Determinants in nuclease specificity of ape1 and ape2, human homologues of Escherichia coli exonuclease III1

doi:10.1006/jmbi.2001.5382 available online at http://www.idealibrary.com on

J. Mol. Biol. (2002) 316, 853±866

Determinants in Nuclease Specificity of Ape1 and Ape2, Human Homologues of Escherichia coli Exonuclease III Masood Z. Hadi{, Krzysztof Ginalski{, Lam H. Nguyen{ and David M. Wilson III* Molecular and Structural Biology Division, Lawrence Livermore National Laboratory P.O. Box 808, L-441, Livermore, CA 94551, USA

Abasic sites and non-conventional 30 -ends, e.g. 30 -oxidized fragments (including 30 -phosphate groups) and 30 -mismatched nucleotides, arise at signi®cant frequency in the genome due to spontaneous decay, oxidation or replication errors. To avert the potentially mutagenic or cytotoxic effects of these chromosome modi®cations/intermediates, organisms are equipped with apurinic/apyrimidinic (AP) endonucleases and 30 -nucleases that initiate repair. Ape1, which shares homology with Escherichia coli exonuclease III (ExoIII), is the major abasic endonuclease in mammals and an important, yet selective, contributor to 30 -end processing. Mammals also possess a second protein (Ape2) with sequence homology to ExoIII, but this protein exhibits comparatively weak AP site-speci®c and 30 -nuclease activities. Prompted by homology modeling studies, we found that substitutions in the hydrophobic pocket of Ape1 (comprised of F266, W280 and L282) reduce abasic incision potency about fourfold to 450,000-fold, while introduction of an ExoIII-like pocket into Ape2 enhances its AP endonuclease function. We demonstrate that mutations at F266 and W280 of Ape1 increase 30 to 50 DNA exonuclease activity. These results, coupled with prior comparative sequence analysis, indicate that this active-site hydrophobic pocket in¯uences the substrate speci®city of a diverse set of sequence-related proteins possessing the conserved four-layered a/b-fold. Lastly, we report that wild-type Ape1 excises 30 -mismatched nucleotides at a rate up to 374-fold higher than correctly base-paired nucleotides, depending greatly on the structure and sequence of the DNA substrate, suggesting a novel, selective role for the human protein in 30 -mismatch repair. # 2002 Elsevier Science Ltd.

*Corresponding author

Keywords: Ape1; Ape2; exonuclease III; abasic endonuclease; mismatch repair

Introduction {These authors contributed equally to this work. Present address: K. Ginalski, Department of Biophysics, Institute of Experimental Physics, Warsaw University, Zwirki i Wigury 93, 02-089 Warsaw, Poland and BioInfoBank, ul. Limanowskiego 24A, 60-744 Poznan, Poland. Abbreviations used: AP, apurinic/apyrimidinic; BER, basic excision repair; EndoIV, endonuclease IV; ExoIII, exonuclease III; L-OddC, dioxolane cytidine; MMS, methylmethane sulfonate; ORF, open reading frame; WT, wild-type. E-mail address of the corresponding author: [email protected] 0022-2836/02/030853±14 $35.00/0

The apurinic/apyrimidinic (AP) site is one of the most frequently formed lesions in DNA, arising as a product of spontaneous hydrolysis of the N-glycosylic bond, mutagen-induced base release, or damaged-base excision by a DNA repair glycosylase.1 Such lesions present both cytotoxic and mutagenic threats to the cell if left unrepaired. To cope with the roughly 10,000 abasic damages formed per genome per day, organisms maintain corrective enzymes called AP endonucleases. These proteins initiate the repair of abasic sites by ®rst locating the lesion in the genome and then incising the phosphodiester backbone immediately 50 to the AP site. Enzymes of the base excision repair (BER) # 2002 Elsevier Science Ltd.

854 pathway subsequently act to remove the abasic fragment, ®ll the nucleotide gap, and seal the nick.2 ± 6 AP endonucleases have been classi®ed into one of two major families after the Escherichia coli proteins exonuclease III (ExoIII or Xth) and endonuclease IV (EndoIV or Nfo). The ExoIII protein of E. coli was originally identi®ed as a 30 -phosphatase capable of activating DNA substrates for polymerase-mediated primer extension.7 Subsequently, ExoIII was shown to exhibit comparable activities as an AP endonuclease, 30 to 50 -exonuclease, and RNase H.3,8 While the biological importance for the latter two of these functions remains unclear, it is evident that the 30 phosphodiesterase and AP endonuclease activities of ExoIII play a major role in maintaining genome integrity and protecting against the mutagenic and cytotoxic effects of various environmental insults, most notably alkylating and oxidizing agents. In humans, the major abasic endonuclease is Ape1, an ExoIII homologue.1 Unlike ExoIII, however, the human protein displays a relatively poor 30 to 50 -exonuclease activity (while exhibiting a robust AP endonuclease activity9,10), as well as a highly selective 30 -phosphodiesterase activity. In particular, Ape1 excises the 30 -oxidative product, 30 -phosphoglycolate, somewhat ef®ciently from gaps and nicks in DNA (yet still with a 100-fold less ef®ciency than when incising at AP sites), but demonstrates little or no activity for these damages when at 30 overhangs or blunt double-strand break ends.11 Recently, Ape1 was found to be the major 30 -nuclease that excises the nucleoside analog dioxolane cytidine (L-OddC), a compound used as a cytotoxic replication chain terminator in anti-cancer and anti-viral treatments.12 Still, it is unknown at what level human Ape1 functions globally in the excision of inappropriate 30 -nucleotides. Moreover, it remains unclear which protein elements of the ExoIII family affect AP endonuclease and/or 30 -nuclease ef®ciency. The recent discovery of yeast Eth1/Apn2 and human Ape2, proteins that maintain signi®cant amino acid sequence similarity to the core nuclease domains of Ape1 and ExoIII, indicates the existence of a new subgroup within the ExoIII family.13 ± 16 This subgrouping was prompted largely by the fact that Ape1 and Ape2 contain distinguishing amino and carboxyl-terminal ends. Yet, despite the highly conserved nature of the nuclease domain, the Ape2-like proteins exhibit relatively poor AP endonuclease and 30 -repair activities.15 ± 17 It therefore seems reasonable that speci®c differences within the active-site pockets of the ExoIII-like proteins give rise to the varying levels of nuclease capacity. Notably, Ape2 maintains all of the critical catalytic residues of Ape1 (i.e. N68, E96, Y171, D210, N212, D283, D308 and H309), which are also conserved in several proteins recently predicted to maintain the four-layered a/b fold of ExoIII and the ``non-speci®c'' nuclease DNaseI.18 This set of

Determinants in Nuclease Speci®city of Ape1 and Ape2

presumably structurally related proteins includes the nuclease of the long interspersed nuclear element-1 (LINE-1 or L1) retrotransposon,19 sphingomyelinases,20 many signaling proteins such as inositol polyphosphate 5-phosphatases,21 yeast carbon catabolite repressor protein, cytolethal distending toxin B, and many proteins with unknown functions. The absolute conservation of the catalytic residues among these proteins and, in many cases, their experimentally demonstrated importance (21,22 and references within), suggests a common, conserved mechanism for cleaving phosphoester bonds, but of vastly different substrates (including phospholipids, nucleic acid and polypeptides). We sought here to ascertain the protein elements that in¯uence the nuclease ef®ciencies of Ape1 and Ape2, and thus indirectly, the substrate speci®city of the structurally related proteins, as well as to explore more thoroughly the role of Ape1 as a 30 -nuclease.

Results The ``hydrophobic'' pocket contributes to the substrate selectivity of the ExoIII family members To identify elements that in¯uence the nuclease capacities of the ExoIII-like proteins, we ®rst generated a multiple sequence alignment of 34 Ape1 (ExoIII)-like and four Ape2-like proteins from different organisms (Figure 1(a)), and then focused on E. coli ExoIII, and the human Ape1 and Ape2 proteins (Figure 1(b)). On the basis of these alignments, and using the known ExoIII and Ape1 crystal structures as templates, a 3D model of the human Ape2 protein was built (see Materials and Methods). These homology modeling studies revealed that Ape2, which has been shown to exhibit a poor AP endonuclease activity,15 lacks the hydrophobic pocket of Ape1 found to pack tightly around the extrahelical abasic sugar moiety in the co-complex crystal structure (see active-site images in Figure 2). This pocket in the Ape1-like proteins is composed of three conserved residues, i.e. F/W/Y, W/M/L and L/I, with an absolute preference for an aromatic amino acid (typically hydrophobic) in the ®rst position, and hydrophobic residues at the other two positions. However, in the Ape2-like subfamily, these positions are occupied by C/V, S/T and L/I, where both the aromatic structure is absent from the ®rst position, and the hydrophobicity of the second position is not maintained. To determine experimentally the role of this pocket in in¯uencing AP endonuclease function, we generated a series of site-speci®c mutants in Ape1 and assessed the capacity of these protein derivatives to recognize and incise abasic-DNA (Figure 3). Notably, an Ape1 protein engineered to possess an Ape2-like active site (i.e. F266C/W280S, L282) displayed a 450,000-fold reduced endonuclease activity; single mutations at either F266 or

Determinants in Nuclease Speci®city of Ape1 and Ape2

855

Figure 1. Amino acid sequence alignment of the ExoIII family. (a) Multiple sequence alignment of Ape1-like (top) and Ape2-like (bottom) proteins from different organisms. Residues conserved in more than half of all sequences are highlighted in black (identical) or grey (similar). Positions of critical residues are marked with an asterisk (*). (b) Alignment of the AP endonuclease domains of Ape1, Ape2 and ExoIII. Identical and similar residues are highlighted as above. Residues of Ape1 that are critical for AP site binding or catalysis are indicated by an asterisk (*). Within this domain, Ape2 displays 29 % and 27 % sequence identity with Ape1 and ExoIII, respectively. Locations of secondary structure elements in Ape1 and ExoIII (as assessed by DSSP60) are marked above and beneath the respective sequences: b-strands are represented as arrows, and a-helices as rectangles. The stretches shown in both (a) and (b) are presented in three dimensions in Figure 2.

856

Determinants in Nuclease Speci®city of Ape1 and Ape2

Figure 2. Active-site comparisons of Ape1, Ape2 and ExoIII. (a) Comparison of the Ape1, ExoIII and Ape2 active sites. Side-chains of Ape1 (green) important for AP site binding and catalysis, and their structural counterparts in ExoIII (cyan) and Ape2 (white), are shown as sticks. The extrahelical AP site of the Ape1-DNA complex24 is shown in orange. (b) Comparison of the Ape1, ExoIII and Ape2 AP site binding pockets. The side-chains of F266, W280 and L282 in Ape1, as well as their structural counterparts in ExoIII and Ape2, are shown as sticks. Solvent-accessible surfaces are indicated in grey for these amino acid residues and in yellow for the abasic sugar moiety. The contact surface areas for the dominant, hydrophobic contacts of Ape1 between Phe266, Trp280, and Leu282, and the extrahelical Ê 2, 28 A Ê 2, and 14 A Ê 2, respectively, as calculated with LPC software. DNA (orange) in the ExoIII deoxyribose are 46 A and Ape2 complex structures was modeled based on the Ape1-DNA crystal complex (see Material and Methods).

W280 resulted in less dramatic reductions (Table 1). A double mutant (F266A/W280I) designed to generate a ``null'' active site in Ape1 similarly exhibited no activity (Figure 3). These data, combined with the observation that an ExoIII W212S mutant (in essence a null mutant) is unable to bind APDNA,23 suggests that at least one active-site aromatic residue is required for ef®cient recognition of AP-DNA by the ExoIII family, perhaps explaining the reduced activity of Ape2. To test this hypothesis, we engineered both an ExoIII-like and Ape1-like hydrophobic pocket into Ape2 (Figure 2(b)). When Ape2 was constructed to possess the hydrophobic pocket of ExoIII (i.e. C260W/S274L), this protein derivative indeed exhibited enhanced AP site repair capacity, as

measured by increased rescue (or increased cell growth along a gradient) of an xth ÿ nfo ÿ mutant bacteria challenged with methylmethane sulfonate (MMS), an alkylating agent8 that produces high levels of cytotoxic AP sites (Figure 4(a)). Yet, when Ape2 was modi®ed to contain the hydrophobic pocket of Ape1 (i.e. C260F/S274W), little if any increased complementation was observed. This apparent discrepancy can likely be explained by computational packing studies (see Materials and Methods). This analysis indicated that only the ExoIII-like pocket (C260W/S274L), and not the Ape1-like pocket (C260F/S274W), would be accommodated in the Ape2 protein, due to local structural constraints around position 274. Consistent with the in vivo complementation studies, pro-

Determinants in Nuclease Speci®city of Ape1 and Ape2 Table 1. Speci®c incision activity of the Ape1 hydrophobic binding site mutants

Ape1 protein WT Ape1 F266Ca W280Sb F266C/W280Sc

Specific activity (pmol minÿ1 mgÿ1)

Fold reduction (relative to WT Ape1)

679,443  67,127 175,413  2134 387  50 1.5  0.3

4 1756 452,962

Puri®ed protein was incubated with 32P-labeled 18 bp duplex DNA containing a single AP site, and the rate of conversion of substrate to incised product was measured as described in Material and Methods. The F266C, W280S and F266C/W280S Ape1 mutants were engineered to either partially or fully recreate the Ape2 active site (see Figure 2(b)). a Previous studies revealed a similar reduction in Ape1 activity with an F266A mutant.25 b Additional mutations at W280 to either I or L (ExoIII-like) resulted in a 700-fold reduction in incision activity (M.H., unpublished results). c Ape1 double mutants F266A/W280A, F266A/W280L, F266A/W280I, F266A/W280S and F266C/W280I displayed a greater than 3750-fold reduction in incision activity (M.H., unpublished results).

tein extracts prepared from double mutant bacteria harboring the Ape2(C260W/S274L) expression construct exhibited substantially enhanced AP endonuclease activity, as compared to cell extracts expressing the vector, an Ape2 single mutant or the Ape2(C260F/S274W) double mutant or (Figure 4(b)). Coomassie blue staining and Western blot analysis of the extracts shown in Figure 4(b) revealed no difference in Ape2 (wild-type or mutant) protein expression (data not shown).

857 These data indicate that the ``hydrophobic'' pocket of the ExoIII family is critical in imparting AP site speci®city and that the speci®c arrangement of residues within this pocket dictates nuclease capacity. Mutations at F266 or W280 in Ape1 enhance its 30 exonuclease activity Due to its tight packing around the extrahelical abasic sugar moiety, the Ape1 hydrophobic pocket (comprised of F266, W280 and L282) was proposed to exclude normal nucleotide bases.24 We reasoned that mutating F266 or W280 to a smaller hydrophobic residue (more similar in nature to the activesite pocket of E. coli ExoIII shown in Figure 2) would permit increased binding of normal DNA nucleotides and result in an altered nuclease speci®city. To further explore the role of this pocket in determining substrate selectivity, we measured the nuclease activity of an F266A Ape1 mutant.25 These studies revealed that highly puri®ed F266A Ape1 protein (Figure 5(a)) exhibits enhanced degradation on an array of DNA substrates (including singlestranded DNA, and duplex DNAs containing either a nick, single nucleotide gap, blunt end or 30 ¯ap), compared to identically puri®ed wild-type (WT) and W267A Ape1 proteins (Figure 5(b)). To con®rm that the enhanced 30 -exonuclease activity was indeed intrinsic to the F266A protein, we constructed a D210N/F266A Ape1 double mutant; residue D210 has been shown to be essential for enzymatic activity, while not adversely affecting DNA-binding activity.26 As shown in Figure 5(c), while the F266A mutant degrades single-stranded

Figure 3. Ape1 mutants engineered to mimic the Ape2 binding pocket have reduced endonuclease activity. (a) SDS-PAGE analysis of puri®ed recombinant Ape1 proteins. Proteins (5 mg as indicated) were separated on a 10 % (w/v) polyacrylamide gel and stained with Coomassie brilliant blue R250. M. Wt., molecular mass standards (in kDa). (b) Incision analysis of Ape1 proteins: 0.014 nM protein (as indicated) was incubated with 100 nM AP-DNA substrate at 37  C for ten minutes (see Materials and Methods). The positions of intact substrate (S) and incised product (P) are indicated. Similar results (representative study shown) were seen in ®ve independent incision studies, using protein from at least three independent puri®cations.

858

Determinants in Nuclease Speci®city of Ape1 and Ape2

Figure 4. Introduction of the ExoIII hydrophobic pocket into Ape2 results in enhanced AP endonuclease function. (a) MMS sensitivity gradient plate test. xth nfo mutant bacteria, alone (i.e. without the pKEN vector) or transformed with the indicated pKEN Ape2 expression construct, were plated along an MMS gradient (the width of the triangle depicts the relative concentration of MMS) and examined for resistance (measured as cell growth). The panel to the left shows unchallenged cells, and the panel to the right shows the gradient challenge. The data shown are representative of four independent experiments. pKEN-Ape1 is the positive control.14 (b) Analysis of AP endonuclease activity. Cell extracts were prepared from E. coli (xth nfo mutant) transformed with an Ape1 or Ape2 protein expression construct (as indicated) and assayed for AP endonuclease activity (see Materials and Methods). Total protein (1 mg) was incubated with 100 nM 50 end-labeled duplex 41F (tagacggatgaataatgagggFagaagttggatttggtagt) DNA for ®ve minutes at 37  C. The data represent mean and standard deviation of conversion of initial substrate to sitespeci®cally incised product. We note that, while AP site incision was increased only mildly in the Ape2 (C260F/ S274W) and wild-type Ape2 extracts, elevated DNA degradation (presumably exonuclease activity) was observed (not shown).

26G DNA, the D210N/F266A double mutant does not. In subsequent experiments, we found that an F266C Ape1 mutant exhibited signi®cantly enhanced 30 to 50 -exonuclease activity (Figure 5(d)). To explore the role of W280 in dictating nuclease speci®city, we measured the activities of W280I, W280L and W280S mutant proteins. Similar to the F266A/C proteins, the W280 Ape1 mutants exhibited increased 30 -exonuclease activity relative to WT on nick DNA substrates, however, this ``enhanced'' activity was far less than that observed with the F266 mutants (Figure 5(d)). The studies reported here support the notion that the hydrophobic pocket of Ape1 in¯uences substrate speci®city. Ape1 functions as a 30 -mismatch nuclease Given that Ape1 can excise nucleotides from DNA in a 30 to 50 manner,9,10 including nucleoside analogs (e.g. L-OddC;12), we explored the ability of the human protein to remove 30 -mismatched nucleotides from two different nick substrates that simulate scenarios of single-nucleotide replacement base excision repair (Figure 6). With substrate 1 (which mimics a situation of ``uracil'' replacement;27), only mismatched T (opposite G) was removed with a greater ef®ciency (about fourfold) than the correctly paired nucleotide C

(Table 2). However, with nick substrate 2 (which mimics a situation of ``8-oxoguanine'' replacement,28 each mismatched nucleotide was excised at a signi®cantly greater rate than the normally base-paired nucleotide G, with mismatched C (opposite C) being removed at a 374-fold higher rate. Notably, in these experiments, it was found that Ape1 removes the 30 -non-mismatched base from nick substrate 1 at a 25-fold greater rate than from nick substrate 2, although both were in a G:C basepair (Figure 6(a)). One possible explanation for this variation is the differences in sequence context around the nick. To test this hypothesis, we synthesized a new DNA substrate, similar to nick substrate 2, but with an A ‡ T-rich sequence (50 -TA-30 or 50 -AT-30 , instead of 50 -GG-30 ) immediately upstream of the 30 -terminal nucleotide (Figure 6(b)). This ``A ‡ T richness'' resulted in an approximately sixfold higher excision rate of correctly paired 30 -terminal nucleotides, suggesting that local DNA sequence does affect WT Ape1 excision activity, as seen above. The ef®ciency at which WT Ape1 removes 30 mismatched nucleotides within a partial duplex DNA context was examined. As summarized in Table 3, WT Ape1 is able to preferentially remove 30 -mismatched nucleotides from a partial duplex substrate, but with less ef®ciency (35 to 93-fold)

859

Determinants in Nuclease Speci®city of Ape1 and Ape2

Figure 5. F266 and W280 Ape1 mutants have enhanced 30 exonuclease activity. (a) Puri®ed recombinant Ape1 proteins. Ape1 proteins (1.5 mg) were fractionated on an SDS/12 % (w/v) polyacrylamide gel and stained with Coomassie brilliant blue R250. Lane 1, protein molecular mass standards (in kDa); lane 2, WT Ape1 protein; lane 3, F266A protein; lane 4, D210N/F266A; and lane 5, W267A. (b) DNA substrate speci®city of F266A Ape1 mutant. Labeled DNA substrate (5 nM as indicated; see Materials and Methods for details) was incubated with 5 nM WT, F266A (F) or W267A (W) Ape1 protein, and then analyzed on a denaturing polyacrylamide gel. Lanes 1, 5, 9, 13, and 17 are the no-protein controls. Lane 18 represents a reaction performed with WT Ape1 protein at 15 nM. Note, using nick DNA substrates labeled at the 30 end of the upstream primer with [a-P32]ddATP (as opposed to above, where the label was at the 50 end), only full-length substrate and ddAMP-labeled mononucleotide products, but no intermediate-sized DNAs, were observed (data not shown). This degradation pattern is consistent with the F266A Ape1 mutant exhibiting an enhanced 30 to 50 exonuclease activity, and not a non-speci®c endonuclease activity. (c) Enhanced 30 -exonuclease activity is intrinsic to the F266A Ape1 mutant. DNA substrate (5 nM as indicated) was incubated with 5 nM WT, F266A (F) or D210N/F266A (DF) Ape1 protein at 37  C for 30 minutes and analyzed as above. Lanes 1, 5, and 9 are the no-protein controls. (d) W280 and F266C Ape1 mutants exhibit enhanced 30 -exonuclease activity. Nick substrate 2 (0.01 nM; see Figure 6) was incubated with 0.02 (lanes 2, 4, 6, 8, 10 and 12) or 0.04 nM (lanes 3, 5, 7, 9, 11 and 13) WT, W280I, W280L, W280S, F266A or F266C Ape1 protein (as indicated) for 30 minutes at 37  C. Lane 1 is the no-protein control. For (b), (c) and (d), (*) represents undigested labeled DNA substrate. All DNA substrates are described in Material and Methods.

than from a nick structure (compared to rates in Table 2). Notably, we were unable to detect any 30 exonuclease degradation (<1 % of substrate digested) of fully complementary or 30 -mismatched blunt-ended DNA substrates (data not shown). Such data are consistent with the experimental observations that Ape1 requires at least three nucleotides of duplex structure 30 of the target site for abasic endonuclease activity,10 and that the Ape1 footprint extends three bases downstream of the AP site on both strands.29 We conclude that the sequence (and thus stability) and duplex nature of DNA surrounding the 30 -terminal nucleotide in¯uences the ef®ciency with which WT Ape1 can degrade DNA.

Discussion We have show here that the ``hydrophobic'' pocket is a critical determinant in the AP site speci®city of the ExoIII-like family and that the weak AP endonuclease activity intrinsic to the Ape2-like proteins15,16 likely stems from ``disruption'' of this pocket. The inability of the Ape2(C260W/S274L) mutant to fully complement the AP endonucleasede®cient bacteria (Figure 4, compare to the Ape1expressing cells) suggests that other elements may affect nuclease capacity. Speci®cally, the three loop regions that impart DNA-binding speci®city to the ExoIII family (which are absent from DNaseI and the L1 nuclease, proteins that lack AP endonuclease activity1,30) differ in amino acid sequence context between Ape1 and Ape2. Furthermore, position N174 of Ape1, the only critical active-site residue besides F266 and W280 that differs

Table 2. Speci®c activities of WT Ape1 at 30 -mismatched DNA substrates Nick substrate DNA sequence

Xa Specific activityb Fold change relative to control a b

1 CGAGCTCGAATTCACTGG

2 TACCGAGCTCGAATTCACTGG TAGAGGATCCCCGCTAGCGGX ATCTCCTAGGGGCGATCGCCCATGGCTCGAGCTTAAGTGACC

TAGAGGATCCCCGCTAGCGGGTAX ATCTCCTAGGGGCGATCGCCCATGGCTCGAGCTTAAGTGACC C (control) 25  6.8

G mismatch 9.6  4.5

A mismatch 17.1  8.2

T mismatch 100.4  16

G (control) <0.9  0.6

A mismatch 109.6  18.8

T mismatch 57.7  7.0

C mismatch 336.4  63.2

1

0.4

0.7

4

1

121

64

374

X Denotes the position where one of the four nucleotides was incorporated in the oligonucleotide to generate the mismatch substrate. The speci®c activity (pmol minÿ1 mgÿ1) was determined from three independent experiments.

861

Determinants in Nuclease Speci®city of Ape1 and Ape2

Figure 6. WT Ape1 removes 30 mismatched nucleotides in nick structures. (a) Time-course experiments using two different 30 -mismatch nick DNA substrates. The top panel is for nick substrate 1 (sequence shown above the panel). The bottom panel is for nick substrate 2. The underlined X indicates the position of the four possible nucleotides. The nucleotide examined is shown above each timecourse run. DNA substrate (5 nM) was incubated with either 0.017 nM (nick substrate 1) or 0.01 nM (nick substrate 2) WT Ape1 protein at 37  C for various times (in minutes) as indicated. (ÿ) Denotes the no-protein control. See Table 1 for determined speci®c activities. (b). The A ‡ T content 50 to the mismatched nucleotide in¯uences WT Ape1 30 -exonuclease activity. DNA substrate (5 nM) was incubated with 0.6 nM WT Ape1 at 37  C for the times (in minutes) indicated above each panel. Shown above the Figure are the three DNA contexts examined surrounding the nick. The speci®c activity (pmol minÿ1 mgÿ1) for the nick DNA substrate displayed on the left is 10.71  1.75; for the substrate displayed in the middle, 1.7  0.9; for the substrate displayed on the right, 25  6.8. (*) The undigested labeled DNA substrate, (*) the digested DNA products. The absence of (*) means that no degraded product was visible.

between the Ape1-like and Ape2-like subfamilies (Figure 1(a)), is not retained. Notably, these positions, as well as the loop domains, are not conserved among the other structurally related proteins. We therefore conclude that the residue content at positions equivalent to N174, F266 and W280 of Ape1, combined with the length and conformation of the key loop regions, are vital elements in imparting substrate selectivity to the core four-layered a/b fold, which maintains a conserved catalytic mechanism. Given the wide range of substrates recognized by this diverse set of proteins,18 and the generally poor in vitro nuclease activities of Ape2 and the weak repair-related

defects of the yeast single mutants,13,17 research aimed at examining the substrate preferences of Ape2 more globally seems warranted. Our ®nding that mutations in the hydrophobic pocket of Ape1 (e.g. F266A/C or W280I/L/S), and not at the proximal residue W267, enhance 30 -exonuclease activity supports the notion that the hydrophobic pocket in¯uences AP site and nucleotide binding. Interestingly, hydrophilic substitutions at residue I614 within the Taq DNA polymerase I hydrophobic dNTP-binding pocket were similarly found to permit binding of larger substrates, such as a bulky ¯uorescent nucleotide,31,32 presumably by widening the recog-

Table 3. Speci®c activities of WT Ape1 at partial double-stranded DNAs with a 30 -mismatch DNA sequence Xa Specific activityb Fold change relative to control a b

TAGAGGATCCCCGCTAGCGGX ATCTCCTAGGGGCGATCGCCCATGGCTCGAGCTTAAGTGACC G (control) 0.04 1

A mismatch 3.1  0.2 78

T mismatch 1.6  0.4 41

C mismatch 3.6  0.5 91

X Denotes the position where one of the four nucleotides was incorporated in the oligonucleotide to generate the substrate. The speci®c activity (pmol minÿ1 mgÿ1) was determined from three independent experiments.

862 nition pocket. Thus, by analogy, mutations at F266 or W280 in Ape1 may permit accommodation of normal nucleotides by relieving the active-site steric hindrance that normally exists. This notion is consistent with the observation that E. coli ExoIII, which possesses a hydrophobic pocket composed of only one aromatic residue, exhibits a relatively powerful 30 -exonuclease activity.8,9,33 By extension, the greater 30 -nuclease activity reported for yeast Eth1/Apn2 (in comparison to its poor AP endonuclease activity17) may be explained by expansion of this pocket. We have shown here that, depending on the sequence and structural context surrounding the 30 -terminal nucleotide, Ape1 exhibits preferential excision of 30 -mismatched nucleotides. The fact that Ape1 removes mismatched nucleotides 35 to 96-fold faster from a nick than from a partial duplex, and 30 -phosphoglycolate residues from a 30 -recessed partial duplex twofold faster than a nick,11 suggests that the chemistry of the target group also in¯uences the 30 -nuclease ef®ciency of Ape1. Morales & Kool34 argued that the rate of 30 end proofreading by the Klenow fragment of DNA polymerase I is governed by the extent of fraying (known to be in¯uenced by the base-pairing and the neighboring sequence content) at the DNA terminal pair. As the A ‡ T content surrounding the substrate 30 -terminal nucleotide in¯uences the ef®ciency with which Ape1 executes excision, it seems reasonable that this rate is affected similarly by DNA fraying. Several other proteins, including Trex1 and p53, have been found to preferentially excise mismatched nucleotides.35 ± 38 However, which protein operates ``when and where'' is unknown. Given the known communication between Ape1 and Polb,39 this ®nding may suggest a novel mechanism in which these proteins act selectively in concert during base excision repair to reduce the in®delity of this DNA polymerase, which lacks detectable 30 -exonuclease proofreading activity.40 Additional biochemical analysis directly comparing the substrate preferences of these 30 to 50 exonucleases and in vivo studies measuring the mutational events of appropriate mutant cell lines will be needed to elucidate the speci®c contributions of these proteins to mismatch nucleotide correction and genome stability.

Materials and Methods Identification of structural determinants of the ExoIII-like AP endonuclease fold To produce a multiple sequence alignment consistent with the general architecture of the ExoIII-like AP endonuclease fold, structural determinants of this fold were ®rst analyzed. Structurally conserved regions, where alignment is meaningful, were identi®ed from the superposition of all available ExoIII (PDB code 1AKO)41 and Ape1 (PDB codes: 1BIX, 1DE8, 1DE9, and 1DEW)41,42 molecular structures, using the Homology module of Insight II (MSI Inc., San Diego, CA, USA). Regions were

Determinants in Nuclease Speci®city of Ape1 and Ape2 considered structurally conserved when pairwise distances between corresponding ExoIII and Ape1 residues Ê . Conservation of speci®c did not exceed, typically, 2.5 A residues and contacts responsible for maintaining tertiary structure, and critical for DNA binding and/or catalysis, were established. PSI-BLAST searches Sequences of the ExoIII family members were collected from the non-redundant protein database using PSI-BLAST searches43 with different inclusion thresholds until pro®le convergence. Complete open reading frames (ORFs) of ExoIII, Ape1 and Ape2, and the AP endonuclease domains of Ape1 and Ape2 alone, were used as the query sequences. The ®nal set of ExoIII-like sequences (Figure 1(a)) did not differ from published data.15 Multiple sequence alignment of the ExoIII-like AP endonuclease domain All AP endonuclease domains of the ExoIII-like proteins were subjected to pairwise sequence similarity checks using the Smith-Waterman algorithm44 implemented in the SSEARCH program.45 Only the sequences that exhibited less than 80 % identity to any other sequence were considered further, to avoid domination by a group of almost identical sequences in the alignment. Initially, the CLUSTAL W program46 was used to generate multiple sequence alignments. Opening and extensions gap penalty were changed systematically, and all of the obtained alignments were inspected for both variability and violation of structural integrity in the structurally conserved regions, especially in the secondary structure elements. The ®nal multiple sequence alignment was obtained by taking, in most cases, the commonest alignment for each region within the context of the structural constraints identi®ed above. Exact positions of gaps in the insertion and deletion regions were adjusted manually to satisfy the structural scaffolds of the ExoIII and Ape1 proteins. Improving alignment quality for the Ape2-like subfamily Because of a low level of sequence similarity and the considerable differences in length between the Ape2-like and Ape1-like endonuclease domains, alignment of these sequences was considered of lower con®dence. For regions within the Ape2-like subfamily that displayed a low level of stability (i.e. were highly dependent on gap penalties), alignments were derived manually, guided by PSIPRED47 secondary structure predictions. While complying with the general requirements of the ExoIII-like AP endonuclease fold, data from pairwise alignments of PSI-BLAST searches were taken into account to generate possible alignments for the Ape2-like proteins. All alignments were then tested for ®tness of Ape2 residue mapping on the structural scaffolds of ExoIII and Ape1, using the 3D evaluation procedure.48 For each alignment, two corresponding molecular models of the Ape2 sequence were built with the Homology module of InsightII using both ExoIII (PDB code 1AKO)41 and Ape1-DNA complex (PDB code 1DEW)24 structures as templates. Backbone conformations were taken from the template structures within the structurally conserved regions

863

Determinants in Nuclease Speci®city of Ape1 and Ape2 and loops of the similar length, and only non-conserved side-chains were substituted with appropriate Ape2 side-chains. Modeling of loops that contained insertion and deletion regions was skipped in this procedure. Models were then evaluated by visual inspection and ProsaII energy pro®les49 for structural consistency, in order to detect improper packing of residues. Such a 3D evaluation procedure enabled selection of a ®nal reliable sequence-to-structure alignment. Ape2 model building Based on the ®nal sequence-to-structure alignment (Figure 1), a model of the Ape2 protein was built with the MODELLER program50 using superimposed ExoIII (PDB code 1AKO)41 and Ape1 (PDB code 1DEW)24 structures as templates. After coordinates were assigned to the Ape2 sequence, side-chains were rebuilt using the SCWRL program with a backbone conformation-dependent rotamer library.51 To preserve conserved contacts, and maximize electrostatic and hydrophobic interactions, the positions of several side-chains were adjusted manually. The ®nal model was subjected to 100 steps of energy minimization with the Discover module of InsightII to remove remaining steric clashes and improve stereochemistry. Energy optimization was performed with the AMBER force®eld52 using steepest and conjugate gradient methods. The overall quality of the modeled structure was checked in detail with the WHAT_CHECK program.53 Analysis of mutations in the AP site binding pocket To identify all residues that take part in forming the recognition pocket for the hydrophobic face of the AP site, detailed analysis of the interatomic contacts between the abasic extrahelical deoxyribose and Ape1 in the cocomplex was performed with LPC software.54 Proposed mutations of this binding pocket were then inspected carefully by replacing the appropriate residues (positions 266 and 280) in the Ape1-DNA complex and (positions 260 and 274) in the modeled structure of Ape2. Importantly, residues at the surface of a protein are generally tolerant to substitution.55 To ensure that all proposed mutations of Ape1 or Ape2 would not disrupt local packing or generate steric clashes with AP DNA, detailed rotamer searches for mutated side-chains were performed using the InsightII and SCWRL programs with a backbone conformation-dependent side-chain rotamer library. The relative orientation of AP DNA in the Ape2 modeled structure was derived from the Ape1DNA complex structure.24 Plasmid constructs Ape1 expression plasmids, pETApeF266A, pETApeF266C, pETApeW280S, pETApeW280I, pETApeW280L and pETApeF266C/W280S were constructed using the PCR overlapping technique as described.26,56 To generate the double mutant Ape1 expression plasmid, pETApeD210N/F266A, pETApeF266A and pETApeD210N were digested with PstI. The appropriate restriction fragments (i.e. a PstI fragment containing the F266A mutation, and the pETApeD210N vector backbone without the PstI fragment) were then puri®ed by agarose gel electrophoresis and ligated.

Ape2 site-speci®c mutant expression constructs, pKENApe2C260F, pKENApe2C260W, pKENApe2S274L, pKENApe2S274W, pKENApe2C260W/S274L and pKENApe2C260F/S274W, were constructed using an overlapping PCR technique. The APE2 coding region was ®rst PCR ampli®ed using CGCGGATCCGGATCAGCATGTTGCGCGTGGTGAGCTGG and CCGGAATTCTCAGCTGGGCCTGCTCCA oligonucleotide primers, and subcloned into the BamHI/EcoRI sites of pGEX-3X (Pharmacia) to generate a GST N-terminal-Ape2 fusion construct, termed pGST-Ape2. The fused ORF was then ampli®ed using CCCAAGCTTATGTCCCCTATACTAG GTTATTGG and CCGGAATTCTCAGCTGGGCCTG CTCCA primers, and cloned into the HindIII/EcoRI sites of pCDNA3.1 (Invitrogen) to create pGST-APE2PCDNA3.1. All site-speci®c mutations were introduced into this plasmid. In brief, two overlapping PCR fragments were generated using primer set CCCAAGCTTATGTCCCCTATACTAGGTTATTGG and CCGGAA TTCTCAGCTGGGCCTGCTCCA and the appropriate site-speci®c mutant primer and . The products from the two PCR reactions were then mixed together (1:10 ratio of ®rst to second PCR product) and used as a template for a third PCR reaction using CCCAAGCTTATGTCCCCTATACTAGGTTATTGG and CCGGAATT CTCAGCTGGGCCTGCTCCA. This product was cloned into the HindIII/EcoRI sites of pCDNA3.1. The sitespeci®c mutant-containing ORF fragments were then ampli®ed with CCGGAATTCATGTCCCCTATACTA GGTTATTGG and CCCAAGCTTTTCAGCTGGGCCTG CTCCA, and cloned into the EcoRI/HindIII sites of pKEN2 and transformed into ExoIII/EndoIV double mutant E. coli for all complementation studies as described.15 All clones were sequenced57 to con®rm the presence of site-speci®c mutations, and no additional errors. Purification of recombinant Ape1 protein All Ape1 mutant proteins were overexpressed in bacteria and puri®ed to >95 % purity as described.58 All mutants (excluding F266C/W280S, which eluted at slightly lower salt concentrations from the S column) had similar chromatographic pro®les, suggesting that general structural integrity is maintained. Protein concentrations were determined as described58 using the theoretical molar extinction coef®cient E280 nm of 53,400 Mÿ1 cmÿ1 calculated for Ape1. Nuclease activity assays To generate abasic DNA substrates, an 18-mer58 or 26-mer29 tetrahydrofuran (F)-containing oligonucleotide (18F or 26F, respectively) was labeled at the 50 -end with [g-32P]ATP and then annealed to a molar equivalent of unlabeled complementary strand (18G, gagtcgtaggacggtgac, or 26G, cgaattctagagggtaccggtgaatt, respectively), with G positioned opposite the abasic lesion. To generate unmodi®ed double-stranded DNA, the 26G oligonucleotide29 was 50 -end labeled and annealed to its complementary oligonucleotide. The GAP, NICK, and 30 FLAP 42 nucleotide doublestranded DNA substrates have been described.59 The sequence of the various 30 -mismatched DNA substrates are shown in the Tables. Labeled double-stranded DNA was puri®ed through a G25 desalting spin column (Pharmacia) according to the instructions of the manufacturer and stored at ÿ20  C. Incision and exo-

864 nuclease assays were performed at 37  C as described,29,58 under conditions optimized for Ape1 nuclease activity and in the linear range of enzymatic function. For AP endonuclease activity, the amount (pmol) of full-length, labeled F-containing DNA converted per minute per milligram of protein to a shorter, incised product was determined. Exonuclease activity was measured as the amount (pmol) of substrate degraded by a single nucleotide per minute per microgram of protein. Protein extracts were prepared from bacterial cells after a one hour, 1 mM IPTG induction. In brief, cells were harvested and resuspended in 50 mM Hepes (pH 7.5), 50 mM KCl, 10 % (v/v) glycerol and 1mg/ml each of leupeptin, chymostatin and aprotenin. Cells were then lysed by sonication with two, ten second bursts. Supernatants were clari®ed by centrifugation, protein concentrations determined and AP endonuclease activity measured as described above.

Acknowledgments We thank Dr Krzysztof Fidelis for his input on the computational component of this study. This work was performed under the auspices of the U.S. Department of Energy by the University of California, Lawrence Livermore National Laboratory under contract no. W-7405Eng-48 and supported by NIH grant CA79056 (to D.M.W.III) and Polish State Committee for Scienti®c Research grant 6P04A03519 (to K. G.).

References 1. Wilson, D. M., III & Barsky, D. (2001). The major human abasic endonuclease Ape1: formation, consequences and repair of abasic lesions in DNA. Mutat. Res. 484, 283-307. 2. Tomkinson, A. E. & Mackey, Z. B. (1998). Structure and function of mammalian DNA ligases. Mutat. Res. 407, 1-9. 3. Wilson, D. M., III, Engleward, B. & Samson, L. (1998). Prokaryotic base excision repair. In DNA Damage and Repair (Nickoloff, J. A. & Hoekstra, M. F., eds), vol. 1, pp. 29-64, Humana Press Inc., Totowa, NJ. 4. Thompson, L. H. & West, M. G. (1999). XRCC1 keeps DNA from getting stranded. Mutat. Res. 459, 1-18. 5. Lindahl, T. (2000). Suppression of spontaneous mutagenesis in human cells by DNA base excisionrepair. Mutat. Res. 462, 129-135. 6. Memisoglu, A. & Samson, L. (2000). Contribution of base excision repair, nucleotide excision repair, and DNA recombination to alkylation resistance of the ®ssion yeast Schizosaccharomyces pombe. J. Bacteriol. 182, 2104-2112. 7. Richardson, C. C. & Kornberg, A. (1964). A DNA phosphatase-exonuclease from Escherichia coli. II. Characterization of the exonuclease activity. J. Biol. Chem. 239, 251-258. 8. Demple, B. & Harrison, L. H. (1994). Repair of oxidative damage to DNA: enzymology and biology. Annu. Rev. Biochem. 63, 915-948. 9. Seki, S., Hatsushika, M., Watanabe, S., Akiyama, K., Nagao, K. & Tsutsui, K. (1992). cDNA cloning, sequencing, expression and possible domain struc-

Determinants in Nuclease Speci®city of Ape1 and Ape2

10.

11.

12.

13.

14.

15.

16. 17.

18. 19.

20.

21.

22.

23.

ture of human APEX nuclease homologous to Escherichia coli exonuclease III. Biochim. Biophys. Acta, 1131, 287-299. Wilson, D. M., III, Takeshita, M., Grollman, A. P. & Demple, B. (1995). Incision activity of human apurinic endonuclease (Ape) at abasic site analogs in DNA. J. Biol. Chem. 270, 16002-16007. Suh, D. ,Wilson D. M., III & Povirk, L. F. (1997). 30 -Phosphodiesterase activity of human apurinic/ apyrimidinic endonuclease at DNA double-strand break ends. Nucl. Acids Res. 25, 2495-2500. Chou, K. M., Kukhanova, M. & Cheng, Y. C. (2000). A novel action of human apurinic/apyrimidinic endonuclease: excision of L-con®guration deoxyribonucleoside analogs from the 30 termini of DNA. J. Biol. Chem. 275, 31009-31015. Johnson, R. E., Torres-Ramos, C. A., Izumi, T., Mitra, S., Prakash, S. & Prakash, L. (1998). Identi®cation of APN2, the Saccharomyces cerevisiae homolog of the major human AP endonuclease HAP1, and its role in the repair of abasic sites. Genes Dev. 12, 31373143. Bennett, R. A. (1999). The Saccharomyces cerevisiae ETH1 gene, an inducible homolog of exonuclease III that provides resistance to DNA-damaging agents and limits spontaneous mutagenesis. Mol. Cell. Biol. 19, 1800-1809. Hadi, M. Z. & Wilson, D. M., III (2000). Second human protein with homology to Escherichia coli abasic endonuclease exonuclease III. Environ. Mol. Mutagen. 36, 312-324. Unk, I., Haracska, L., Johnson, R. E., Prakash, S. & Prakash, L. (2000). Apurinic endonuclease activity of yeast Apn2 protein. J. Biol. Chem. 275, 22427-22434. Unk, I., Haracska, L., Prakash, S. & Prakash, L. (2001). 30 -Phosphodiesterase and 30 ! 50 exonuclease activities of yeast Apn2 protein and requirement of these activities for repair of oxidative DNA damage. Mol. Cell. Biol. 21, 1656-1661. Dlakic, M. (2000). Functionally unrelated signalling proteins contain a fold similar to Mg2‡-dependent endonucleases. Trends Biochem. Sci. 25, 272-273. Feng, Q., Moran, J. V., Kazazian, H. H., Jr & Boeke, J. D. (1996). Human L1 retrotransposon encodes a conserved endonuclease required for retrotransposition. Cell, 87, 905-916. Matsuo, Y., Yamada, A., Tsukamoto, K., Tamura, H., Ikezawa, H., Nakamura, H. & Nishikawa, K. (1996). A distant evolutionary relationship between bacterial sphingomyelinase and mammalian DNase I. Protein Sci. 5, 2459-2467. Whisstock, J. C., Romero, S., Gurung, R., Nandurkar, H., Ooms, L. M., Bottomley, S. P. & Mitchell, C. A. (2000). The inositol polyphosphate 5-phosphatases and the apurinic/apyrimidinic base excision repair endonucleases share a common mechanism for catalysis. J. Biol. Chem. 275, 3705537061. Beernink, P. T., Segelke, B. W., Hadi, M. Z., Erzberger, J. P., Wilson, D. M., III & Rupp, B. (2001). Two divalent metal ions in the active site of a new crystal form of human apurinic/apyrimidinic endonuclease, Ape1: implications for the catalytic mechanism. J. Mol. Biol. 307, 1023-1034. Shida, T., Kaneda, K., Ogawa, T. & Sekiguchi, J. (1999). Abasic site recognition mechanism by the Escherichia coli exonuclease III. Nucl. Acids Symp. ser. 42, 195-196.

865

Determinants in Nuclease Speci®city of Ape1 and Ape2 24. Mol, C. D., Izumi, T., Mitra, S. & Tainer, J. A. (2000). DNA-bound structures and mutants reveal abasic DNA binding by APE1 and DNA repair coordination. Nature, 403, 451-456. 25. Erzberger, J. P., Barsky, D., Scharer, O. D., Colvin, M. E. & Wilson, D. M., III (1998). Elements in abasic site recognition by the major human and Escherichia coli apurinic/apyrimidinic endonucleases. Nucl. Acids Res. 26, 2771-2778. 26. Erzberger, J. P. & Wilson, D. M., III (1999). The role of Mg2‡ and speci®c amino acid residues in the catalytic reaction of the major human abasic endonuclease: new insights from EDTA-resistant incision of acyclic abasic site analogs and site-directed mutagenesis. J. Mol. Biol. 290, 447-457. 27. Singhal, R. K., Prasad, R. & Wilson, S. H. (1995). DNA polymerase beta conducts the gap-®lling step in uracil-initiated base excision repair in a bovine testis nuclear extract. J. Biol. Chem. 270, 949-957. 28. Hill, J. W., Hazra, T. K., Izumi, T. & Mitra, S. (2001). Stimulation of human 8-oxoguanine-DNA glycosylase by AP-endonuclease: potential coordination of the initial steps in base excision repair. Nucl. Acids Res. 29, 430-438. 29. Nguyen, L. H., Barsky, D., Erzberger, J. P. & Wilson, D. M., III (2000). Mapping the protein-DNA interface and the metal-binding site of the major human apurinic/apyrimidinic endonuclease. J. Mol. Biol. 298, 447-459. 30. Cal, S., Tan, K. L., McGregor, A. & Connolly, B. A. (1998). Conversion of bovine pancreatic DNase1 to a repair endonuclease with high selectivity for abasic sites. EMBO J. 17, 7128-7138. 31. Patel, P. H., Suzuki, M., Adman, E., Shinkai, A. & Loeb, L. A. (2001). Prokaryotic DNA polymerase I: evolution, structure, and ``base ¯ipping'' mechanism for nucleotide selection. J. Mol. Biol. 308, 823-837. 32. Patel, P. H., Kawate, H., Adman, E., Ashbach, M. & Loeb, L. A. (2001). A single highly mutable catalytic site amino acid is critical for DNA polymerase ®delity. J. Biol. Chem. 276, 5044-5051. 33. Seki, S., Ikeda, S., Watanabe, S., Hatsushika, M., Tsutsui, K., Akiyama, K. & Zhang, B. (1991). A mouse DNA repair enzyme (APEX nuclease) having exonuclease and apurinic/apyrimidinic endonuclease activities: puri®cation and characterization. Biochim. Biophys. Acta. 1079, 57-64. 34. Morales, J. C. & Kool, E. (2000). Importance of terminal base pair hydrogen-bonding in the 30 -end proofreading by the Klenow fragment of DNA polymerase I. Biochemistry, 39, 2626-2632. 35. Huang, P. (1998). Excision of mismatched nucleotides from DNA: a potential mechanism for enhancing DNA replication ®delity by the wild-type p53 protein. Oncogene, 17, 261-270. 36. Hoss, M., Robins, P., Naven, T. J., Pappin, D. J., Sgouros, J. & Lindahl, T. (1999). A human DNA editing enzyme homologous to the Escherichia coli DnaQ/MutD protein. EMBO J. 18, 3868-3875. 37. Bakhanashvili, M. (2001). Exonucleolytic proofreading by p53 protein. Eur. J. Biochem. 268, 2047-2054. 38. Mazur, D. J. & Perrino, F. W. (2001). Excision of 30 termini by the trex1 and trex2 30 to 50 exonucleases: characterization of the recombinant proteins. J. Biol. Chem. 276, 17022-17029. 39. Bennett, R. A., Wilson, D. M., III, Wong, D. & Demple, B. (1997). Interaction of human apurinic endonuclease and DNA polymerase beta in the base

40. 41.

42.

43.

44. 45.

46.

47. 48.

49. 50. 51.

52.

53. 54.

55.

56.

excision repair pathway. Proc. Natl Acad. Sci. USA, 94, 7166-7169. Tanabe, K., Bohn, E. W. & Wilson, S. H. (1979). Steady-state kinetics of mouse DNA polymerase beta. Biochemistry, 18, 3401-3406. Mol, C. D., Kuo, C. F., Thayer, M. M., Cunningham, R. P. & Tainer, J. A. (1995). Structure and function of the multifunctional DNA-repair enzyme exonuclease III. Nature, 374, 381-386. Gorman, M. A., Morera, S., Rothwell, D. G., de La Fortelle, E., Mol, C. D., Tainer, J. A. et al. (1997). The crystal structure of the human DNA repair endonuclease HAP1 suggests the recognition of extra-helical deoxyribose at DNA abasic sites. EMBO J. 16, 65486558. Altschul, S. F., Madden, T. L., Schaffer, A. A., Zhang, J., Zhang, Z., Miller, W. & Lipman, D. J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucl. Acids Res. 25, 3389-3402. Smith, T. F. & Waterman, M. S. (1981). Identi®cation of common molecular subsequences. J. Mol. Biol. 147, 195-197. Pearson, W. R. (1991). Searching protein sequence libraries: comparison of the sensitivity and selectivity of the Smith-Waterman and FASTA algorithms. Genomics, 11, 635-650. Thompson, J. D., Higgins, D. G. & Gibson, T. J. (1994). CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-speci®c gap penalties and weight matrix choice. Nucl. Acids Res. 22, 46734680. Jones, D. T. (1999). Protein secondary structure prediction based on position-speci®c scoring matrices. J. Mol. Biol. 292, 195-202. Venclovas, C., Ginalski, K. & Fidelis, K. (1999). Addressing the issue of sequence-to-structure alignments in comparative modeling of CASP3 target proteins. Proteins Suppl. 3, 73-80. Sippl, M. J. (1993). Recognition of errors in threedimensional structures of proteins. Proteins: Struct. Funct. Genet. 17, 355-362. Sali, A. & Blundell, T. L. (1993). Comparative protein modelling by satisfaction of spatial restraints. J. Mol. Biol. 234, 779-815. Bower, M. J., Cohen, F. E. & Dunbrack, R. L., Jr (1997). Prediction of protein side-chain rotamers from a backbone-dependent rotamer library: a new homology modeling tool. J. Mol. Biol. 267, 12681282. Weiner, S. J., Kollman, P. A., Case, D. A., Singh, U. C., Ghio, C., Alagona, G. et al. (1984). A new force ®eld for molecular mechanical simulation of nucleic acids and proteins. J. Am. Chem. Soc. 106, 765-784. Hooft, R. W., Vriend, G., Sander, C. & Abola, E. E. (1996). Errors in protein structures. Nature, 381, 272. Sobolev, V., Sorokine, A., Prilusky, J., Abola, E. E. & Edelman, M. (1999). Automated analysis of interatomic contacts in proteins. Bioinformatics, 15, 327-332. Funahashi, J., Takano, K., Yamagata, Y. & Yutani, K. (2000). Role of surface hydrophobic residues in the conformational stability of human lysozyme at three different positions. Biochemistry, 39, 14448-14456. Ausubel, F. M. (1997). Editor of Current Protocols in Molecular Biology, John Wiley and Sons, New York, NY.

866

Determinants in Nuclease Speci®city of Ape1 and Ape2

57. Wilson, D. M., III, Carney, J. P., Coleman, M. A., Adamson, A. W., Christensen, M. & Lamerdin, J. E. (1998). Hex1: a new human Rad2 nuclease family member with homology to yeast exonuclease 1. Nucl. Acids Res. 26, 3762-3768. 58. Hadi, M. Z., Coleman, M. A., Fidelis, K., Mohrenweiser, H. W. & Wilson, D. M., III (2000). Functional characterization of Ape1 variants identi®ed in the human population. Nucl. Acids Res. 28, 3871-3879.

59. Lee, B. I. & Wilson, D. M., III (1999). The RAD2 domain of human exonuclease 1 exhibits 50 to 30 exonuclease and ¯ap structure-speci®c endonuclease activities. J. Biol. Chem. 274, 37763-37769. 60. Kabsch, W. & Sander, C. (1983). Dictionary of protein secondary structure: pattern recognition of hydrogen-bonded and geometrical features. Biopolymers, 22, 2577-2637.

Edited by J. Miller (Received 18 July 2001; received in revised form 18 October 2001; accepted 4 January 2002)