- Research Article
- Open Access
Molecular dynamics simulation of the opposite-base preference and interactions in the active site of formamidopyrimidine-DNA glycosylase
© The Author(s). 2017
- Received: 12 November 2016
- Accepted: 20 April 2017
- Published: 8 May 2017
Formamidopyrimidine-DNA glycosylase (Fpg) removes abundant pre-mutagenic 8-oxoguanine (oxoG) bases from DNA through nucleophilic attack of its N-terminal proline at C1′ of the damaged nucleotide. Since oxoG efficiently pairs with both C and A, Fpg must excise oxoG from pairs with C but not with A, otherwise a mutation occurs. The crystal structures of several Fpg–DNA complexes have been solved, yet no structure with A opposite the lesion is available.
Here we use molecular dynamic simulation to model interactions in the pre-catalytic complex of Lactococcus lactis Fpg with DNA containing oxoG opposite C or A, the latter in either syn or anti conformation. The catalytic dyad, Pro1–Glu2, was modeled in all four possible protonation states. Only one transition was observed in the experimental reaction rate pH dependence plots, and Glu2 kept the same set of interactions regardless of its protonation state, suggesting that it does not limit the reaction rate. The adenine base opposite oxoG was highly distorting for the adjacent nucleotides: in the more stable syn models it formed non-canonical bonds with out-of-register nucleotides in both the damaged and the complementary strand, whereas in the anti models the adenine either formed non-canonical bonds or was expelled into the major groove. The side chains of Arg109 and Phe111 that Fpg inserts into DNA to maintain its kinked conformation tended to withdraw from their positions if A was opposite to the lesion. The region showing the largest differences in the dynamics between oxoG:C and oxoG:A substrates was unexpectedly remote from the active site, located near the linker joining the two domains of Fpg. This region was also highly conserved among 124 analyzed Fpg sequences. Three sites trapping water molecules through multiple bonds were identified on the protein–DNA interface, apparently helping to maintain enzyme-induced DNA distortion and participating in oxoG recognition.
Overall, the discrimination against A opposite to the lesion seems to be due to incorrect DNA distortion around the lesion-containing base pair and, possibly, to gross movement of protein domains connected by the linker.
- DNA glycosylase
- Molecular dynamics
- Opposite-base specificity
- Reaction mechanism
Formamidopyrimidine-DNA glycosylase (Fpg or MutM) is a bacterial DNA repair enzyme that removes several abundant oxidized bases from DNA. The principal substrate bases of Fpg are 8-oxoguanine (oxoG), 2,6-diamino-4-oxo-5-formamidopyrimidine (fapyG) and 2,4-diamino-5-formamidopyrimidine (fapyA) [1, 2] but the enzyme also can recognize several dozens of other damaged purines and pyrimidines [3–10]. By excision of a damaged base, Fpg initiates base excision repair (BER), which engages AP endonucleases, a DNA polymerase and a DNA ligase to restore the integrity of DNA [11, 12].
The activity of Fpg towards oxoG has attracted much attention due to abundance and biological importance of this lesion, induced in DNA by oxidative metabolism byproducts, oxidative stress, and ionizing radiation . Steric and electrostatic repulsion between the substituent at C8 and the sugar–phosphate atoms effectively pushes oxoG towards the syn conformation, in which oxoG forms a Hoogsteen pair with A [14, 15]. Misincorporation of A by DNA polymerases, in the absence of repair, leads to a G → T transversion after the second round of replication.
Systems for repair of oxoG have been found in all cellular organisms. The tendency of oxoG to form pairs with both C and A presents a challenge to its repair: both oxoG:C and oxoG:A pairs must be converted into G:C pairs. This requirement is not trivial since a simple removal of oxoG from an oxoG:A mispair would generate a G → T transversion after the repair. This problem is circumvented by repair of oxoG:A pairs in two sequential rounds of BER . The non-damaged (but inappropriately incorporated) A is removed first and replaced with C, and the resulting oxoG:C pair is then repaired through the excision of oxoG. In E. coli, the mutagenic potential of oxoG is counteracted by three enzymes, Fpg, MutT, and MutY, collectively known as a “GO system”. Fpg excises oxoG from oxoG:C pairs but has little activity towards oxoG:A substrates to prevent G → T transversions [1, 17]. Another DNA glycosylase, MutY, specifically excises A from A:oxoG mispairs. If G in DNA is oxidized to oxoG, it will inevitably be paired with C and will be removed by Fpg. If, on the other hand, A is incorporated during replication opposite an unrepaired oxoG, the resulting oxoG:A mispair will be a substrate for MutY but not for Fpg. The repair synthesis then has a chance to incorporate C opposite oxoG .
Mean r.m.s.d. values of the models and their standard deviations (Å) over the last 8 ns of the runs
1.50 ± 0.06
1.58 ± 0.07
1.69 ± 0.07
1.51 ± 0.05
1.75 ± 0.05
1.43 ± 0.06
1.66 ± 0.06
1.45 ± 0.06
1.68 ± 0.06
1.53 ± 0.06
1.41 ± 0.06
1.74 ± 0.06
1.37 ± 0.10
1.55 ± 0.08
1.60 ± 0.13
1.58 ± 0.11
1.89 ± 0.06
1.57 ± 0.08
2.16 ± 0.06
1.86 ± 0.07
1.66 ± 0.12
1.61 ± 0.06
1.44 ± 0.10
1.92 ± 0.13
Molecular dynamics simulations (10 ns) were performed using the BioPASED molecular dynamics modeling software  using the AMBER ff99 force field with BioPASED modifications and EEF1 analytical implicit solvent model , with an integration time step of 1 fs. The system was gradually heated from 10 K to 300 K during 50 ps and equilibrated at this temperature (the heating time was 150, 200, and 250 ps in the repeat runs of the PRO-GLH models). A classic molecular dynamics trajectory was generated in the NVT ensemble with harmonic restraints of 0.001 kcal/A2 for the protein and water atoms, 0.25 kcal/A2 for the atoms of the terminal nucleotides, and 0.0025 kcal/A2 for the rest of the DNA atoms. Coordinates of each atom of the system were saved each 2 ps, thus producing a trajectory size of 5000 snapshots. The trajectories were analyzed using MDTRA , a part of the BISON package . Trajectories were compared using moving MWZ method  with bins of 50 snapshots. Statistically significant differences in parameters between different models were estimated using F-test, with false discovery rate method (Benjamini–Hochberg procedure) employed to correct for multiple comparisons . Hydrogen bonds were searched using MDTRA . Structures were visualized and rendered using VMD , RasMol  and PyMOL (Schrödinger, Portland, OR).
pH dependence of Fpg activity
Eco-Fpg was purified as described . Oligonucleotides were synthesized in-house from commercially available phosphoramidites (Glen Research, Sterling, VA). An oligonucleotide 5′-CTCTCCCTTCXCTCCTTTCCTCT-3′ (X = oxoG) was 32P-labeled using polynucleotide kinase (SibEnzyme, Novosibirsk, Russia) and γ[32P]ATP (PerkinElmer, Waltham, MA) following the manufacturer’s protocol and annealed to a complementary strand placing C or A opposite oxoG. The reactions (20 μl) included 25 mM sodium phosphate buffer (H3PO4/NaH2PO4, NaH2PO4/Na2HPO4, and Na2HPO4/Na3PO4 conjugate pairs spanning the range of pH 4.0–9.0), 100 nM duplex oligonucleotide substrate, and either 2 nM (steady-state experiments) or 500 nM Fpg (single-turnover experiments). The reaction was allowed to proceed for 1 min either at 37 °C with 2 nM Fpg or at 0 °C with 500 nM Fpg, and was stopped by adding 20 μl of formamide/EDTA gel loading dye and heating at 95° for 3 min. The products were separated by electrophoresis in 20% polyacrylamide gel/8 M urea and quantified by phosphorimaging using a Molecular Imager FX system (Bio-Rad Laboratories, Hercules, CA). Three independent experiments have been performed. Calculation of pK a for Pro1 and Glu2 were done using PROPKA v3.1 . Circular dichroism spectra were recorded on a JASCO J-600 CD spectrometer (JASCO Analytical Instruments, Tokyo, Japan) at 25 °C in 30 mM Na phosphate with a 1-nm step.
A taxonomically balanced sample of 124 bacterial Fpg sequences (limited to two sequences per taxonomic family) was constructed by protein BLAST search  in the NCBI non-redundant protein sequences database using Eco-Fpg as a query, filtered for conservation of the N-terminal Pro–Glu dipeptide and C-terminal zinc finger, and clipped from the absolutely conserved N-terminal Pro to the absolutely conserved Gln after the fourth Cys of the zinc finger as described [50, 51]. Alignment of multiple sequences and neighbor-joining tree construction was performed using Clustal Omega . Hierarchical analysis of conservation of physicochemical properties in the alignments was done using AMAS  with 5% atypical residues allowed; the results are reported as conservation numbers (C n ).
General model considerations
Selection of the starting structure
Currently, the Protein Data Bank holds 56 released X-ray structures of Fpg, belonging to four bacterial species and sampling several points along the reaction coordinate [18–33]. Our selection of the starting structure for MD was guided by the following considerations. First, it should contain DNA with the damaged base still in place, residing in the enzyme’s active site. Second, minimal deviation from the wild-type enzyme recognizing oxoG should be present. Third, the structure should have good resolution (<2.0 Å), with as few as possible residues missing.
Based on these considerations, we have chosen 1XC8, the 1.95-Å structure of wild-type Lla-Fpg bound to DNA containing a non-cleavable carbacyclic fapyG analog (carba-fapyG ) as a starting model (Fig. 1a). In carba-fapyG, a methylene group substitutes for O4′, and the damaged base, fapyG, is different from oxoG only by the absence of a bond between N9 and C8 of the purine heterocycle O4′ . The Lla-Fpg is nearly identical to Eco-Fpg in its selectivity for C vs A as the opposite base .
OxoG glycosidic angle
Besides 1XC8, the structures of Fpg bound to DNA with an extrahelical damaged base include Lla-Fpg bound to DNA containing carbacyclic N5-benzyl-fapyG (3C58 ), carbacyclic oxoG (4CIS ) or 5-hydroxy-5-methylhydantoin (2XZF, 2XZU ) and Bst-Fpg bound to DNA containing oxoG (1R2Y) or 5,6-dihydrouracil (1R2Z) . With the exception of 1R2Y and 4CIS, the damaged bases in the structures are quite different from oxoG. In the 1R2Y structure, oxoG is present in DNA, and the cleavage is prevented by changing the absolutely conserved catalytic Glu2 residue into Gln . In this structure, oxoG is often stated to be in the syn conformation in the active site, yet its χ angle (101°) is actually in the anti domain (namely, in its border range, so-called “high syn”) (Fig. 1d). On the contrary, oxoG opposite C in B-DNA is usually stated to be anti as it forms regular Watson–Crick bonds [55, 56], yet its χ angle in the crystal structure (–55°, ) is actually in the syn domain. Carba-fapyG in 1XC8 is in the high anti range (χ = –64°), and only in 4CIS, carba-oxoG is unambiguously syn (χ = 27°, Fig. 1d). Moreover, oxoG in 4CIS does not form the same set of hydrogen bonds with the active site as in 1R2Y. The possibility of conformation artifacts induced by E2Q mutation has been amply discussed in the literature [23, 38, 57–61]. Therefore, we have chosen 1XC8, which straddles the syn/anti border (Fig. 1d) as our starting model and allowed the conformation to drift into the most preferable χ range during MD.
Opposite base glycosidic angle
While the oxoG:A pair in B-DNA exists as oxoG(syn):A(anti), this does not mean that the same conformations will be observed in the complex with Fpg, since the hydrogen bonds within the mispair are lost upon oxoG eversion, and the conformation of the nucleotides is governed largely by their interaction with the protein residues. The most relevant example is given by another oxoG mispair, oxoG:G, which adopts the conformation oxoG(syn):G(anti) in the B-DNA duplex  but the G opposite to the lesion flips into syn and forms two strong hydrogen bond with an Arg residue when this duplex is bound to Bst-Fpg . Therefore, in addition to the oxoG:C pair, we have constructed two series of models with oxoG:A, with A in either anti or syn conformation to fully explore the range of possible dynamics of Fpg–oxoG:A complex.
Although MD in explicit solvent is common nowadays, recent advances in implicit solvent models revived the popularity of this alternative [63–65]. The major advantages of implicit solvent over the explicit one are speed, better estimates of solvation and folding energy, wider coverage of conformational space and more accurate account for pH and residue ionization. The latest versions of Poisson–Boltzmann, generalized Born and hybrid implicit/explicit models are comparable with explicit solvent-based calculations with respect to agreement with experimental free energy data [63–65]. Although the acceleration of conformation sampling afforded by implicit solvent strongly depends on the modeled system, direct comparisons show a 7–10-fold increase for a system with several conformational transitions . Since our primary interest was to sample a wide range of conformations available for the Fpg–substrate complexes, we have chosen a hybrid model combining an implicit solvent with explicit strongly bound water molecules; such approaches retain the advantages of implicit methods but significantly improve quality of protein–DNA interface models .
Protonation state of the catalytic dyad
The ionizable groups of Pro1 and Glu2 directly participate in the enzymatic reaction. Mechanistically, the nucleophilic attack by Pro1 at C1′[oxoG] requires N[Pro1] to carry a lone electron pair (Fig. 1b). On the other hand, opening of the deoxyribose ring involves protonation of its O4′, which is near Oε2 of Glu2 (Fig. 1b); quantum mechanics/molecular mechanics (QM/MM) simulations show that O4′ protonation provides a low-barrier path to glycosidic bond cleavage by Fpg and its eukaryotic functional analog, OGG1 [33, 36, 68]. From several structures Fpg–DNA complexes, it has been suggested that the proton is shuttled from N[Pro1] to Oε2[Glu2], perhaps through a network of crystallographic water molecules present in the active site [19, 26]; this possibility was also favored by QM/MM analysis . However, no attempt to estimate pK a of Pro1 and Glu2 has been reported in the literature. It is possible that a mixture of Pro1/Glu2 ionization states exists in the active centers of different Fpg molecules at physiological pH; although only one of them (PRN-GLH) is permissive for the reaction chemistry, the path to this state may go through other states. Therefore, we have performed MD of the full system for four ionization states of the Pro1–Glu2 catalytic dyad: PRN-GLH, PRN-GLU, PRO-GLH, and PRO-GLU (see Methods).
Overall model characterization
The root mean square deviation (r.m.s.d.) with respect to the backbone of the starting structure of the complex was calculated every 2 ps. R.m.s.d. values of all the models increased rapidly during the first 500 ps of the dynamics and stabilized at approximately 1.6 Å (see Table 1 and Additional file 1: Fig. S1). Overall r.m.s.d. of all models was similar; however, in the active center (defined as all protein residues with at least one atom within 4 Å of oxoG nucleotide or the opposite C/A nucleotide, plus three nucleotide pairs centered on the oxoG), the r.m.s.d. of the C models was significantly lower than in the A models (p < 10–4). The DNA backbone displayed higher mobility than the protein backbone: average r.m.s.d. of the DNA residues was greater by 0.54–0.98 Å depending on the model. The overall complex conformation was stable along the whole trajectory with a radius of gyration ~20 Å for each model (20.17 ± 0.04 to 20.30 ± 0.05 Å). The angle of DNA kink was also stable (55° ± 2° to 63° ± 2°, depending on the model).
To test the consistence of results between independent runs, we have selected three models (PRO-GLH-C, PRO-GLH-Aa, and PRO-GLH-As) and performed three additional simulations with each one, to the total of nine additional simulations, using different heating times (150, 200, and 250 ps) to provide different conditions for the start of the production run. Then the resulting four trajectories for each model (one original and three new) were compared. The r.m.s.d. values of individual runs were similar (1.0–1.5 Å over the last 8 ns, Additional file 1: Fig. S1). The inter-run r.m.s.d. were expectedly higher (1.9–2.2 Å, Additional file 1: Fig. S1) but still did not show significant divergence of the models. Stable hydrogen bonds, including model-specific ones, were well consistent across the four runs (Additional file 1: Fig. S5B); the 90% cut-off of the mean occurrence identified as stable all Watson–Crick bonds and 79% of the main-chain bonds observed in the 1XC8 crystal structure.
Pro–Glu catalytic dyad
Arrangement of the reacting groups in the models
Key distances and angles around the reacting C1′ atom of oxoG
Distance N[Pro1]… C1′[oxoG0], Å
Distance Oε2[Glu2]… O4′[oxoG0], Å
Angle C1′[oxoG0]… N[Pro1]… Cδ[Pro1], degrees
Angle O4′[oxoG0]… C1′[oxoG0]…N[Pro1], degrees
# of snapshots with optimal geometry
3.73 (3.41–4.04) a
1077 (788) b
Defining the “optimal geometry” as N[Pro1]…C1′[oxoG] distance < 4 Å, Oε2[Glu2]…O4′[oxoG0] distance < 4.5 Å, and C1′[oxoG0]…N[Pro1]…Cδ[Pro1] and O4′[oxoG]…C1′[oxoG]…N[Pro1] within 20° of the ideal values, we have sampled the population of the zone with all four parameters optimal (Table 2). All A(anti) models showed the optimal geometry very rarely. For C and A(syn) models, PRO-GLH was most populated, followed by PRN-GLH. Interestingly, PRN models were more selective towards C vs A(syn). Other sensible definitions of “optimal” Pro1 and Glu2 distances (e. g., the lowest quartile of the respective distance population), also showed C models spending more time in the optimal conformation than A models. The preference of C models for the optimal geometry was also evident in the repeat runs of the PRO-GLH models (Table 2).
pKa estimate of the catalytic dyad residues
To get an independent estimate of the protonation state of Pro1 and Glu2, we have used PROPKA, an empirical algorithm based on the spatial proximity of charged residues . In addition to our starting structure, we have considered several other PDB structures of Fpg from different species (Additional file 2: Table S1). In all cases, pK a of Pro1 was notably lowered (by 0.35–2.99 units) compared with the reference pK a of N-terminal Pro, while pK a of Glu2 was considerably higher (by 1.54–3.64 units) than the reference pK a of the internal Glu side chain. Similar pK a changes were reported for phage T4 endonuclease V, another DNA glycosylase employing the N-terminal amino group and a Glu carboxyl as a catalytic dyad . Interestingly, structures of free Fpg and Fpg bound to undamaged DNA with the sampled base still intrahelical displayed more acidic pK a for Glu2, suggesting that this group may be specifically activated upon eversion of the damaged nucleotide. Although PROPKA considers the influence of nucleic acid ligands on amino acid ionization potential only approximately, it is nevertheless clear that in Fpg, Pro1 is considerably more acidic, and Glu2, more basic than expected.
pH profile of Fpg activity
Since Pro1 has to be deprotonated for the reaction, the rising activity vs pH plot with a single inflection may be explained by this deprotonation and suggests that the equilibrium ionization state of Glu2 is not rate-limiting. As discussed below, Glu2 may be conveniently protonated by a water molecule trapped in the active site.
Other interactions of the catalytic dyad
Pro1 did not form stable interactions within the active site in the models, consistent with the available structural information on the pre-catalytic Fpg complexes. In contrast, in 1XC8 and other known structures of Fpg, Glu2 accepts two hydrogen bonds from the amides of Ile172 and Tyr173 (or their counterparts in other Fpgs). These bonds were stable in all our simulations independently of the protonation state of Glu2. Several available structures (1K82, 1L1T, 1L1Z [19, 20]) strongly suggest that these two bonds hold the carboxylic group of Glu2 in a position suitable for interaction with a nearby water molecule, and, later in the reaction, with O4′ of the damaged nucleotide that becomes a hydroxyl after base excision and sugar ring opening. Importantly, a water molecule (see the section “Water molecules in the Fpg–DNA complex”) was the only stably interacting group other than Ile172 and Tyr173, and this interaction was not affected by the protonation state of Glu2.
Fpg interactions with oxoG and the opposite base
Stability of oxoG in the base-binding pocket
In the high syn 1R2Y model of Bst-Fpg, four consecutive main chain amide nitrogens belonging to the Thr220–Tyr224 loop form a crown around O6 of oxoG, positioned at distances and angles suitable for hydrogen bond formation (Fig. 5c). Surprisingly, even though the oxoG base is rotated nearly 180° from its position in 1R2Y, the same set of contacts is maintained by the homologous loop Ser218–Tyr222 of Lla-Fpg (Fig. 5b, d). This observation agrees well with the literature data on simulation of Bst-Fpg with oxoG forced into the anti conformation  and with the same pattern of contacts to O6 in the 1XC8 structure of Lla-Fpg . Notably, the “distinguishing” bond between the main chain carbonyl of Ser220 and pyrrolic N7 of oxoG, seen in Bst-Fpg 1R2Y structure but absent from Lla-Fpg 1XC8, was not observed in our simulations.
Interactions and dynamics of the opposite base
In all reported structures of Fpg bound to DNA with the fully everted damaged nucleotide, specific recognition of C opposite to the lesion is governed by two hydrogen bonds from Arg109 after its insertion into the DNA void: Nε[Arg109]–O2[C(0)] and Nη2[Arg109]–N3[C(0)]. If G substitutes for C, Nε and Nη2 of the inserted Arg form slightly suboptimal bonds with N7 and O6, respectively, of the G base in the syn orientation, whereas T in place of C retains a bond with Nε[Arg109] but experiences a clash between two hydrogen bond donors, N3[T] and Nη2[Arg109] .
Despite the geometry of the As models is less disturbed in comparison with the Aa models, oxoG:A experimentally is still a poor substrate for Fpg. A comparison of equilibrium binding, steady-state and pre-steady state kinetics of the E. coli enzyme [4, 76] suggests that the Fpg–oxoG:A complex forms quickly but then is much slower to proceed to the catalytically competent conformation than the Fpg–oxoG:C complex, leading to a ~15-fold higher apparent K M (14 nM for oxoG:C vs 190 nM for oxoG:A ; 8.7 nM for oxoG:C vs 150 nM for oxoG:A ). At the same time, the observed effect of A on k cat is minor [4, 76]. Since K M reflects the population of the last pre-catalytic state, it is tempting to suggest that only oxoG:A(syn) may be capable of attaining the catalytically competent conformation, thus necessitating the anti–syn transition in the course of the productive reaction with oxoG:A and partly explaining its slow progression with this substrate where A is initially anti [14, 15].
Aromatic wedge-induced distortion
Specific distant interactions in Fpg–DNA complexes
Model-specific hydrogen bonds
In order to select out inter- and intramolecular interactions specific for oxoG:C, we have searched for hydrogen bonds that existed (i. e., had an energy > 1.2 kcal/mol) in > 1% of the snapshots. Around 600 such hydrogen bonds were found in each model. In all models, less than 50% of the found bonds existed for more than 90% of the snapshots, and less than 25% of the found bonds existed in less than 25% of the snapshots (Additional file 1: Fig. S5A, B). The former class may be considered to represent stable, functionally important hydrogen bonds, whereas the latter one is most likely due to conformational fluctuations. Therefore, all detected hydrogen bonds were first analyzed with respect to their occurrence in these categories (≤90% vs > 90% and ≤ 25% vs > 25%). Pearson’s mean square contingency coefficients (φ) for pairwise comparison between different models showed no significant contribution of protonation state or substrate into the overall distribution of bonds in the high- and low-stability categories.
Fpg regions with C-specific bonds outside the active site
The most prominent opposite-base-specific feature in the protein structure was a cluster at the start of the C-terminal domain immediately next to the interdomain linker (residues Glu134–Phe140). Most of the amino acid residues there engaged in multiple bonds, forming a network, which existed in two stable configurations. In one, which was statistically significantly more often observed in A models, Thr136 formed two bonds with Glu134, one with Asp139, and one with Phe140 (N[Thr136]–O[Glu134], Oγ[Thr136]–O[Glu134], N[Asp139]–Oγ[Thr136], N[Phe140]–Oγ[Thr136]), and a N[Tyr137]–Oε1/Oε2[Glu138] was present. A completely different set of bonds was characteristic of C models (Oγ[Thr136]–Oε1/Oε2[Glu138], N[Asp139]–Oε1/Oε2[Glu138], N[Phe140]–O[Thr136]). As a result, the Glu134–Phe140 loop adopted different conformations in the C and A models (Fig. 8). Importantly, the conservation of Fpg sequence is quite high in this region (Additional file 1: Fig. S7), underlying its functional significance despite its position well away from the active site.
The only other region of known functional importance where consistently different bonds existed was the β-hairpin zinc finger, a structural motif in Fpg involved in major groove tracking and lesion recognition  (Fig. 8). Several C/A-specific hydrogen bonds were scattered in the β-sandwich domain around the C-terminal end of the long α-helix αA, which carries the catalytic Pro–Glu dyad at the other end (Fig. 8). The functional significance of this region is not clear; most C/A specific residues here are located in surface loops and are not conserved (Additional file 1: Fig. S7).
Fpg regions with A(syn) and A(anti)-specific bonds outside the active site
In addition, we have searched for bonds specific for A models in different (anti or syn) conformations of the orphaned A (Additional file 1: Fig. S6C). Most of the differences were encountered between protein and DNA, and within DNA, reflecting the conformational changes inflicted by introducing the disfavored A base. The protein residues affected by the conformation of the orphaned nucleotide showed little overlap with the C/A-specific interactions. The most prominent Aa/As-specific contacts were formed by Tyr29/Arg31 and His91/Lys110, two elements that coordinate the phosphates flanking the orphaned A, and Lys155 that contacts DNA a few nucleotides away from the lesion but is important for Fpg activity . The Glu134–Phe140 C/A-specific linker-adjacent region showed no significant difference between Aa and As models.
Water molecules in the Fpg–DNA complex
Dynamics of structural water in Fpg
The structure of Lla-Fpg–DNA complex, 1XC8, contains the total of 397 water molecules. However, only 22 of those reside at the protein–DNA interface and only seven are buried at it (i. e., have < 10% solvent exposure). The structures of Fpg–DNA complexes from different species, as well as the structures of the homolog of Fpg, Eco-Nei, in a complex with DNA , suggest that several water molecules form a tight network of bonds in the enzyme’s active site that may serve to shuttle protons during the concerted cleavage of three bonds catalyzed by Fpg.
We have explicitly modeled the seven water molecules buried at the protein–DNA interface and determined whether they form hydrogen bonds with two or three Fpg or DNA donors or acceptors at the same snapshot. Such water bridges, if persistent, may indicate an important role of water in structure maintenance or reaction mechanism. There were no significant differences between models or between groups of models in the number of water bridges. One particular pair of acceptors, Oε1/Oε2[Glu76] and O8[oxoG0], was consistently found bridged by two water molecules in 8 of 12 models. In several models, such multiple water-mediated connections existed between the non-bridging phosphate oxygens of oxoG0 and T+1 and between O2P[oxoG0] and Nη1[Arg109] but their occurrence was much less common. No donor/acceptor triplets were connected by multiple bridges.
Possible role of structural water molecules in Fpg mechanism
The identified stable triplets are suggestive of an important role of water in the mechanism of action of Fpg. The water molecule trapped between Oε1/Oε2[Glu2], Oε1[Glu5] and N2[oxoG0] is located at the position suitable for proton transfer to Glu2, required for the protonation of O4′ of oxoG nucleotide; water-mediated proton transfer to Glu2 was earlier proposed on structural reasons [19, 26]. In the GLH models, this water stably donated a bond only to the unprotonated Oε1[Glu2] (73% bond occurrence averaged over all GLH models, compare with 5% for the bond to Oε2) but when Glu2 was charged, Oε2 accepted this hydrogen with a higher frequency (64% and 32% bonds to Oε1 and Oε2, respectively) (Fig. 9b). In the GLH-Aa models, the protonated Oε2 showed a tendency to donate a hydrogen bond to the water molecule rather than accept one (23% in PRN-GLH-Aa, 58% in PRO-GLH-Aa, 0–7% in other GLH models), consistent with poor substrate properties of anti A. It should be mentioned that in QM/MM analysis of fapyG excision by Fpg this water molecule was inhibitory to the reaction, preventing the protonation of O4′ by neutral Glu2  and should be displaced from its crystallographic position after donating a proton to Oε2.
The water molecule bridging the protein residues with the phosphates of T+1 and oxoG0 may be important for distorting the DNA duplex. Notably, the distance between the phosphorus atoms P[T+1] and P[oxoG0] is significantly shorter than in the regular B-DNA in all models. This pinching of the phosphates around T+1, together with wedging of Phe111 and insertion of Met75 and Arg109, assists in kinking the DNA axis by ~60° and eversion of the damaged nucleotide.
The tightly coordinated two-water bridge to O8[oxoG0] presents an intriguing conundrum. On one hand, water-mediated recognition of this unique carbonyl would be an attractive mechanism of direct oxoG sensing in the active site pocket. On the other hand, Glu76, which in our models participates in the water coordination, is present only in a small branch of the Fpg family tree consisting of two closely phylogenetically related groups, Bacilli and Mollicutes (which include L. lactis and G. stearothermophilus), while in all other Fpg sequences this position is occupied by Ser/Thr with very rare exceptions (Additional file 1: Fig. S9, Additional file 3). In the structure of Bst-Fpg, the presence of Glu76 stabilizes the everted oxoG in the high syn conformation through hydrogen bonding with N2[oxoG], whereas its in silico replacement with Ser reverts the preferred χ angle to the anti domain . In Eco-Fpg, Ser74 and Lys217 correspond to Glu76 and Arg220 of Lla-Fpg, and Lys217 forms a direct hydrogen bond with O8[oxoG] . Obviously, there are several ways by which Fpg enzymes can employ the residues at these positions to the effect of recognizing the exocyclic oxygen at C8 either directly or indirectly.
The opposite-base selectivity of Fpg and some other DNA glycosylases (eukaryotic OGG1 and TDG, bacterial MutY and Mug, etc.) is extremely important for the prevention of mutations in the course of DNA repair. Analysis of the causes of this selectivity is complicated due to the paucity of structures of DNA glycosylases bound to substrates with disfavored opposite bases. In the case of Fpg, no structure containing oxoG:A, the biologically relevant disfavored mispair, is available. Our modeling effort was mostly undertaken to analyze possible structural features of such a complex and reveal those that could explain low activity of Fpg on oxoG:A substrates.
As our models suggest, introduction of A opposite to oxoG indeed distorts the protein–DNA interface within ±2 base pairs around the lesion site, outside of which DNA exists as a normal duplex. Arg-109 and Phe-111, two residues that Fpg inserts into DNA to sharply kink it and maintain oxoG everted from the base stack, tended to withdraw if A was opposite to the lesion, indicating that the pre-catalytic complex of Fpg with oxoG:A is inherently unstable. Interestingly, although the oxoG:A mispair adopts oxoG(syn):A(anti) conformation in free DNA, our models showed that upon Fpg binding and oxoG eversion, the orphaned A is more stable as a syn conformer, engaged both in hydrogen bonding with Arg-109 and in base stacking. We speculate that Fpg binding to oxoG(syn):A(anti) may be energetically disadvantageous and require rotation of the A base around the glycosidic bond for rare events of base excision; direct test of this hypothesis would require solving the structure of Fpg–DNA(oxoG:A) complex or stopped-flow kinetics with a series of fluorescent reporter bases incorporated next to A, in which case the anti–syn transition may be expected to be observed in the fluorescent traces.
Analysis of model-specific hydrogen bonds unexpectedly revealed a cluster of highly conserved residues next to the interdomain linker of Fpg, which adopted alternative conformations when C or A was in the opposite strand. This cluster is remote from what is usually considered the active site of Fpg; however, it is packed against a helix–two-turn–helix motif that is present in all Fpg family members and partly forms the DNA-binding groove. Of note, it has been shown that in Nei, a homolog of Fpg specific for oxidized pyrimidines, a structural rearrangement of the linker and the region adjacent to it induces productive DNA binding . Thus, our models add weight to a hypothesis of indirect readout by DNA glycosylases, which states that recognition of damaged bases is not limited to formation of specific bonds but greatly relies on the differences in energetics and dynamics of protein and DNA parts that may be far away from the moiety being recognized.
Structural and kinetic data together with QM/MM modeling of Fpg favor the reaction chemistry that combines a nucleophilic attack at C1′ of oxoG by N[Pro1] residue and protonation of O4′ of oxoG by Oε2[Glu2] [33, 35, 36]. The latter step is important since it affords a ~60 kcal/mol lower barrier to glycosidic bond breakage compared to base protonation as the leaving group activation . Such mechanism requires Pro1 to be in the unprotonated, and Glu2, in the protonated state immediately before the reaction, implying that both these residues should change their preferred protonation state. Our measurements of the pH dependence of Fpg activity suggest that only one group is ionized in a pH-dependent manner, in which case it is consistent with Pro1 N-terminal secondary amine losing a proton at increasing pH. Consequently, the ionization state of Glu2 in the Fpg–DNA complex shows no evidence of being pH-dependent, which means that the assembled active site is capable of protonating Glu2, possibly using a water molecule as a proton shuttle. The arrangement of the reacting atoms is only consistent with the reaction stereochemistry with SN2 displacement of O4′ as the first step, in agreement with the QM/MM data . Since the substitution of Gln for Glu2 inactivates the enzyme, which rules out simple hydrogen bonding as the primary function of Glu2, the mechanistic implication of our results is that Glu2 has to be deprotonated again later in the reaction, likely by the nascent alkoxide O4′, and contribute its charge to the stabilization of the transition state of the departing oxoG base. In the QM/MM simulation, several consecutive acts of proton transfer between Oε2[Glu2], O4′[oxoG], N[Pro1], and O8[oxoG] allow the enzyme to lower the highest barrier in the reaction from 71 kcal/mole (as with direct oxoG protonation path) to 13 kcal/mole relative to the lesion recognition complex ; a similar energy gain was calculated for fapyG excision .
Finally, our modeling concerned only the pre-catalytic complex of Fpg–DNA. It is now clear that the selectivity of DNA glycosylases is not determined exclusively by interactions in their pre-catalytic complexes, the structures of which are relative easy to establish by X-ray crystallography, but also relies on several kinetic gates along the full recognition pathway, including primary encounter and damaged base eversion. Future modeling of the early steps of recognition of oxoG-containing pairs will add clarity to our understanding of the opposite-base discrimination by Fpg.
The modeling was performed on an NKS-30T cluster at the SB RAS Supercomputing Center.
The work was supported by Russian Foundation for Basic Research (grant 17-04-01761-a). The funding body had no role in the design of the study and collection, analysis, and interpretation of data and in writing the manuscript.
Availability of data and materials
The trajectories generated in the current study are available from the corresponding author on reasonable request. All results are presented in the main text and additional supporting files.
DOZ and YNV have designed the study. AVP has carried out the MD simulations. AVE has performed biochemical experiments. AVP and YNV have contributed custom software. AVP, YNV and DOZ have participated in the analysis and interpretation of MD trajectories, and writing of the manuscript. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
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