©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Role of Glutamic Acid 988 of Human Poly-ADP-ribose Polymerase in Polymer Formation
EVIDENCE FOR ACTIVE SITE SIMILARITIES TO THE ADP-RIBOSYLATING TOXINS (*)

(Received for publication, August 18, 1994; and in revised form, November 28, 1994)

Gerald T. Marsischky (§) Brenda A. Wilson (¶) R. John Collier (**)

From the Department of Microbiology and Molecular Genetics, Harvard Medical School, Shipley Institute of Medicine, Boston, Massachusetts 02115

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

Sequence similarities between the enzymatic region of poly-ADP-ribose polymerase and the corresponding region of mono-ADP-ribosylating bacterial toxins suggest similarities in active site structure and catalytic mechanism. Glu of the human polymerase aligns with the catalytic glutamic acid of the toxins, and replacement of this residue with Gln, Asp, or Ala caused major reductions in synthesis of enzyme-linked poly-ADP-ribose. Replacement of any of 3 other nearby Glu residues had little effect. The Glu mutations produced similar changes in activity in the carboxyl-terminal 40-kDa catalytic fragment fused to maltose-binding protein: E988Q and E988A reduced polymer elongation >2000-fold, and E988D 20-fold. Smaller changes were seen in chain initiation. The mutations had little effect on the K of NAD, indicating a predominantly catalytic function for Glu. The results support the concept of similar active sites of the polymerase and the ADP-ribosylating toxins. Glu may function in polymer elongation similarly to the toxins' active site glutamate, as a general base to activate the attacking nucleophile (in the case of the polymerase, the 2`-OH of the terminal adenosine group of a nascent poly-ADP-ribose chain).


INTRODUCTION

Poly-ADP-ribose polymerase (PARP), (^1)a DNA-binding protein found in most eukaryotes, is activated by DNA strand breaks in vitro or in vivo to make long, branched glycosidic polymers of ADP-ribose from NAD(1, 2, 3, 4, 5) . Much of the polymer is covalently attached to PARP itself(6) , but certain other proteins such as histones have also been found to have poly-ADP-ribose attached(7) . Although PARP has been suggested to have a role in DNA repair, much remains to be learned about its physiological functions.

The polymerase consists of a single polypeptide chain (113 kDa for the human enzyme, hPARP) and contains three functional domains(8) . The amino-terminal domain binds DNA and has two zinc fingers that are required for recognizing DNA strand breaks and activating the enzyme (9, 10, 11) . The carboxyl-terminal catalytic domain poly-ADP-ribosylates the central, so-called automodification domain(8) . The automodification domain has been predicted to have a leucine zipper (12) and may be responsible for the dimerization of PARP(13) . A carboxyl-terminal 40-kDa tryptic fragment of PARP containing the catalytic domain has ADP-ribosylation activity that is unaffected by DNA and approximately equal in magnitude to that of the intact enzyme in the absence of DNA(14) .

The ADP-ribosylation reactions catalyzed by PARP bear resemblances to those catalyzed by the ADP-ribosylating bacterial toxins, such as diphtheria and cholera toxins(15, 16) . All of these enzymes catalyze transfer of the ADP-ribose moiety of NAD to a substrate protein. In the toxins, the site of attachment of ADP-ribose is typically a specific side chain on a host cell regulatory protein, whereas PARP initiates poly-ADP-ribose chains by ADP-ribosylating acidic side chains on its own automodification domain. Following the initiation step, PARP effects chain elongation by ADP-ribosylating the 2`-OH group of terminal adenosines of poly-ADP-ribose chains, and it catalyzes branch formation by ADP-ribosylating the 2`-OH group of internal ribose moieties within the poly-ADP-ribose chain. PARP also shares with toxins the ability to catalyze NAD hydrolysis, a reaction of no known physiological significance for these proteins. Like some ADP-ribosylating toxins, the polymerase requires a macromolecular cofactor for enzymic activation. Both cholera toxin (16, 17) and Pseudomonas aeruginosa exoenzyme S (18) are activated by specific host proteins, whereas PARP is activated by DNA.

The ADP-ribosylation activity of many, and perhaps all, ADP-ribosylating toxins is dependent on a key catalytic glutamic acid residue. Photoaffinity labeling with NAD first identified Glu as an active site residue in diphtheria toxin (DT) (19, 20) , and functionally equivalent glutamic acids were subsequently found in other toxins, Glu in P. aeruginosa exotoxin A(21, 22) , Glu in pertussis toxin(23, 24, 25) , and Glu in the Clostridium limosum C3-like ADP-ribosyltransferase(26) . Where tested, mutations of these residues severely affected ADP-ribosyltransferase activity and toxicity(23, 25, 27, 28, 29, 30, 31) , while causing little change in substrate binding. Thus, a largely catalytic role is indicated for the active site glutamic acids. The crystallographic structures of exotoxin A(32) , Escherichia coli heat-labile toxin(33) , DT(34) , and pertussis toxin (35) have shown strong similarities in the active site fold among the ADP-ribosylating toxins and the presence of glutamic acid in the position corresponding to Glu of DT.

Domenighini et al.(36) reported sequence similarities between the enzymatic region of PARP and the catalytic domains of several of the ADP-ribosylating toxins, in particular DT and P. aeruginosa exotoxin A. Although the percent identity was low, nearly all of the active site residues of these toxins were conserved in PARP, including the active site glutamic acid, corresponding to Glu of hPARP. To test the possibility that Glu has a role in catalyzing formation of poly-ADP-ribose and to probe the similarities in active site structure between PARP and the toxins, we have made a series of site-directed replacements for this and certain other residues in hPARP, expressed the mutant proteins in E. coli, purified them, and characterized their changes in activity.


EXPERIMENTAL PROCEDURES

Materials

Reagents used were from Sigma unless otherwise noted. Aldrich supplied 3-aminobenzamide. Histones and NAD were purchased from Boehringer Mannheim, [alpha-P]NAD (800 Ci/mmol) was from DuPont NEN, and amylose resin and Factor Xa was from New England Biolabs. Snake venom phosphodiesterase was a gift from Miyoko Ikejima.

Bacterial Strains and Plasmids

The lon E. coli strain ME8417 was a gift of Miyoko Ikejima. The expression plasmid for poly-ADP-ribose polymerase, pTP(37) , consists of the complete cDNA for the human fibroblast poly-ADP-ribose polymerase under transcriptional control of the trp promoter in a pBR322-derived vector. The expression plasmid pMAL-C2 was purchased from New England Biolabs (Beverly, MA).

Molecular Biology

Standard molecular biology techniques were performed according to protocols described(38) . Site-directed mutants of hPARP were created using the Amersham Corp. oligonucleotide-directed in vitro mutagenesis system, version 2, using the following oligonucleotide primers: E988Q, CTATATAACCAATACATTGTC; E988D, CTATATAACGATTACATTGTC; E988A, CTATATAACGCGTACATTGTC; E931Q, GGAAACATGTATCAACTGAAGCACGC; E923Q, CCTGTTGGGACAAGTTGCCCTTG; E883Q, GATAGCCCCGCCTCAAGCGCCCGTG; and H862A, GATTGCTGTGGGCCGGGTCCAGG, according to the instructions of the manufacturer. All mutants were verified by DNA sequencing. For construction of pMBP40, DNA encoding the carboxyl-terminal portion of PARP in the plasmid pTP was amplified by polymerase chain reaction using the oligonucleotide primers 5`-MALs (5`-CCCTCTAGAAGTACTGTAAATCCTGGCACCAAGTCC-3`) and 3`-MALa (5`-CCCACTAGTCCCGGGGTCGACCACCGGGTGTGACTCGGCTACCTC-3`), and then cloned into the vector pMAL-c2 (New England Biolabs).

Partial Purification of Full-length PARP

E. coli crude extracts were prepared according to Ikejima et al.(37) , diluted to 0.2 M NaCl in buffer A (buffer A: 0.1 M Tris-Cl, pH 7.5, 0.05 M NaHSO(3), 5 mM EDTA, 5 mM dithiothreitol), and bound in batch to phosphocellulose P11 at 4 °C for 4 h. A column was poured with the slurry, and PARP was eluted with a linear gradient of 0.2-1.0 M NaCl in buffer A. Peak fractions were pooled, divided into aliquots, and stored at -80 °C.

Purification of MBP40 Fusion Proteins: Crude Extracts

Overnight cultures of TB1 (harboring pMAL-c2 derivative plasmids) in 10 ml of L-broth containing 100 mg/ml ampicillin were used to inoculate 500 ml of the same medium. Production of the fusion protein was induced by the addition of 1 mM IPTG for 4 h in cultures grown to mid-log phase. Cells were harvested and resuspended in 25 ml of lysis buffer (30 mM NaCl, 50 mM NaH(2)PO(4), pH 7.0, 0.25% Tween 20, 10 mM EDTA, 10 mM EGTA, 10 mM beta-mercaptoethanol) and frozen overnight at -20 °C. Cell suspensions were then thawed in a room temperature water bath and disrupted by sonication. NaCl (5 M) was added to a final concentration of 0.5 M, and crude extracts were prepared by centrifugation at 9000 times g for 30 min. Crude extract (25 ml) was bound in batch to 5-20 ml of amylose resin (New England Biolabs) which had been previously washed with column buffer (0.5 M NaCl, 50 mM NaH(2)PO(4), pH 7.2, 10 mM EDTA, 10 mM beta-mercaptoethanol), at 4 °C for 6 h. A column was poured with the slurry, which was then washed with 5-10 volumes of column buffer. The fusion protein was eluted with column buffer containing 50 mM maltose, and the pooled fractions were concentrated and dialyzed using Centriprep-30 ultrafiltration units (Amicon). The concentrated pools were then aliquoted and stored at -80 °C. Proteolytic nicking with factor Xa was performed in column buffer for 2 h at a ratio of 100:1 amylose-purified MBP40 to Factor Xa.

ADP-ribosylation Assays

PARP reactions (total volume 100 µl) were incubated at 25 °C in 100 mM Tris-Cl, pH 8.0, containing 100 µM [P]NAD (5 Ci/mmol), 25 µg/ml DNA, 10 mM MgCl(2), and 5 mM dithiothreitol. ADP-ribosylation assays were terminated by spotting samples onto 10% trichloroacetic acid/ether- (w/v) saturated 3MM filter paper. Unincorporated [P]NAD was removed with three 5% trichloroacetic acid washes of 15 min each. Samples were washed once with 100% methanol and dried before quantitation in 2 ml of fluor by scintillation counting.

Hydrolysis of the E988Q PARP-(mono-ADP-ribose) Bond with Hydroxylamine

Phosphocellulose-purified E988Q PARP (20 µg) was ADP-ribosylated under standard conditions with 50 µM [P]NAD (5 Ci/mmol) and precipitated by addition of an equal volume of 40% trichloroacetic acid. The trichloroacetic acid pellet was resuspended in 0.1 M HEPES pH 7.0, 0.5% SDS, and aliquots were incubated with an equal volume 2 M NH(2)OH, pH 7.0, for various time points. Reactions were terminated by spotting onto trichloroacetic acid/ether-saturated 3MM filter paper and unincorporated counts removed with repeated 5% trichloroacetic acid washes. Residual radiolabel (unreleased ADP-ribose groups) was measured by scintillation counting.

Gel Quantification of ADP-ribose Products

The method was based on a protocol from Aboul-Ela et al.(39) . ADP-ribosylation assays were terminated by addition of an equal volume of 40% trichloroacetic acid (w/v). Samples were precipitated on ice for 30 min. Trichloroacetic acid pellets were collected by microcentrifugation for 10 min at 4 °C, washed once with 20% trichloroacetic acid, reprecipitated with 20% trichloroacetic acid after being dissolved in formic acid, washed once with 100% ethanol at -20 °C, and dried in a vacuum centrifuge. The dried pellets were then incubated with 25 µl of 0.1 N NaOH, 10 mM EDTA at room temperature for 15 min to hydrolyze the ADP-ribose from protein. The samples were neutralized with HCl, MgCl(2) was added to 10 mM, and digestion with snake venom phosphodiesterase was performed for 5 h. The reaction was terminated by heating to 37 °C for 5 min, and the samples were run on a 15 or 20% polyacrylamide gel in Tris borate-EDTA buffer(38) . Radiolabel was then quantified using a Phosphor-Imager (Molecular Dynamics) and ImageQuant software (Molecular Dynamics).

Poly-ADP-ribose Chain Length Determination

Two-dimensional thin layer chromatography was performed essentially according to Keith et al.(40) . The chromatograms were quantified using a PhosphorImager and ImageQuant software. Average chain length, and number of chains/enzyme were calculated according to Kawaichi et al.(41) .


RESULTS

The report of Domenighini et al.(36) led us to construct a refined alignment of the catalytic region of known vertebrate and insect PARP sequences with that of DT and P. aeruginosa exotoxin A (Fig. 1) (42) . An alignment in which most of the major active site residues of these toxins were conserved, including Glu and His (corresponding to Glu and His, respectively, of DT) was achieved with introduction of only a minimal number of gaps. Besides Glu, the 3 other glutamic acids in the catalytic domain of hPARP (Glu, Glu, and Glu) are conserved in the polymerase from various species, but not in the toxins.


Figure 1: Amino acid sequence alignment of PARP from various species with diphtheria toxin and exotoxin A from P. aeruginosa. Toxin alignment(22) : DTA, DT fragment A; ETA, P. aeruginosa exotoxin A. PARP alignment: PARP.hum, human(67) ; PARP.mus, mouse(68) ; PARP.rat, rat (partial sequence)(69) ; PARP.bov, bovine(70) ; PARP.chk, chicken(71) ; PARP.xen, Xenopus laevis (EMBL: XLPARPG, accession no. Z12139 (B. M., Saulier-le Drean, personal communication); PARP.sar, Sarcophaga peregrina(72) ; PARP.dro, Drosophila melanogaster(12) . The PARP alignment is based on an alignment generated with PILEUP (Wisconsin Genetics Computer Group,(73) ). For each sequence group, identities to the consensus sequence are shown as dots (.). Gaps inserted to allow alignment within sequence groups are shown as dashes(-). Gaps inserted to allow alignment between sequence groups are designated by slashes (). Residues shared between PARP (including at least six of the eight PARP sequences) and one or both of the toxin sequences are designated by shading. Asterisks (*) represent toxin active site residues. Human PARP residues altered by site-directed mutagenesis are indicated by triangles (), and residues mutated by Simonin et al.(14, 58) are designated by V.



Using oligonucleotide-directed mutagenesis, we replaced Glu with aspartic acid, glutamine, and alanine, Glu, Glu, and Glu with glutamine, and His with alanine. Initially, we tested extracts of the lon E. coli strain ME8417 expressing the mutant PARP proteins for ability to incorporate radiolabel from [P]NAD into trichloroacetic acid-insoluble material during a 5-min incubation. The 3 Glu mutants showed a reduction in incorporation of 45- to 1100-fold, with E988A showing the greatest reduction in incorporation and E988Q the least (Table 1). The H862A mutation also caused a major reduction, comparable to that of E988A. In contrast, the E931Q, E923Q, and E883Q mutants displayed little or no change in activity.



The mutant forms of PARP were purified by chromatography on phosphocellulose, and the products formed by each during incubation with [P]NAD were characterized. The radiolabeled products were cleaved from the enzyme with weak base, fractionated by electrophoresis on SDS-polyacrylamide gels, and analyzed by autoradiography with a PhosphorImager (Fig. 2). In contrast to the wild-type protein, in which large amounts of high molecular weight polymer were formed, the E988Q mutant incorporated only mono-ADP-ribose (trace amounts of low molecular weight oligo-ADP-ribose could be detected in the 2 h sample, but only by careful inspection of the autoradiograph). Mono-ADP-ribose was also the major product of the E988A mutant, but small amounts of low molecular weight oligo-ADP-ribose (n leq 5) were seen at late time points. Among the 3 Glu mutants characterized, only the E988D mutant made long poly-ADP-ribose chains. Polymer elongation occurred at a greatly reduced rate in this mutant, however, and full-length polymer was seen only after prolonged incubation (1-2 h, in contrast to wild-type PARP, which produced full-length polymer within 5 min). Processive elongation by the E988D mutant is implied by the fact that long, rather than short, oligomer chains predominate at late time points, despite this mutant's low overall activity. The products made by the control mutants (E931Q, E923Q, and E883Q) were indistinguishable from those made by wild-type PARP (data not shown). The H862A mutant made a mixture of mono- and short oligo-ADP-ribose chains (n leq 10).


Figure 2: ADP-ribosylation product time courses of phosphocellulose-purified PARP. A, PARP (2 µg) was incubated with [P]NAD under standard assay conditions in a volume of 120 µl. At time intervals (noted in min), 20-µl aliquots were withdrawn and trichloroacetic acid-precipitated. Samples were then hydrolyzed with 0.1 N NaOH, 20 mM EDTA, neutralized, and run on 15% polyacrylamide, 0.1% SDS Tris-borate EDTA gels as described under ``Experimental Procedures.'' The dried gels were analyzed using a PhosphorImager. B, plot of PhosphorImager scanning data from samples in Fig. 3A. Wild-type (bullet), E988Q (Delta), E988D (box), and E988A (circle). Inset shows Glu mutant data at higher resolution.




Figure 3: Susceptibility of the E988Q PARP-(mono-ADP-ribose) bond to hydrolysis by hydroxylamine. Phosphocellulose-purified E988Q PARP (20 µg) was ADP-ribosylated under standard conditions with 50 µM [P]NAD, and precipitated by addition of an equal volume 40% trichloroacetic acid. The trichloroacetic acid pellet was resuspended in 0.1 N HEPES pH 7.0, 0.5% SDS and aliquots were mixed with an equal volume 2 M NH(2)OH pH 7.0 (bullet), or H(2)O (circle), and incubated for various time points. Reactions were terminated by spotting to trichloroacetic acid/ether-saturated paper and unincorporated counts removed with repeated 5% trichloroacetic acid washes. Residual (unreleased) ADP-ribosylation was measured by scintillation counting.



The E988Q mutant exhibited rapid initial incorporation of mono-ADP-ribose, differing little from the wild-type protein (Fig. 2B). This suggested little effect of the mutation on initiation, despite the large effect on elongation. In contrast, with the E988A mutant there was a major defect in initiation as well as elongation. The E988D mutant showed an intermediate level of activity, with defects in both initiation and elongation. Like the wild-type enzyme, all of the Glu mutants were dependent on DNA for ADP-ribosylation activity.

The identity of the E988Q product as mono-ADP-ribose was established by its comigration with authentic [P]ADP-ribose in polyacrylamide gels (Fig. 2) and its comigration with unlabeled ADP-ribose on two-dimensional thin layer chromatograms (data not shown). In addition, digestion of the product with snake venom phosphodiesterase yielded P-labeled AMP, as determined by two-dimensional thin layer chromatography (data not shown). The bond linking the E988Q mono-ADP-ribose product to PARP was sensitive to hydrolysis by hydroxylamine at pH 7.0 (Fig. 3), consistent with the ester bond reported to link poly-ADP-ribose to the wild-type enzyme(39) . Approximately two mono-ADP-ribosyl groups/PARP monomer were incorporated by the E988Q mutant at saturation, which is approximately equal to the number of poly-ADP-ribose attachment sites we found with wild-type PARP. In E. coli, a 90 kDa form of the enzyme resulting from aberrant chain initiation at an internal methionine is produced in addition to the wild-type 113 kDa form. Both forms were ADP-ribosylated to an equal extent in the E988Q mutant, although the 90 kDa form has been shown to have only basal ADP-ribosylating activity(37) . The value of approximately two poly-ADP-ribose chains (or mono-ADP-ribose groups, for the E988Q mutant) per monomer differs from an earlier report of approximately 15 sites of polymer attachment on PARP(41) . The basis of this discrepancy is unknown.

We introduced the Glu mutations into the minimal catalytic fragment of PARP in order to study the effect of these replacements on enzymic activity in the absence of other factors that might complicate analysis (e.g. activation by DNA, or the presence of the 90 kDa form). This fragment comprises the carboxyl-terminal 40 kDa of the 51-kDa PARP catalytic domain(8, 14) and has ADP-ribosyltransferase activity that is independent of DNA and approximately equal in magnitude to that of the intact enzyme in the absence of DNA (about 1/500 that of the fully activated enzyme; 14). To facilitate purification of the fragment, we attached maltose-binding protein (MBP) to its amino terminus using the polymerase chain reaction (43, 44) . Cleavage of the resulting fusion protein, termed MBP40 (82 kDa) at a factor Xa protease site at the fusion junction yielded the expected MBP fragment of 42 kDa and PARP fragment of 40 kDa (data not shown). Fortuitously, the MBP40 fusion protein proved to be 10-fold more active than the 40-kDa catalytic fragment alone, due to the ability of the PARP catalytic domain to ADP-ribosylate the MBP portion of the fusion protein. The intact MBP40 fusion protein was therefore routinely used instead of the 40-kDa fragment in basal activity measurements of the Glu mutants.

Following initial studies demonstrating that the effects of the Glu mutations in MBP40 were similar to those in hPARP, we determined the K(m) for NAD for each mutant form of MBP40. The K(m) for NAD of the PARP catalytic fragment has been reported to be roughly 50 µM(14) , and published values for intact PARP range between 50 and 100 µM NAD (37, 45) As shown in Table 2, none of the Glu mutants showed major alterations in K(m). Values for the catalytic coefficient, k/K(m), were dramatically reduced relative to wild-type MBP40, however. The E988Q and E988D mutants were decreased by 35- and 50-fold, respectively, and the E988A by 400-fold. These results are consistent with the notion that Glu plays a catalytic role in formation of poly-ADP-ribose. It should be noted that the values obtained were calculated from total ADP-ribose incorporation under the assumption of equal K(m) values for polymer initiation and elongation. There may be a difference in these values, however, since the K(m) for NAD of the mono-ADP-ribosylating (elongation deficient) E988Q mutant appears to be slightly lower than that of the wild-type enzyme.



Catalysis of the initiation reaction for poly-ADP-ribose synthesis, in which (ADP-ribose)-protein esters are formed, could conceivably differ from that of the elongation and branching reactions, which produce polyglycosides of ADP-ribose. We therefore characterized the products of the MBP40 fusion proteins more fully in order to explore the role(s) of Glu in polymer elongation and initiation further and to seek information about its possible role in polymer branching. Wild-type and mutant MBP40 fusion proteins were incubated with [P]NAD, and the trichloroacetic acid-precipitable fraction was collected, washed, and hydrolyzed with base. After neutralization, the products were treated with snake venom phosphodiesterase and separated by two-dimensional thin layer chromatography. Measurements of label in phosphoribosyl-AMP (main chain product), AMP (product of the protein-distal chain terminus), and diphosphoribosyl-AMP (branch product), provided the basis for estimating chain length, number of polymer chains/enzyme, and degree of branching, according to the method of Kawaichi et al.(41) .

The Glu replacements caused only modest effects on the rate of initiation, as judged by the number of polymer chains/enzyme (Table 3). The E988Q mutant was reduced in initiation only 3-fold, and the E988D and E988A mutants by 20- and 30-fold, respectively. These results are consistent with the kinetics of polymer formation by the full-length mutant PARP proteins (see Fig. 2). Chain elongation, on the other hand, as estimated from the phosphoribosyl-AMP product, was dramatically reduced in the E988Q and E988A mutants (2800- and 2200-fold, respectively), but decreased only 20-fold by the E988D substitution. Estimates of branching efficiency from the diphosphoribosyl product could not be obtained from the E988Q and E988A proteins because they made only short polymer and in trace amounts. In the E988D mutant, branch, as a proportion of total poly-ADP-ribose synthesized, was only slightly lower than in the wild-type enzyme (0.5 and 0.8% for the mutant and wild-type, respectively).




DISCUSSION

The findings that both conservative and nonconservative replacements for Glu caused major reductions in elongation of poly-ADP-ribose chains in PARP (and the mutation of His to Ala caused a similar reduction) are consistent with the suggestion that PARP and the ADP-ribosylating toxins have similar active site structures and folds. The alignment alone provides substantial support for this proposal (Fig. 1). In DT, which appears to have the closest sequence similarity to PARP among the toxins, His, Tyr, Tyr, and Glu, which are near each other within the NAD site, are conserved in the alignment with hPARP as His, Tyr, Tyr, and Glu, respectively. All of these residues are identical in PARP from various species, and there is also significant conservation among neighboring residues, particularly near the histidine, the first tyrosine, and the glutamate. In the crystallographic structure of DT, His, Tyr, and Tyr are believed to form the nicotinamide subsite, and Glu has been shown to be important in catalyzing ADP-ribose transfer(48) . In addition, Phe in hPARP, which is also conserved among diverse species, aligns with Trp of DT, another determinant of NAD affinity(47) .

In this study we focused on Glu of hPARP because its putative homolog in DT, Glu, plays a major catalytic role and appears to be the only functional active site residue universally conserved among the ADP-ribosylating toxins. The E988Q and E988D mutations demonstrated the importance of the presence and precise spatial location of a side chain carboxylate at this site for poly-ADP-ribose chain elongation. Thus, the Gln mutant was almost devoid of elongation activity (2800-fold reduction), whereas the Asp mutant was reduced only about 20-fold. These mutations had relatively little effect on chain initiation, however, a 3-fold reduction for the Gln and 20-fold for the Asp, indicating a different, and less important, function for Glu in initiation. The Ala substitution caused reductions in elongation and initiation comparable to those of the E988Q and E988D mutants, respectively.

In DT (and by implication, aeruginosa exotoxin A) ADP-ribosylation of the diphthamide residue of EF-2 proceeds via an ordered sequential mechanism in which NAD must bind to the active site cleft of the catalytic fragment before the second substrate EF-2 can bind and the reaction take place(48) . Replacing Glu of DT with Gln, Asp, or Ser showed that this residue is essential for the ADP-ribosylation of EF-2, but relatively unimportant for NAD-glycohydrolysis(46) . On the basis of kinetics measurements for the two reactions, together with knowledge of the K(d) for NAD and the stereochemistry of the ADP-ribose-protein linkage, the ADP-ribosylation and NAD-glycohydrolase reactions are believed to proceed via a direct SN(2) displacement mechanism(15, 49, 50) . In this model, the nicotinamide group of NAD would be displaced by the incoming nucleophile, namely the diphthamide moiety of EF-2 for the ADP-ribosylation reaction or water for NAD-glycohydrolysis. The dramatic reduction in ADP-ribosylation activity that occurs when Glu is mutated may be explained by the carboxyl group of Glu serving as a general base that activates the incoming nucleophile, diphthamide(50) . This mechanism is consistent with the fact that mutation of Glu has little effect on NAD affinity or the NAD-glycohydrolase reaction. In other ADP-ribosylating toxins, the homologs of Glu of DT are assumed to serve similar functions in activating the incoming nucleophile that serves as the attachment point for ADP-ribose.

For hPARP we propose that Glu functions in a similar manner to catalyze elongation of poly-ADP-ribose chains (Fig. 4) by hydrogen bonding to properly position and to activate, through its action as a general base, the 2`-OH of the terminal adenosine group of a nascent poly-ADP-ribose chain. This activated species would carry out the nucleophilic attack on the nicotinamide-ribose bond of NAD to form an (ADP-ribose)-(ADP-ribose) glycoside with an alpha linkage (see below). Similar catalytic and positioning roles through hydrogen bonding have been proposed for active site carboxyl groups in other systems(51, 52, 53) . Consistent with this prediction, the E988D mutant was the only mutation at position 988 that retained significant polymer elongation activity. Trace elongation activity of the E988A mutant might result from the uncovering of another acidic group near the PARP active site by the smaller alanine side chain, as observed in the case of an alanine substitution of a catalytic glutamic acid of ricin, a toxin with endoglycosidase activity (54) .


Figure 4: Model for the role of Glu in poly-ADP-ribose synthesis. A, initiation of poly-ADP-ribose. Glu facilitates nucleophilic attack on the nicotinamide-ribose bond by hydrogen bonding to, and positioning, the substrate (automodification domain) acidic side chain. B, elongation of poly-ADP-ribose. Glu catalyzes the nucleophilic attack on the nicotinamide-ribose bond by activating the 2`-OH of the terminal adenosine of a polymer chain. MOD, PARP automodification domain; CAT, PARP catalytic domain.



In PARP one would predict a priori that the precise role of Glu in the initiation reaction is different from its role in the elongation reaction, given that acidic side chain(s) on the automodification domain serve as sites of chain attachment. Glu could serve in initiation by hydrogen bonding to position these substrate acidic groups of the automodification domain in the PARP active site cleft, thereby facilitating nucleophilic attack by these groups on the nicotinamide-ribose bond. The substrate carboxyl groups presumably function as intrinsic nucleophiles, and being predominantly ionized at neutral pH, they would not require activation by Glu (Fig. 4). This proposal is consistent with the relatively small (3-fold) reduction in initiation activity seen in the E988Q mutant and somewhat larger effects seen with the other Glu mutants. Glutamine has been shown to activate incoming nucleophiles such as water via hydrogen bonding in other instances(50) . Significantly, PARP also hydrolyzes NAD (NAD-glycohydrolase activity) concomitant with polymer synthesis, which may reflect a competition between the initiation of new polymer chains and hydrolysis of NAD by water.

The notion that PARP and the ADP-ribosylating toxins share a common mechanism of catalysis is supported by, among other factors, the stereochemistry of the ADP-ribosylation products. All ADP-ribosylating toxins examined cause inversion of configuration at the anomeric carbon of the nicotinamide ribose, from beta-linkage in NAD to alpha between the ADP-ribose and the substrate protein(49) . Similarly, the (ADP-ribose)-(ADP-ribose) glycosidic bonds made by PARP are alpha in configuration(55, 56, 57) . The configuration of the ester linkage between the initial ADP-ribose of poly-ADP-ribose and the protein is not known. It has been proposed that an SN(2) mechanism may account for this inversion in the case of DT(49) . Although it is possible that an enzyme-catalyzed inversion of configuration can occur by an SN(1) mechanism, there are few examples, presumably because of steric and electronic considerations regarding the substrate nucleophile and leaving groups(58) . Additionally, NAD-glycohydrolases, most of which use an SN(1) mechanism, are additionally able to catalyze base exchange and methanolysis of NAD, whereas neither PARP nor the toxins catalyze either reaction. Finally, poly-ADP-ribose chains are elongated in a manner that is analogous to mono-ADP-ribosylation by the toxins: that is, by sequential addition of new ADP-ribose residues to the protein-distal termini of growing polymer chains(59, 75) . (^2)A proximal growth model proposed earlier (60) is apparently incorrect.

After this study was initiated, another model for PARP structure and function was put forth by Simonin et al.(14, 61) , in which PARP was proposed to resemble a family of glutamic acid dehydrogenases, in particular the Clostridium symbosium glutamic acid dehydrogenase, whose crystal structure was recently determined(62) . The homology reported between PARP and the glutamate dehydogenases is limited, however, and many gaps were used in aligning PARP residues with those conserved among dehydrogenases. Although these authors found that mutations of several residues conserved between PARP and the dehydrogenases have an effect on the ADP-ribosylation activity of PARP (14, 61) , a toxin model for PARP structure can also explain the results of these same mutations. For instance, 2 of the PARP residues mutated by Simonin et al., Lys and Asp, are located in important parts of a toxin-like structure. Lys is conserved with the toxins, and mutations of this residue in DT (Arg) result in a loss of activity. (^3)Likewise, although Asp is not conserved with the toxins, mutations at this position also result in a loss of activity (Trp of DT; 47). Our finding that an alanine substitution for His in hPARP results in a large loss of ADP-ribosylation activity is consistent with results obtained with substitutions of His in DT(63) . Conversely, the dehydrogenase model does not predict a role for Glu, which would be on the outer edge of a beta-strand, away from the active site cleft. If this were the true physical context of Glu, it would be more difficult to rationalize a large, and specific, effect on polymer synthesis of a glutamine replacement.

It should be noted that the toxins do not use a Rossman fold to bind NAD, and the structure of the C. symbosium glutamate dehydrogenase, or any other dehydrogenase, does not resemble the structure of the catalytic domain of either DT or P. aeruginosa exotoxin A. A crystallographic structure of the PARP catalytic domain will be required to show which physical model is correct. The presence of Glu in the active site cleft of PARP would be strong evidence in favor of a mechanism of ADP-ribosylation similar to that of the ADP-ribosylating toxins.

If PARP is ultimately shown to have an active site fold similar to that of the ADP-ribosylating toxins, this will raise interesting questions regarding the evolutionary lineages of the various ADP-ribosyltransferases. The ADP-ribosyltransferases are likely to be an ancient class of enzymes, as evidenced by the regulation of nitrogen metabolism by ADP-ribosylation in the bacterium Rhodospirillum rubrum(64) and the poly-ADP-ribosylating activities reported in dinoflagellates (65) and a thermophilic archaebacterium, Sulfobolus sulfataricus(66) . While the ADP-ribosylating toxins may have evolved directly from precursor ADP-ribosyltransferases in bacteria, it has been suggested, on the basis of the strong specificity of diphtheria toxin for the diphthamide residue of elongation factor-2, that such toxins may have originated from the ``capture'' and modification of eukaryotic genes encoding endogenous regulatory proteins(74) . Additional sequence data from eukaryotic and prokaryotic ADP-ribosyltransferases will aid in resolving such questions.


FOOTNOTES

*
This work was supported by National Institutes of Health Grants AI 22021 and AI 22848. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
This work represents research performed in partial fulfillment of requirements for the Ph.D. degree in the Department of Molecular Biology and Microbiology, Sackler School of Biomedical Sciences, Tufts University School of Medicine. Present address: Dept. of Cellular and Molecular Biology, Dana-Farber Cancer Institute, Boston, MA 02115.

Present address: Dept. of Biochemistry and Molecular Biology, Wright State University School of Medicine, Dayton, OH 45435.

**
To whom correspondence should be addressed; Tel.: 617-432-1930; Fax: 617-432-0115; collier{at}warren.med.harvard.edu.

(^1)
The abbreviations used are: PARP, poly-ADP-ribose polymerase; hPARP, the human polymerase; DT, diphtheria toxin; MBP, maltose-binding protein.

(^2)
G. T. Marsischky, M. Ikejima, and R. J. Collier, manuscript in preparation.

(^3)
H. Fu, S. R. Blanke, and R. J. Collier, manuscript in preparation.


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