Structural basis of photosensitivity in a bacterial
Abigail I. Nasha,1, Reginald McNultyb,1, Mary Elizabeth Shillitob, Trevor E. Swartzc,2, Roberto A. Bogomolnic,
Hartmut Lueckeb,3, and Kevin H. Gardnera,3
aDepartments of Biochemistry and Pharmacology, University of Texas Southwestern Medical Center, 5323 Harry Hines Boulevard, Dallas, TX 75390–8816;
bCenter for Biomembrane Systems, Department of Molecular Biology and Biochemistry, Department of Physiology and Biophysics, Department of
Information and Computer Science, University of California, Irvine, CA 92697–3900; and
California, Santa Cruz, CA 95064
cDepartment of Chemistry and Biochemistry, University of
Edited by J. Clark Lagarias, University of California, Davis, CA, and approved April 22, 2011 (received for review January 11, 2011)
Light-oxygen-voltage (LOV) domains are blue light-activated sig-
naling modules integral to a wide range of photosensory proteins.
Upon illumination, LOV domains form internal protein-flavin ad-
ducts that generate conformational changes which control effector
function. Here we advance our understanding of LOV regulation
with structural, biophysical, and biochemical studies of EL222, a
light-regulated DNA-binding protein. The dark-state crystal struc-
ture reveals interactions between the EL222 LOV and helix-turn-
helix domains that we show inhibit DNA binding. Solution biophy-
sical data indicate that illumination breaks these interactions,
freeing the LOV and helix-turn-helix domains of each other. This
conformational change has a key functional effect, allowing
EL222 to bind DNA in a light-dependent manner. Our data reveal
a conserved signaling mechanism among diverse LOV-containing
proteins, where light-induced conformational changes trigger acti-
vation via a conserved interaction surface.
allosteric regulation ∣ photosensing ∣ PER-ARNT-SIM domain
proteins frequently contain effector domains whose activity is
regulated by specialized sensory domains sensitive to various sti-
muli. One widely distributed class of such sensory domains is the
PAS (PER-ARNT-SIM) family, whose members typically regulate
protein/protein interactions in response to changing environmen-
tal cues (1). A subset of PAS domains, called light-oxygen-voltage
(LOV) domains, use flavin cofactors to detect changes in blue
light intensity or redox state (2). LOV domains are found in reg-
ulatory proteins for phototropism (3), seasonal gene transcription
(4), bacterial stress responses (5, 6), and many other diverse bio-
logical responses. Within these pathways, LOV domains control a
wide range of effector domains, including kinases, F boxes, and
DNA-binding domains (7). Recently, these natural proteins have
been joined by engineered LOV fusions that confer in vitro and in
vivo LOV-based photoregulation to a range of protein targets
This raises the question: How can a class of light-regulated
domains with similar tertiary structures control such a wide vari-
ety of effectors? What is clear is that LOV domains all share
similar architectures and photochemical responses to illumina-
tion, harnessing the energy of incoming blue light photons to
form a covalent adduct between the Sγ sulfur on a conserved
cysteine residue and the C4a carbon of a flavin cofactor (12, 13).
Formation of this bond generates structural changes that propa-
gate to the domain surface, altering the interactions of the core
LOV domain with intra- or interprotein partners (14–18). For ex-
ample, structural studies on Avena sativa phototropin 1 LOV2
(AsLOV2) demonstrated light-induced unfolding of the Jα-helix
located C-terminal to the canonical LOV domain (15). Similarly,
the Neurospora crassa VIVID protein reorients an N-terminal
nvironmental sensory proteins play a crucial function for
cellular adaptation in response to changing conditions. These
α-helix, β-strand extension of its LOV domain upon illumination
(18). In both cases, the external structures interact with the
β-sheet surface of the LOV domain, suggesting a site for signal
propagation common between them. The functional importance
of regulated interactions at this site have been validated by the
ability of point mutations on the β-sheet or interacting effector
surfaces to decouple changes in effector activity from adduct
formation (18, 19).
Among the known LOV-containing proteins are several tran-
scription factors, such as the zinc-finger containing N. crassa
white collar-1 (WC-1) (20) and the algal basic leucine zipper
AUREOCHROMEs (21). Although light controls the binding
of these proteins to DNA, the mechanism(s) of this regulation
is not understood at a molecular level. Here we address thisshort-
coming by examining how a LOV domain directly regulates DNA
binding, establishing the generality of LOV signaling. Our studies
focus on EL222, a 222 amino acid protein isolated from the mar-
ine bacterium Erythrobacter litoralis HTCC2594. In addition to
an N-terminal LOV domain, EL222 also contains a C-terminal
helix-turn-helix (HTH) DNA-binding domain representative of
LuxR-type DNA-binding proteins (22). Combining regulatory
models from a diverse group of LOV-based photosensors (15)
and LuxR-type proteins (23), we hypothesized that the EL222
N-terminal LOV domain represses DNA-binding activity of the
C-terminal domain in the dark, and that this inhibition would be
released with blue light illumination.
Dark-State Crystal Structure of EL222 Suggests Mode to Inhibit DNA
Binding. As an initial step to examining this model, we solved the
2.1-Å resolution crystal structure of EL222 in the dark state
(Table S1), observing interactions between the LOV and HTH
domains consistent with our hypothesis (Fig. 1). The EL222 struc-
ture contains both of the two expected domains, an N-terminal
Author contributions: A.I.N., R.M., H.L., and K.H.G. designed research; A.I.N., R.M., M.E.S.,
H.L., and K.H.G. performed research; A.I.N., R.M., M.E.S., T.E.S., and R.A.B. contributednew
reagents/analytic tools; A.I.N., R.M., H.L., and K.H.G. analyzed data; and A.I.N., R.M., H.L.,
and K.H.G. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factor amplitudes of EL222 have
been deposited in the Protein Data Bank, www.pdb.org (PDB ID code 3P7N) and NMR
chemical shifts with the BioMagResBank, www.bmrb.wisc.edu (accession no. 17640).
1A.N. and R.M. contributed equally to this work.
2Present address: Early Stage Pharmaceutical Development, Genentech, Inc., South San
Francisco, CA 94080.
3To whom correspondence may be addressed. E-mail: firstname.lastname@example.org or Kevin.Gardner@
This article contains supporting information online at www.pnas.org/lookup/suppl/
www.pnas.org/cgi/doi/10.1073/pnas.1100262108PNAS ∣ June 7, 2011 ∣ vol. 108 ∣ no. 23 ∣ 9449–9454
α/β LOV domain and a C-terminal all-helical HTH domain. A
single FMN chromophore was observed within the LOV domain,
orienting the critical isoalloxazine C4a atom only 3.9 Å from the
cysteine (Cys) 75 Sγ atom that is expected to form the photoche-
mical adduct. The LOV domain is followed by a C-terminal
Jα-helix as observed in other LOV structures (15, 24), but here
serves as an interdomain linker that associates more closely with
the HTH effector domain rather than docking onto the LOV
β-sheet surface as in AsLOV2 (15). This arrangement allows
the EL222 LOV β-sheet surface to directly interact with the
4α-helix and 1α-2α loop of the HTH domain. This β-sheet inter-
face is analogous to that used by other LOVand PAS domains to
bind their effectors (25) (Fig. S1), burying approximately 700 Å2
of surface area between the EL222 LOV and HTH domains.
Notably, we observed differences in the relative arrangement
of the LOV and HTH domains in the two molecules of EL222
found in the asymmetric unit, due to the translation of the
HTH domain by approximately 2.5 Å parallel to the axis of helix
4α (Fig. S1E). Although this translation slightly alters the parti-
cular interactions between domains (Table S2), both molecules
still fundamentally use a similar mix of hydrophobic and polar
contacts at the LOV/HTH interface (Fig. 1B). The plasticity of
this interface is consistent with a signaling role, poised for the
facile conversion between conformations via allosteric change
within the LOV domain (26). As in the structures of NarL
and DosR (27, 28), the regulatory LOV domain of EL222
contacts the HTH dimerization helix (4α), but it does not also
directly contact with the DNA-binding recognition helix (3α)
as observed with the regulatory domains of these other structures.
Such structural comparisons supported our hypothesis that
EL222 fails to bind DNA in the dark by both sequestering the
likely dimerization interfaces (LOV β-sheet; HTH 4α-helix)
and the LOV domain interfering with HTH-DNA interac-
tions (Fig. S2).
Photoactivation of EL222 Leads to Adduct Formation and Domain-
Scale Rearrangements. Turning from structure to function, we
examined light-induced changes in the visible absorbance spec-
trum to establish that EL222 can undergo LOV photochemistry.
As expected for a flavin-containing LOV domain, we observed
significant absorbance around 450 nm with vibrational fine struc-
ture (29) (Fig. 2A). This absorbance diminished significantly after
illuminating samples, with three isosbestic points at 330, 384, and
407 nm, consistent with formation of the Cys-FMN adduct. After
ceasing illumination, we observed subsequent dark-state recovery
of the characteristic absorbance profile with first order exponen-
tial kinetics (τ ¼ 25.5 s for 25°C, pH 6.0).
Having established the photosensitivity of EL222, we probed
the ability of adduct formation to generate large-scale conforma-
tional changes using limited proteolysis. Both dark and lit EL222
treated with chymotrypsin demonstrated an initial cleavage re-
moving the N-terminal His6-tag within the first 5 min (Fig. 2B),
but exhibited different behavior with extended incubation times.
Dark-state samples underwent little additional proteolysis, con-
sistent with a well-folded, compact protein. In contrast, lit-state
samples were more quickly and extensively proteolyzed, with little
full-length protein remaining intact after 60 min. Notably, chymo-
trypsin treatment of lit-state samples generated stable fragments,
one of which was consistent with an intact LOV domain (Fig. 2B,
species C). Mass spectrometry established that this fragment was
generated by cleavage within the interdomain Jα-helical linker at
Met159, which packs against the HTH 4α-helix in the dark-state
structure. These data, together with our observation of protease-
resistant fragments in dark conditions, suggest that light-induced
conformational changes increase the accessibility of the Jα-linker
via reorientation of the ordered LOVand HTH domains. This is
supported by limited differences between CD spectra recorded
under dark and lit conditions (Fig. S3).
NMR Studies of EL222 Photoactivation Establish Long-Range Light-
Induced Conformational Changes. To probe these light-induced
changes at higher resolution, we used solution NMR spectro-
scopy. Using a combination of triple resonance and NOESY data,
we assigned15N,13C, and1H chemical shifts of EL222 in the dark
state (81% of the backbone, 40% of the side chain). TALOS ana-
lyses of these chemical shifts (30), combined with through-space
1H-1H NOE data, let us confirm that EL222 has very similar sec-
ondary and tertiary structures in the crystal and solution states.
Notably, solution measurements confirmed that EL222 is mono-
meric under these conditions. Upon illumination, we observed
chemical shift and peak intensity changes at many backbone
and side chain positions as observed in15N∕1H heteronuclear sin-
gle quantum coherence (HSQC) (Fig. 3A) and13C∕1H HSQC
spectra (Fig. S4). Such changes reflect alterations in the local
electronic environments around NMR-active nuclei. Critically,
all spectra maintained comparable chemical shift dispersion in
the dark and lit states, consistent with LOV photochemistry
inducing a domain reorientation, but not unfolding as observed
in AsLOV2 (15).
To identify which sites experienced significant changes, we
compared15N∕1H HSQC spectra recorded under dark and lit
conditions (Fig. 3a), using15N∕1H Scotch exchange spectroscopy
to assign lit-state chemical shifts by correlating dark-state15N
shifts with lit-state1H shifts (31). From the 109 pairs of dark- and
lit-state chemical shifts unambiguously assigned with this analysis,
we established that chemical shift changes occur throughout the
length of the protein (Fig. 3B). Although clusters of perturbed
residues in the LOV domain likely report on adduct-induced con-
figurational changes in the FMN chromophore and resulting con-
formational changes in the surrounding protein, we also clearly
observed long-range (>15 Å from the flavin C4a atom) effects at
sites outside the LOV domain as well. These include changes in
reveals extensive LOV-HTH interactions predicted to
inhibit HTH DNA-binding activity. (A) Overview of
EL222 structure, highlighting locations of the LOV
(blue) and HTH (gold) domains and the Jα-helix
(bronze) connecting the two. The LOV domain binds
to the HTH domain using the LOV β-sheet surface,
(B) Expansion of the LOV/HTH interface as observed
in chain A of the EL222 structure, as indicated by
the boxed region in A. To bind the LOV domain,
the HTH domain presents the 1α-2α linker and 4α-
helix, the latter of which typically provides a dimeri-
zation interface for DNA-bound HTH domains. Thus
sequestered, the 4α-helix is unable to participate in
HTH/HTH interactions observed in many DNA-bound
The dark-state crystal structure of EL222
www.pnas.org/cgi/doi/10.1073/pnas.1100262108Nash et al.
the N-terminal A’α helix that precedes the LOV domain, plus
multiple residues in the HTH domain that are significantly
shifted (Δδ > 0.05 ppm; Fig. 3C). These include several residues
in the 4α-helix, including Leu213, Arg215, Ile216, and Glu219,
plus sites in the 1α-2α (Leu178) and 3α-4α (Lys208, Thr209)
loops. All of these residues are proximal to the LOV β-sheet, sup-
porting our limited proteolysis findings that localized light-driven
cysteinyl adduct formation triggers structural alterations beyond
the LOV domain itself and fully throughout EL222. In addition,
the significant perturbation of residues at the interface between
the domains further supports an interdomain reorientation upon
blue light illumination.
To complement this view from chemical shift changes, we used
NMR-based measurements of backbone amide deuterium ex-
change rates to establish light-induced changes in domain struc-
ture and stability. We obtained these data by resuspending
uniformly15N-labeled samples in D2O-containing buffer, moni-
toring exchange by loss of intensity in consecutively recorded
15N∕1H HSQC spectra. As we have not assigned the lit-state
chemical shifts,2H exchange measurements under illumination
relied on duty-cycling the sample between the dark and lit states,
using assigned dark-state spectra to measure rates. Converting
these exchange rate data into protection factors (32), we found
that numerous sites across the protein exchanged very slowly in
the dark state, consistent with stable hydrogen bonding as ex-
pected from regular secondary structure (Fig. 3D). Many amides
within the LOV domain β-sheet surface are very well protected as
expected for PAS domains (15, 33) and specific residues within
the first and fourth helices of the HTH (1α and 4α) appear re-
fractory to exchange. Upon illumination, these highly protected
regions showed an overall decrease in protection factor, sugges-
tive of distortion in the LOV structure as previously observed in
AsLOV2 (15). The fact that these sites remained protected from
exchange overall is consistent with both the LOV and HTH do-
mains remaining stably folded, and with light inducing a separa-
tion or relative reorientation of the LOV and HTH domains as
suggested by limited proteolysis and chemical shift analyses.
Photoactivation of EL222 Promotes DNA-Binding Activity. These
light-induced structural changes imply a corresponding func-
tional change, which we presumed to be a light-activated DNA-
binding activity, given our data above and the domain architec-
ture of EL222. Without a preestablished biological role of this
protein, we started without any validated DNA-binding site. To
address this issue, we used a candidate-based approach, assuming
that EL222 might be autoregulatory and bind to a DNA sequence
upstream of its own coding sequence. Scanning through the 350-
bp region located 5′ to the start of EL222 translation with a series
of 21 overlapping 45-bp candidate sequences tested, none bound
EL222 as assessed by gel shift assays conducted under dark con-
ditions. However, all of the candidate sequences bound EL222
under illumination at or above 70-μM protein (Fig. S5A), suggest-
ing light-dependent activation of nonspecific DNA binding.
Titrating to lower protein concentrations, we found two se-
quences that bound EL222 at concentrations as low as 7 μM
(Fig. 4 for results of one of these sequences, oligomer 1). In both
instances, DNA binding only occurred when the protein:DNA
mix was incubated under white light. Binding was cooperative
with respect to protein concentration, with a Hill coefficient of
approximately four, suggesting that a pair of dimers bound within
this 45-bp section. No binding occurred under dark-state condi-
tions, even at protein concentrations capable of nonspecific DNA
binding in the light. Protein previously exposed to bright light,
then allowed to recover to dark state overnight at 4 °C also de-
monstrated the same minimal residual DNA-binding activity as
protein that was not exposed to light, indicating the activity is re-
versible and light dependent (Fig. S5B). From these data, we can
conclude that EL222 demonstrates light-dependent DNA-bind-
ing activity. Although the DNA sequence used in these gel shift
experiments bound with the highest affinity of all sequences
tested, we suspect that this is not an optimal binding sequence
for EL222 based on the affinities of similar HTH-containing pro-
teins for their cognate DNA sequences (34, 35). Nevertheless,
these data suggest that this DNA sequence retains its utility
for assaying protein activity in future structural and/or functional
Taken together, our data demonstrate that conformational
changes propagate through the LOV domain upon illumination,
disrupting inhibitory LOV-HTH interactions mediated by the
LOV β-sheet. To test this, we mutated several sites to constitu-
14 35146164 220
with domain-level reorganization. (A) UV-visible absorbance spectra of
EL222, showing the expected absorbance near 450 nm with fine structure
from protein-FMN interactions for dark-state EL222 (black). Illumination in-
duces covalent adduct formation with a loss of absorbance above 400 nm
(red), which gradually returns in the dark with spontaneous adduct decay
(orange through purple, spectra recorded approximately every 5 s). The rate
of dark-state recovery was determined by fitting the absorbance at 450 nm
following illumination. (Inset) Data shown in black and fit to first-order
exponential in red. (B) SDS-PAGE analysis of chymotrypsin limited proteolysis
experiments shows kinetics of degradation are affected by illumination, with
the Jα-helical linker becoming more accessible upon illumination. Signifi-
cantly populated species include His6-EL222 (14–222) (A), EL222 (14–222)
(B), and EL222 (14–156, corresponding to an isolated LOV domain) (C).
Photochemical formation of Cys-FMN adduct in EL222 is correlated
Nash et al. PNAS
June 7, 2011
tively break the LOV/HTH dark-state interaction and generate
proteins locked in the DNA-binding conformation. One of these
mutations, L120K, targeted a hydrophobic patch between the
β-sheet surface of the LOV domain and the HTH 4α-helix
(Fig. S6A). Gel filtration chromatography established that this
mutant is a monomer in solution, as is wild-type EL222
(Fig. S6B). Gel shift assays conducted under dark-state condi-
tions demonstrated that EL222 L120K bound DNA with similar
affinity to wild type under lit-state conditions (Fig. S6C). Limited
proteolysis of L120K using chymotrypsin showed little difference
between the protein in the dark or lit state (Fig. S6D), with both
resembling the lit state of wild-type protein. These results suggest
that the L120K mutation forces EL222 into a lit-state-like struc-
ture that constitutively binds DNA.
Within the context of regulation of HTH-containing proteins, our
data are consistent with the EL222 LOV domain inhibiting DNA
binding in the dark state via interactions with the HTH 4α-helix
and several interhelical loops (Figs. 1, 3, and 5). Disruption of
these interdomain contacts by light-induced conformational
changes in the LOV domain (or mutagenesis of residues at
the LOV/HTH interface) induces DNA-binding activity. A simi-
lar regulatory model is used by other two-domain response
regulator proteins, including the Escherichia coli nitrite/nitrate
response protein NarL. In this case, transfer of a phosphate group
to the N-terminal receiver domain disrupts inhibitory contacts of
this domain with the C-terminal LuxR-type HTH domain, allow-
ing dimerization and DNA binding (27, 36, 37). Studies of
response regulator proteins from NarL and other LuxR family
members indicate that their regulatory domains also contact the
HTH 1α-2α loop and 4α-helix (28, 36, 37), similar to EL222.
Although this aspect of regulation shows strong parallels between
NarL and EL222, we note that they are activated quite differ-
ently. In contrast with the intramolecular mechanism that we
describe for EL222, NarL activation is entirely dependent on a
separate sensor protein (NarQ or NarX) that detects an en-
vironmental signal (nitrate or nitrite) (38, 39) and initiates an
intermolecular phosphotransfer to NarL. Finally, although the
combination of N-terminal sensory and C-terminal HTH DNA-
EL222 acquired under dark (black) or lit (red) conditions show light-induced changes in peak location and intensity. (B) Chemical shift difference analysis of
15N∕1H HSQC spectra shown in Fig. 3A indicate significant changes occurring in both domains, including the HTH 1α-2α loop, 3α-4α loop, and 4α-helix located at
the interface with the LOV domain. Secondary structure elements as indicated by the NMR data and X-ray structure are indicated. (C) Mapping values from
Fig. 3B onto the dark-state crystal structure illustrates the pattern of chemical shift perturbations at the interdomain interface. Side chains are indicated for 1α-
2α loop, 3α-4α loop, and 4α-helix residues in the HTH domain with15N∕1H chemical shift changes upon illumination. (D)2H exchange protection factor analyses
(32) of EL222 conducted in the dark (black) and lit (red) states show similar protection, but to a lower overall degree upon illumination, consistent with
reorganization of two ordered domains. Protection factors >106are lower bound estimates because these sites did not sufficiently exchange for robust fitting
of the time-dependent peak intensity changes.
Solution NMR data suggests EL222 undergoes light-induced rearrangement of two ordered domains. (A) Superposition of15N∕1H HSQC spectra of
www.pnas.org/cgi/doi/10.1073/pnas.1100262108 Nash et al.
binding domains may suggest that EL222 resembles response
regulators that directly detect diffusible small ligands in the cell
(35, 40), we note that some of these proteins may likely be
controlled through ligand-induced protein folding (35) rather
than covalent bond formation as seen in NarL and EL222.
Our results also further validate a conserved aspect of LOV
domain and, more generally, PAS domain signaling via the
β-sheet surface. Many PAS and LOV domains use this surface for
hetero- or homodimerization, whereas others bind different
N- and C-terminal segments that are essential to signaling
(15, 18, 25). Some of these interactions can be modulated by co-
factors within the PAS/LOV domain, providing a ligand-regulated
environmental switch. EL222 extends this paradigm by demon-
strating that fully folded effector domains can bind to this surface,
harnessing conformational changes within the LOV domain to
rearrange the LOV-effector complex (without unfolding the
effectors, as seen with the isolated Jα-helix in AsLOV2; ref. 15).
Notably, these effector domains have different structures but
appear to work through a common mechanism involving the
β-sheet, potentially explaining how a single type of sensory do-
main can regulate a diverse group of effectors (7). Such informa-
tion is particularly useful for both understanding naturally
occurring LOV-regulated proteins and engineering light-regu-
lated systems. These currently include LOV fusions to small
GTPases, metabolic enzymes, DNA-binding domains, and other
enzymes (8–10). All of these designed proteins have taken advan-
tage of the well-characterized signaling mechanism of AsLOV2,
including the PA-Rac1 light-activated GTPase (8). This fusion
protein tethers the photosensory LOV domain closely to the ef-
fector GTPase when the Jα-helix is bound by the LOV domain,
inhibiting enzymatic activity. With the knowledge of the broader
principles provided here by EL222, such engineering may well be
extended to an even larger range of target effectors as part of the
rapidly growing toolbox of “optogenetic” tools (41) that offer pre-
cise spatial and temporal control of protein activity in vitro and
Protein Expression and Solution Characterization. EL222 protein samples were
obtained using standard E. coli heterologous expression and affinity purifi-
cation methods as detailed in SI Methods. Thin layer chromatography estab-
lished that EL222 bound FMN, not FAD or riboflavin. Additional solution
characterization included UV-visible absorbance spectroscopy (60 μM sample;
Varian Cary 50 spectrophotometer), CD spectroscopy (15 μM sample; AVIV
62DS), and limited proteolysis (1∶43 wt∶wt ratio of chymotrypsin∶EL222).
Photoexcited adduct-containing states were generated using a photographic
flash (UV-vis absorbance, CD) or filtered mercury lamp (limited proteolysis).
Crystallographic Structure Determination. Crystals of EL222 were grown using
the hanging drop method, using equal volumes of 8 mg∕mL EL222 (1–222)
and a reservoir of 20% (wt∕vol) PEG 8K, 0.1 M MOPS (pH 7.5), 0.1 M ammo-
nium acetate. X-ray diffraction data were collected from a single crystal on
beam line 7-1 at Stanford Synchrotron Radiation Laboratory. The structure
was solved by four-step molecular replacement using PHASER (42), with in-
dependent search models for the LOV and HTH domains (without a Jα-helix
for the LOV domain). The structure of the Jα-interdomain helix was built
manually as supported by difference density. The initial model of EL222
was subjected to iterative cycles of model building with COOT (43) and sub-
sequent refinement with REFMAC5 (44) and PHENIX (45). Final R and Rfree
values were 26.3% and 32.9%, respectively, with further statistics of the
refinement available in Table S1.
Solution NMR Studies. Solution NMR data were collected at University of
Texas Southwestern using Varian 600 and 800 MHz spectrometers equipped
with cryogenically cooled probes and laser illumination as previously de-
scribed (15), with samples between 250–650 μM. NMR data were processed
using NMRPipe (46) and analyzed with NMRView (47). Backbone and limited
side-chain chemical shift assignments of dark-state EL222 were obtained
using1H-CH3(V/I/L), U-2H,13C,15N-labeled protein and a combination of
2H-modified triple resonance and NOESYexperiments. Lit-state chemical shift
differences were determined using15N∕1H Scotch data to correlate dark- and
lit-state chemical shifts (31), whereas lit-state2H exchange rates were deter-
mined using interleaved dark/lit acquisition (15).
DNA-Binding Studies. DNA-binding activity was assessed using gel shift assays
using32P-labeled dsDNA 45-bp oligonucleotide fragments of DNA located
to the 5′ end of the EL222 gene, as detailed in SI Methods. Gel shift results
presented in Fig. 4 used one of these fragments (oligomer 1, genomic posi-
tion 983532–983577), using a photographic flash to generate the photoex-
cited adduct state.
ACKNOWLEDGMENTS. We thank Brian Zoltowski, Giomar Rivera-Cancel, and
Laura Motta-Mena for assistance with data collection and analysis, and
further thank all members of the Gardner laboratory for constructive
comments provided on this manuscript. This work was supported by grants
20 701014 15 16 18 1912
strates no observable DNA binding to the 45-bp dsDNA oligomer 1 under
dark-state conditions at protein concentrations up to 70 μM. (B) Following
illumination, cooperative DNA binding is observed to the same 45-bp dsDNA
oligomer used for dark-state conditions.
EL222 is a light-activated DNA-binding protein. (A) EL222 demon-
able of binding DNA as the LOV domain sequesters the HTH 4α-helix and has
steric conflicts with DNA if it could bind in monomeric form. The photoche-
mical formation of a cysteinyl/flavin adduct in the LOV domain generates
conformational changes that release inhibitory LOV/HTH interactions and
expose the 4α helix, likely with concomitant changes in the interdomain
LOV/HTH linker. The freed 4α-helix is then free to participate in HTH homo-
dimerization upon binding DNA substrates, as observed in other HTH/DNA
complexes, potentially also involving LOV/LOV interactions between EL222
Model for EL222 activation by blue light. In the dark, EL222 is incap-
Nash et al.PNAS
June 7, 2011
from the National Institutes of Health (R01 GM081875 to K.H.G., R01 AI07000
to H.L., T32 GM008297 supporting A.I.N.), National Science Foundation
(0843662 to R.A.B.), The Robert A. Welch Foundation (I-1424 to K.H.G.),
and a University of California, Irvine Chancellor’s Fellowship to H.L. The
E. litoralis genome sequence data was provided by Stephen Giovannoni’s
laboratory (Oregon State University, Corvallis, OR) and The J. Craig Venter
Institute with grant support from The Gordon and Betty Moore Foundation
Microbial Genome Sequencing Project.
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www.pnas.org/cgi/doi/10.1073/pnas.1100262108Nash et al.
Nash et al. 10.1073/pnas.1100262108
Cloning, Expression,and Purificationof EL222for Crystallographic Stu-
dies. DNA containing full-length EL222, a light-regulated DNA-
binding protein, was cloned into pET/100D TOPO (Invitrogen)
and expressed in Escherichia coli BL21(DE3) cells. Cells were
grown in LB-AMP at 37°C in the dark. Upon reaching A600
0.6–0.8, expression was induced with 1 mM IPTG. After 2 h in-
duction, cells were centrifuged at 4;400 × g for 15 min and sub-
sequently lysed with a French Pressure Cell containing 20 mM
Tris (pH 7.5), 1 mM PMSF, 300 mM NaCl, and one tablet of
Complete™ EDTA-free (Roche) protease inhibitor. Lysates were
centrifuged at 39;800 × g for 1 h before purifying the protein
using FPLC at 4 °C in the dark. First, supernatant was loaded
onto a CO2þ-Talon column with buffer A [20 mM (Tris pH)
7.5, 30 mM NaCl, 10% glycerol]. Elution was performed with buf-
fer B [500 mM imidazole, 20 mM Tris (pH 7.5), 30 mM NaCl,
10% glycerol]. Next, protein was loaded onto a Hitrap Q FF
anion exchange column equilibrated with buffer C [20 mM Tris
(pH 7.5), 30 mM NaCl, 10% glycerol, 1 mM DTT] and eluted
with a linear gradient mixing in buffer D [20 mM Tris (pH 7.5),
1 M NaCl, 10% glycerol, 1 mM DTT]. Protein was then dialyzed
into 20 mM Tris (pH 7.5), 30 mM NaCl, 10% glycerol and con-
centrated to 16 mg∕mL.
Crystallization. Crystallization of purified protein was first
achieved with the Hauptman-Woodward Institute (HWI) robot
(1). We are grateful to Tina Veatch from the HWI for manually
checking that the crystals in the black and white images were
actually colored yellow, consistent with flavin-bound protein.
Subsequently, the protocol was adjusted for hanging drop vapor
diffusion. Drops contained 2 μL of 20% (wt∕vol) PEG 8 K, 0.1 M
MOPS (pH 7.5), 0.1 M ammonium chloride from the reservoir,
and 2 μL of 8 mg∕mL protein in 20 mM Tris (pH 7.5), 300 mM
NaCl. Drops also contained 0.1 M guanidine HCl and 1% glycer-
ol as an additive. Trays were kept in the dark at 16 °C. Crystals
took up to 2 mo to grow. Crystals were harvested under red light
and transferred stepwise to a reservoir solution containing 5%,
10%, and 15% glycerol as a cryoprotectant prior to flash-cooling
with liquid nitrogen.
X-Ray Data Collection. X-ray diffraction data were collected remo-
tely at beam line 7-1 at the Stanford Synchrotron Radiation
Laboratory using a Q315 CCD area detector from Area Detector
Systems Corporation. Two separate datasets, 180° and 360°, were
collected on a single crystal with an oscillation width of 1°. The
exposure time was 20 s per image. The data from both runs were
indexed and integrated using the program d*trek (2) and subse-
quently scaled and merged to 2.1 Å. For data reduction and
refinement statistics, see Table S1.
Structure Solution and Refinement. Search models were generated
with the program SWISS-MODEL (3) based on Protein Data
Bank (PDB) entries 2PR5 and 1YIO for the light-oxygen-voltage
(LOV) domain and helix-turn-helix (HTH) domain, respectively.
The amino acid sequence of each model was substituted with that
of EL222. A four-step molecular replacement search was
performed with the program PHASER (4). Space groups P2
and P21were considered for these sequential six-dimensional
searches, the latter being the correct choice. First, two copies
of the LOV domain were placed, followed by a search for two
copies of the LuxR domain, resulting in an initial R factor of
49.1%. To improve the quality of the phases, solvent flattening
was performed with the program DM (5). The resulting map
was used to build a more accurate model from scratch using
the program BUCCANEER (6). Subsequently, density not mod-
eled by Buccaneer was built manually using the program COOT
(7). Positional and isotropic B-factor refinement was performed
iteratively using the program REFMAC5 (8). A final round of
refinement was done with PHENIX (9). The final R factor
and Rfreewere 26.3% and 32.9%, respectively. Model geometry
was analyzed using the program PROCHECK (10). Coordinates
have been deposited with the PDB/Research Collaboratory for
Structural Bioinformatics (accession code 3P7N).
Differences Between the Two Molecules in the Asymmetric Unit. The
main-chain heavy atom rmsd, computed with the program SSM
(11), between the two independent copies in the asymmetric unit
(molecules A and B) is 1.23 Å, significantly larger than the values
between their individual LOV domains (0.52 Å) and their indi-
vidual HTH domains (0.81 Å). This indicates a rigid-body motion
of LOV vs. HTH domains between the two copies of the protein
in the asymmetric unit (Fig. S1E), specifically a relative transla-
tion of one HTH domain by about half a helix pitch (2.5 Å) par-
allel to the axis of helix 4α. Although this results in some
differences in interdomain interactions (Table S2), both mole-
cules still involve the same residues in making a mix of hydropho-
bic and polar contacts at the LOV/HTH interface (Fig. 1B).
Using PISA (12) to analyze the interfaces between the LOV
and HTH domains (disregarding the two Jα-helices bridging pairs
of domains), we found that the LOV/HTH interface buries 714
and 612 Å2of surface area for molecules A and B, respectively
(Table S2). Consistent with the smaller surface area, molecule B
also has a smaller number of hydrogen bonds at the LOV/HTH
interface (eight for molecule A, four for B) and is predicted to
have a slightly smaller solvation free energy gain for complexing
the LOVand HTH domains (−6.8 kcal∕mol for A, −6.6 kcal∕mol
for B) (Table S2) (12). Finally, we also noted differences between
the main-chain temperature factors of the two molecules (37 Å2
for A, 46 Å2for B), albeit with the same pattern of lower and
higher values along the length of the protein (Fig. S7).
Cloning, Expression, and Purification of EL222 for NMR and Functional
Studies. DNA encoding full-length EL222 and a 13-residue
N-terminal truncation (residues 1–222 and 14–222, respectively)
were subcloned into the expression vector pHis-Gβ1-Parallel1, a
derivative of the pHis-Parallel1 vector (13, 14). Mutagenesis was
carried out using QuikChange II XL from Stratagene according
to manufacturer’s instructions. E. coli were transformed and
grown in either LB broth or M9 minimal media as described
for individual experiments below. Cultures were grown at 37°
C to an A600of 0.6–0.9 and then induced at 20°C in the dark
by addition of 0.5 mM IPTG. After 16 h induction, cells were
centrifuged and the resulting pellets resuspended in 50 mM Tris,
100 mM NaCl, pH 8.0 buffer at 4 °C and lysed by sonication.
Lysates were clarified by centrifugation at 48;000 × g for 30 min.
The resulting supernatant was loaded on a Ni2þ-nitrilotriacetate
(NTA) column, allowing for rapid affinity purification of His-Gβ1
tagged proteins by gradient elution, mixing in the same buffer
containing additional 500 mM imidazole. After exchanging the
protein-containing fractions into 50 mM Tris, pH 8.0 buffer,
the His-Gβ1 tag was cleaved using 1 mg His6-Tobacco Etch Virus
(TEV) protease (15) per 30 mg of fusion protein. Proteolysis re-
actions were incubated overnight at 4°C and stopped using a
Ni2þ-NTA column to remove the His6-Gβ1 tag and His6-TEV
Nash et al. www.pnas.org/cgi/doi/10.1073/pnas.11002621081 of 9
protease. Postcleavage, the resulting protein contains only three
vector-derived residues, GEF at the N-terminus. TEV-cleaved
protein was loaded onto a MonoQ column to separate small
remaining impurities. Highly pure protein was eluted with a lin-
ear gradient mixing in 50 mM Tris, 1 M NaCl pH 8.0 buffer,
exchanged into 50 mM sodium phosphate, 100 mM NaCl pH
6.0 buffer, and concentrated to a final protein concentration
of 100–250 μM.
Thin LayerChromatographyAnalysisof FlavinContentin EL222.Given
that LOV domains are commonly associated with either FMN or
FAD as their flavin cofactor, we experimentally determined
that EL222 preferentially associates with FMN using thin layer
chromatography (TLC) analysis. Samples of full-length EL222
(1–222) were exchanged into H2O and concentrated to approxi-
mately 250 μM. A 2× volume of ethanol was added to the sample
and incubated for 2 min in a boiling water bath. The denatured
protein sample was blotted onto a silica gel-based TLC plate, with
standard samples of FMN, FAD, and riboflavin (250 μM each) in
separate lanes. Separation proceeded in a 12∶3∶5 n-butanol:
acetic acid: H2O solvent system, and the resulting bands were
visualized with a UV lamp. This analysis showed that recombi-
nantly expressed EL222 only associates with FMN, with no sig-
nificant incorporation of FAD or riboflavin.
ble absorbance spectra were measured on a Varian Cary Series 50
spectrophotometer from 250–550 nm. Dark-state spectra were
obtained on 60-μM samples exposed only to red light for the pre-
ceding 24 h, whereas lit-state spectra were obtained immediately
after exposing the sample to illumination from a photographic
flash. Kinetic experiments monitored the return of the A450signal
following illumination. Data points were fitted using a first-order
exponential to obtain the reported time constant (τ).
Limited Proteolysis. Full-length (1–222) EL222 samples which re-
tained an N-terminal His6tag were used for limited proteolysis,
providing an internal control for proteolytic activity between the
His6tag and the LOV domain. A 1∶43 ratio (wt∕wt) of chymo-
trypsin to protein was used in a single volume, with subsequent
samples collected from a common stock. Samples collected for
each time point were stopped by the addition of SDS loading buf-
fer containing 25% glycerol and visualized on 20% SDS-PAGE
gel. For mass spectrometry analysis, reactions were stopped with
1% trifluoroacetic acid. Dark-state experiments were conducted
under dim red light, whereas lit-state experiments were per-
formed under constant irradiation produced by an Oriel mercury
lamp (model number 66902) at 50 mW power with a broadband
blue-green filter (Oriel no. 51970).
Circular Dichroism Spectroscopy. A total of 500 μL of 15 μM full-
length EL222 was used for each CD experiment, using a 1-mm
path length cell. Dark-state spectra were collected under com-
plete darkness, whereas lit-state spectra were recorded following
exposure to photographic flash. CD data were collected using a
wavelength range from 195 to 260 nm at 12 °C with 1.5-nm band-
width and 3 s averaging time. For lit-state spectra, data collection
was paused every 9 s and the sample reexposed to a photographic
flash to ensure lit-state protein. Final data were generated from
an average of three repeats.
Solution NMR Spectroscopy.All of the following NMR experiments
utilized uniformly15N,13C-labeled EL222(14–222) protein, gen-
erated from bacteria grown in M9 minimal media containing
1 g∕L of15NH4Cl and 3 g∕L of13C6glucose. Solution NMR
experiments were performed on Varian Inova 600 and 800 MHz
spectrometers equipped with cryogenically cooled probes at
25 °C, using NMRPipe (16) for data processing and NMRview
(17) for analysis. Lit-state15N∕1H and13C∕1H heteronuclear sin-
gle quantum coherence (HSQC) spectra were acquired with a
488-nm, 200-mW Coherent Sapphire laser. The output from this
laser was focused into a 10-m long, 0.6-mm diameter quartz fiber
optic. The other end of the fiber was placed into the bottom of a
coaxial insert tube (Wilmad) inside a 5-mm NMR sample tube,
allowing the illuminated tip to be immersed in protein solution
without contamination. Power level measurements were con-
ducted prior to every experiment to establish the efficiency of
coupling the laser output to the fiber optic, and all power levels
reported here are those measured at the end of the fiber. Lit-state
15N∕1H and13C∕1H HSQC spectra were recorded by preceding
each transient in the experiment with a 50-mW, 120-ms laser
pulse during the 1.06-s delay between transients (14).
Backbone chemical shift assignments were acquired with
2H-modified triple resonance data on a uniformly15N,13C,2H-
labeled sample. This sample was prepared with M9 media for
protein growth composed of D2O, 1 g∕L15NH4Cl, and 3 g∕L
13C6glucose as the sole carbon source for U-2H∕15N∕13C
protein (14–222). The purified protein was buffer exchanged into
50 mM MES, 100 mM NaCl pH 6.0 buffer and concentrated to
250 μM. Spectra were taken at 25 °C on a Varian Inova 600 MHz
spectrometer fitted with a triple resonance cryoprobe. Assign-
ment of the backbone and Cβ NMR resonances was achieved
with the following 3D NMR experiments modified for use on
highly deuterated samples: HNCA (18), HN(CO)CA (19),
HNCACB (19), HN(CO)CACB (19). Side-chain13C and1H re-
sonances were assigned with15N∕13C-edited NOESY spectra on
protein grown in M9 minimal media for U-15N∕13C protein. Ad-
ditional assignments were obtained using (H)C(CO)NH-total
correlation spectroscopy (20) on perdeuterated protein with pro-
tonated methyl groups at the valine γ, leucine δ, and isoleucine δ1
positions. This protein was produced by growing E. coli in D2O
based M9 minimal media containing both15NH4Cl and13C6glu-
cose, supplemented 30 min prior to induction with 80 mg∕L
α-ketovalerate and 50 mg∕L α-ketobutyrate (21).
To measure deuterium exchange rates on EL222 using solution
NMR methods, a 550-μL sample of 600 μM15N-labeled protein
(14–222) in 50 mM sodium phosphate (pH 6.0), 100 mM NaCl,
pH 6.0 buffer was lyophilized. Immediately prior to acquiring
spectra, the lyophilized sample was rehydrated with 550 μL D2O
and placed in the magnet.15N∕1H HSQC spectra were acquired
as an automated series of 79–90 datasets, each taking approxi-
mately 20 min to complete. Lit-state spectra were obtained using
a 50-mW laser pulse as described above. The lit-state spectra
were recorded interleaved with dark-state spectra, letting us take
advantage of the chemical shift assignments in the dark state.
Protection factors were then calculated based on standard meth-
ods (22), with a correction for the lit-state rates given the inter-
conversion between the dark and lit states.
DNA Binding as Assessed by Electrophoretic Mobility Shift Assay. We
designated 45-bp lengths of DNA within 350 bp of the gene start
site to use in a gel shift assay. Oligonucleotides (Integrated DNA
Technologies) were staggered to cover all possible binding sites in
this region. Lyophilized DNA was resuspended in 50 mM Tris,
100 mM NaCl, 2 mM MgCl2, 12% glycerol, pH 8.0 buffer.
Reverse complementary pairs were annealed by heating 40 μL
of 100 pmol∕μL DNAto 95 °C in a heat block for 5 min., followed
by slow cooling to room temperature over 1 h. Following anneal-
ing, 100 ng of DNA were labeled with32P ATP using T4 polynu-
cleotide kinase in a 20 μL reaction volume. Unincorporated
32P was purified away using ProbeQuant G50 Microcolumns
(GE Healthcare). Concentrations were estimated assuming
80% recovery from column. Unlabeled protein (both 1–222
and 14–222) was purified and concentrated to 250 μM. Reaction
conditions included 0.01 mg∕mL BSA, 0.02 mg∕mL deoxyino-
sine/deoxycytosine, 0.04 ng∕μL
32P-labeled DNA (1.45 nM),
Nash et al. www.pnas.org/cgi/doi/10.1073/pnas.11002621082 of 9
and varying concentrations of protein in 50 mM Tris, 100 mM
NaCl, 2 mM MgCl2, 12% glycerol, pH 8.0 buffer. Protein was
the final addition to each reaction followed by brief centrifuga-
tion and incubation in ice slush for 30 min. For dark-state reac-
tions, samples were prepared under dim red light and incubated
in a covered ice bucket, whereas lit-state reactions were con-
ducted under bright white light illumination in a clear glass bea-
ker with periodic exposure to camera flash. After incubation,
samples were separated on a 5–10% TAE gels at 4°C for 2 h
at 100 V. Samples allowed to recover after illumination were
flashed in the absence of DNA, then stored in the dark at 4°C
for 24 h to allow complete recovery of the dark-state conforma-
tion. Again, this was conducted under either dim red light or
bright white light. Gels were dried with a heated slab gel dryer
fitted with vacuum pump for 1 h and developed using FujiFilm
FLA-5100 imaging system following exposure to PhosphorIma-
ging plates. Quantification of percent DNA bound was per-
formed using FujiFilm MultiGauge v2.3 software (FUJIFILM
Medical Systems USA, Inc.).
These surveys of EL222 binding to sequences within the
350 bp located 5′ of the start of EL222 translation indicated
two sequences that exhibited higher affinity binding: oligomer
1 (genomic base pairs 983532–983577), GGTAGGATCCATC-
mer 2 (genomic base pairs 983647–983692), GGCCCCGAGGT-
Experimental data presented in the main text and Fig. S5
utilized oligomer 1; Fig. S5 also utilized a lower affinity oligomer
from elsewhere in the EL222 promoter (base pairs 983469–
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A EL222: LOV-4α
B AsLOV2: LOV-Jα
C VVD: LOV-aα/bβ
D YtvA: LOV-LOV
E EL222: LOV-4αvs. AsLOV2: LOV-Jα
aligned to a common view, presenting the β-sheet surface toward the reader. Effector regions located outside the canonical LOV domain are shown in bright
pink, with regions of the LOV domain surface located within 5 Å of those effectors colored in magenta. This alignment shows that all of these structures share
the use of the LOV β-sheet surface to interact with their effectors, despite the diversity ofsurfaces they provide, including predominantly α-helical surfaces from
(A) 4α-helix of EL222 HTH domain and (B) Jα-helix of Avena sativa phototropin 1 LOV2 (AsLOV2) (1, 2), (C) mixed α/β surface from Vivid (VVD) (3), and
(D) predominantly β-sheet surface from the YtvA LOV–LOV homodimer (4). We emphasize that this same interface is used by both of the EL222 molecules
observed in the crystallographic asymmetric unit, despite the translation of the HTH domains by approximately 2 Å along the direction of the C-terminal
4α-helix. This is demonstrated in E by a superposition of LOV domains from EL222 and AsLOV2 (gray) and indicating the 4α-helices from EL222 molecules
A and B (green and blue, respectively) with the Jα-helix of AsLOV2 (magenta). This demonstrates that all of these helices associate with similar positions
on the LOV domain surface, but with slightly different orientations (relative approximately 35° rotation between EL222’s 4α and AsLOV2’s Jα-helices).
Consistent with this movement, the main-chain atom rmsd between all residues in the two EL222 molecules (1.23 Å) is higher than that between the isolated
LOV (0.52 Å) and the HTH domains (0.81 Å). Interactions within 5 Å of the C-terminal helix of molecule A with its LOV domain are highlighted magenta.
LOV domains utilize a common β-sheet interface for interacting with a wide range of effectors. Structures of four LOV-containing proteins have been
1 Harper SM, Neil LC, Gardner KH (2003) Structural basis of a phototropin light switch. Science 301:1541–1544.
2 Halavaty A, Moffat K (2007) N- and C-terminal flanking regions modulate light-induced signal transduction in the LOV2 domain of the blue light sensor phototropin 1 from Avena
sativa. Biochemistry 46:14001–14009.
3 Zoltowski BD, et al. (2007) Conformational switching in the fungal light sensor Vivid. Science 316:1054–1057.
4 Möglich A, Moffat K (2007) Structural basis for light-dependent signaling in the dimeric LOV domain of the photosensor YtvA. J Mol Biol 373:112–126.
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of TraR/DNA complex (1), with TraR dimer shown as green and red monomers. (B) Superposition of one TraR molecule (green) and EL222 (gold, blue) by least-
squares fitting of the HTH domains of both proteins (shown with solid ribbons, whereas the regulatory domains are partially transparent). (C) Ribbon diagram
of EL222 alone (in orientation shown in B) demonstrates that the LOV and HTH domains are oriented in the dark-state crystal structure of EL222 in a mode
inconsistent with DNA binding without major domain rearrangement. In particular, the LOV domain clashes with DNA and occludes the HTH 4α-helical
LOV-HTH interactions observed in the EL222 crystal structure are predicted to interfere with HTH dimerization and DNA binding. (A) Crystal structure
1 Vannini A, et al. (2002) The crystal structure of the quorum sensing protein TraR bound to its autoinducer and target DNA. EMBO J 21:4393–4401.
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[θ]mrw (deg cm2 dmol-1) * 104
at 208 and 222 nm suggest well-folded protein with significant α-helical character in the dark-state (black), consistent with the crystal structure of this protein.
Following illumination, little change was observed in the secondary structure characteristics of the protein (red).
Circular dichroism spectra of dark- and lit-state EL222 indicate minimal change in secondary structure content upon illumination. The double minima
0.0 1.0 2.0
1.92 2.02 2.12 2.22
(red) conditions. Analysis of the methionine Cϵ region located near 2 ppm1H, 15 ppm13C (and validated by changes in peak sign in constant time13C∕1H HSQC
spectra) indicated a perturbation in one Met Cϵ methyl group upon illumination. Because all the methionine residues in EL222 are located outside the canonical
LOV domain, this supports the long-range propagation of light-induced chemical shift and structural changes.
Light-induced chemical shift changes in the methyl region of13C∕1H HSQC spectra of EL222. Overlaid spectra were acquired under dark (black) or lit
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0.100 5 10 20 70
0.1 5 10 20 70
highest specificityaverage/low specificity
A Differential affinity of DNA sites for EL222
B Reversibility of DNA binding post-illumination
dark-state binding light-state binding
25 12025 12µM0
25 12025 12µM
permanently triggered by illumination. (A) Comparison of the EL222 concentration dependence of EMSA band shifts for a higher affinity site (oligomer
1, GGTAGGATCCATCGGGCAGTGCGGTCAGCGGCATGCCGGCAGCAG, genomic base pairs 983532–983577) with a lower affinity one located elsewhere in
the EL222 promoter (ACAGCAATTGCAATGGTGCCGCGAGGGCTGTGAACTACCTGTTGC, genomic base pairs 983469–983513) reveals an approximate fivefold
difference in binding affinity under these conditions. (B) DNA binding requires illumination and is not permanently induced by exposure to light. Comparison
of EMSA assays of samples that have not been exposed to significant blue light (“pristine”) and those that have been illuminated then allowed to recover in the
dark for 24 h at 4°C (“recovered”) show that neither bind the high affinity oligomer 1 significantly in the dark (maximum DNA binding ¼ 2–4% at 12 μM
EL222). Both proteins bind DNA avidly and equivalently when illuminated, with approximately 100% DNA bound under the same condition.
EL222 binds with differential affinities to different DNA sequences in the EL222 promoter and does so in a light-dependent manner that is not
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14 16 20
LOV domain (blue) and points down toward 4αof the HTH (gold). (B)The L120K mutant appears to be monomeric (black), similarto wild type (red), as shown by
comparable retention times in Superdex 75 gel filtration chromatography. The elution volumes for three standards are shown: ovalbumin, 43 kDa; chymo-
trypsin A, 25 kDa; RNase A, 13.7 kDa. (C) Electrophoretic mobility shift assays show L120K binds DNA (oligomer 1) in the dark with comparable affinity as wild-
type EL222 in the light. However, L120K appears to have higher order binding at higher protein concentrations (labeled new complex) that wild type does not
demonstrate. (D) Limited proteolysis with chymotrypsin in the dark and lit states reveals little change in enzyme accessibility with both states behaving similarly
to the wild-type lit-state protein.
A point mutation predicted to disrupt LOV-HTH interactions in EL222 leads to constitutive DNA binding. (A) L120K is located on the Hβ-strand of the
represented as a heat map (low, blue; high, red, ranging from 20 to 86 Å2). The average B factor for molecule B (46.3 Å2) is considerably higher than that of
molecule A (37.1 Å2). In both molecules, the HTH domain has a higher average B factor than the corresponding LOV domain and, in turn, the Jα-linker helix has
a higher B factor than either the LOV or HTH domains.
Differences in temperature factors between the two EL222 molecules in the asymmetric unit. B factors of molecule A and B (A and B, respectively) are
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Table S1. Crystallographic data reduction and refinement statistics for Download full-text
the EL222 dark-state structure
Unit cell parameters
Matthews coefficient, Å3∕Da
No. of molecules in the asymmetric unit
Solvent content, %
36.93 - 2.10 (2.18 - 2.10)
Resolution range, Å
No. of reflections
Avg B factor, Å2
Rmsd bond lengths ðÅÞ∕angles, °
36.93 - 2.10
Ramachandran plot, %
Most favorable regions
No. of protein atoms
No. of water molecules
Note, values in parentheses are for the highest resolution shell (2.18–2.10 Å).
Table S2. Molecular interactions are different at the LOV-HTH domain interface for the two
molecules in the asymmetric unit
Buried interface area, Å2
ΔGi, kcal∕molΔGiP valueHB SB
EL222 molecule A without Jα
EL222 molecule B without Jα
There is a difference in the number of hydrogen bonds (HB) for molecules A (8) and B (4) and no salt bridges
(SB) are present at the interface for EL222. The computed solvation free energy gains (ΔGi) of the domain/
domain interactions are favorable for molecules A (−6.8 kcal∕mol) and B (−6.6 kcal∕mol), both of which are
significantlylower thanthat ofAvena sativa
(−15.3 kcal∕mol). The ΔGi P values (P < 0.5) of all interactions support that the interface surfaces are
interaction specific. All information in this table was computed using the PISA (Protein Interfaces, Surfaces,
and Assemblies) service at the European Bioinformatics Institute (http://www.ebi.ac.uk/msd-srv/prot_int/
1 Krissinel E, Henrick K (2007) Inference of macromolecular assemblies from crystalline state. J Mol Biol 372:774–797.
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