From the Department of Biochemistry and Biophysics, University of
North Carolina School of Medicine,
Chapel Hill, North Carolina 27599-7260
DNA photolyases repair pyrimidine dimers via a
reaction in which light energy drives electron donation from a
catalytic chromophore, FADH
, to the dimer. The
crystal structure of Escherichia coli photolyase suggested
that the pyrimidine dimer is flipped out of the DNA helix and into a
cavity that leads from the surface of the enzyme to FADH
.
We have tested this model using the Saccharomyces
cerevisiae Phr1 photolyase which is >50% identical to E. coli photolyase over the region comprising the DNA binding
domain. By using the bacterial photolyase as a starting point, we
modeled the region encompassing amino acids 383-530 of the yeast
enzyme. The model retained the cavity leading to FADH
as
well as the band of positive electrostatic potential which defines the
DNA binding surface. We found that alanine substitution mutations at
sites within the cavity reduced both substrate binding and
discrimination, providing direct support for the dinucleotide flip
model. The roles of three residues predicted to interact with DNA
flanking the dimer were also tested. Arg452 was found to be
particularly critical to substrate binding, discrimination, and
photolysis, suggesting a role in establishing or maintaining the dimer
in the flipped state. A structural model for photolyase-dimer interaction is presented.
 |
INTRODUCTION |
Pyrimidine bases absorb strongly in the UV region and are highly
susceptible to photochemical reactions that alter their structures. In
DNA, cyclobutane pyrimidine dimers
(CPDs)1 and
pyrimidine-pyrimidone (6-4) photoproducts are the most frequent and
biologically significant products of these reactions. These lesions are
lethal and mutagenic and must be repaired to ensure cell survival and
genetic stability. DNA photolyases repair CPDs and (6-4) photoproducts
via reactions in which near UV or visible light provides the energy for
bond rearrangements which restore the pyrimidines to their undamaged
state. Two types of DNA photolyases have been recognized with regard to
substrate specificity as follows: cyclobutane dipyrimidine photolyases
and (6-4) photolyases (see Ref. 1 for a recent review). Each type of
enzyme recognizes and efficiently repairs a single type of damage (2,
3). The cyclobutane dipyrimidine photolyases, hereafter referred to as
CPD photolyases, are the subject of this report.
Understanding how the CPD photolyases efficiently repair pyrimidine
dimers in DNA entails answering the following two questions: how do the
enzymes recognize pyrimidine dimers specifically in the midst of a vast
excess of nondamaged bases, and how do the enzymes catalyze dimer
photolysis? Photolysis involves two noncovalently bound chromophores,
reduced FAD and a second chromophore which, depending upon the source
of the enzyme, is either folate or deazaflavin (4). Absorbance of a
photon of photoreactivating light subsequent to substrate binding
initiates electron donation by enzyme-bound FADH
to one
of the bases in the dimer (4, 5). The photon may be absorbed either
directly by FADH
or, more often, by the second
chromophore which transfers energy to the flavin chromophore (1). Both
electron transfer and energy transfer are highly efficient processes,
resulting in a quantum yield for the overall photolysis reaction of
0.6-1.0 for CPDs containing thymine or uracil (6,
7).2
CPD photolyases are structure-specific enzymes that display binding
discrimination comparable to that seen for sequence-specific DNA-binding proteins. Studies on the Escherichia coli and
Saccharomyces cerevisiae enzymes have shown that the
equilibrium association constant for
(cis,syn)-CPDs in DNA is approximately
109 M
1, whereas the association
constant for nondamaged DNA is 103
M
1 (6, 8-10). The affinity for
(trans,syn)-pyrimidine dimers in DNA and U<>U dimers
in RNA is only about 10-fold greater than that for nondamaged DNA;
nevertheless, once bound, these lesions are photolyzed efficiently (7,
11). Thus the presence of a cyclobutane dimer, the geometry of the
bases in the dimer, and the absence of a 2'-OH on the sugar phosphate
backbone are determinants of binding specificity. Important recognition
elements are also found in DNA flanking the dimer. In particular,
ethylation of the first phosphate 5' to the dimer and 3-4 phosphates
3' to the dimer in the lesion-containing strand inhibits binding (12, 13). At least some of these interactions contribute to binding specificity as shown by the fact that discrimination between
dimer-containing and undamaged oligonucleotides decreases as the
substrate is shortened (6). In addition, mutations in the yeast Phr1
photolyase have been identified which simultaneously decrease substrate
discrimination and alter interactions with DNA phosphates surrounding
the dimer (9). These results imply that the structure of the flanking sugar-phosphate backbone is uniquely altered by the dimer.
A structural basis for the efficiency of the photolysis reaction and
for specific substrate recognition has been provided by the crystal
structure of E. coli photolyase (14). The polypeptide chain
is folded into an amino-terminal
/
domain and a carboxyl-terminal helical domain with the folate cofactor nestled into a shallow cleft
between the two domains. The flavin chromophore lies deeply buried in
the center of the helical domain, with direct access to solvent limited
to a cavity leading from the edge of the isoalloxazine ring of flavin
to the surface. The cavity lies in the center of a trace of positive
electrostatic potential that runs along the flat face of the helical
domain, and both the dimensions of the cavity and the asymmetric
distribution of hydrophobic and polar residues within the cavity are
appropriate to accommodate a pyrimidine dimer. These features of the
photolyase structure led Park and co-workers (14) to propose that
E. coli photolyase binds DNA along the trace of positive
electrostatic potential and that in the enzyme-substrate complex the
dimer is flipped out of the DNA helix and into the cavity leading to
FAD. Here we report the results of mutational studies designed to test
the model for photolyase binding using the yeast Phr1 photolyase. Phr1
and E. coli photolyases contain identical chromophores and
are 50% identical in primary sequence over the region encompassing
most of the proposed DNA binding surface and the cavity leading to
flavin. Our results provide the most detailed picture yet of
interactions at the photolyase-DNA interface, support the dinucleotide
flip model, and suggest that interactions between DNA and residues
inside and outside of the cavity contribute to binding affinity, to
substrate discrimination, and to maintaining the dimer in the flipped
state.
 |
EXPERIMENTAL PROCEDURES |
Protein Modeling--
Modeling was performed using the program
FRODO (PSX, version 6.6) on an Evans and Sutherland PS300 Graphics
system. The E. coli photolyase crystal structure (see Ref.
14; Protein Data Base code 1dnp) was used as the starting point for
modeling the structure of the DNA-binding site of yeast photolyase. The region of the
-helical domain of E. coli photolyase
encompassing helices 11 through 18 (amino acids 273-419) contains most
of the residues predicted to interact with DNA in and surrounding the cavity leading to the flavin chromophore (14). Within this region the
overall sequence identity between the two enzymes is 50% (Fig. 1). To model this region, the 73 nonidentical amino acids in the E. coli photolyase sequence
were replaced with the corresponding amino acids from the S. cerevisiae Phr1 sequence (15). Each new amino acid was subjected
to a modeling and refinement protocol. First, the amide nitrogen,
carbon, and amide carbon of each amino acid were placed in the position
of the previous E. coli amino acid. The yeast amino acid
side chain was then rotated about its first dihedral rotation angle
(
1) to produce an alignment devoid of steric clash.
Because
1 displays a strong tendency to assume values
near 60, 180, and 300°, each new side chain was examined first in
each of these positions. The preferred alignment was at a
1 value that placed the yeast side chain near the
E. coli side chain and did not produce any steric clashes
(within 2.5 Å). At positions 285, 291, and 368 (Phr1 positions 395, 401, and 478), alternative values for
1 had to be
chosen. A similar protocol was used to model the dihedral angle
2 between the
and
carbons on Gln, Glu, Lys, Arg,
Met, Phe, Tyr, and Trp side chains. Again, the preferred dihedral
values were those that placed the yeast side chain near the
corresponding E. coli side chain. Steric clashes at
positions 307, 308, and 411 (Phr1 positions 417, 418, and 521)
prohibited placing these side chains in the preferred positions. The
program REFINE, written by Dr. Jan Hermans (University of North
Carolina, Chapel Hill), was then used to refine the geometry (bond
lengths, bond angles, and dihedral angles) for the S. cerevisiae model. Individual amino acids were first refined, then
the entire structure was analyzed over the course of 10 cycles. After
10 cycles, the cumulative coordinate shift was determined, and further
modeling was performed until this shift value decreased to less than
5.0 Å. An identical approach was used to replace Arg226,
which lies near the rim of the cavity leading to flavin, with the
equivalent yeast residue, Lys330.

View larger version (37K):
[in this window]
[in a new window]
|
Fig. 1.
Alignment of E. coli and S. cerevisiae photolyases over the region modeled for the yeast Phr1
photolyase. Numbering above and below the
sequences refer to the positions in the E. coli and yeast
enzymes, respectively. Identities are indicated by *. Sites of
interaction (direct or water-mediated hydrogen bonds) between the
apoenzyme and the flavin ( ) and folate ( ) chromophores in the
E. coli crystal structure are marked. Amino acids selected
for alanine substitution mutagenesis (yeast amino acids
Lys383, Glu384, Trp387,
Arg452, Phe494, and Gln514) are
shown enclosed in black boxes. Lys330 was also
modeled and mutated.
|
|
Construction of PHR1 Mutants--
The entire PHR1
coding region and approximately 500 bp of 3'-flanking sequence were
subcloned from pCB1241/recon (9) into XmnI-PstI-digested pMal-c2 (New England Biolabs),
yielding pGBS424. Oligonucleotides used in the construction of the
various mutants are listed in Table I,
and the locations of the introduced mutations and relevant restriction
sites are shown in Fig. 2. To facilitate construction of the K330A mutant, PCR mutagenesis was used to introduce
a SmaI site into PHR1 at nucleotides 684-689
(relative to the first PHR1 ATG; Fig. 2), yielding plasmid
pGBS425. The amino acid sequence of photolyase was not changed in this
construction. Alanine substitution mutations were introduced at
Lys330, Glu384, Arg452,
Phe494, and Gln514 using two-primer
(Lys330, Arg452) or four-primer PCR mutagenesis
(Lys383, Glu384, Phe494, and
Gln514) and pGBS425 as template. Vent DNA polymerase (New
England Biolabs) was used in the first PCR stage (25 cycles). For
4-primer PCR, the first stage PCR products were purified on 3% GTG
agarose gels (FMC Bioproducts), recovered in sterile water, and used
for the second PCR reaction in conjunction with appropriate outside
primer pairs (Table I). Amplification was carried out for 15 cycles using Taq polymerase (Life Technologies, Inc.). Purified PCR
products were digested with appropriate restriction endonucleases and
cloned into pGBS425 which had been digested with the same set of
enzymes as follows: KpnI and XbaI for K383A and
E384A, XbaI and PstI for F494A and Q514A,
KpnI and XbaI for R452A, and SmaI and
KpnI for K330A (Fig. 2). In each case the nucleotide
sequence of the amplified region was verified to ensure that no
additional mutations were introduced by the PCR. The photolyase bearing
three substitutions (K330A/E384A/F494A) was constructed by subcloning
restriction fragments containing the E384A and the F494A mutations into
the K330A plasmid using unique restriction sites surrounding each mutation. To express PHR1 without any attached fusion
protein, the ~2.4-kilobase pair BglII-PstI
fragment from each mutant was subcloned into similarly digested
pCB1241/recon. Construction of mutant W387A has been described
previously (9).

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 2.
Map of the restriction sites used in
constructing the PHR1 mutations. The direction of
transcription of PHR1 is indicated by the arrow.
Restriction sites and their locations relative to the first base in the
PHR1 open reading frame are shown above the
line, and the sites of PCR-generated mutations are shown
below the line.
|
|
Photolyase Purification and Spectral Characterization--
All
photolyases were overexpressed from pCB1241/recon derivatives (no
maltose binding protein fusion) in E. coli strain
CSR603F'lacIQ (Phr
) and purified
as described previously to greater than 95% purity as judged by
Coomassie Blue staining (9). The protein concentration and chromophore
content for each protein preparation were determined by spectroscopy
from 220 to 700 nm (9). The spectra were closely examined for the
presence of peaks near 450, 480, 580, and 625 nm, which are indicative
of oxidation of the reduced flavin chromophore to the blue neutral
radical or FADox (16).
Quantitation of Specific and Nonspecific Equilibrium Association
Constants (KA, KS, KNS)--
A 43-bp
DNA substrate containing a centrally located thymine dimer was prepared
for these studies as shown in Oligonucleotide 1.
A (cis,syn)-thymidine dimer (a gift from Xiaodong
Zhao and Aziz Sancar) was synthesized and incorporated into the top
strand oligonucleotide shown using standard oligonucleotide coupling protocols (17). Full-length oligonucleotide was purified from a
denaturing polyacrylamide gel, quantitated by absorbance at 254 nm, and
labeled at the 5' end using T4 polynucleotide kinase and
[
-32P]ATP (>6,000 Ci/mmol). The bottom strand
oligonucleotide was quantitated by absorbance at 254 nm, and the two
oligonucleotides were mixed at a concentration of 500 nM
dimer oligonucleotide and 1500 nM bottom strand
oligonucleotide in 20 mM Tris-HCl (pH 8.4), 50 mM KCl, heated at 90 °C for 5 min, and then allowed to anneal by slow cooling to room temperature. This procedure resulted in
incorporation of >90% of the dimer oligonucleotide into
double-stranded DNA. Oligo(dT)18 (Sigma) dissolved in 10 mM Tris (pH 8.0), 1 mM EDTA was used as
competitor for nonspecific binding studies.
For each photolyase preparation the fraction of photolyase molecules
active in DNA binding was determined by titrating a fixed concentration
of enzyme (2-3 × KD) with increasing amounts of substrate, as described previously (9). Electrophoretic mobility
shift assays (EMSAs) were used to separate bound and free substrate.
Radioactivity present in the bound and free DNA bands was quantitated
using an Ambis Radioanalytic Imager (Ambis Systems). The equilibrium
association constants (KA) of the various
photolyases for the dimer-containing substrate were determined by
titrating substrate (5 × 10
9 M) with
enzyme (9). Control reactions without enzyme were used to calculate the
amount of background smearing of the DNA in each gel. The slope of a
Eadie-Scatchard plot (ES/St × Ef
versus ES/St) of these data yielded
KA (where ES = bound DNA concentration;
St = total substrate concentration; and Ef = free enzyme concentration). The binding affinity for nonspecific DNA
(KNS) was measured by titrating the enzyme-dimer
oligonucleotide complex (70% of substrate bound in the absence of
competitor) with cold oligo(dT)18 over the nucleotide
concentration range of 1 × 10
4 M to
5 × 10
3 M. The x intercept
of a plot of 1/ES versus concentration of oligo(dT)18 yielded
(1/KNS) (18).
KS, the intrinsic specific association constant of
photolyase for the dimer, was determined from the relationship
Kobs = KS(1 + [D]KNS) where D= molar
concentration of nondimer nucleotides in the 43-bp substrate (9). Each
KA, KNS, and active molecule
determination was performed at least 3 times.
Ethylnitrosourea Interference--
End-labeled double-stranded
dimer-containing substrate was ethylated at phosphate groups (19) as
follows. 100 µl of ethanol saturated with ethylnitrosourea (Sigma)
were added to 4 pmol of labeled substrate dissolved in 100 µl of
sodium cacodylate (pH 8.0). Following incubation at 50 °C for 1 h, ethylnitrosourea was removed by seven ethanol precipitations in 0.5 M CH3CO2NH4 with 25 µg of carrier RNA. The DNA was washed with 95% ethanol after the
final precipitation, dissolved in 0.1 mM EDTA, and
recovered substrate was quantitated by scintillation counting.
Electrophoretic mobility shift assays, in a volume of 100 µl, were
performed using the conditions described above. 5 × 10
9 M substrate was incubated with sufficient
photolyase to produce 60% binding as judged by EMSA. Following EMSA,
the bound and free DNAs were cut out of the gel, and the DNA was eluted
by overnight agitation in 0.5 M
CH3CO2NH4, 1.0% SDS, filtered
through glass wool, extracted with phenol and ether, and precipitated
in ethanol. Following a second ethanol precipitation and ethanol wash,
the samples were lyophilized to dryness and then suspended in 15 µl of 10 mM NaPO4 (pH 7.0), 1 mM EDTA.
The DNA was cleaved at ethylated phosphates by addition of 2.5 µl of
1 M NaOH followed by incubation at 90 °C for 30 min.
Fifteen µl of urea dye (6 M urea, 0.25% bromphenol blue)
were added, and aliquots were loaded onto a 12% denaturing acrylamide
gel alongside sequence ladders (A + G) (20). Band intensity was
quantitated using a Molecular Dynamics PhosphorImaging System. To
account for differences in loading, counts were normalized using as
reference a band outside of the photolyase-binding site. The
ethylnitrosourea interference pattern for each photolyase was
determined twice.
In Vitro Repair Assay--
Labeled 43-bp dimer-containing
substrate was incubated in a volume of 300 µl with a sufficient
concentration of wild-type or substituted photolyase to produce greater
than 70% binding as measured by EMSA. Following a 20-min incubation in
the absence of photoreactivating light, the binding reactions were
placed in individual compartments in a 12-well microtiter plate and
irradiated with 365 nm light (General Electric BLB) at a fluence rate
of 2 J/m2/s (UV Products UV Radiometer with a 365-nm
detector). Following each 25 J dose, a 25-µl aliquot was removed and
loaded onto an EMSA gel. The amount of repair (increase in free
substrate) following each dose was quantitated taking into account the
amount of free substrate present prior to photoreactivation. The
quantum yield for each photolyase was calculated as described
previously (5, 9). Because the light source was not monochromatic, we
have expressed the quantum yield for each enzyme relative to the
quantum yield for the wild-type enzyme, rather than as an absolute
value. A minimum of two experiments were performed for each
photolyase.
 |
RESULTS |
Protein Modeling--
By using the crystal structure of E. coli photolyase (14) as a starting point, we modeled the structure
of yeast Phr1 photolyase over the region encompassing residues 383 through 532 (E. coli photolyase residues 273-422; Figs. 1
and 3). The validity of this approach is
supported both by the high degree of sequence conservation in these
regions of the enzymes (50% identity with no gaps; Fig. 1 and see Ref.
15), which suggests a similar fold, and by the results obtained during
the modeling. Of the 73 nonidentical residues replaced during modeling,
only 6 residues (Phr1 residues 395, 401, 417, 418, 478, and 521)
produced steric clashes when modeled using the torsion angles of the
equivalent E. coli residue. Each of these residues could,
however, be modeled using alternative standard torsion angles. Most of
these residues are solvent-exposed and none lie near the proposed DNA
binding surface or the flavin-binding site; therefore it is unlikely
that an incorrect choice of torsion angle for these residues would
affect either the overall accuracy of the model or the structure of
either the DNA-binding site or the flavin-binding site. Overall the
structures of the enzymes over the modeled region are highly similar to
one another (Fig. 3), as well as to the recently solved
Aspergillus nidulans photolyase structure (21). The latter
enzyme exhibits 70% sequence identity to the E. coli
photolyase within the modeled region (21).

View larger version (107K):
[in this window]
[in a new window]
|
Fig. 3.
Structures of the DNA binding domains of
E. coli and yeast photolyase. Shown on the
left is a space-filling model of E. coli
photolyase (EPL) derived from the crystal structure. The
view looks into the cavity that leads from the surface to
FADH , which is shown in yellow. The folate
chromophore, which is mostly hidden behind the protein, is shown in
green. The region of EPL that was used to model the
structure of the yeast photolyase (YPL) DNA binding domain
is shown in cyan. On the right, the EPL pattern
and the modeled YPL structure are enlarged to show the high degree of
similarity between the structures. The band of positively charged amino
acids that extend lengthwise across the faces of the structures is
shown in dark blue.
|
|
Two important results relevant to this study emerged from the modeling
of the yeast enzyme. (i) The structure of the flavin-binding site is
conserved. Within the modeled region there are 6 amino acids that
interact with FAD (14), 5 of which are identical to those found in the
E. coli enzyme (Fig. 1). The single amino acid substitution
(Trp338(E. coli)
Tyr448(S.
cerevisiae)) places the Tyr448 side chain OH within
H-bonding distance of the same FAD phosphate contacted by the ring
nitrogen of Trp338 (data not shown). Of the seven
FAD-contacting residues that lie outside of the modeled region (14), 5 are identical in both enzymes and one of the nonidentical amino acids
(Arg236(E. coli)
Gly340(S.
cerevisiae)) is predicted to make contact only through backbone substituents. The same substitution occurs in the A. nidulans photolyase structure where the FAD contact is conserved
(21). Based upon the conservation of the cofactor-binding site, we
conclude that the orientation of FADH
is likewise
retained. (ii) The crucial features of the proposed DNA binding surface
of the E. coli enzyme (14) are conserved in yeast photolyase
(Fig. 3). This conservation is seen most strongly for residues lining
the cavity leading from the surface to the flavin chromophore. The
three substitutions, Lys383(S. cerevisiae)
Asn273(E. coli), Phe494(S.
cerevisiae)
Trp384(E. coli), and
Lys330(S. cerevisiae)
Arg226(E. coli), conserve the asymmetric
distribution of polar and hydrophobic residues in the cavity. The band
of positive electrostatic potential extending out from the cavity is
also conserved and is augmented by several substitutions:
Lys383(S. cerevisiae)
Asn273(E. coli), Lys516(S.
cerevisiae)
Glu406(E. coli), and
Gln503(S. cerevisiae)
Ala393(E. coli). The structural similarity of
the proposed DNA binding surfaces of the yeast and E. coli
enzymes is consistent with the fact that the footprints made by the
enzymes on dimer-containing DNA are essentially identical (12).
Based upon the results of the modeling study, we selected
Phe494, Glu384, Lys330,
Lys383, Arg452, and Gln514 of
PHR1 as targets for alanine substitution mutagenesis. The first three residues lie within the active site cavity and are predicted to interact with a pyrimidine dimer flipped into the cavity,
but they do not appear to interact directly with FAD. This latter
observation is crucial to the interpretation of DNA binding data
because mutations that destabilize flavin binding usually lead to
unfolding of the enzyme.2 Lys383,
Arg452, and Gln514 lie outside of the cavity
and along the region of positive electrostatic potential on the
proposed DNA binding surface (14). Based upon preliminary docking
experiments (data not shown), these residues are likely to interact
with the DNA flanking the dimer.
Effects of Ala Substitutions on Substrate Binding and
Discrimination--
Wild-type and alanine-substituted photolyases
described above were purified and characterized by UV spectroscopy.
Photolyase from the previously reported mutant W387A (9) was also
purified and used for comparative purposes in all of the studies
described below. For all enzyme preparations, both the folate and
flavin chromophores were present in approximately equimolar (0.9-1.0) stoichiometry with the apoenzyme, and neither the flavin blue neutral
radical nor oxidized flavin were detected (data not shown). Thus the
overall structure of the enzyme was not perturbed in the mutants, the
flavin-binding site was intact, and the normal oxidation state of the
flavin chromophore was retained. The integrity of the flavin-binding
site and retention of the normal oxidation state of the flavin
chromophore are particularly noteworthy. The dinucleotide flip model
predicts that the dimer interacts with both the adenine and
isoalloxazine rings of flavin (14), and photolyase lacking flavin does
not bind pyrimidine dimers with measurable affinity (22). Furthermore,
although the redox state of flavin does not alter DNA binding,
oxidation of the flavin chromophore to the blue neutral radical reduces
the quantum yield of photolysis by an order of magnitude and enzyme
containing oxidized FAD is inactive in photolysis (5, 22).
The equilibrium binding affinities of the photolyases for the 43-base
pair substrate containing a single pyrimidine dimer were determined by
EMSA. Each of the substituted photolyases displayed reduced
affinity for the dimer (Table II). The
KA values for enzymes bearing single substitutions
in the active site cavity varied widely. Substitution at
Lys330, Phe494, or Glu384 produced
a 40-60% decrease in affinity, whereas substitution of
Trp387 decreased binding 16-fold. These results suggest
that individually Lys330, Phe494, and
Glu384 contribute little to the overall binding energy,
whereas Trp277 makes a major contribution. That these
residues do indeed participate in binding was evident from the
KA of the photolyase in which all three amino acids
were substituted simultaneously with alanine. The reduction in binding
affinity exhibited by the triple mutant (K330A/F494A/E384A) was
comparable to that seen with the W387A mutant. Phe494 and
Glu384 in particular are deeply recessed into the cavity
and cannot contact any residue in normal B-DNA. In addition, both
E. coli and yeast photolyase bind pyrimidine dimers in
single-stranded DNA with an affinity similar to that seen with
double-stranded DNA (6), which rules out binding to an extrahelical
base in the complementary strand as a binding determinant. Therefore
the reduced binding affinity exhibited by these mutants strongly
supports the dinucleotide flip model for photolyase binding.
View this table:
[in this window]
[in a new window]
|
Table II
Equilibrium binding constants for photolyase-DNA interaction and
relative quantum yields for the photolysis reaction
|
|
Substitutions outside of the active site cavity produced larger
decreases in affinity (Table II). The KA for
interaction between dimer-containing DNA and the K383A mutant was
decreased approximately 7-fold, whereas KA values
for R452A and Q514A decreased 21- and 29-fold, respectively. The large
decrease in binding affinity seen with these mutants, compared with the smaller decreases seen with most of the cavity mutants, argues that
much of the free energy of binding comes from interactions between
photolyase and DNA flanking the dimer rather than from direct
interaction with the dimer. This is consistent with previous observations suggesting that photolyase recognizes not only the dimer
but also DNA structural components flanking the dimer (9, 12, 13).
The ability of a DNA-binding protein to recognize its specific target
among an excess of nonspecific binding sites is determined by the ratio
of the binding constants KS/KNS (specific binding/nonspecific binding) known as the discrimination ratio. Among the photolyases bearing single substitutions in the cavity, only the W387A mutant displayed a large reduction in
discrimination ratio (Table II). Once again synergy was observed with
the triple mutant more severely compromised in substrate discrimination
than any of the single cavity mutants including the W387A mutant. Two of the single substitutions outside of the cavity produced larger decreases in discrimination ratio than did any of the substitutions (single or multiple) inside the cavity (Table II). Particularly noteworthy is the R452A substitution which produced a 1.7-fold increase
in KNS and 20-fold reduction in
KS. Among the 10 Phr1 substitution mutants now
characterized (Ref. 9 and this work), this is the only mutant that
exhibits an increase in KNS. Clearly
Arg452 is a major determinant of binding specificity.
Ethylation Interference Studies--
Interactions between
photolyase and DNA phosphates surrounding the dimer contribute to both
specific and nonspecific substrate binding (9). Ethylation of DNA
phosphates interferes with these interactions either by eliminating a
negative charge or by steric interference and is therefore a useful
method for probing interactions at the binding interface. The results
of ethylation interference studies on the mutant photolyases and
wild-type enzyme are shown in Figs. 4 and
5. In previous studies we demonstrated
that ethylation of the first phosphate 5' to the dimer and the first
through the fourth phosphates 3' to the dimer interferes with binding
by wild-type photolyase, whereas ethylation of the intradimer phosphate
has no effect (9, 12). Interference at the fourth phosphate 3' to the
dimer is generally weak, and in the experiments reported here was not
clearly discernible. In all other respects the ethylation interference
pattern shown for wild-type photolyase in Fig. 4 is identical to those
reported previously. In contrast, each of the mutants displayed changes
in the ethylation interference pattern consistent with alterations in
the binding interface.

View larger version (157K):
[in this window]
[in a new window]
|
Fig. 4.
Ethylnitrosourea (ENU)
interference pattern for the wild-type and substituted
photolyases. Substrate containing a thymine dimer and
32P-labeled at the 5' end of the dimer-containing strand
was treated with ethylnitrosourea, incubated with photolyase, separated
into free and bound fractions by EMSA, cleaved at modified phosphates,
and analyzed on a 12% denaturing polyacrylamide gel. A representative
autoradiogram is shown. A + G denotes an A + G sequencing
ladder. Photolyases are indicated by the mutant name wild type
(WT) or K330A/E384AF/F494A (T.M.). For
each photolyase, B is the bound fraction from an EMSA
reaction, and F is the free (unbound) fraction. The 1st
phosphate 5' to the dimer and the 1st, 2nd, and 3rd phosphates 3' to
the dimer are indicated by between the WT lanes. The
band used to normalize the number of counts in each lane is indicated
by the white diamond.
|
|

View larger version (31K):
[in this window]
[in a new window]
|
Fig. 5.
Quantitation of the ethylation interference
data shown in Fig. 4. For each band analyzed, the fraction
(freecpm/boundcpm) for the wild-type enzyme was
subtracted from the same fraction for the mutant enzyme, and the result
was multiplied by 100. The legend within the figure refers to positions
of the phosphates relative to the pyrimidine dimer. The dashed
lines above and below the x axis indicate
20% change, which is the minimum considered to be significant and
reproducible.
|
|
Among the active site cavity mutants, two distinct patterns of altered
ethylation interference were apparent (Fig. 5). Photolyases W387A and
K330A exhibited decreased interference at all sites, suggesting that
interactions between the enzymes and phosphates in the dimer-containing
strand have changed along the entire binding interface. Despite this
general similarity, the two photolyases displayed distinctive
differences in the interference pattern at specific phosphates
indicating that the protein-DNA interface is uniquely altered in each
mutant. Furthermore the relative magnitude of these effects is
consistent with the relative decrease in KA. Mutants
E384A, F494A, and the triple mutant (K330A/E384A/F494A) displayed
increased interference at the second or third phosphate 3' to the dimer
and, in the cases of E384A and the triple mutant, decreased
interference at one or both phosphates flanking the dimer. Thus it
appears that, in this second class of mutant, loss of interactions
within the active site cavity attenuates photolyase-DNA phosphate
interactions at sites adjacent to the dimer and enhances dependence on
interactions at the 2nd and 3rd phosphates 3' to the dimer. Overall the
changes in ethylation interference patterns seen with these mutants
indicate that residues within the active site cavity play a role in
orienting the dimer-containing strand along the entire length of the
binding interface.
Ethylation interference patterns similar to those obtained for the
active site cavity mutants were also observed when residues outside of
the cavity were altered (Fig. 5). K383A exhibited a general decrease in
interference at all sites, similar both qualitatively and
quantitatively to the pattern seen with the W387A mutant. In the
modeled structure the side chain of Lys383 packs against
the edge of the Trp387 indole ring, and therefore changing
one of these residues to Ala may affect the position of the other. The
ethylation interference pattern of the R452A mutant resembled that of
the triple mutant (K330A/E384A/F494A) suggesting that, despite its
surface location, Arg452 plays a key role in orienting the
dimer in the active site cavity. This interpretation is supported by
the results of quantum yield experiments discussed below. The Q514A
mutant displayed an unusual pattern in that interference increased at
all sites. The simplest explanation is that one or more strong
nonphosphate contacts have been lost in the mutant and that, as a
result, interactions with phosphates contribute relatively more to the
overall binding energy.
The Effects of Substitution Mutations on Quantum
Yield--
Substitutions that affect the electronic environment within
the active site, the positioning of the dimer within the active site,
or the equilibrium between the flipped and nonflipped state potentially
alter the efficiency of photolysis. Therefore we determined the quantum
yield for photolysis by each substituted enzyme and compared it to the
quantum yield of the wild-type enzyme. Each of the purified proteins
was bound to dimer-containing substrate, and the complex was
photoreactivated with 365 nm light in increments of 25 J/m2. The amount of DNA repaired by photolyase was
quantified by electrophoretic mobility shift assay. Under the
experimental conditions employed,
70% of the substrate was bound
prior to exposure to limiting photoreactivating light. Thus multiple
cycles of binding and photoreactivation by a single enzyme molecule do
not contribute significantly to the results obtained.
The quantum yield value obtained for each photolyase relative to
wild-type is shown in Table II. Among the substitutions within the
cavity, K330A and F494A had little or no effect on the quantum yield.
In contrast, E384A, the triple substitution mutant, and W387A displayed
60-80% reductions in quantum yield relative to the wild-type enzyme.
The Glu384 side chain lies in the floor of the active site
cavity (Fig. 6), and its negative charge
may assist in directing electron transfer from FADH
to
the dimer and away from solvent. The further reduction in quantum yield
seen with the triple substitution mutant may reflect both loss of this
effect and, consistent with the DNA binding defect, reorientation of
the dimer in the active site. Trp387 is predicted to make
van der Waals contact with the dimer (Ref. 14 and Fig. 6); its loss may
permit reorientation of the dimer in the binding pocket or alter the
equilibrium between the flipped and nonflipped state. The most
surprising result of the quantum yield study was the 60% decrease in
quantum yield seen with the R452A mutant. Arg452 lies
outside of the active site cavity, and thus alanine substitution was
not expected to affect the electronic environment within the active
site. Indeed, none of the other substitutions outside of the cavity
significantly changed the quantum yield (Table II). An attractive
explanation is that interaction between Arg452 and DNA
residues flanking the dimer plays a crucial role in establishing or
stabilizing the dimer in the flipped state.

View larger version (84K):
[in this window]
[in a new window]
|
Fig. 6.
The modeled active site and DNA binding
surface of S. cerevisiae photolyase with a pyrimidine dimer
docked in the active site cavity. Amino acids examined by
mutagenesis in this and a previous study (9) are shown in
blue. Met455, which is also predicted to
interact with the dimer but has not been mutated, is shown in
orange; FADH is shown in yellow;
the folate chromophore is shown in green, and the dimer is
shown in red. To show more clearly the relationship of the
dimer bases to residues in the cavity, the structure has been rotated
approximately 25° around the y axis relative to the view
shown in Fig. 3. The entire modeled region is shown.
|
|
 |
DISCUSSION |
A Model for Dimer Binding by Photolyase--
Approximately 50% of
the total energy of enzyme-substrate complex formation between
photolyase and dimer-containing DNA comes from interaction between the
enzyme and elements of the dimer (7, 12, 13). If, as proposed by Park
et al. (14), the dimer resides within the cavity leading
from the flavin chromophore to the surface of the enzyme, interactions
with residues lining the cavity should contribute significantly to the
binding energy. The results reported here provide direct experimental
confirmation of this prediction. Alanine substitution at each of four
sites (Lys330, Glu384, Phe494, and
Trp387) within the active site cavity reduces substrate
binding. Because these residues lie too deeply buried in the pocket to
interact with normal B-DNA, the only explanation for these results is
that either the enzyme or the substrate undergoes a dramatic structural alteration that places these amino acids within interacting distance with DNA. A large scale conformational change in the enzyme is highly
unlikely. With the exception of Lys330, all of the cavity
residues probed here lie within helices that are held firmly in place
by multiple interactions with neighboring structural elements, and
movement of these helices toward the surface would disrupt multiple
interactions with the flavin chromophore. These considerations,
combined with the structural and chemical complementarity of the active
site to the dimer, provide strong evidence that the cavity in
photolyase identified by Park et al. (14) is indeed the
binding site for an extrahelical pyrimidine dimer.
A model of a pyrimidine dimer docked in the active site cavity of the
yeast Phr1 photolyase is shown in Fig. 6. This model is based upon the
crystal structure of a thymine dimer (23) and the modeled yeast
photolyase structure; until the structure of a pyrimidine dimer in
double-stranded DNA is solved at atomic resolution, this model provides
a useful context for interpreting the results of this and previous
structure-function studies (9, 24). The 5'
3' placement of the
dimer is predicated upon our observation that the enzyme interacts more
extensively with DNA residues 3' to the dimer and upon the locations of
mutations that reduce DNA binding (this work and Refs. 9, 12, and 13). In this model the 5' base in the dimer is involved in
-
stacking interactions with Trp387; the methyl group of the 5' base
is sandwiched into a hydrophobic pocket between Trp387 and
Phe494; the methyl group of the 3' base is involved in
hydrophobic interactions with Phe494; N-3 of both bases are
within hydrogen bonding distance of O-
of Glu384, and
the phosphate 5' to the dimer is within hydrogen bonding distance of
the side chains of Lys330 and Lys383. Other
potential interactions involving the dimer include a hydrogen bond
between the exocyclic amine of the adenine base in FAD and O-4' of the
5' dimer, and a "hydrogen bond" (25) between the sulfur in
Met455 and the methyl group of the 3' base.
Arg452, Gln514, Arg507, and
Lys517 lie on the surface of the enzyme immediately outside
of the active site cavity and are positioned to interact with DNA
residues 3' to the dimer. Alanine substitution at each of these sites
reduces the affinity of the enzyme for dimer-containing DNA (this work and Ref. 9). This model, in conjunction with the data reported here and
in our previous work (9), provides insight into the mechanisms used by
the enzyme to discriminate between different types of bases in dimers
and between nondamaged and damaged DNA.
Substrate Recognition and Discrimination--
A number of
interactions within the active site cavity are predicted to contribute
to base discrimination. Phe494 interacts with methyl groups
on both thymidines in the dimer, and loss of these interactions should
decrease binding. Consistent with this prediction, the affinity of
E. coli photolyase for dUU dimers in DNA is approximately
3-fold lower than for TT dimers (7), and as we have shown here, alanine
substitution of Phe494 results in a similar decrease in
binding. An 8-fold decrease in binding affinity (compared with TT
dimers) has been reported for C-containing dimers (7). According to the
model, this likely reflects not only loss of interactions with
Phe494 but also loss of hydrogen bonds between FAD and O-4'
of the 5' thymine in the dimer and between Glu384 and the
protonated N-3 of both thymines. In addition steric clash between the
exocyclic amine group of C and the exocyclic amine of adenine in FAD is
likely to constrain the approach of C-containing dimers to FAD. This
may contribute to the 20-fold lower quantum yield for photolysis of CC
dimers relative to TT dimers (7). Overall the structure of the active
site cavity favors binding of TT dimers, which are the predominant
cyclobutane-type dimers formed following UV (26).
Interactions with residues within the active site cavity also
contribute to discrimination between dimer and nondimer DNA. Trp387 is the single most important "discrimination"
residue identified here and likely contributes to substrate
discrimination in two ways. (i) Because the two bases in the dimer are
covalently linked by the rigid cyclobutane ring, parallel alignment of
the 5' base and Trp387 side chain simultaneously orients
both bases thereby establishing the full repertoire of interactions
within the active site cavity. (ii) Stacking interactions between
Trp387 and the 5' base lower the energetic cost of dimer
flipping by partially compensating for base stacking interactions that
are disrupted when the dimer is moved out of the helix. It should be
noted that most of the interactions in the active site cavity can also
occur when two nondimerized pyrimidines are flipped into the helix.
This suggests that much of the binding specificity conferred by
interactions within the active site cavity arises from differences in
the energy cost of flipping two unlinked bases versus a
dimer. Several studies suggest that the presence of a dimer in a DNA
helix weakens stacking interactions with adjacent bases and hydrogen
bonding with the partners of the dimerized bases (27-29), and thus the
energetic cost of flipping a dimer out of the helix and into the active
site cavity should be lower than the cost of flipping two noncovalently
linked pyrimidines. The free energy contributed by interactions between
cavity residues and the dimer is sufficient to stabilize the dimer in
the flipped state but is not sufficient to stabilize two noncovalently
linked pyrimidine bases flipped out of the helix. This line of
reasoning suggests that for the W387A mutant and the triple mutant
(K330A/E384A/F494A) the equilibrium between the flipped and nonflipped
dimer is shifted. Presumably flipping of the dimer is accompanied by
repositioning of phosphates immediately surrounding the dimer, as has
been seen for other flipped nucleotides (30, 31). Both the triple
mutant and the W387A mutant display large decreases in ethylation
interference at these sites, consistent with loss of the phosphate
positioning that is unique to the flipped state. We emphasize that
decreased ethylation interference in these mutants is a secondary
effect of loss of contacts within the active site cavity. The decrease in quantum yield seen with these mutants may likewise reflect a
decrease in the equilibrium between the flipped and nonflipped state.
Lys383, Arg452, and Gln514 lie
outside of the active site cavity, and substitutions at each of these
sites reduces both overall binding affinity as well as the ability of
photolyase to discriminate between dimer-containing and nondamaged DNA.
According to the model these residues contact regions immediately 5' to
the dimer (Lys383) or 3' to the dimer (Arg452
and Gln514). Both molecular modeling studies and NMR
experiments suggest that in dimer-containing DNA the helix is distorted
at the nucleotide 5' to the dimer and the first and second nucleotides
3' to the dimer (27, 28, 32). Thus there is an excellent correlation between the DNA sites that promote dimer-specific recognition and the
location of photolyase residues that contribute to specific binding. As
might be expected for loss-of-contact mutants at these sites, alanine
substitution at Lys383 or Gln514 decreases both
specific and nonspecific binding with the larger effect being seen on
specific binding. In contrast, alanine substitution at Arg452
increases nonspecific binding as well as decreases specific
binding. This could reflect introduction of a new interaction that
interferes with binding or, more likely in our opinion, loss of the
Arg452 side chain which interferes with binding to nondimer
DNA. It is interesting to note that this mutant also exhibits a
profound defect in the quantum yield for repair and a substantial and
specific decrease in ethylation interference at phosphates immediately surrounding the dimer. These are characteristics shared with the Trp387 mutant and the triple mutant (K330A/E384A/F494A) and
suggest that, like these mutants, Arg452 may play an
important role in maintaining the dimer in the flipped orientation. We
note that several DNA-binding proteins that "flip" nucleotides out
of the helix do so by inserting one or more side chains into the helix,
thereby displacing the base (31, 33). Whether this is the role of
Arg452 must await the cocrystal structure.
Evolutionary Implications--
Two classes of CPD photolyases have
been identified based upon sequence homology (34). Most microbial
photolyases, including the E. coli, A. nidulans,
and S. cerevisiae enzymes, are members of class I and share
25-43% sequence homology, whereas most photolyases from higher
eukaryotes are members of class II and share 38-72% homology. Because
the homology between enzymes in different classes is only 10-17%, it
is pertinent to ask whether enzymes in these two classes employ common
mechanisms to recognize and repair pyrimidine dimers. While direct
information on the three-dimensional structure of class II enzymes is
lacking, several lines of evidence support a common mechanism. All of
the enzymes require reduced FAD to carry out photolysis (4, 34). The
M. thermoautotrophicum photolyase, the only representative
of class II enzymes that has been characterized extensively with
respect to DNA binding, contacts the same phosphates surrounding the
dimer as do the yeast and E. coli enzymes (35). Finally,
despite the low level of overall homology between the two classes of
enzymes, there is striking conservation of amino acids lining the
active site cavity, and to a lesser extent of amino acids that interact
with flanking DNA residues (Table III).
Even more surprising is the sequence conservation between residues in
the active site cavity of the class I photolyases and the (6-4)
photolyases (Table III). Evidence has been presented suggesting that
the (6-4) photolyases also flip their substrates out of the DNA helix
(36). However, the structure of the binding site must be sufficiently
different to exclude pyrimidine dimers that are not efficiently bound
by (6-4) photolyases (2, 37). Nevertheless, of the five residues within the cavity that are predicted to contact the dimer in CPD photolyases, four are conserved in the (6-4) photolyases. This is consistent with
the proposed evolution of the class I CPD and (6-4) photolyases from a
common ancestral gene (38, 39) and further suggests that the
different binding specificities of the enzymes entail surprisingly few
changes in the active site cavity. The cocrystal structures of
these two types of photolyases should provide insight into the crucial
interactions required for such discrimination.
We thank Dr. Hengming Ke for assistance with
the protein modeling; Dr. Hee-Won Park for providing the structure of
the pyrimidine dimer; Drs. Aziz Sancar and Xiaodong Zhao for dimer
substrate and useful discussions; and Kalpana Kasala for technical
assistance.