apyrimidinic endonuclease1

doi:10.1006/jmbi.2000.3653 available online at http://www.idealibrary.com on

J. Mol. Biol. (2000) 298, 447±459

Mapping the Protein-DNA Interface and the Metal-binding Site of the Major Human Apurinic/Apyrimidinic Endonuclease Lam H. Nguyen, Daniel Barsky, Jan P. Erzberger and David M. Wilson III* Molecular and Structural Biology Division, Lawrence Livermore National Laboratory P.O. Box 808, L-441 Livermore, CA 94551, USA

Apurinic/apyrimidinic (AP) endonuclease Ape1 is a key enzyme in the mammalian base excision repair pathway that corrects AP sites in the genome. Ape1 cleaves the phosphodiester bond immediately 50 to AP sites through a hydrolytic reaction involving a divalent metal co-factor. Here, site-directed mutagenesis, chemical footprinting techniques, and molecular dynamics simulations were employed to gain insights into how Ape1 interacts with its metal cation and AP DNA. It was found that Ape1 binds predominantly to the minor groove of AP DNA, and that residues R156 and Y128 contribute to protein-DNA complex stability. Furthermore, the Ape1-AP DNA footprint does not change along its reaction pathway upon active-site coordination of Mg2‡ or in the presence of DNA polymerase beta (polb), an interactive protein partner in AP site repair. The DNA region immediately 50 to the abasic residue was determined to be in close proximity to the Ape1 metal-binding site. Experimental evidence is provided that amino acid residues E96, D70, and D308 of Ape1 are involved in metal coordination. Molecular dynamics simulations, starting from the active site of the Ape1 crystal structure, suggest that D70 and E96 bind directly to the metal, while D308 coordinates the cation through the ®rst hydration shell. These studies de®ne the Ape1-AP DNA interface, determine the effect of polb on the Ape1-DNA interaction, and reveal new insights into the Ape1 active site and overall protein dynamics. # 2000 Academic Press

*Corresponding author

Keywords: AP endonuclease; Ape1; base excision repair; DNA binding; metal coordination

Introduction In order to maintain genetic integrity, organisms have developed various means to repair damaged DNA. The base excision repair (BER) pathway typically involves the removal of a single damaged nucleotide or baseless site from the DNA (reviewed by Mol et al., 1999). Apurinic/apyrimidinic (AP) sites are formed from spontaneous hydrolysis of the N-glycosyl bond, from attack of Present address: J. P. Erzberger, Department of Molecular and Cellular Biology University of California, Berkeley CA 94720, USA. Abbreviations used: BER, base excision repair; AP, apurinic/apyrimidinic; OH
, hydroxyl radical; F, tetrahydrofuran; WT, wild-type; D, distance. E-mail address of the corresponding author: [email protected] 0022-2836/00/030447±13 $35.00/0

bases by free radicals, or by the action of repair enzymes called DNA N-glycosylases which remove damaged or unconventional bases (reviewed by McCullough et al., 1999). Ape1, the major mammalian AP endonuclease, is an essential component of the BER pathway (Xanthoudakis et al., 1996). Ape1, in the presence of Mg2‡, cleaves the phosphodiester bond immediately 50 to an AP site, generating a 30 OH group and a 50 deoxyribose moiety (reviewed by Demple et al., 1994). The single residue gap can then be ®lled by DNA polymerase beta (polb) (reviewed by Wilson, 1998), and the remaining 50 -deoxyribose phosphate group excised by its phosphodiesterase activity (Matsumoto & Kim, 1995). The ®nal nick is sealed by DNA ligase I or an XRCC1-DNA ligase III complex (Prasad et al., 1996; Caldecott et al., 1994). # 2000 Academic Press

448

AP DNA Interactions and Divalent Metal Coordination of Ape1

Although Ape1 and polb do not form a stable protein-protein complex (Dimitriadis et al., 1998), polb does bind Ape1-AP DNA binary complexes to form a higher-order ternary complex, and the phosphodiesterase activity of polb is accelerated by the presence of Ape1 (Bennett et al., 1997). polb also forms a complex with XRCC1 (Caldecott et al., 1996; Kubota et al., 1996) or mammalian DNA ligase I (Prasad et al., 1996). In addition, polbb, the XRCC1 N-terminal domain, and a gapped DNA substrate can form a higher-ordered ternary complex in vitro (Marintchev et al., 1999). These data suggest coordination of various enzymatic steps in the BER pathway. Besides AP endonuclease activity, Ape1 acts as a 30 -phosphodiesterase, removing lesions resulting from oxidative damage of DNA such as 30 -phosphoglycolates (Suh et al., 1997), and has both 30 to 50 exonuclease and RNAseH activities (reviewed by Demple & Harrison, 1994; Rothwell & Hickson, 1997). It is noteworthy that these activities of Ape1 are relatively poor in comparison to those exhibited by its bacterial counterpart, exonuclease III (ExoIII). AP endonucleases are classi®ed into two families based on amino acid sequence homology to Escherichia coli ExoIII or endonuclease IV (EndoIV) (reviewed by Demple & Harrison, 1994). Ape1 is homologous to ExoIII. The three-dimensional structures of Ape1, ExoIII, EndoIV, and the EndoIV-AP DNA complex have been determined by X-ray crystallography (Mol et al., 1995; Gorman et al., 1997; Hos®eld et al., 1999). EndoIV inserts sidechains into the DNA base stack through the minor groove, compresses the DNA backbone, bends the DNA 90  , and promotes double-nucleotide ¯ipping to sequester the extrahelical AP site into the enzyme catalytic pocket (Hos®eld et al., 1999). The molecular details of the Ape1-AP DNA interactions are not fully known, yet biochemical studies have shed signi®cant light on its repair reaction. Based on kinetic and binding studies of Ape1 and its mutants, the reaction pathway, with a minimal number of complexes, is as follows: Ape1 binds speci®cally to AP DNA in the absence of Mg2‡ to form a stable intermediate complex (Wilson et al., 1997). This complex is then converted to a catalytically competent complex in the presence of Mg2‡. Catalysis subsequently occurs, resulting in cleavage of the phosphodiester bond immediately 50 to the AP site and the formation of a protein-product complex. This complex then dissociates, releasing Ape1 from nicked AP DNA (Lucas et al., 1999). Product dissociation appears to be Mg2‡ concentration-dependent (Masuda et al., 1998b). Ape1 requires at least four base-pairs 50 and three base-pairs 30 of an AP site for incision activity (Wilson et al., 1995). For the AP strand, methylation of guanine residues located one or three base-pairs 50 of the AP site, or ethylation of phosphate groups two or three positions 30 of the AP site prevented Ape1-AP DNA binary complex formation. While no phosphate ethylation interference

was detected for the complementary strand, methylation at two base-pairs 50 , or one or three base-pairs 30 of the AP site impaired Ape1 binding (Wilson et al., 1997). These data provided the ®rst information of how Ape1 engages its target substrate. Here, we re®ne our understanding of the molecular interactions of Ape1 and AP DNA, examine the effect of polb and Mg2‡ on the DNA structure of Ape1-AP DNA binary complexes, map the Ape1 metal-binding site in terms of proximity to the DNA substrate, and provide direct evidence of the amino acid residues involved in metal coordination.

Results Ape1 protects six to seven bases on either DNA strand around the AP site and the presence of polb b does not change the footprint The purity of the Ape1 proteins used in this work is shown in Figure 1(a). To determine how Ape1 interacts with AP DNA, we employed the chemical footprinting reagent hydroxyl radical (OH
). Due to their small size, OH
are useful probes for studying DNA contacts at high resolution (Dixon et al., 1991). OH
cleave the DNA directly by attacking the deoxyribose ring (Hertzberg & Dervan, 1984; Balasubramanian et al., 1998). The double-stranded DNA used in this study was a 26 bp duplex with tetrahydrofuran (F), an abasic site analog (Wilson et al., 1995), near the center (Figure 1(b)). As shown in Figure 2, Ape1 protects seven bases on the strand containing F and six bases on the complementary strand. The F residue is located in the center of the protected region on the abasic strand. The guanine base opposite the F residue is strongly protected, whereas the surrounding bases are less protected by Ape1 from OH
-mediated cleavage. There is a hypersensitive band within the footprint that migrates at the same position on the gel as the Ape1 cleavage product. A simple interpretation is that this ``hypersensitivity'' is the result of Ape1 incision due to residual enzymatic activity even in the presence of EDTA. Consistent with this interpretation, at the necessarily high levels of Ape1 used in the footprinting assays, Ape1 incised an amount of labeled AP DNA (1 %) similar to that present in the hypersensitive band. In addition, as shown below, such hypersensitivity is not observed within the footprint of the catalytically inactive Ape1 mutant D210N. A D210N mutation reduces incision activity by 25,000-fold, without affecting the speci®c DNAbinding activity of Ape1, indicating a critical role for this residue in the catalytic reaction (Erzberger & Wilson, 1999). Using the OH
footprinting approach, we examined whether the presence of polb, which has been shown to form a ternary complex with Ape1 and AP DNA (Bennett et al., 1997), causes any change

AP DNA Interactions and Divalent Metal Coordination of Ape1

Figure 1. Protein and AP DNA substrate reagents used in this work. (a) Puri®ed Ape1 mutant proteins. Ape1 proteins (1.0 mg) used in this work were fractionated on an SDS/12 % polyacrylamide gel and stained with Coomassie blue dye. Lane 1, wild-type Ape1; lane 2, D210N mutant; lane 3, D308A; lane 4, D70R, lane 5, D210N/D308A double mutant; lane 6, E96Q; lane 7, H309S; lane 8, N68A; lane 9, R156Q; lane 10, Y128A, and lane 11, D210N/D70A double mutant. The protein molecular mass standards (in kDa) are indicated on the right. (b) The duplex DNA substrate. 26F is 50 -AATTCACCGGTACCFTCTAGAATTCG-30 , 26G is the complementary strand where a G is positioned directly opposite F. F is the tetrahydrofuran residue, a synthetic abasic site analog (Wilson et al., 1995). The arrow indicates the phosphodiester linkage incised by Ape1.

in the footprint. As shown in Figure 2, we cannot detect any difference in the footprint pattern of Ape1-AP DNA complexes in the presence of polb, indicating that polb does not bind elsewhere to the DNA, and that either polb binds directly to Ape1 without changing the Ape1-DNA interactions or replaces exactly some of the Ape1-DNA contacts. Ternary complexes were observed by gel retardation assays (Bennett et al., 1997; data not shown). Ape1 mutants Y128A and R156Q have reduced AP DNA binding activity The non-speci®c nuclease DNaseI displays structural similarity to the ExoIII family of proteins (Mol et al., 1995). Comparison of the crystal structures of Ape1 and the DNaseI-DNA complex led to a proposed model for the Ape1-AP DNA binary complex (Gorman et al., 1997). In this model, Y128 and R156 residues of Ape1 are implicated in DNA contacts. To test this prediction and to better de®ne the DNA-binding interface of Ape1, we con-

449

Figure 2. Ape1 binds in the minor groove and DNA polymerase b does not cause a change in the Ape1speci®c footprint. (a) The OH
footprint of Ape1-AP DNA complex. Lanes 1 to 7 are samples with labeled 26F strand; lanes 8 to 14 are with labeled 26G strand. Lanes 1 and 14 are the no cleavage agent controls. Lanes 2 and 13 are the no protein controls with 10 mM Fe(AS)2. Lanes 3 and 12 are the no protein controls with 5 mM Fe(AS)2. Lanes 4 to 11 are with 10 mM Fe(AS)2. Lanes 4 and 11 are reactions with 160 nM Ape1; lanes 5 and 10, 160 nM Ape1 and 580 nM polb; lanes 6 and 9, 160 nM Ape1 and 1.74 mM polb; lanes 7 and 8, 1.74 mM polb. (b) Summary of the OH
footprint data. The ®lled vertical bars above each base indicate protection from cleavage by OH
in solution. The height indicates the relative strength of footprint protection as determined by Phosphorimager scans of three independent experiments.

structed Y128A and R156Q Ape1 mutants and asked if there was a loss in DNA-binding af®nity by gel retardation assays. As shown in Figure 3, both the Y128A and R156Q mutations resulted in a >100-fold reduced DNA-binding capacity (with a corresponding reduction in speci®c incision activity of fourfold and 70-fold, respectively), consistent with the involvement of these residues in AP DNA complex stability. Sites of AP DNA in proximity to the metal-binding site of Ape1 To gain additional information regarding the topography of Ape1-AP DNA binary complex, we determined which bases of the AP DNA substrate are located near the metal-binding site of Ape1 by employing an Fe2‡-cleavage assay (Mustaev et al.,

450

AP DNA Interactions and Divalent Metal Coordination of Ape1

were obtained with D210N as observed with wildtype (WT) Ape1 protein and 26G-labeled duplex AP DNA substrates (data not shown). As shown in Figure 4, the strongest ironmediated cleavage signals on the F-containing strand or complementary strand are all 50 to the F residue. The addition of Mg2‡ reduced these ironpromoted cleavages, indicating that Mg2‡ and Fe2‡ are competing for the same metal-binding site in Ape1 (Figure 4). The iron cleavage signals also decreased in the absence of the reducing agent dithiothreitol (DTT; data not shown), consistent with the DNA being cleaved by a OH
mechanism (Zaychikov et al., 1996). We conclude that the metal is located immediately upstream of the AP residue prior to catalysis, consistent with the metal ion being involved in cleavage of the phosphodiester bond immediately 50 to the AP site.

Figure 3. Ape1 mutants Y128A and R156Q have reduced repair activity. (a) DNA-binding activity of WT and mutant Ape1 proteins. Positions of protein-DNA complexes (C) and unbound AP DNA duplex substrates (S) are indicated on the left: 5 nM 26F duplex DNA probe was incubated with increasing amounts of Ape1 protein. Lane 1 is the no protein control. Lanes 2 to 5 are reactions with WT Ape 1. Lanes 6 to 10 are with R156Q Ape1 mutant. Lanes 11 to 15 are with Y128A Ape1 mutant. Lanes 2, 6, and 11, 5 nM respective Ape1 protein; lanes 3, 7, and 12, 15 nM; lanes 4, 8, and 13, 45 nM; lanes 5, 9, and 14, 135 nM; lanes 10 and 15, 405 nM. (b) Incision activity assay of WT and mutant Ape1 proteins. Lane 1 is the no protein control. Lanes 2 to 7 are reactions with WT Ape 1. Lanes 8 to 13 are with R156Q Ape1 mutant. Lanes 14 to 19 are with Y128A Ape1 mutant. Lanes 2, 8, and 14, 0.03 nM respective Ape1 protein; lanes 3, 9, and 15, 0.09 nM; lanes 4, 10, and 16, 0.27 nM; lanes 5, 11, and 17, 0.81 nM; lanes 6, 12 and 18, 2.43 nM; lanes 7, 13, and 19, 7.3 nM. Substrate, S; incised product, P. A representation of three independent experiments is shown.

1997). OH
generated by the liganded iron under aerobic conditions degrade biopolymers (such as DNA or protein) with a diffusion-limited rate within an estimated range of 1 nm. Since Fe2‡ was able to support the incision activity of Ape1 at the excess protein to DNA ratios (at least 6:1) used in our binding and footprinting assays (data not shown), we performed the metal-cleavage studies with the catalytically inactive D210N mutant (Erzberger & Wilson, 1999). This both reduced the amount of incised background product generated and allowed us to determine the DNA bases in close proximity to the metal in the ternary complex (Ape1-AP DNA-Fe2‡) prior to incision. Similar iron cleavage patterns

Figure 4. Sites of AP DNA in proximity to the metalbinding site of Ape1. (a) Iron-mediated cleavages of AP DNA. Lanes 1 to 4 are double-stranded DNA substrate with the labeled 26F strand. Lanes 5 to 7 are the labeled 26G DNA substrate. Each reaction has 5 nM AP DNA and contains 20 mM Fe(AS)2. Lanes 1 and 7 are the no protein controls. Lanes 2 and 6 are reactions with 30 nM Ape1 D210N. Lane 3 reaction is the same as that of lane 2 except 10 mM MgCl2 was added as a metal competitor. Lanes 4 and 5 are reactions done with 30 nM Ape1 D210N mutant and 150 nM DNA polb. (b) Summary of the cleavage sites on the AP DNA substrate. Relative cleavage intensity of each base due to iron is proportional to the length of the vertical bars. Hatched horizontal bars beneath the top F-containing DNA strand shows the minimum bases 50 and 30 of the F residue required for Ape1 incision (Wilson et al., 1995).

AP DNA Interactions and Divalent Metal Coordination of Ape1

451

Effects of amino acid mutations on the metal-binding function of Ape1 In the crystal structure of Ape1, a samarium ion Ê of residues E96, D70, D308, is located within 6 A H309, and N68 (Gorman et al., 1997), all of which are capable of coordinating a cationic metal. To examine which residues among these affect Ape1 metal-binding activity, we determined the ironmediated cleavage signals of corresponding mutant proteins. We reasoned that if an amino acid is directly or indirectly involved in metalbinding, then appropriate mutation of this residue would cause a decrease in the binding of Fe2‡, thereby reducing the iron-mediated cleavage signal of the complementary strand of duplex AP DNA substrates. We assayed only the complementary strand because several Ape1 mutants, excluding D210N and H309S, display signi®cant catalytic activity at the saturating protein concentrations used in these experiments. When compared to the D210N Ape1 mutant, H309S and N68A did not cause a decrease in ironpromoted cleavages (Figure 5(a)). Conversely, E96Q, D308A, and D70R, all exhibited a substantial reduction in the iron cleavage signal, with D70R and D308A showing the most severe reduction. Even though the D308A mutant has been shown to exhibit a modest twofold decrease in AP DNA af®nity (Erzberger & Wilson, 1999), increasing the D308A protein concentration by threefold does not lead to a corresponding threefold increase in iron cleavage signal. Thus, D308A is at saturating levels relative to the DNA substrate in this assay and the decrease in signal is not due to a decrease in the DNA equilibrium binding constant. E96Q, D70R and N68A mutants have an AP DNA-binding af®nity nearly identical with WT (Erzberger & Wilson, 1999; data not shown). The speci®c AP endonuclease activities of E96Q, D70R, and N68A Ape1 mutants are reduced 2200, 27, and 600-fold, respectively, relative to that of WT Ape1 (Erzberger & Wilson, 1999; data not shown). We also constructed double mutants containing D210N with D70A or D308A to allow us to assay both DNA strands for iron-promoted cleavage. We constructed the double mutant with D70A instead of D70R because alanine is a more subtle mutation. Both double mutants showed a severe decrease in iron-speci®c cleavage on both strands of F-containing DNA duplexes (Figure 5(b)), consistent with the results for the D308A and D70R single mutants (Figure 5(a)). Both double mutants were found to bind F DNA substrates as well as D210N and WT Ape1 protein (data not shown). D70R and D308A Ape1 mutants exhibit reduced activity in low Mg2‡ Previous characterization of D70R and D308A mutants revealed only a minor reduction in enzymatic activity in buffers containing 10 mM MgCl2 (Erzberger & Wilson, 1999). The results of the Fe2‡-

Figure 5. The effect of Ape1 mutations proximal to the metal-binding site. (a) The effect of mutations of amino acid residues in proximity to the metal-binding site on iron-mediated cleavage of the labeled 26G DNA strand. Each reaction has 5 nM AP DNA. Lane 1 is the no iron control. Lane 2 is the no protein control. Lane 3 is reaction with 30 nM D210N mutant; lane 4, 30 nM D308A; lane 5, 30 nM D70R; lane 6, 30 nM E96Q; lane 7, 30 nM N68A; lane 8, 30 nM H309S; and lane 9, 90 nM D308A. (b) Mutations D70A and D308A reduce iron-mediated cleavage of both the 26F strand and its complement 26G. Lanes 1 to 5 are with labeled 26F strand, and lanes 6 to 9 are with labeled 26G complementary strand. Each reaction has 5 nM AP DNA. Lane 1 is the no iron control reaction. All other lanes are reactions with 20 mM Fe(AS)2. Lanes 2 and 6 are the no protein controls. Lanes 3 and 7 are reactions with 30 nM D210N Ape1 mutant; lanes 4 and 8, 30 nM D210N/ D70A double mutants; and lanes 5 and 9, 30 nM D210N/D308A. The arrows on the left and/or right side indicate the positions of four strongest iron cleavage signals.

cleavage studies led us to assess the effect of Mg2‡ concentration on the incision activity of these mutants, as previous studies revealed that an E96A mutant is more sensitive to reduced Mg2‡ concentrations relative to WT (Barzilay et al., 1995). At high levels of Mg2‡ (5 mM), D70R and D308A Ape1 mutants cleaved AP DNAs nearly as well as WT Ape1 protein. However, at Mg2‡ concentrations of 1 mM and less, the activity of the

452

AP DNA Interactions and Divalent Metal Coordination of Ape1

mutants was sharply reduced in comparison to WT protein (Figure 6). This result further supports a role for D70 and D308 in metal coordination. Molecular dynamics simulations show flexible loop regions and suggest a potential mechanism for metal coordination To gain new insights into the overall Ape1 protein dynamics, active-site residue interactions, and metal-binding, all-atom molecular dynamics simulations were performed. Two simulations of the WT human Ape1 protein, F-sim and E-sim (which differ methodologically in that they employ either a force-shifted cutoff or a particle mesh Ewald summation to calculate the electrostatic interactions, respectively), were executed. Both 500 ps simulations were started from the crystal structure reported by Gorman et al. (1997), except that the Sm3‡ cation was replaced with Mg2‡ as described in Materials and Methods. The average deviation of backbone heavy atoms (non-hydrogen) for the entire protein molecule converged within 100 ps, Ê and yielding an overall RMSD of 1.07(0.05) A Ê 1.11(0.04) A over the last 400 ps for E-sim and Fsim, respectively. This result suggests that the protein maintains structural integrity and similar dynamics throughout both simulations. The RMSD values for all heavy atoms (backbone and side-chains) Ê (E-sim) and are slightly larger at 1.48(0.05) A Ê (F-sim), as would be expected when 1.61(0.05) A factoring in typical side-chain movements. Predictably, the highest motion occurs primarily on the protein surface (Figure 7). Most notably, three regions (one that includes residue R177, one that includes N229, and one that includes residue M270), which appear to be involved in AP site rec-

Figure 6. The AP endonuclease activity of D70R and D308A mutants is hypersensitive to low Mg2‡ concentrations: 10 nM labeled AP DNA and 0.34 nM (0.13 ng) (Hang et al., 1997) wild-type, D70R, or D308A Ape1 were incubated together at 37  C for ten minutes. Lane 1 is the no protein control (ÿ). Lanes 2 to 5 are reactions with wild-type Ape1; lanes 6 to 9, D70R Ape1 mutant; and lanes 10 to 13, D308A Ape1 mutant. Concentration of MgCl2 in mM (0.1, 0.5, 1, or 5) is indicated above each lane. Substrate, S; incised product, P.

ognition (Gorman et al., 1997; Cal et al., 1998), show the most dramatic side-chain movements. In fact, the backbone of the M270-containing region is the most ¯exible section of the entire protein backbone, showing twice the average RMS ¯uctuations. We suggest that these ¯exible loop regions may undergo conformational shifts during speci®c DNA binding. Relevant to the biochemical studies here, we examined in detail the active-site catalytic residues surrounding the metal cation. Our analysis is presented for the E-sim only, since it showed slightly smaller deviations from the crystal structure than the F-sim. The duration of binding to or molecular coordination of an ion is expressed as the probability (Pdist) that the bond exists at any given time, based on a maximum distance criterion denoted by dist. Such bonds can also be expressed by the average distance (D) between the two atoms (closest non-hydrogen atoms) throughout the simulation. A carbonyl oxygen atom of E96 binds Mg2‡ Ê with P2.4 AÊ ˆ 1.0 (Figure 8), at D ˆ 1.83(0.05) A while the other carbonyl oxygen atom of E96 is Ê with hydrogen bonded to Y171 (D ˆ 3.06(0.61) A 2‡ P3.2 AÊ ˆ 0.69). The Mg is also coordinated by D70 Ê , P2.4 AÊ ˆ 1.0) and four active-site (D ˆ 1.83(0.04) A Ê, water molecules (all with D ˆ 2.0(0.1) A P2.4 AÊ ˆ 1.0). Thus, Mg2‡ is bound tightly by both the carboxylate residues of D70 and E96 and a shell of four water molecules. This binding occurred after rapid rearrangements of the water molecules and amino acid residues in the crystal structure, yet is so tight that it cannot be said to have sampled all possible arrangements. D308 does not bind to the ion directly, but rather the ®rst hydration shell of Mg2‡ (relative to Mg2‡, Ê , P5 AÊ ˆ 0.92) as shown in Figure 8. D ˆ 4.3(0.4) A This binding is considerably less tight than direct ion coordination, and one noteworthy difference of the F-sim is the ability of D308 to ``¯ip-out'' into Ê , P5 AÊ ˆ 0.6 for the Fthe solvent (D ˆ 5.8(1.7) A sim). While N68 more consistently coordinates the Ê, ®rst solvation shell of Mg2‡ (D ˆ 4.2(0.2) A P5 AÊ ˆ 1.0) than D308, this is likely a result of N68 simply being more rigidly positioned within the active-site, since the biochemical studies do not support a major role for this residue in metal coordination. D210 rarely reaches the ®rst solÊ , P5 AÊ ˆ 0.0), vation shell of Mg2‡ (D ˆ 6.6(0.2) A consistent with the biochemical data indicating no prominent role for D210 in metal coordination. No dramatic DNA conformational change was detected in the Ape1-DNA binary complex upon the addition of Mg2‡ To analyze whether there is any change in DNA substrate conformation upon metal coordination, but prior to phosphate hydrolysis, we determined the OH
footprint of the catalytically inactive Ape1 mutant, D210N, with and without a divalent metal ion. We reasoned that Ape1 forms the ®rst Ape1AP DNA complex in the presence of EDTA, and

AP DNA Interactions and Divalent Metal Coordination of Ape1

453

Figure 7. The solvent-accessible surface of the non-hydrogen atoms of Ape1 (stereo image), depicting atom positions averaged over a 500 ps molecular dynamics simulation (E-sim). The color ordering, blue, green, yellow and rust, indicates the degree of motional ¯uctuations (RMSF) from largest to smallest. The magnesium ion within the active site is colored red. The three ¯exible loop regions harboring residue R177 or N229 (to the right) or encompassing residues Y269, M270, M271 and N272 (to the left) are shown. These Figures were created within VMD (Humphrey et al., 1996), using SURF (Varshney et al., 1994) to calculate the surfaces.

that D210N would form a catalytically competent complex in terms of metal coordination (Figure 5). As shown in Figure 9, the Ape1 D210N footprint does not change upon the addition of Mg2‡. Furthermore, the D210N footprint in the presence of Mg2‡ is indistinguishable from the footprint of WT Ape1 (Figure 2; with the exception of the hypersensitivity, which likely represents residual cleavage activity seen with the WT protein). We conclude that there is no major DNA structural change when the Ape1-AP DNA complex is converted to a Mg2‡-coordinating ternary complex.

Discussion Ape1 interactions with AP DNA It has been shown that the abasic deoxyribose sugar ring or an extrahelical base are not essential components in AP site recognition by Ape1 (Wilson et al., 1995; Erzberger et al., 1998; Erzberger & Wilson, 1999). The molecular dynamics simulations performed here reveal that Ape1 possesses three ¯exible loop domains (Figure 7) that are important for speci®c binding to AP site-containing

Figure 8. In the same orientation as Figure 7 is shown a detailed view of the Ape1 active site, prior to the simulation (left) or captured after 25 ps of dynamics (right). The Figure to the left indicates the positions of Ape1 residues within the crystal structure without hydrogen atoms (Gorman et al., 1997). Four water oxygen atoms (navy blue) and the carbonyl oxygen atoms of D70 and E96 coordinate the magnesium ion (green sphere) uninterruptedly throughout Ê (left) and 2.4 A Ê (right) are shown as broken black the simulation (right). All electrostatic interactions of less than 3 A lines.

454

AP DNA Interactions and Divalent Metal Coordination of Ape1

Figure 9. No major DNA conformational change is detected upon the coordination of Mg2‡. Lanes 1 to 5 are with labeled 26F strand; lanes 6 to 10 are with labeled 26G strand. Lanes 1 and 10 are the no cleavage agent controls. Each reaction contains 4 nM AP DNA duplex. Lanes 2 and 9 are the no protein controls with 10 mM Fe(AS)2. Lanes 3 and 8 are the no protein controls with 5 mM Fe(AS)2. Lanes 4 to 7 are with 10 mM Fe(AS)2. Lanes 4 and 7 are reactions with 160 nM D210N Ape1 with 5 mM EDTA; lanes 5 and 6, 160 nM Ape1 and 5 mM MgCl2 instead of 5 mM EDTA. Complexes of AP DNA-Ape1-Mg2‡ were formed prior to the 30 second treatment with the footprint reagents. The vertical bars on the left and right side of the Figure indicate the DNA region protected from cleavage by OH
generated by the Fe(EDTA) complex.

DNA substrates (Gorman et al., 1997; Cal et al., 1998). While the structure of AP DNA has been shown to deviate only slightly from unmodi®ed DNA, NMR and computational studies have found kinking, melting, and structural ¯uctuations around the abasic site, indicating a unique ¯exibility local to the AP lesion (Coppel et al., 1997; Ayadi et al., 1999; Barsky et al., 2000). Similar increased ¯exibility is also observed with gapped DNA (Roll et al., 1998), an oligo substrate that is bound by Ape1 (Masuda et al., 1998a). However, nicked

DNA, which retains an unkinked B-DNA conformation with only slight distortions at the lesion site, is not bound by Ape1. Thus, Ape1 may be probing, using its unique adjustable loop domains, for speci®c DNA structures and/or increased DNA backbone ¯exibility. Since AP DNA is thermodynamically less stable than normal B-DNA (Gelfand et al., 1998), Ape1 may induce speci®c DNA distortions that permit binding to AP DNA. In fact, previous footprinting studies revealed that Ape1 promotes hypersensitivity to the cleavage agent Cu-1,10-phenanthroline at the abasic residue (Wilson et al., 1997). However, the hypersensitive band in the footprinting studies presented here (Figure 2) appears to be the result of Ape1 incision, even in the presence of EDTA. Furthermore, when using potassium permanganate (KMnO4) as a probe (Sasse-Dwight, 1991), we did not observe any Ape1-enhanced reactivity of thymine in AP DNA substrates, even when thymine was placed in the complementary strand immediately across from the AP site (L.N. and D.M.W.III, unpublished observations); KMnO4 strongly oxidizes unpaired or distorted pyrimidines in DNA, reacting much more strongly with thymine than cytosine residues (Sasse-Dwight, 1991). These data suggest that Ape1 does not cause a large melting of duplex DNA upon binding, yet does not exclude the possibility that a more subtle DNA conformational change occurs. The ability of Ape1 to promote cleavage in EDTA at acyclic AP site analogs, which display increased backbone ¯exibility, and not cyclic AP structures, suggests that Ape1 alone (i.e. in the absence of Mg2‡) can induce structural rearrangements in DNA necessary for hydrolysis (Erzberger & Wilson, 1999). The groove in which Ape1 is located can be identi®ed by analysis of the DNA protection pattern (Dixon et al., 1991). If the protected region on each strand is offset by two to three base-pairs in the 30 direction, the protein-DNA interaction takes place in the minor groove. In contrast, if binding occurs in the major groove, then the protected regions on each strand are offset in the 50 direction. The results of the footprinting experiments presented here are consistent with Ape1 binding predominantly in the minor groove of AP DNA (Figure 10). The other major AP endonuclease, E. coli EndoIV, also binds to the minor groove of AP DNA (Hos®eld et al., 1999), suggesting that this mode of complex formation may be a common mechanism used by such repair endonucleases. Thus, damage-speci®c proteins may recognize unique structural conformations or deformities in the minor groove of damaged DNA, in a manner similar to the way in which architectural proteins recognize unique minor groove structural distortions of sequence-speci®c DNA (reviewed by Bewley et al., 1998). Although Ape1 binds primarily in the minor groove, the protein also contacts AP DNA through major groove interactions as determined by previous methylation interference studies (Wilson

AP DNA Interactions and Divalent Metal Coordination of Ape1

455

Figure 10. The Ape1-DNA interface. Hydroxyl radical footprinting studies revealed that Ape1 protects an area of the AP DNA (left) of about seven bases on the abasic strand (blue) and six bases on the complementary strand (orange), with the F residue (gold) located in the center of the protected region. The arrow indicates the site of Ape1 incision. It is noteworthy that previous ethylation interference studies (Wilson et al., 1997) indicate phosphate contacts two or three positions 30 of the abasic site on the F-containing strand. In addition, methylation interference experiments revealed base contacts both in the minor and major grooves (not shown). Site-directed mutagenesis has revealed a role for speci®c amino acid residues of Ape1 in AP DNA complex stability (right), unveiling the DNAbinding groove of Ape1 (see the text for discussion). The ¯exible loop domains (harboring R177, N229 and M270) are indicated for orientation. The magnesium ion is indicated by a red sphere.

et al., 1997). Perhaps major groove interactions are most critical during the initial stages of AP DNA complex formation (i.e. during recognition, prior to the formation of the speci®c binary complex), as signi®cant major groove contacts were not observed with the pre-formed Ape1-AP DNA complexes here. Alternatively, certain methylations may have prevented Ape1-DNA complex formation indirectly by either modifying neighboring base-protein contacts or by causing a more general conformational change in the DNA substrate that interferes with Ape1 association (Yang & Carey, 1995). Ape1-polb b coordination No obvious difference in the Ape1 protein footprint was detected upon the addition of polb, a protein that forms a ternary complex with Ape1 and AP DNA (Bennett et al., 1997). Thus, polb does not appear to alter Ape1-speci®c interactions with AP DNA or to interact with DNA directly upon binding the Ape1-AP DNA binary complex, although polb interactions with DNA ends (where OH
footprinting resolution is poor) cannot be excluded. We propose that transient association of polb with this complex is mediated through protein-protein interactions that are dependent on conformational changes in Ape1 that occur upon

binding AP DNA, since polb-Ape1 binary complexes have not been observed in the absence of DNA (Bennett et al., 1997; Dimitriadis et al., 1998). Whether speci®c substrate conformational changes occur during the repair reaction of Ape1 that promote subsequent polb DNA binding and its 2-deoxyribose-5-phosphate lyase activity (Bennett et al., 1997; Xu et al., 1998) will need to be determined. The Ape1 repair steps Upon location of the AP site, the ®rst stable Ape1-AP DNA binary complex is formed (Wilson et al., 1997). Based on the crystal structure of the DNaseI-DNA complex (Lahm & Suck, 1991; Weston et al., 1992), structurally equivalent residues in Ape1, Y128 and R156, were predicted to interact with DNA and mediate binary complex assembly (Gorman et al., 1997). When these two residues were mutated, there was a decrease in both protein-DNA complex stability and AP endonuclease activity, consistent with the prediction that these residues are involved in AP DNA-binding. N212, F266, W267, H309 and Y171 have also been shown to affect Ape1-AP DNA complex stability (Rothwell & Hickson, 1996; Erzberger et al., 1998; Erzberger & Wilson, 1999; Masuda et al., 1998a). The combination of the footprinting and

456

AP DNA Interactions and Divalent Metal Coordination of Ape1

site-directed mutagenesis studies provides an emerging picture of the Ape1-AP DNA interface (summarized in Figure 10). The ®rst speci®c binary complex is then converted to the catalytically competent complex in the presence of Mg2‡ (Lucas et al., 1999). Using the high-resolution OH
footprinting technique, no major rearrangements of the DNA structure were detected upon the coordination of Mg2‡, suggesting that there are no large conformational changes in DNA upon metal-binding. The lack of major conformational changes in the DNA upon metal coordination is not unprecedented. Structural and biochemical studies of the restriction endonuclease EcoRI indicate that there is no major conformational change induced in DNA upon the addition of Mg2‡ to the protein-DNA complex (reviewed by Jen-Jacobson, 1997). However, there are structural rearrangements required for catalysis in the immediate vicinity of the scissile phosphodiester bond (reviewed by Jen-Jacobson, 1997), as seems to be the case for Ape1 (Erzberger & Wilson, 1999). Protein conformational changes are also likely necessary after Mg2‡ binding for steric complementarity and ef®cient catalysis to occur. Residue D308 is likely to be necessary for such tertiary structural changes in Ape1, since with the D308A mutant, a conformational shift involving metalbinding became an experimentally observable, rate-limiting step in the kinetic pathway (Lucas et al., 1999). The observation that D308 is capable of rotating out of the active-site (in F-sim only) may provide evidence that this residue is involved in recruiting or releasing the metal cation to or from (or correctly orienting the metal within) the catalytic pocket of Ape1. A Fe2‡ cleavage technique (Mustaev et al., 1997) was employed to map the sugars of the AP DNA substrate proximal to the metal-binding site of Ape1 and to identify residues of Ape1 that are involved in metal coordination. The sugars on both DNA strands located immediately upstream of the AP site were found to be cleaved, with the strongest signal at the sugar immediately 50 to the AP residue. These data are consistent with the metal being involved in the hydrolysis of the phosphodiester bond 50 to the lesion. We found that H309S or N68A mutations in Ape1 did not cause a decrease in iron cleavage signals, indicating no major or a less signi®cant involvement for these residues in metal coordination. However, mutating D70, E96, and D308 caused a substantial decrease in the intensity of the iron cleavage signals. Furthermore, as observed with the E96A mutant (Barzilay et al., 1995), D70R and D308A exhibited hypersensitivity to low Mg2‡ concentrations in comparison to WT Ape1. These results are consistent with other studies that have suggested a role for E96 and D308 in metal-binding (Barzilay et al., 1995; Masuda et al., 1998b; Lucas et al., 1999). Our results are the ®rst to implicate D70 in metal coordination, and may suggest that Ape1 main-

tains a unique metal-binding site (which may play a role in substrate speci®city) as this aspartate residue is not conserved in ExoIII or DNaseI (see the discussion by Erzberger & Wilson, 1999). Molecular dynamics simulations provided a visualization of WT Ape1 active-site dynamics and suggest that E96 and D70 bind directly to the metal ion, while D308 interacts with the metal ion indirectly through the ®rst hydration shell. From the results presented here, it seems likely that these three residues (and not N68, D210, or H309), in cooperation with three active-site water molecules (Figure 8), form the primary pre-incision metalbinding site for orienting the target scissile phosphate. Additional roles for these amino acids or for the metal ion in nucleophile generation or leaving group stabilization, and the number of metals within the active-site, cannot be discerned from our studies here. It is noteworthy that the mutagenic pattern of D210, where a histidine substitution has a less dramatic negative effect than an alanine mutation on Ape1 catalytic activity (Erzberger & Wilson, 1999), is consistent with a role for this residue in stabilizing the negatively charged, post-incision leaving group, either through proton donation or metal chelation. Further studies are clearly needed, and the exact role of the metal cation in the dissociation step (Masuda et al., 1998b) will need to be determined. The data presented here extend our current working knowledge of how Ape1 engages DNA, communicates with polb during BER, and coordinates its catalytically critical divalent metal co-factor. During the review of this paper, a report describing three crystal structures of Ape1 and DNA was published (Mol et al., 2000). This study provides direct evidence that the ¯exible loop regions described here are central to AP site-speci®c binding and that Ape1 may be a structure-speci®c nuclease that detects and productively binds DNA that adopts a kinked conformation and can present a ¯ipped-out AP site to a selective recognition pocket.

Material and Methods Buffers and reagents Reagents were purchased from Sigma unless otherwise indicated. Restriction enzymes were purchased from New England Biolabs. Radio-labeled nucleotides were from Amersham. Spectrophotometric-grade glycerol was obtained from Fisher. All pH values were determined at 21  C. Oligos were purchased from Operon Biotechnologies. Oligos were further puri®ed by denaturing 20 % polyacrylamide gel electrophoresis prior to use (Sambrook et al., 1989). Plasmid constructions Site-directed mutagenesis was performed as described (Erzberger & Wilson, 1999). To generate a D210N/ D308A double mutant Ape1 expression construct, plasmids containing an APE1 gene with either D210N or

AP DNA Interactions and Divalent Metal Coordination of Ape1 D308A mutation (Erzberger & Wilson, 1999) were digested with PstI. Appropriate restriction fragments were then puri®ed by agarose gel electrophoresis and ligated. To construct an APE1 D210N/D70A gene, plasmids containing a D210N or D70A mutation (Erzberger & Wilson, 1999) were digested with AlwNI, and appropriate DNA fragments were gel-puri®ed and ligated. Plasmid constructs were sequenced as described by Wilson et al. (1998). Purification of recombinant proteins All Ape1 proteins were overexpressed in bacteria and puri®ed as described (Erzberger & Wilson, 1999), except a gel-®ltration chromatography step was added for the D210N double mutants (see above). Brie¯y, following ion-exchange chromatographies (Erzberger & Wilson, 1999), Ape1 protein fractions were pooled, concentrated by 80 % (w/v) ammonium sulfate, and then fractionated on a BioRad BioSil SEC 125-5 gel ®ltration column (7.8 mm  300 mm) in 50 mM NaHepes (pH 7.5), 5 % glycerol (w/v), 0.1 mM EDTA, and 0.1 mM DTT. The ¯ow rate was 0.25 ml/minute. Size markers used for calibration are thyroglobulin (670 kDa), gamma globulin (158 kDa), ovalbumin (44 kDa), myoglobin (17 kDa), and vitamin B-12 (1.35 kDa). Proteins were detected by ultraviolet absorbance at 280 nm. Ape1 proteins were dialyzed overnight against 50 mM Tris-HCl (pH 7.9), 50 mM KCl, 20 % glycerol, 1 mM PMSF and 0.1 mM DTT, and stored at ÿ70  C. Proteins were more than 95 % pure as determined by Coomassie blue staining of SDS/polyacrylamide gels (Figure 1(a)). All mutants had similar chromatographic pro®les, suggesting structural integrity has been maintained. For polb puri®cation, protein extracts from BL21(lDE3) bacteria harboring pT7Polb (a generous gift from Dr Stuart Linn, University of California, Berkeley) were generated from IPTG-induced 2 l culture as described above for Ape1, except that the resuspension buffer was 50 mM Tris (pH 8.0), 0.1 mM EDTA, 1 mM DTT, 5 % glycerol (buffer PC) containing 400 mM KCl. In addition, PMSF was added to a ®nal concentration of 1 mM prior to sonication. After centrifugation, clari®ed extracts were applied to a Q20 column pre-equilibrated with buffer PC containing 400 mM KCl. Flowthrough material was collected, diluted with an equal volume of buffer PC, and applied to an S10 column pre-equilibrated with buffer PC containing 200 mM KCl. Protein was eluted with a linear gradient of KCl from 200 mM to 800 mM. Polb protein was detected by SDS-PAGE and Coomassie blue staining (>95 % pure). Incision and binding assays Nuclease and gel retardation assays were done as described by Erzberger & Wilson (1999). Hydroxyl radical footprinting protection assay Either oligo 26F or 26G (Figure 1(b)) was labeled at the 50 -end and annealed to a molar equivalent of unlabelled complementary strand (Erzberger & Wilson, 1999). Double-stranded DNA was then puri®ed through a G25 desalting spin column (Pharmacia) according to the instruction of the manufacturer, and stored at ÿ70  C. Binary Ape1-DNA complexes were formed by incubating on ice, 0.1 pmol of the labeled DNA fragment with at least 3 pmol of various Ape1 proteins in 50 ml of

457

50 mM KHepes (pH 7.5), 1 mM DTT, 100 mg/ml BSA, 50 mM KCl, and 0.01 % (v/v) Triton X-100 for 20 minutes with either 5 mM EDTA or 5 mM MgCl2. The reactions were then ``footprinted'' by adding 4 ml of 50 mM sodium ascorbate, 4 ml of 3.2 % H2O2, and 4 ml of 125 mM Fe(EDTA)ÿ2 at 4  C for 30 seconds (Dixon et al., 1991). The reactions were stopped with 100 ml of 60 mM thiourea. The samples were extracted with an equal volume of phenol/chloroform/isoanyl alcohol (25:24:1, by vol.), and precipitated with 0.1 vol. of 10 M LiCl, 10 mg of glycogen and 300 ml of 100 % (v/v) ethanol. The samples were resuspended in 7 ml of 50 % formamide, heated at 80  C for ®ve minutes, and analyzed on a 20 % (w/v) polyacrylamide, 7 M urea denaturing gel. Visualization of the labeled substrate was achieved using a Molecular Dynamics (Sunnyvale, CA) STORM 860 Phosphorimager and quantitative analysis was performed using Molecular Dynamics ImageQuant v1.11 software. The locations of protected bases were mapped relative to various chemical sequencing ladders which were generated as described by Kassavetis et al. (1989). Permanganate reactivity was done as described by Sasse-Dwight (1991) except the reactions were performed on ice instead of at 37  C. Metal site proximity cleavage assay Binary Ape1-DNA complex was formed as described above except without EDTA in a 20 ml volume. The cleavage reactions were initiated by adding 20 mM ferrous ammonium sulfate (Fe(AS)2) and the reactions were continued for ten or 20 minutes at 37  C. The reactions were extracted with phenol, precipitated with ethanol, fractionated on a 7 M urea/20 % (w/v) denaturing polyacrylamide gel, and analyzed as above. Computer simulations Molecular dynamics simulations of Ape1 were performed beginning with the protein structure derived from crystallography (Gorman et al., 1997; RCSB PDB #1BIX). Ape1 was simulated under constant pressure and temperature (NPT) by the strong coupling method of Nose and Hoover, as implemented in CHARMM26 (Brooks et al., 1983; Feller et al., 1995) version 26b2 (Department of Chemistry, Harvard University). The primary (cubic) cell of this periodic system measures about Ê on a side and contains 21,571 atoms, including 60 A 5734 water molecules, only 208 of which were present in the original crystal structure. The image cells contribute Ê another 22,000 atoms to the simulation. Finite, 12 A force-shifted cutoffs (F-sim) or Particle-Mesh Ewald summation (E-sim) were used to handle the electrostatic interactions. We have employed each of these electrostatic approximations in two otherwise identical simulations, denoted F-sim and E-sim. The simulations required about six weeks for each 500 ps simulation on a single DEC Alpha workstation. Protonation states of the titratable residues have been chosen to be consistent with pH 7.5, where the protein is most active (Kane & Linn, 1981). To more closely model in vivo conditions, the samarium ion present in the active-site of the Ape1 crystal was replaced with magnesium, the preferred metal cofactor of the human enzyme (Barzilay et al., 1995). The three other samarium ions from the crystal structure were discarded to reduce protein self-interaction. The charged residues and Mg2‡ gave rise to a net charge of ‡3 for which three chloride

458

AP DNA Interactions and Divalent Metal Coordination of Ape1

Ê away from ions were added to the solvent, initially 15 A the protein, to keep the system neutral (especially important in studies employing Ewald summations). Ê was judged too far for a A sulfur-sulfur distance of 3.5 A disul®de bond (C93-C208).

Acknowledgments This work was carried out under the auspices of the US Department of Energy by Lawrence Livermore National Laboratory under contract number W-7405ENG-48 and supported by an NIH grant (CA79056) to DMWIII. We thank Ms Tina Xi and Dr Harvey Mohrenweiser for sequencing support, and Dr Ian McConnell for critical inputs.

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Edited by J. M. Miller (Received 21 December 1999; received in revised form 15 February 2000; accepted 24 February 2000)