak, and a slight ∼2 nm shift of the 450 nm peak (Sakai et al. 2001). At-Phot1-LOV2 and phy3-LOV2 show the typical dome shape, whereas Clamydomonasphot-LOV1 (Kottke et al. 2002), At-Phot1-LOV1, At-Phot2-LOV2, and the oat phot1-LOV2-C39A mutant show the f lat top feature. In oat phot1-LOV2, the f latness of thespectrum seems to ref lect a decrease in the environment polarity (C39 is more polarthan C39A, Figure 14.1) (Corchnoy et al. 2003). Replacement of a cysteine with a morepolar serine (C39S) has negligible effect on the UV-A absorption band, but it de-creases f luorescence quenching (see below). This difference suggests a stronger po-lar interaction by cysteine as compared to serine. The stronger polar interaction bythe cysteine residue could be explained by differences in the position of the sidechains in the mutants, or alternatively by the presence of the net charge of a thiolate,S–, a partially ionized thiol or perhaps involvement of a charge transfer complex. Al-though these perturbations of the short wavelength band must ref lect effects of localfields on the chromophore electronic polarizability, the available structural data for


Figure 14.1 FMN vibronic structure evidentfor chromophore in LOV-domain bindingpocket. UV-A absorption band of LOV do-mains is sensitive to the position, ionization

state, and/or residue at position 39. Also, notedifference in 370 nm absorption band betweenChlamydomonas phot-LOV1 and wild-type oatphot1-LOV2.

309

phy3-LOV2 and Chlamydomonas phot-LOV1 domains provides no information ongroups involved in these specific interactions.

At room temperature, differences in the vibronic structure of the 370 nm peak ofChlamydomonas phot-LOV1 and phy3-LOV2 are evident [phy3-LOV2 absorption issimilar to that of oat phot1-LOV2 (Kennis et al. 2003) (Salomon et al. 2000) (Figure14.1)]. This could ref lect differences in the location of the sulfur of cys39 with respectto the FMN chromophore and/or differences in the binding-site electrostatics. The X-ray structure of the Chlamydomonas Phot-LOV1 was solved to 1.9 Å (Fedorov et al.2003). In comparisons of the X-ray crystal structures of Chlamydomonas phot-LOV1(Fedorov et al. 2003) and phy3-LOV2 (Crosson et al. 2001), the major discrepancy isin the position of the sulfur group of cys39. In Chlamydomonas phot-LOV1, the cys-teine appears in two conformations, whereas in phy3-LOV2, the cysteine appears ina single conformation. However, the structure of Chlamydomonas phot-LOV1 wassolved at liquid nitrogen temperatures, whereas that of phy3-LOV2 was solved atroom temperature. It should be noted that phy3-LOV2 also displayed the cysteine intwo conformations when solved at 77 K (Crosson, Chapter 15). The presence of theseconformers alone does not explain the subtle, but significant, spectral differencesthat must originate from different chromophore environments. These different en-vironments could account for their different reactivities, quantum yields, and cyclingtimes (Kasahara et al. 2002). Therefore, it may not be appropriate to generalize thephotochemical mechanisms proposed for a particular LOV domain to all LOV do-mains.

14.2.2.2 Chromophore Fluorescence and Raman ScatteringFree FMN has a relatively high f luorescence quantum yield and low efficiency forphotochemical reactions in solution (Weber 1950). In an efficient system, photo-chemistry has to compete efficiently with both radiative (f luorescence) and thermalrelaxation (internal conversion, IC) processes. Fluorescence efficiency of free FMNapproaches 26% (Weber 1950), whereas in the protein binding pocket, it is around13% (Kennis et al. 2003). Although protein–chromophore interactions listed aboveand binding-site electrostatics may have an effect on f luorescence yield, a major con-tribution to the decrease in f luorescence of FMN in the chromoprotein seems to bethe competing process of intersystem crossing (ISC) into the f lavin triplet statecaused by the presence of the cysteine sulfur atom in the vicinity of the f lavin chro-mophore [see below, (Holzer et al. 2002; Kennis et al. 2003)]. The f luorescence quan-tum yield of phy3 LOV2 is 0.13 (Kennis et al. 2003) and that of Chlamydomonas phot-LOV1 is 0.17 (Kennis et al. 2003). The LOV domains’ f luorescence quantum yield issensitive to both the chromophore binding-site electrostatic environment and thepresence of the sulfur of cys39 (Salomon et al. 2000; Swartz et al. 2001).

The inf luence of binding-site electrostatics is evident from the relative marked in-crease in f luorescence yield upon substitution of the cys39 for a serine (0.13 to about0.19) and alanine (0.13 to about 0.24) (Swartz et al. 2001). The increased spin orbitcoupling in presence of sulfur is expected to enhance the S1→T1 transition by the ISCprocess, decreasing the f luorescence yield. The triplet yield depends on both the ISCrate and the internal conversion (IC) rate. The ISC rate increase for either oat phot1-


310

LOV2 vs. free FMN (Kennis et al. 2003) or oat phot1-LOV2 vs. oat phot1-LOV2C39A(Schuttrigkeit et al. 2003) has been reported to be about 2.4. There is disagreement inthe reported values for triplet yield in the presence and absence of sulfur. The quan-tum yield of triplet formation (ΦT) for free FMN has been reported to be about 0.6(Kennis et al. 2003; Schuttrigkeit et al. 2003). There was no measurable increase intriplet formation in phy3-LOV2 (ΦT = 0.6) (Kennis et al. 2003). However, in oat phot1-LOV2, ΦT was reported as 0.83 [consistent with an earlier report of 0.88 (Swartz et al.2001)], an increase from 0.68 for oat phot1-LOV2C39A (Schuttrigkeit et al. 2003). Thereported discrepancy in ΦT between oat phot1-LOV2 and phy3-LOV2 originates fromdifferent values assigned to the IC rates. A low triplet yield of 0.26 reported for theChlamydomonas phot-LOV1 domain is in sharp contrast to these high values in oatphot1-LOV2 (Islam et al. 2003). These discrepancies may ref lect variability in thesample preparation and/or differences in fusion proteins used [calmodulin bindingprotein (CBP), maltose binding protein (MBP), and histidine-tagged protein] or thedifferent LOV domains studied.

Both the absorption spectrum of the ground state (see above) and f luorescence da-ta suggest that differences not apparent in the structural data must exist in the envi-ronment directly interacting with the FMN chromophore of various LOV domainsstudied to date. It is arguable from these data that there may be subtle positional dif-ferences in the cys39 sulfur in various LOV domains. An additional factor not ad-dressed is that FMN in the LOV domains is restricted in a rigid binding pocket whereconformational strain may affect the evolution of the system in the excited state, mod-ifying the various relaxation channels, including f luorescence. The LOV domains’f luorescence emission shows vibronic structure that is absent in the free FMN in so-lution (Kennis et al. 2003). The vibronic structures of the f luorescence emission andexcitation spectra ref lect vibrational progressions in the ground and excited states, re-spectively. In general, identical progressions (mirror images) are indicative of smallor no changes in the molecular coordinates of the atomic nuclei between ground andexcited state. In oat phot1-LOV2, the progressions for f luorescence emission and ex-citation are about 1070 cm–1 and 1200 cm–1, respectively. This difference is indicativeof changes in the nuclear configurations in the excited state (Schuttrigkeit et al.2003).

Fluorescence yield is affected by electrostatic interactions in the excited state. Ion-ic electrostatic potential provides the strongest and longest range interaction. There-fore, both location and ionization state of the sulfur of cys39 could be expected to in-f luence the f lavin f luorescence yield. In mercuric ion reductase, a cysteine sulfur islocated slightly closer to isoalloxazine C(4a) (Müller 1991) than in phy3-LOV2(Crosson et al. 2001) (2.7 Å vs. 4.2 Å,) and its ionization state is the dominant factorin controlling the f luorescence yield of the chromophore at low pH (Miller et al.1990). It was first suggested by f luorescence pH titrations that the oat phot1-LOV2sulfur of cys39 existed predominantly as a thiolate (S–) at neutral pH (Swartz et al.2001). The FMN f luorescence in oat phot1-LOV2 is independent of pH in the range4–10. At high pH, there is nearly complete f luorescence quenching upon titration ofthe f lavin N3 with an apparent pK around 10.5. At low pHs (3–4) there is a markedincrease in the f luorescence of oat phot1-LOV2 (yield increases 0.13 to 0.20). As in


311

the mercuric ion reductase system (Miller et al. 1990), this result was interpreted as areduction in f luorescence quenching of the FMN upon protonation of the sulfur thi-olate. It was suggested that cys39 was in a proton equilibrium with a putative basewith an apparent pK around 4–4.5. This equilibrium would yield a ground state con-taining a Cys39 thiolate as the dominant species at neutral pH. In the reductase sys-tem, the f luorescence of FAD is fully quenched by the neighboring thiolate anion,while in the LOV2 domain, because of the longer distance, a thiolate effect would notexpected to be as large.

This interpretation is not consistent with the reported disappearance of an S-H vi-bration (negative difference band) at around 2500 cm–1 upon formation of the cys-teinyl adduct in phy3-LOV2 (Iwata et al. 2002) and Chlamydomonas phot-LOV1 (Ata-ka et al. 2003). Loss of an S–H bond is expected when a protonated ground state sul-fur group (thiol) forms an adduct with the FMN. Band assignment was confirmed bythe expected shift to lower frequencies caused by H/D isotopic replacement whenmeasurements were carried out in D2O. The amplitude of the negative SH vibrationband would be expected to be pH dependent in a thiol/thiolate equilibrium in whichthe thiolate is the reactive species, because the thiol would ionize as thiolate is con-sumed in the reaction. The lack of pH dependence of the FTIR S–H band betweenpH 7 and pH 10 (77–250 K) argues against such a pK equilibrium in phy3-LOV2 (Iwa-ta et al. 2003). Based on the apparent pH independence of the S–H difference bandamplitude, it was suggested that the phy3-LOV2 C39 thiol has a pKa higher than 10,inferring that it is buried in a hydrophobic environment but able to equilibrate withbulk deuterons (Iwata et al. 2003).

The fusion protein used for this study (Iwata et al. 2003) is larger than that used forthe X-ray crystallography work, with 20 and 40 additional amino acids on the N ter-minus and C terminus, respectively, and a CBP tag. It should be noted that LOV2 fu-sion proteins used in different laboratories contain varying amounts of native se-quence and specific tags that may or may not have been cleaved, in addition to the do-main sequence used for two available X-ray crystallographic studies of LOV domains(Crosson et al. 2001; Fedorov et al. 2003). These additional components may inf lu-ence the solution structure, molecular folding, solvent accessibility of specificresidues, and more important the stabilization of conformations favoring differentpositions of the reactive cysteine. Adduct formation occurs at cryogenic temperatures[phy3 LOV2, (Iwata et al. 2003); oat phot1-LOV2, (T.E. Swartz and R.A. Bogomolni un-published; M. Gauden, S. Crosson, I. Van Stokkum, R. van Grondelle, K. Moffat andJ. Kennis unpublished)]. At these low temperatures, only 36% and 64% of the groundstate converts into the adduct at 77 K and 100 K, respectively (Iwata et al. 2003), anda temperature-dependent amplitude of the S–H band is attributed to an increase inthe extinction coefficient for that absorption band at lower temperatures (Iwata et al.2003). These observations point to the importance of molecular dynamics and otherprocesses in the reaction mechanism.

Pre-resonance Raman of the Chlamydomonas phot-LOV1 (Ataka et al. 2003) and oatphot1-LOV2 (J. Ciorciari and R.A.Bogomolni unpublished) ground states obtainedwith 752 nm and 785 nm Raman probe beams, respectively, show spectra very simi-lar to that of free FMN, consistent with non-covalent binding of the FMN within the


312

LOV domain pocket. Hydrogen bonding of FMN in the protein causes slight shifts insome of the bands, confirming spectroscopically hydrogen bonds seen in the crystalstructure (Ataka et al. 2003). The pre-resonance Raman spectrum of the ground stateis dominated by chromophore vibrational bands but shows some contribution of theprotein amide vibrations in the 1600–1700 cm–1 range that overlap with chromophorecarbonyl bands (Ataka et al., 2003; J. Ciorciari and R.A. Bogomolni unpublished).

14.2.2.3 Chromo-peptide Optical ActivityThe circular dichroism (CD) spectrum of the oat phot1-LOV2 ground state in the nearUV-visible regions shows negative bands that coincide with absorption bands of thechromophore (Salomon et al. 2000; Corchnoy et al. 2003). This optical activity is notpresent in the ground state spectrum of free FMN in aqueous solution (S. Corchnoyand R. A. Bogomolni unpublished), suggesting that FMN in LOV2 is in a protein-asymmetric environment. In the mid-UV region, there are CD bands in the aromat-ic region, perhaps contributed by tryptophan and high-energy chromophore bands.The far UV CD spectrum shows a positive band around 193 nm and negative bandsat 208 nm and 224 nm, indicative of the presence of alpha-helical secondary struc-ture, and the overall spectrum is consistent with the secondary-structure contentshown in the X-ray crystal structure (Corchnoy et al. 2003).

14.3Photochemistry

14.3.1Photocycle Kinetics and Structure of its Intermediates

To date, almost a dozen LOV domains have been shown to undergo similar chro-mophore structural changes during their photocycles. However each has distinct ki-netic characteristics and quantum yields and some have truncated photocycles (Sa-lomon et al. 2000; Kasahara et al. 2002; Kottke et al. 2002; Losi et al. 2002; Imaizumiet al. 2003; Schwerdtfeger et al. 2003). Early studies of the photochemistry were car-ried out in the E coli-expressed oat phot1-LOV2 domain (Salomon et al. 2000; Swartzet al. 2001). Photoreactions of other FMN-binding LOV domains studied afterwardswere shown to be qualitatively similar. Following light absorption, the LOV domainsundergo a photocycle characterized by a series of transient photo-intermediates(Swartz et al. 2001; Kottke et al. 2002; Losi et al. 2002). Presently, two excited statesand one metastable thermal intermediate have been kinetically resolved in the LOVdomain photocycle. Brief ly, at room temperature and slightly alkaline pH, light ab-sorption at around 450 nm promotes the So→S1 transition to the FMN singlet excit-ed state that decays with a half-time around 2 ns into a red-absorbing species that ab-sorbs maximally at 660 nm (Kennis et al. 2003). This species, labeled LOV2T

660, hasbeen shown to be an FMN triplet state (Figure 14.2 A) (Swartz et al. 2001; Kottke etal. 2002; Losi et al. 2002; Kennis et al. 2003). LOV2T

660 decays in microseconds into thesecond metastable intermediate, which absorbs maximally at 390 nm and is labeled


313

LOVS390 (Swartz et al. 2001). The LOVS

390 intermediate thermally relaxes back to theground state, LOVD

447 (Figure 14.2 B).

14.3 Photochemistry

Figure 14.2 (A) difference absorption spectraof oat phot1-LOV2 after excitation with a477-nm laser pulse. Spectra collected at 0.03,0.13, 0.33, 1.0, 3, 10, and 100 µs after light ex-citation. (B) Time-resolved light-induced ab-

sorption changes for oat phot1-LOV2 between1 and 100 s after light excitation. Spectra aretaken every 15 s. Arrows indicate spectralchanges with time.

314

Global kinetic analysis of laser f lash-induced absorption changes of oat phot1-LOV2 (actinic wavelength 477 nm, 4-ns pulse) measured from 30 ns to 100 µs givesa single decay time constant of 2 µs in this time window, and isosbestic points indi-cate a single resolvable, spectroscopically identifiable species (Swartz et al. 2001). Thedata were globally fit by assuming a linear kinetic scheme with two intermediates(LOV2T

660 and LOVS390) and the spectra of both species was calculated (Swartz et al.

2001) (Figure 14.3). Because the measured amount of the initial bleach of the groundstate is half that of the amount of LOVS

390 formed, a simultaneous 1:1 split decay ofLOV2T

660 back to the ground state LOV2D447 and forward to LOVS

390 was included in thekinetic scheme, resulting in a calculated time constant of 4 µs for each decay direc-tion (Swartz et al. 2001). The calculated LOV2T

660 spectrum closely matches that of thetriplet state of FMN (Figure 14.3) (Swartz et al. 2001). Measurements on the fem-tosecond timescale show direct conversion of the FMN triplet state from the singletexcited state in about 2–3 ns (Kennis et al. 2003). Since no other intermediates areseen on this time scale, the triplet state forms directly from the excited singlet statevia intersystem crossing (ISC) (Kennis et al. 2003; Schuttrigkeit et al. 2003). Furtheranalysis of the triplet-state spectrum suggests a mixture of species in which the FMNN5 exists as an equilibrium of both protonated and unprotonated forms (Kennis et al.2003).

The LOVS390 species that is formed from the triplet state decays spontaneously in

the dark back to the ground state. The rate of return of the ground state varies froma few seconds to many minutes (Salomon et al. 2000; Kasahara et al. 2002; Losi et al.


Figure 14.3 Ground-state oat phot1-LOV2spectrum and calculated intermediate spectra.

LOVL660 is the triplet state. The LOVS

390 statecontains the cysteinyl adduct.

315

2002; Schwerdtfeger et al. 2003), with some LOV domains not returning to theground state at all (Imaizumi et al. 2003). This long-lived metastable intermediate in-volves the formation of a protein-FMN covalent bond. Specifically an S–C bond isformed between the sulfur of cys39 and the C(4a) carbon of FMN (Figure 14.4). Be-cause this bond forms in microseconds and decays in seconds, continuous illumina-tion of a LOV2 sample with blue light converts most of the sample into LOVS

390; thisproperty has conveniently allowed for structural and spectroscopic studies of theLOVS

390 intermediate. Conformation of the C–S bond in LOVS390 was obtained in this

way by X-ray crystallography, and NMR (Salomon et al. 2001; Crosson et al. 2002; seeCrosson, chapter 15). Formation of the cysteine-C(4a) bond during the photocyclewas also inferred from difference infrared-absorbance spectroscopy (FTIR). Three vi-brational bands occurring at 1580 cm–1, 1550 cm–1, and 1350 cm–1, that are stronglycoupled to the stretching of the C(4a)–N5 double bond, disappear upon formation ofLOV2S

390 (Swartz et al. 2002; Ataka et al. 2003), and two new bands form at 1516 cm–1

and 1536 cm–1 (Swartz et al. 2002). Normal mode calculations on a simplified modelC(4a) thio-adduct predict the disappearance of the former bands and the appearanceof these two bands on adduct formation (Swartz et al. 2002). They are associated withsymmetric and antisymmetric stretching modes involving all three rings of an isoal-loxazine containing an sp3 hybridized carbon at position C(4a). Thus, FTIR results al-so show formation of a bond at the C(4a) position (Swartz et al. 2002; Ataka et al.2003). The light-induced CD changes in the 250–500 nm range show the appearanceof a large positive band at 270–290 nm upon formation of LOV2S

390. This band coin-cides with a higher energy FMN absorption band around 270 nm. The large optical

14.3 Photochemistry

Figure 14.4 Oat phot1–LOV2 reaction scheme.

316

activity associated with this band is consistent with the formation of an sp3 chiralC(4a) center in the LOV2S

390 adduct (Salomon et al. 2000; Corchnoy et al. 2003).Both circular dichroism (CD) and difference FTIR have demonstrated protein con-

formational perturbations associated with the photocycle (Swartz et al. 2002; Ataka etal. 2003; Corchnoy et al. 2003; Iwata et al. 2003). Numerous positive and negativeFTIR bands are observed in the protein amide regions (Swartz et al. 2002; Ataka et al.2003; Iwata et al. 2003), and CD difference spectra show transient loss in alpha he-licity (Corchnoy et al. 2003). The magnitude and character of these protein confor-mational changes depend on the size and sequence of the specific LOV constructsused for the studies. In particular, these structural changes become more apparent inlarger constructs containing additional protein segments on the C-terminal end ofthe LOV2 domain. In a larger construct, difference 3D NMR has revealed protein per-turbations of an amphipathic alpha-helix C-terminal from the LOV2 domain (Harperet al. 2003; see Crosson, Chapter 15), the direction in which signal propagation is ex-pected in the full-length chromoprotein. However, the physiological significance ofthis structural change is not clear because these are truncated phototropin molecules.In the full construct, the relevant structural change may propagate further withoutmajor changes in such an intervening segment.

14.3.2Photo-backreaction

It was inferred that in Chlamydomonas phot-LOV1 the inability to fully bleach theground state was due to a photo-backreaction from LOVS

390 to LOVD447 (Kottke et al.

2002). This photo-backreaction was directly measured in phy3-LOV2 and occurs in100 ps (Kennis et al. 2004), with quantum yield estimated to be between 0.2 and 0.3(Kennis et al. 2004). It is unknown if this UV-A induced back-reaction is physiologi-cally relevant.

14.4Reaction Mechanisms

14.4.1Adduct Formation

14.4.1.1 Proton Transfer Reaction Initiates Triplet State DecayThe rate of adduct formation is 5 times slower in D2O than in H2O, confirming thatthe rate-limiting step between triplet state and adduct formation is a proton-transferreaction (Corchnoy et al. 2003). The triplet state (LOVT

660) in wild-type oat phot1-LOV2, decays simultaneously back to LOVD

447, the ground state, and LOVS390 in both

D2O and H2O. The observed forward decay of LOVT660, to LOVS

390 is five times slowerin D2O (Corchnoy et al. 2003). Analysis of the data shows that the forwards/backward1:1 split ratio for its decay is unchanged in D2O and that the backward decay to theground state is also slowed down 5-fold. This analysis suggests that both decay reac-


317

tions of the triplet (forward to the adduct state and back to the ground state) are ratelimited by a single proton-transfer reaction. The magnitude of the deuterium resultis suggestive of a primary isotope effect (Melander et al. 1980). It is reasonable thatthis primary isotope effect is protonation of N5 (see mechanism below); however, atthis time there is no explicit data to support this hypothesis.

Mutation of cysteine 39 to an alanine (LOV2C39A) results in a truncated photocy-cle as compared to the wild type. Because these proteins are missing the reactive cys-teine, they do not form the LOVS

390 intermediate; however, after a short laser f lash,these proteins do form the LOVT

660 intermediate. This intermediate LOV2C39AT660

decays directly back to the ground state (LOV2C39AD447) in 72 µs, one order of mag-

nitude slower than wild type (Swartz et al. 2001). The presence of a sulfur atom in theWT oat phot1-LOV2 domain may contribute to the triplet decay presumably by in-creasing the rate of spin f lipping due to spin orbit coupling (Swartz et al. 2001). Notunexpectedly, the decay of LOV2C39AT

660 back to the ground state, LOV2C39AD447,

does not show a deuterium effect (Corchnoy et al. 2003).Several alternate reaction pathways for the formation of the adduct following initial

activation to the triplet state have been proposed. They include direct transfer of a pro-ton from the cysteine thiol to N5 of the triplet state followed by formation of the S-Cbond (Crosson et al. 2001), excited-state proton transfer to N5 preceding triplet for-mation followed by reaction of a thiolate with FMN C(4a) (Kennis et al. 2003), in-volvement of a f lavosemi-quinone free radical and reaction with a sulfur radical (Bit-tl et al. 2003; Kay et al. 2003; Kottke et al. 2003), and proton transfer to the triplet FMNfollowed by reaction of a C(4a) carbo-cation with a cys39 thiolate (Swartz et al. 2001).Available evidence is insufficient to establish the correct mechanism. Common to allmechanisms is the redistribution of charge around the FMN N5-C(4a) double bondthat occurs in the FMN triplet state, resulting in an increase of basicity of N5 (Song1968; Fedorov et al. 2003; Neiss et al. 2003). In the ionic recombination mechanism,protonation of N5 either from the C39 thiol (Crosson et al. 2001; Kennis et al. 2003)or from an as yet unidentified acid group (Swartz et al. 2001), draws electronic den-sity from the N5-C(4a) double bond, leaving C(4a) as a reactive carbo-cation that is at-tacked by the ionized sulfur of cysteine 39, forming the f lavin-cysteinyl adduct.

Kennis et al. (2003) proposed that proton transfer from the C39 thiol occurs in theexcited state with transient thiolate formation, whereas Swartz et al. (2001) have in-voked the existence of a ground-state thiolate and a putative proton-acceptor residuethat donates a proton to N5, initiating the split decay of the triplet state to eitheradduct or ground state. Vibrational-spectroscopy evidence that the sulfur of cysteineexists as a thiol has led to the argument that the proton originates directly from thecysteine SH. Although the proton could come from another group, no other acidgroups have been identified in the vicinity of the chromophore. The crystal structure,however, shows two water molecules that could conduct a proton from a residue thatis not directly interacting with the chromophore. The C39A mutants do produce aneutral semiquinone radical under high light intensities (see below) (Kay et al. 2003).Formation of the neutral semquinone requires donation of a hydrogen atom from anas yet unidentified group.

14.4 Reaction Mechanisms

318

It is generally accepted that in thiol proteases, the cysteine in the catalytic site whichforms a thiolate-imidazolium ion pair is separated by about 4 Å from the imidazolering of a histidine (Bergmann et al. 1997). The pKa values of imidazole and thiol insolution are around 6 and 8.5 respectively. In the thiol proteases, these pKa valuesshift to 8 and 4.5, respectively, with the thiol becoming a stronger acid than the imi-dazole (McGrath 1999). The pKa of FMN N5 in water is ∼0 in the ground state andjumps to about pKa ∼5 in the triplet state. In phy3-LOV2, the sulfur of cys 39 is 3.7 Åfrom N5 and 4.2 Å from C(4a) (Crosson et al. 2001). It would not be surprising thatthey would interact electrostatically and modulate each other’s pKa values in both theground state and triplet state. The expectation is that the N5 pKa is increased and thethiol pKa is decreased due to the interaction. It should also be noted that hydrogenbonding has been shown to modulate the pKa of N5 in the excited state (Yagi et al.1980). Presumably, the pKa values in the triplet state would approach values consis-tent with a spontaneous proton transfer.

The radical-pair mechanism involves formation of a f lavin semiquinone from theFMN triplet state in a one-electron photoreduction that may be followed by a protontransfer to give an anionic or neutral f lavin radical (Figure 14.5). In this scenario, thesulfur donates either an electron (and then the proton) or a hydrogen atom resultingin formation of a sulfur radical and neutral semiquinione radical. The adduct wouldform by recombination of the two radicals. The oat phot1-LOV2 triplet state absorp-tion spectrum shows no evidence of a semiquinone (Swartz et al. 2001). It is possiblethat the triplet state is the rate limiting step, and that the rapid rate of disappearanceof the semiquinone would not allow its transient accumulation and its detection inthe absorption changes. Kay et al. (Kay et al. 2003) point out that protonation of theFMN triplet state is not necessarily the rate-limiting step in a radical-pair mecha-nism, whereas protonation of the triplet state should be a rate-limiting step in an ion-ic mechanism. The ionic mechanism would allow adduct formation in the tripletstate, whereas the reaction-pair mechanism would require the reaction in a singletstate (antiparallel spins recombine to form a bond) (Kay et al. 2003).

The radical recombination mechanism proposed for adduct formation was basedon the detection of a f lavine-semiquinone radical in a C39A mutant of oat phot1-LOV2 [labeled C450A in that work (Kay et al. 2003)]. In the absence of sulfur, the pho-toreaction produces significant amounts of the neutral f lavo-semiquinone that ab-sorb maximally around 600 nm. The mechanism proposed for this reaction involvesformation of the triplet state, followed by proton and electron transfers from uniden-tified donor(s). In wild type oat phot1-LOV2, the proposed mechanism conservesthese features and simply substitutes the thiol for the unidentified donor (Kay et al.2003). At physiological light intensities, the formation of a neutral f lavosemiquinonewas not observed in LOV2C39A (Swartz et al. 2001), but it was observed at very highlight intensities (T. E. Swartz and R.A. Bogomolni unpublished). At similar high lightintensities, a small fraction of wild-type oat phot1-LOV2 also forms a neutral semi-quinone (T.E. Swartz and R.A. Bogomolni unpublished). At these light intensities,most of the sample forms the adduct and undergoes a fully reversible photocycle re-turning to the ground state in tens of seconds. The long-lived semiquinone forms viaa branched reaction pathway. It returns to the ground state in tens of minutes to


319

hours, presumably re-oxidized by atmospheric oxygen (T.E Swartz and R.A. Bogo-molni unpublished). Figure 14.5 summarizes the various reaction mechanisms pro-posed for adduct formation.

It should be mentioned that although the Chlamydomonas phot-LOV1 photocyclecontains intermediates similar to those of oat phot1-LOV2, it involves a more com-plex kinetic scheme. The Chlamydomonas phot-LOV1 triplet state decays forwardwith two time constants, one of 800 ns and one of 4 µs; it has been suggested that thetriplet state exists as a mixture of two species with different decay times (Kottke et al.2002). However, because their spectral properties are nearly identical, it is not possi-ble to discriminate between the two (Kottke et al. 2002). In contrast to oat phot1-LOV2, Chlamydomonas phot-LOV1 does not show an appreciable return to theground state from the triplet state. The isotope deuterium effect has not been re-ported on the Chlamydomonas phot-LOV1 adduct formation or decay.

14.4.2Adduct Decay

The mechanism of dark recovery of the ground state LOV2D447 from LOVS

390, which re-quires breakage of a carbon sulfur-bond, remains a mystery. Synthetic C4a-sulfuradduct model compounds have been used for structural studies (Müller 1991), andthe C(4a)-Sulfur covalent bond is strong (typically greater than 200 kJ mol–1) and sta-ble in aqueous media. Breakage of this bond in the protein environment must involvea catalytic process provided by specific protein residues in the FMN binding pocket.In oat phot1-LOV2, the rate of adduct decay shows only a slight decrease at lower pH[apparent pK around 6.5 (Corchnoy et al. 2003)], but in Chlamydomonas phot-LOV1there is a marked pH dependence in which the decay rate increases several fold be-tween pH 8 and pH 3 with an apparent pK around 5–6 (Kottke et al. 2002), indicat-ing that it is base-catalyzed. These different pH sensitivities demonstrate another ex-ample of the differences in chemical behavior between LOV domains. Abstraction ofthe N(5) proton (acid–base catalysis) could be the event that initiates the back reac-tion. The dark recovery rate in oat phot1-LOV2 is 3 times slower in D2O than in H2O,suggesting that a proton-transfer reaction is the rate-limiting step (Swartz et al. 2001).

Because the deuterium effect is only a factor of three, the rate-limiting step in theback reaction is perhaps not a primary isotope effect and could ref lect breaking of ahydrogen bond rather than a proton transfer. Because the pKa of N5 is modulated byhydrogen bonding on the f lavin (Yagi et al. 1980), perhaps the variability of relaxationin different LOV domains is a measure of this hydrogen bonding. Kinetics of CDchanges in the far UV indicate that the protein returns to the ground state with thesame kinetics as the chromophore and with identical kinetic deuterium isotope ef-fects suggesting concerted events and a common rate-limiting proton-associatedevent (Corchnoy et al. 2003). The X-ray structure fails to suggest a suitable catalyticbasic group in the vicinity of N5. The back reaction has been measured in almost adozen different LOV domains, with rates varying from seconds to minutes to no re-turn of the ground state (Salomon et al. 2000; Kasahara et al. 2002; Kottke et al. 2002;Losi et al. 2002; Imaizumi et al. 2003; Schwerdtfeger et al. 2003). The sequences of

14.4 Reaction Mechanisms

320

these various LOV domains present no clear insight into the varying relaxation rates.Difference FTIR spectra show hydrogen-bonding perturbations associated with thetwo buried waters that are in close proximity to the chromophore (Iwata et al. 2003),suggesting that these waters are possibly involved in the photochemistry. They mayform a proton-conducting channel to a basic group not in the immediate vicinity ofthe chromophore, a mechanism that has been found in other proteins. It has beensuggested that mediation of the structure of a conserved salt bridge occurs during thephotocycle and results in a transmitted signal (Crosson et al. 2003).

14.4Future Perspectives

The LOV domains are the photosensitive module of larger proteins that control vary-ing physiological responses. The studies reviewed here have been carried out in iso-lated fusion chromoproteins expressed in heterologous systems (E. coli). Thereforethe photochemical behavior and molecular changes observed in these systems do notprovide information on the important mechanism of intramolecular signal trans-duction in the native chromoproteins. Neither the structure nor the aggregation stateof full-length phototropin is known. Measurements on the aggregation state of LOV1(M. Salomon, personal communication) suggest phot1 forms a dimer raising the pos-


Figure 14.5 Proposed reaction mechanisms for adduct formation.

321

sibility of cis vs. trans signal transduction. In addition, the native photoreceptor ismembrane bound. It is unknown if the protein is post-translationally modified witha membrane anchor or binds a membrane protein and if these components affectphotochemistry. The photocycle kinetics of full-length phototropin expressed in in-sect cells (Kasahara et al. 2002) differ from those of isolated LOV domains but re-semble those of peptides containing both LOV1 and LOV2 domains (Kasahara et al.2002). The effect of phosphorylation on photochemistry is also unknown. Thesequestions will only be answered when native protein is isolated from plants.

References

References

Ataka, K., P. Hegemann, et al. (2003). Biophys.J. 84(1): 466–474.

Bergmann, E. M., S. C. Mosimann, et al.(1997). J. Virol. 71(3): 2436–2448.

Bittl, R., C. Kay, et al. (2003). Biochemistry42(28): 8506–8512.

Briggs, W. and J. Christie (2002). Trends PlantSci. 7(5): 204–210.

Corchnoy, S., T. Swartz, et al. (2003). J. Biol.Chem. 278(2): 724–731.

Crosson, S. and K. Moffat (2001). Proc. Natl.Acad. Sci. USA 98(6): 2995–3000.

Crosson, S. and K. Moffat (2002). Plant Cell14(5): 1067–1075.

Crosson, S., S. Rajagopal, et al. (2003). Bio-chemistry 42(1): 2–10.

Fedorov, R., I. Schlichting, et al. (2003). Bio-phys. J. 84(4): 2474–2482.

Harper, S., L. Neil, et al. (2003). Science301(5639): 1541–1544.

Holzer, W., A. Penzkofer, et al. (2002). Pho-tochem. Photobiol. 75(5): 479–487.

Imaizumi, T., H. Tran, et al. (2003). Nature426(6964): 302–306.

Islam, S., A. Penzkofer, et al. (2003). Chem.Phys. 291(1): 97–114.

Iwata, T., D. Nozaki, et al. (2003). Biochemistry42(27): 8183–8191.

Iwata, T., S. Tokutomi, et al. (2002). J. Am.Chem. Soc. 124(40): 11840–11841.

Kasahara, M., T. Swartz, et al. (2002). PlantPhysiol. 129(2): 762–773.

Kay, C., E. Schleicher, et al. (2003). J. Biol.Chem. 278(13): 10973–10982.

Kennis, J., S. Crosson, et al. (2003). Biochem-istry 42(12): 3385–3392.

Kennis, J. T. M., I. H. M. van Stokkum, et al.(2004). J. Am. Chem. Soc. 126(14): 1412–1413.

Kottke, T., B. Dick, et al. (2003). Biochemistry42(33): 9854–9862.

Kottke, T., J. Heberle, et al. (2002). Biophys. J.84(2): 1192–1201.

Losi, A., E. Polverini, et al. (2002). Biophys. J.82(5): 2627–2634.

McGrath, M. E. (1999). Annu. Rev. Biophys. Bio-molec. Struct. 28: 181–204.

Melander, L. and W. Saunders (1980). Reactionrates of isotopic molecules. New York, Wiley.

Miller, S. M., V. Massey, et al. (1990). Biochem-istry 29(11): 2831–41.

Müller, F. (1991). Chemistry and biochemistry off lavoenzymes. Boca Raton, CRC Press.

Neiss, C. and P. Saalfrank (2003). Photochem.Photobiol. 77(1): 101–109.

Sakai, T., T. Kagawa, et al. (2001). 98(12): 6969–74.

Salomon, M., J. Christie, et al. (2000). Biochem-istry 39(31): 9401–9410.

Salomon, M., W. Eisenreich, et al. (2001). Proc.Natl. Acad. Sci. U. S. A. 98(22): 12357–12361.

Schuttrigkeit, T., C. Kompa, et al. (2003).Chem. Phys. 294(3): 501–508.

Schwerdtfeger, C. and H. Linden (2003). EmboJ. 22(18): 4846–4855.

Song, P. S. (1968). Photochem. Photobiol. 7(3):311–3.

Stanley, R. J. (2001). Antioxid. Redox Signal.3(5): 847–866.

Swartz, T., S. Corchnoy, et al. (2001). J. Biol.Chem. 276(39): 36493–36500.

Swartz, T., P. Wenzel, et al. (2002). Biochem-istry 41(23): 7183–7189.

Van der Horst, M. A. and K. J. Hellingwerf(2004). Accounts Chem. Res. 37(1): 13–20.

Weber, G. (1950). Biochem. J. 47: 114–121.Yagi, K., N. Ohishi, et al. (1980). Biochemistry

19(8): 1553–7.

Documents

Handbook of Photosensory Receptors (BRIGGS:PHOTORECEPTORS O-BK) || LOV-Domain Photochemistry