JBC Biosymposia, Inc.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M602673200 on August 9, 2006

J. Biol. Chem., Vol. 281, Issue 40, 29525-29532, October 6, 2006
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
281/40/29525    most recent
M602673200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Adhikari, S.
Right arrow Articles by Roy, R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Adhikari, S.
Right arrow Articles by Roy, R.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Magnesium, Essential for Base Excision Repair Enzymes, Inhibits Substrate Binding of N-Methylpurine-DNA Glycosylase*

Sanjay Adhikari, Jeffery A. Toretsky, Linshan Yuan, and Rabindra Roy1

From the Department of Oncology, Lombardi Comprehensive Cancer Center, Georgetown University Medical Center, Washington, D. C. 20057

Received for publication, March 22, 2006 , and in revised form, August 8, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
N-Methylpurine-DNA glycosylase (MPG) initiates base excision repair in DNA by removing a wide variety of alkylated, deaminated, and lipid peroxidation-induced purine adducts. MPG activity and other DNA glycosylases do not have an absolute requirement for a cofactor. In contrast, all downstream activities of major base excision repair proteins, such as apurinic/apyrimidinic endonuclease, DNA polymerase beta, and ligases, require Mg2+. Here we have demonstrated that Mg2+ can be significantly inhibitory toward MPG activity depending on its concentration but independent of substrate type. The pre-steady-state kinetics suggests that Mg2+ at high but physiologic concentrations decreases the amount of active enzyme concentrations. Steady-state inhibition kinetics showed that Mg2+ affected Km, but not Vmax, and the inhibition could be reversed by EDTA but not by DNA. At low concentration, Mg2+ stimulated the enzyme activity only with hypoxanthine but not ethenoadenine. Real-time binding experiments using surface plasmon resonance spectroscopy showed that the pronounced inhibition of activity was due to inhibition in substrate binding. Nonetheless, the glycosidic bond cleavage step was not affected. These results altogether suggest that Mg2+ inhibits MPG activity by abrogating substrate binding. Because Mg2+ is an absolute requirement for the downstream activities of the major base excision repair enzymes, it may act as a regulator for the base excision repair pathway for efficient and balanced repair of damaged bases, which are often less toxic and/or mutagenic than their subsequent repair product intermediates.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cellular DNA is continuously exposed to endogenous or exogenous chemical or physical agents that induce DNA lesions. DNA base damage threatens genomic stability and cellular viability. Multiple DNA repair pathways exist in all organisms, from bacteria to humans, to preserve the integrity of the genome (1). If not repaired, damaged bases could be mutagenic (2) and/or cause cell death by blocking DNA replication (3). In all organisms, repair of DNA-containing small adducts, as well as altered and abnormal bases, occurs primarily via the base excision repair (BER)2 pathway, beginning with cleavage of the base by a DNA glycosylase (1, 2). Mechanistically, DNA glycosylases are categorized into two classes: mono- and bifunctional DNA glycosylases. Monofunctional DNA glycosylases, such as N-methylpurine-DNA glycosylase (MPG) and uracil-DNA glycosylase, use an activated water molecule as a nucleophile to generate an apurinic or apyrimidinic (AP) site in DNA. Bifunctional DNA glycosylases/AP lyases, such as NTH1 and OGG1, use an activated amino group (Lys) or imino group (Pro) as the nucleophile to create a Schiff base intermediate that coordinates base removal and subsequent strand incision (AP lyase) 3' to the AP site (4, 5). The mammalian MPG is known to excise at least 17 structurally diverse modified bases from DNA (6). These lesions include 3-alkylpurines, 7-alkylguanine, 1,N6-ethenoadenine ({epsilon}A), N2,3-ethenoguanine, and hypoxanthine (Hx), all of which are purine derivatives (7–12). Moreover, the base alterations are located in both the major and minor grooves of duplex DNA. Its orthologs in Escherichia coli (AlkA) and yeast (MAG) have an overlapping although not identical substrate range. Nonetheless mammalian MPG and E. coli AlkA do not share significant sequence similarity or structural homology (13, 14), despite this functional similarity and the fact that 3-methyladenine is a preferred substrate for both. MPG excises {epsilon}A and Hx more efficiently than AlkA and MAG (11), but unlike AlkA, it cannot excise O2-alkylpyrimidines (15, 16) and oxidized bases such as 5-formyluracil and 5-hydroxymethyluracil (17) from DNA. MAG also does not excise O2-methylthymine (6, 18).

The role of the magnesium (Mg2+) ion on DNA glycosylases is not well established. Notably, intranuclear Mg2+ concentration is highly variable. Depending on conditions, its may vary up to 75 mM. Unlike the normal cells, tumor cells have a higher Mg2+ content in the nucleus (19, 20). It is reported that 5–10% of total Mg2+ is in a free state in the cells (21). A recent report suggests that Mg2+ reduces the efficiency of the base excision and strand incision activities of hOGG1 on DNA containing 8-oxoG under single turnover conditions (22); however, the reduction was more pronounced for the AP-lyase activity. The Schiff base formation between hOGG1- and 8-oxoG-containing DNA was abrogated in the presence of Mg2+. These results suggest that hOGG1 operates mainly as a monofunctional glycosylase under physiologic concentrations of Mg2+ (22). There is a growing list of DNA glycosylases that cleave the damaged base depending on the base opposite to the damaged one. For example, for base discrimination, human endonuclease III (hNTH1) depends strongly on Mg2+ (23). However, Mg2+ has also been shown to stimulate the turnover of thymine-DNA glycosylase (24). Mg2+ acts as a cofactor of many enzymes involved in oxidative phosphorylation, nucleic acids and protein synthesis, and mitotic activity of normal cells. The adult human body contains 21–28 g of Mg2+. As a nutritional element the daily requirement of Mg2+ (200–700 mg) is crucial (25).

In the present study, we have demonstrated that Mg2+ is not required for MPG activity. However, Mg2+ can inhibit MPG activity significantly, at physiologically relevant concentrations, by abrogating its substrate binding ability without any effect on its catalytic chemistry. Because Mg2+ is an absolute requirement for the downstream activities of the major BER enzymes (26), it may act as a regulator for the BER pathway for efficient and balanced repair of damaged bases, which are often less toxic and/or mutagenic than their subsequent repair product intermediates.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Purification of Recombinant Mouse MPG—The mouse MPG wild type and the deletion mutant were purified as described previously (27).

Oligonucleotide Substrate Preparation—A 50-Mer oligonucleotide substrate containing Hx and {epsilon}A, with the sequence 5'-TCGAGGATCCTGAGCTCGAGTCGACGXTCGCGAATTCTGCGGATCCAAGC-3' (where X represents {epsilon}A or Hx), was purchased from Operon Technologies (Alameda, CA) and Gene Link (Hawthorne, NY). The complementary oligonucleotide containing an A opposite {epsilon}A or Hx was synthesized by the Recombinant DNA Laboratory Core Facility at the University of Texas Medical Branch (Galveston, TX). The oligonucleotides were purified on a sequencing gel. The Hx or {epsilon}A oligonucleotide was labeled at the 5'-end using T4 polynucleotide kinase and [32P]ATP and annealed to a complementary oligonucleotide to prepare a 32P end-labeled duplex oligonucleotide as described previously (27).

MPG-mediated Excision Activity Assay—The MPG proteins, full-length (1.1–12.5 nM) and truncated (N{Delta}100C{Delta}18, 3.2 nM), were incubated individually with 5' 32P-labeled {epsilon}A- or Hx-containing duplex oligonucleotide substrates (1 nM) in the presence of different divalent metal ions, EDTA, EGTA, control duplex oligo or in various combinations for 10 min at 37 °C in an assay buffer (25 mM HEPES-KOH, pH 7.9, 0.5 mM dithiothreitol, 150 mM NaCl, and 10% glycerol) in a total volume of 20 µl. The reaction was stopped by inactivating the enzyme at 75 °C for 5 min. The products containing the AP sites were then quantitatively cleaved into smaller fragments by incubating them with a large excess amount (100 ng) of AP endonucleases at 37 °C for 10 min after the concentration of Mg2+ was adjusted to 5 mM (27).

The reaction mixture was then mixed with 40 µl of loading buffer containing 1x DNA dye (diluted from blue-orange 6x loading dye; Promega, Madison, WI), 85% formamide, and 0.03 N NaOH and heated at 95 °C for 5 min. The samples were resolved by electrophoresis at 60 °C using Criterion gel (BioRad) containing 20% polyacrylamide and 7 M urea. Radioactivity in the incised oligonucleotides was quantified by exposing the gel to x-ray films and measuring the band intensities using an imager (Chemigenius Bioimaging System) with quantification software (Syngene Inc., San Diego, CA).

DNA Binding Studies Using Surface Plasmon Resonance—In search of a mechanism of modulation of MPG activity by Mg2+, we examined the MPG-Hx binding using a Biacore 1000 (Biacore, Uppsala, Sweden). A 50-mer duplex oligonucleotide containing an Hx or adenine (no modification) at the 26th position from the 5'-end of one strand was used for measuring enzymeoligo interaction. Oligos were biotinylated and immobilized on streptavidin-coated Biacore chips. Then N{Delta}100C{Delta} 18 MPGs at various concentrations (0–400 nM) were injected, and the surface plasmon resonance units were measured over 30 min. There is an advantage to using the truncated MPG (N{Delta}100C{Delta}18) over the full-length protein for binding assays; the truncated protein is slower (75% less active) in the Hx cleavage reaction, albeit with an inhibitory effect similar to Mg2+ (data not shown). Thus it was convenient to establish the equilibrium substrate binding condition with the truncated protein. The binding kinetics for oligonucleotides containing Hx or no modification was established with a series of MPG concentrations. The Langmuir isotherms at various protein concentrations allowed us to calculate the kinetic binding parameters based on on/off rates and protein concentrations. Then we tested the effect of various concentration of Mg2+ on MPG (125 nM) binding toward Hx or unmodified oligonucleotide.

Single Turnover (STO) Kinetic Study—The enzymes (12.5 nM) were incubated individually with 1 nM 5' 32P-labeled Hx-containing duplex oligonucleotide substrates at 37 °C in an assay buffer (25 mM HEPES-KOH, pH 7.9, 100 µg/ml nuclease-free bovine serum albumin, 0.5 mM dithiothreitol, 150 mM NaCl, and 10% glycerol) in a total volume of 100 µl. Aliquots of 5 µl were taken out at different time points (0–20 min) and heat-inactivated at 80 °C in a preheated microcentrifuge tube. Then we tested the Mg2+ effect at concentrations of 0–500 µM. The products containing the AP sites were quantitatively cleaved into smaller fragments followed by resolving on denaturing gels; radioactivity in the incised oligonucleotide was also quantified as described for the activity assay.

Burst Analysis—The enzymes (5 nM) were individually incubated with a 5' 32P-labeled hypoxanthine-containing duplex oligonucleotide (50 nM) at 37 °C under conditions similar to those described for the STO kinetic study.

Steady-state Kinetics—The enzyme (12.5 nM) was incubated with a 5' 32P-labeled Hx-containing duplex oligonucleotide (0–60 nM) substrates for 5 min at 37 °C under assay conditions similar to those described above. The reaction products were also analyzed and quantified as described for the activity assay.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Effect of Mg2+ and Other Divalent Ions on MPG Activity—The activity of purified full-length MPG was measured in the presence of 0–5 mM MgCl2 concentration using Hx as a substrate. Although Mg2+ showed a slight stimulatory effect with Hx (70–125%) at micromolar concentrations, a higher concentration of Mg2+ (1–5 mM) inhibited MPG reaction significantly (Fig. 1, A and B). Mg2+ inhibited the reaction, but an equivalent amount of EDTA could rescue nearly full activity of MPG (Fig. 2). EGTA did not have an effect on the inhibition. However, EGTA as a specific chelator had a significantly higher affinity for Ca2+ than for Mg2+, and thus the results were in accordance with expectations. Based on these findings, it appears that the Mg2+-mediated inhibition of MPG is reversible. Mg2+ has been shown to modulate the double helix structure of DNA (26). Mg2+ may modify the DNA structure and thus inhibit the binding of MPG to the DNA substrate. We tested this possibility by adding an increasing amount of undamaged oligonucleotide containing adenine in place of Hx. The results showed that unlike EDTA, DNA could not rescue MPG activity from Mg2+-mediated inhibition; rather increasing the DNA concentration inhibited the activity of MPG (Fig. 3). DNA-mediated inhibition of activity is very common for MPG and other DNA glycosylases (26). Nevertheless, it is apparent that to affect activity, Mg2+ modulates MPG protein but not the substrate DNA. Next, we tested the effect of other divalent metals on MPG-Hx enzymatic reaction (Table 1). Zn2+ and Mn2+ had a pronounced inhibitory effect, whereas Ca2+ was the poorest inhibitor. The IC50 values for these metal ions in inhibiting MPG were as follows: Zn2+ < Mn2+ < Mg2+ < Ca2+. Because MPG has broad substrate specificity, we tested DNA substrates other than Hx for Mg2+-mediated modulation of the MPG activity and found that Mg2+ could inhibit the MPG excision activity for {epsilon}A-containing oligonucleotides. However, no stimulation was observed by Mg2+ in excision of {epsilon}A by MPG. As expected, EDTA rescued MPG activity toward {epsilon}A excision from Mg2+ inhibition (data not shown). These results demonstrated that Mg2+ can be slightly stimulatory or significantly inhibitory toward MPG activity depending on its concentration and substrate type.


View this table:
[in this window]
[in a new window]
  		TABLE 1
Modulation of product formation by various divalent metal ions in MPG-mediated Hx cleavage reaction

 


Figure 1
View larger version (16K):
[in this window]
[in a new window]
  		FIGURE 1.
A, modulation of product formation by Mg2+ in MPG-mediated Hx cleavage reaction. MPG (3.25 nM) was reacted with a 50-mer 32P-labeled Hx-containing oligonucleotide (1 nM) at 37 °C for 10 min. The details of the reaction conditions are described under "Materials and Methods." B, data obtained in A and a separate set of experiments using 1.1 nM MPG under reaction conditions similar to those in A were plotted. Data represent mean values with standard error derived from three independent experiments.

 


Figure 2
View larger version (28K):
[in this window]
[in a new window]
  		FIGURE 2.
Effect of EDTA on Mg2+-mediated inhibition. MPG (1.1 nM) was reacted with Hx-containing oligonucleotide (1 nM) under conditions similar to those described for Fig. 1 with the exception of the addition of EGTA and different EDTA concentrations. Data represent mean values with standard error derived from three independent experiments.

 
Mechanism Analysis for Inhibition of MPG by Mg2+—Although Zn2+ appears to be the most important inhibitor among the divalent metal ions tested, we studied the mechanism of Mg2+ inhibition because Mg2+ is an absolute requirement for the rest of the downstream enzymes in the BER pathway. To elucidate this mechanism, we tested the major steps of the reaction process. We performed pre-steady-state kinetics to analyze the mechanisms of Mg2+ inhibition of MPG activity. Pre-steady-state kinetic analysis provides the opportunity to identify the intermediate step(s) that might be affected by Mg2+. Fig. 4 illustrates the basic intermediate steps of the MPG reaction. In general, we and others have showed that DNA glycosylases such as MPG bind substrates and generate a Michaelis-Menten complex in which an enzymatic reaction or catalysis occurs to cleave the scissile glycosidic bond (7, 28–30). Then the enzyme dissociates from the product, the AP sites, and turns over to bind a new molecule of substrate. All of the reaction rates at transient steps are slow for MPG and in general for all of the DNA glycosylases tested thus far (7, 28–30). We analyzed the effect of Mg2+ on substrate binding using surface plasmon resonance in Biacore. We took advantage of the slow reaction rates of MPG and measured the effect of Mg2+ on the glycosidic bond cleavage (catalysis) step by STO kinetics and the product dissociation step by multiple turnover reaction conditions. In addition, the multiple turnover conditions allowed us to measure the active enzyme concentration that might be affected by Mg2+.


Figure 3
View larger version (11K):
[in this window]
[in a new window]
  		FIGURE 3.
Effect of DNA on Mg2+-mediated inhibition. MPG (1.1 nM) was reacted with Hx-containing oligonucleotide (1 nM) under conditions similar to those shown for Fig. 1 with the exception of the addition of different concentrations of duplex control DNA of the same sequence as the lesioned DNA. Data represent mean values with standard error derived from three independent experiments.

 


Figure 4
View larger version (5K):
[in this window]
[in a new window]
  		FIGURE 4.
Proposed model for reaction mechanism of MPG. The steps and the notations used to indicate the intermediate steps of MPG reaction are as follows: E, the enzyme MPG; S1, the substrate with the modified base ({epsilon}A or Hx); P1, the excised modified base; P2, the oligo substrate with AP site after the base is excised. The rate constants are: KD, the binding constant between substrate and enzyme; kchem, the catalytic constant at the chemistry step; and kpd, the product dissociation constant.

 
Surface Plasmon Resonance (SPR) to Analyze Substrate Binding—The 50-mer oligonucleotide containing Hx was biotinylated and immobilized on streptavidin-coated chips to a density of 550 resonance units (data not shown). Unmodified oligonucleotide with a similar sequence was used as a control. Using SPR one can follow real-time DNA-protein interactions compared with other commonly used methods, such as a gel shift assay. We established the binding conditions for oligonucleotides containing Hx or no modification by injecting truncated MPG at 0–400 nM. The SPR was monitored and the resonance units plotted against time (Fig. 5, A and B). The Langmuir isotherms at various protein concentrations allowed us to calculate the kinetic parameters based on on/off rates and protein concentrations. We found that the KD values (dissociation constants) for unmodified and Hx-containing oligonucleotides were 176 and 2.5 nM, respectively, indicating that MPG binds both the oligonucleotides, albeit with ~70-fold higher affinity for Hx. Then we tested the effect of Mg2+ on MPG binding toward Hx-containing oligonucleotide and found that Mg2+ increased the affinity between MPG and Hx at a lower concentration and significantly inhibited the interaction at its higher concentrations in a dose-dependent manner (Fig. 5C). These results suggest that the modulation of substrate binding by Mg2+ altered the overall activity of MPG.

STO Kinetics—Prompted by the observation that Mg2+ can inhibit MPG, probably through modulation of substrate binding, we tested whether Mg2+ affects any of the catalytic intermediate steps other than the binding step. We conducted STO kinetics with full-length MPG proteins to measure the kchem (28–30). The reaction was performed at substrate and enzyme concentrations of 1 and 12.5 nM, respectively. Data were analyzed using the first-order rate equation,

Formula(Eq. 1)
where A0 represents the amplitude of the exponential phase and kobs is the observed rate constant associated with the reaction process. Under the STO conditions ([E] >> [S]), all of the substrate molecules should remain bound by enzymes. The binding step should not affect the rate of product formation, and hence, under these conditions kobs can be considered as kchem. Under three different concentrations of Mg2+, the kchem was similar (0.32 ± 0.1) for MPG-Hx reactions (Fig. 6), indicating that Mg2+ does not affect the chemistry step of MPG activity.

Burst Analysis—Next we measured the rate of product release (Kpd) and active enzyme concentration available for the reaction (28, 31). The extremely slow turnover rate of MPG during excision of Hx provided the opportunity to perform burst analysis under reaction conditions of [S] >> [E], where the substrate and enzyme concentration were 50 and 5 nM, respectively. The data were fit to the equation,

Formula 2(Eq. 2)
Although the curves were biphasic, indicating the presence of pre-steady state and steady state, the burst periods were unusually long (5 min). Generally, the burst rate, which is actually the rate of enzyme reaction during the pre-steady state of burst analysis, is similar to the rate of chemical reaction (kchem) at pre-steady state under STO conditions. Indeed, the times required to complete the first cycle (burst) of product generation in burst analysis (Fig. 7) and attain the saturation of product formation in STO kinetic analysis (Fig. 6) are similar. This slow reaction rate made the burst phase sluggish. The slow reaction rate provided the opportunity to measure pre-steady-state kinetics without using stop flow.


Figure 5
View larger version (19K):
[in this window]
[in a new window]
  		FIGURE 5.
A and B, Langmuir isotherm of MPG binding to 50-mer biotinylated oligonucleotides containing adenine (A) or Hx (B) using Biacore 1000. Binding kinetic parameters were obtained using various MPG concentrations (0–400 nM). C, effect of Mg2+ on MPG binding to Hx-containing oligonucleotide. The DNA-protein interactions were carried out using 125 nM MPG in the presence of different Mg2+ concentrations (0–5 mM).

 


Figure 6
View larger version (20K):
[in this window]
[in a new window]
  		FIGURE 6.
A, effect of Mg2+ on MPG-Hx reaction under single turnover conditions. The reaction was performed using substrate and enzyme concentrations of 1 and 12.5 nM, respectively, as described under "Materials and Methods." B, effect of Mg2+ on pre-steady-state kinetic parameters under single turnover conditions. Data derived from A were analyzed using the first-order rate equation [P]t = A0{1 – exp(–kobst)} as described under "Results." kchem is the catalytic constant at the chemistry step.

 
Plots of product concentration (Pt) versus time (t) were analyzed using Equation 2 to determine the kinetic parameters, A0 (amplitude of the burst), and kss (slope of the linear phase). In terms of the microscopic rate constants in Fig. 5, A0 and kss can be defined by the following,

Formula 3(Eq. 3)
and

Formula 4(Eq. 4)
respectively, where k' is the effective rate constant for the overall pre-steady-state process, E + S1->E·P2, before the product (P2) is released from the enzyme. Fig. 7 clearly shows that k' >> kpd, which simplifies Equations 3 and 4 to

Formula 4(Eq. 5)
and

Formula 4(Eq. 6)


Figure 7
View larger version (22K):
[in this window]
[in a new window]
  		FIGURE 7.
A, effect of Mg2+ on MPG-Hx reaction under multiple turnover conditions. The reaction was performed using substrate and enzyme concentrations of 50 and 5 nM, respectively, as described under "Materials and Methods." B, effect of Mg2+ on pre-steady-state kinetic parameters under multiple turnover conditions. Data derived from A were analyzed using the rate equation [P]t = A [1 – exp(K*t)] + L*t (also described under "Results)." A, active MPG concentration; K ({approx}kchem), burst constant; L/A (= kpd), microscopic product dissociation rate constant.

 
The curve fit provided the amplitude of the burst (A0), which is related to the active enzyme concentration involved in the glycosylase reaction. Although, we used similar amounts of protein for all reactions, we found that with increasing Mg2+ concentration, active MPG concentration became significantly limiting (Fig. 7B). The microscopic rate constant for product release (kpd) was evaluated from the relationship

Formula 4(Eq. 7)
where kss was determined from the slopes of the linear portions of the plots (Fig. 7). The kpd value (0.016 ± 0.001 min–1) for wild-type MPG is extremely low and is apparently rate-limiting in the MPG-mediated multistep reaction process. Slow turnover and product inhibition are the hallmarks of most of the DNA glycosylases (27). However, Mg2+ could significantly decrease the active protein in the reaction, but it did not show any significant alteration in product release rate. On the basis of these findings, it appears that Mg2+ directly binds to the MPG protein, modulates its substrate DNA binding ability, and thus affects its overall enzymatic activity.

Effect of Mg2+ on Steady-state Kinetics—As apparent from pre-steady-state kinetics that Mg2+ perhaps inhibits MPG by direct binding, we further investigated the inhibition mechanism by using steady-state kinetics. Increasing doses of Mg2+ increased Km but did not affect Vmax significantly (Table 2) suggesting again the abrogation of substrate binding of MPG by Mg2+.


View this table:
[in this window]
[in a new window]
  		TABLE 2
Effect of Mg2+ on steady-state kinetics

The reaction conditions are as described under "Materials and Methods."

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, we have demonstrated that Mg2+ can be modulatory or significantly inhibitory toward MPG activity depending on its concentration and substrate type. Zn2+ is a more potent inhibitor than Mg2+. Moreover, the cell nucleus contains a multitude of metals, which might affect MPG activity as well; however, from the BER viewpoint it was intriguing to elucidate the mechanism of Mg2+ inhibition, as apparently Mg2+ is an absolute cofactor for most if not all of the downstream BER enzymes. Unlike those enzymes, MPG and other DNA glycosylases do not require Mg2+ or any other cofactor for damage recognition and/or excision (26). Serendipitously, we discovered Mg2+ as a strong inhibitor of MPG while attempting to develop a concerted MPG-cleavage assay incorporating APE and the appropriate concentration of Mg2+. The latter is required for APE activity for the incision and strand break at AP sites, which are generated after MPG-mediated base excision. Mg2+ inhibited OGG1 AP-lyse activity without appreciable effects on its glycosylase activity (22). Here we found that Mg2+ can inhibit the glycosylase activity of a monofunctional DNA glycosylase, MPG.

Notably, K+ or Na+ was shown to be exchanged by Mg2+ in a specific binding pocket of heat shock IV protease from Haemophilus influenzae and also affected its enzymatic activity (32). The available MPG-substrate cocrystal structure had single Na+ in a defined pocket (33). Our results show that Mg2+ can inhibit the activity of MPG in a specific manner by abrogating its substrate binding. It could be achieved by disrupting the structural integrity of the catalytic pocket via an exchange with Na+ in the Na+-binding pocket and by conformational changes. However, it was not possible to perform the reciprocal experiment, such as testing whether Na+ added exogenously could rescue Mg2+-mediated inhibition of MPG activity, because 2.5 mM Mg2+ inhibited 80–90% of the MPG activity in the presence of 150 mM Na+, which suggests an apparently much higher affinity of MPG for Mg2+. Extremely high concentrations of Na+ are needed to replace Mg2+ in MPG. Further structural studies of MPG in the presence of Mg2+ are required to understand the exact mechanism of Mg2+-mediated MPG inhibition. It is notable that MPG in a substrate-bound complex cannot be crystallized in the presence of Mg2+. Lau et al. (33) could not observe Mg2+ in their substrate-bound cocrystal structure, although they used high concentrations of Mg2+ during the crystallization process; indeed, our results showed that Mg2+ inhibits substrate binding, which favors their observation. Burst analysis showed that Mg2+ decreases active enzyme concentration. Alternatively one can predict that other than these cations, the anion (Cl–) may have an effect on MPG activity. We have previously shown that MPG requires 150 mM NaCl for optimum activity. We used the same NaCl concentration in all our reactions for this study. That means that 150 mM Cl– was present in the assay mixture and that the further addition of 2.5 mM MgCl2 should increase Cl– by an additional 5 mM. Obviously this increase is negligible compared with the existing Cl– ions. More importantly, 2.5 mM MgCl2 inhibited 80–90% of the MPG activity (Fig. 1). If the anion (Cl–) is crucial for MPG inhibition, one could expect a similar inhibitory effect, because all cationic metals used in this study had the same anionic counterpart. However, our results demonstrate that this was not the case. The cation, not the anion, inhibited MPG activity. We also found that Mg2+ at a low concentration induces a slight stimulation in MPG activity, which could be due to a more optimal conformation of the substrate or to the protein itself. A similar salt-induced stimulation was observed with human endonuclease III enzyme (23). Because stimulation of MPG was only observed with Hx, not {epsilon}A, it is tempting to predict that DNA conformation near the damaged base or the adduct itself, rather than the MPG protein, was affected by low Mg2+ to cause stimulation in activity.

This is the first demonstration of an inhibitor for the enzyme MPG, a DNA glycosylase, with recognition and cleavage specificity for a wide variety of structurally diverse alkylated, deaminated, and etheno DNA adducts. Despite the importance of DNA repair glycosylases in the initiation of repair of mutagenic and toxic DNA base adducts inflicted by endogenous and exogenous sources, such as oxidative and nitrosative stress, replication errors, and cigarette smoke, there are not many well characterized inhibitors for this important class of BER enzymes. UGI (uracil-DNA glycosylase inhibitor), a viral protein, has been shown to inhibit uracil-DNA glycosylase, which excises uracils in DNA generated from replication misincorporations (34). Speina et al. (35) screened a battery of compounds including various base analogs and tryptophan pyrolysate (Trp-P-1), a mutagenic DNA intercalator heterocyclic amine found in cooked food, which targets a number of E. coli and human DNA glycosylases. The 2-thioxanthine was effective for E. coli Fpg protein and was shown to inhibit its excision activity whereas Trp-P-1 inhibits multiple DNA glycosylases such as AlkA, TagA, MPG, Nth, and Fpg by altering the DNA secondary structures (35).

In cellular systems, at physiologically relevant concentrations, Mg2+ is not genotoxic but is required to maintain genomic stability. In addition to its stabilizing effect on DNA and chromatin structure, Mg2+ is an essential cofactor in almost all enzymatic systems involved in major transactions of DNA, including replication and repair. To our knowledge, Mg2+-mediated enzyme inhibition is not common. For most cases Mg2+ acts as a cofactor. Furthermore, as an essential cofactor in nucleotide excision repair, BER, and mismatch repair pathways, Mg2+ is required for the removal of DNA damage generated by environmental mutagens, endogenous processes, and DNA replication. Intracellular Mg2+ concentrations are highly regulated, and Mg2+ acts as an intracellular regulator of cell cycle control and apoptosis (25). Mg2+ concentration varies in the nucleus (19). Therefore, intranuclear concentrations of Mg2+ may change in response to DNA damage. This phenomenon and its effect on repair pathways are yet to be elucidated.

The in vivo consequences of the variations in MPG activity with changes in Mg2+ concentration are not known. Although reliable figures are not available for changes in the amount of free Mg2+ in cells after exposure to DNA-damaging agents, MPG might, under some circumstances of low Mg2+ concentration, exhibit efficient binding of damaged DNA substrate and cleavage to generate AP sites. The delayed effect of increased Mg2+ concentration allows APE to bind AP sites without cleavage and protect cells from the deleterious effect of AP sites. AP sites are potentially more mutagenic and toxic than any of the MPG substrate-damaged bases. In fact deliberate MPG overexpression could not protect cells against alkylating agents; rather, it induced hypersensitivity and hypermutability because of increased production of AP sites and repair imbalances (36–38). Therefore, to ensure the complete and balanced repair, under some circumstances, Mg2+ concentration might increase after MPG action for activation of APE and, subsequently, other BER enzymes such as DNA polymerase beta and ligases. The high inhibitory Mg2+ concentration of MPG might also ensure the inhibition of repair imbalances. Thus the homeostasis and the changes in Mg2+ concentration might act as regulators for the BER pathway to ensure efficient and balanced repair of base damage and prevent genomic instability and error-free survival. How such modulation of Mg2+ might regulate BER in the cells in response to DNA damage remains to be investigated.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants CA 92306 (to R. R.) and CA 88004 (to J. A. T.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: Lombardi Comprehensive Cancer Center, LL level, S-122, 3800 Reservoir Rd., N.W., Georgetown University Medical Center, Washington, D. C. 20057. Tel.: 202-687-7390; Fax: 202-687-1068; E-mail: rr228{at}georgetown.edu.

2 The abbreviations used are: BER, base excision repair; AP, apurinic/apyrimidinic; MPG, N-methylpurine-DNA glycosylase; APE, apurinic/apyrimidinic endonuclease; OGG, 8-oxoguanine-DNA glycosylase; Hx, hypoxanthine; {epsilon}A, ethenoadenine; STO, single turnover. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Tadahide Izumi from LSU Stanley S. Scott Cancer Center for providing purified APE protein. We thank Dr. Sujata Choudhury of Georgetown University and Dr. Tapan Biswas of Harvard Medical School for critically reading the paper. We also thank Ellen McLaughlin and Karen Howenstein for expert editorial and secretarial help.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Lindahl, T. (1993) Nature 362, 709–715[CrossRef][Medline] [Order article via Infotrieve]
  2. Friedberg, E. C., Walker, G. C., and Siede, W. (1995) DNA Repair and Mutagenesis, p. 59, American Society for Microbiology, Washington, D.C.
  3. Larson, K., Sharn, J., Shenkar, R., and Strauss, B. (1985) Mutat. Res. 233, 211–218
  4. Breimer, L., and Lindahl, T. (1984) J. Biol. Chem. 259, 5543–5548[Abstract/Free Full Text]
  5. Hatchet, Z., Kow, Y. W., Purmal, A. A., Cunninghum, R. R. P., and Wallace, S. S. (1994) J. Biol. Chem. 269, 18814–18820[Abstract/Free Full Text]
  6. Singer, B., and Hang, B. (1997) Chem. Res. Toxicol. 10, 713–732[CrossRef][Medline] [Order article via Infotrieve]
  7. Roy, R., Biswas, T., Hazra, T. K., Roy, G., Grabowski, D. T., Izumi, T., Srinivasan, G., and Mitra, S. (1998) Biochemistry 37, 580–589[CrossRef][Medline] [Order article via Infotrieve]
  8. Roy, R., Brooks, C., and Mitra, S. (1994) Biochemistry 33, 15131–15140[CrossRef][Medline] [Order article via Infotrieve]
  9. Roy, R., Kennel, S. J., and Mitra, S. (1996) Carcinogenesis 17, 2177–2182[Abstract/Free Full Text]
  10. O' Connor, T. R. (1993) Nucleic Acids Res. 21, 5561–5569[Abstract/Free Full Text]
  11. Saparbaev, M., and Laval, J. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 5873–5877[Abstract/Free Full Text]
  12. Dosanjh, M. K., Roy, R., Mitra, S., and Singer, B. (1994) Biochemistry 33, 1624–1628[CrossRef][Medline] [Order article via Infotrieve]
  13. Labahn, J., Scharer, O. D., Long, A., Ezaz-Nikpay, K., Verdine, G. L., and Ellenberger, T. E. (1998) Cell 86, 321–329
  14. Lau, A. Y., Scharer, O. D., Samson, L., Verdine, G. L., and Ellenberger, T. (1998) Cell 95, 133–157
  15. Lindahl, T., Sedgwick, B., Sekiguchi, M., and Nakabeppu, Y. (1988) Annu. Rev. Biochem. 57, 133–157[CrossRef][Medline] [Order article via Infotrieve]
  16. McCarthy, T. V., Karran, P., and Lindahl, T. (1984) EMBO J. 3, 545–550[Medline] [Order article via Infotrieve]
  17. Bjelland, S., Birkeland, N. K., Benneche, T., Volden, G., and Seeberg, E. (1994) J. Biol. Chem. 269, 30489–30495[Abstract/Free Full Text]
  18. Bjoras, M., Klungland, A., Johansen, R. F., and Seeberg, E. (1995) Biochemistry 34, 4577–4582[CrossRef][Medline] [Order article via Infotrieve]
  19. Kroeger, H., and Trosch, W. (1974) J. Cell. Physiol. 83, 19–25[CrossRef][Medline] [Order article via Infotrieve]
  20. Lukacs, G. L., Zs-Nagy, I., Steiber, J., Gyori, F., and Balazs, G. (1996) Scanning Microsc. 10, 1191–1200[Medline] [Order article via Infotrieve]
  21. Li-Smerin, Y., Levitan, E. S., and Johnson, J. W. (2001) J. Physiol. 533, 729–743[Abstract/Free Full Text]
  22. Morland, I., Luna, L., Gustad, E., Seeberg, E., and Bjørås, M. (2005) DNA Repair 4, 381–387[Medline] [Order article via Infotrieve]
  23. Eide, L., Luna, L., Gustad, E., Henderson, P. T., Essigmann, J. M., Demple, B., and Seeberg, E. (2001) Biochemistry 40, 6653–6659[CrossRef][Medline] [Order article via Infotrieve]
  24. Waters, T. R., Gallinari, P., Jiricny, J., and Swann, P. F. (1999) J. Biol. Chem. 274, 67–74[Abstract/Free Full Text]
  25. Bronzetti, G., Croce, C. D., and Davini, T. (1995) J. Environ. Pathol. Toxicol. Oncol. 14, 197–204[Medline] [Order article via Infotrieve]
  26. Izumi, T., Wiederhold, L. R., Roy, G., Roy, R., Jaiswal, A., Bhakat, K. K., Mitra, S., and Hazra, T. K. (2003) Toxicology 15, 43–65[CrossRef]
  27. Roy, R., Biswas, T., Lee, J. C., and Mitra, S. (2000) J. Biol. Chem. 275, 4278–4282[Abstract/Free Full Text]
  28. Liu, X., and Roy, R. (2002) J. Mol. Biol. 321, 265–276[CrossRef][Medline] [Order article via Infotrieve]
  29. Porello, S. L., Leves, A. E., and David, S. S. (1998) Biochemistry 37, 14756–14764[CrossRef][Medline] [Order article via Infotrieve]
  30. Abner, C. W., Lau, A. Y., Ellenberger, T., and Bloom, L. B. (2001) J. Biol. Chem. 276, 13379–13387[Abstract/Free Full Text]
  31. Strauss, P. R., Beard, W. A., Patterson, T. A., and Wilson, S. H. (1997) J. Biol. Chem. 272, 1302–1307[Abstract/Free Full Text]
  32. Sousa, M. C., and Mckay, D. B. (2001) Acta Crystallogr. Sect. D 57, 1950–1954[CrossRef][Medline] [Order article via Infotrieve]
  33. Lau, A. Y., Wyatt, M. D., Glassner, B. J., Samson, L. D., and Ellenberger, T. (2000) Proc. Natl. Acad. Sci. U. S. A. 5, 13573–13578
  34. Bennett, S. E., and Mosbaugh, D. W. (1992) J. Biol. Chem. 267, 22512–22521[Abstract/Free Full Text]
  35. Speina, E., Ciesla, J. M., Graziewicz, M., Laval, J., Kazimierczuk, Z., and Tudek, B. (2005) Acta Biochim. Pol. 52, 167–178[Medline] [Order article via Infotrieve]
  36. Sobol, R. W., Kartalou, M., Almeida, K. H., Joyce, D. F., Engelward, B. P., Horton, J. K., Prasad, R., Samson, L. D., and Wilson, S. H. (2003) J. Biol. Chem. 278, 39951–39959[Abstract/Free Full Text]
  37. Posnick, L. M., and Samson, L. D. (1999) J. Bacteriol. 181, 6763–6771[Abstract/Free Full Text]
  38. Rinne, M. L., He, Y., Pachkowski, B. F., Nakamura, J., and Kelley, M. R. (2005) Nucleic Acids Res. 33, 2859–2867[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
S. Adhikari, A. Uren, and R. Roy
Dipole-Dipole Interaction Stabilizes the Transition State of Apurinic/Apyrimidinic Endonuclease Abasic Site Interaction
J. Biol. Chem., January 18, 2008; 283(3): 1334 - 1339.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
S. Adhikari, A. Uren, and R. Roy
N-terminal Extension of N-Methylpurine DNA Glycosylase Is Required for Turnover in Hypoxanthine Excision Reaction
J. Biol. Chem., October 12, 2007; 282(41): 30078 - 30084.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
281/40/29525    most recent
M602673200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Adhikari, S.
Right arrow Articles by Roy, R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Adhikari, S.
Right arrow Articles by Roy, R.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2006 by the American Society for Biochemistry and Molecular Biology.