DNA contains the instructions required to make proteins and other molecules in cells. DNA is made of four building blocks called deoxyribonucleotides, which are in turn made from molecules called ribonucleotides by enzymes known as ribonucleotide reductases (RNRs for short). RNR enzymes are responsible for maintaining a good balance in the levels of the different deoxyribonucleotides in cells, which is essential for DNA to be made and repaired correctly.

Previous work has shown that each RNR can act on all four ribonucleotides. However, these enzymes become more selective for certain ribonucleotides depending on which deoxyribonucleotide is most common within the cell. For example, when a deoxyribonucleotide called dGTP is plentiful, it binds to a so-called “specificity site” on the enzyme and alters the shape of the enzyme’s active site. This then means that a ribonucleotide called ADP will bind in preference to the other ribonucleotides. However, it was not clear how the binding of deoxyribonucleotides to the enzyme influences the shape of the active site.

Zimanyi et al. used a technique called X-ray crystallography to determine the three-dimensional structures of a bacterial RNR enzyme when it is bound to all four different combinations of deoxyribonucleotides and ribonucleotides. In the absence of nucleotides, the active site adopts a shape that resembles an open barrel. However, when RNR is bound to a deoxyribonucleotide at the specificity site and a ribonucleotide at the active site, the barrel clamps down, bringing the specificity site and the active site closer together. Additionally, a loop of the protein interacts with each of the deoxyribonucleotides in a different way and communicates their identity directly to the active site, which rearranges itself to hold on to the corresponding preferred ribonucleotide.

Zimanyi et al.’s findings provide an explanation for how RNRs can select between ribonucleotides so that they produce a good balance of deoxyribonucleotides in cells. This will inform future efforts to develop molecules that inhibit RNRs, which may have the potential to be used to treat bacterial infections or to kill cancer cells.

Deoxyribonucleotides, the building blocks for DNA biosynthesis, are produced in the cell from ribonucleotide precursors by ribonucleotide reductase (RNR) (Figure 1). Three classes of RNRs are known, categorized by the cofactor they use to generate a protein radical required for catalysis. The best characterized of all RNRs is the class Ia enzyme from Escherichia coli that employs a di-iron-tyrosyl-radical cofactor to initiate chemistry and requires two dimeric protein subunits for enzymatic activity. The α₂ subunit contains two (β/α)₁₀ barrels, which house the active sites at the barrel centers (Eriksson et al., 1997; Uhlin and Eklund, 1994), and the β₂ subunit utilizes a largely helical secondary structure to house the radical cofactor (Sjöberg and Reichard, 1977) (Figure 1B–C). As a central controller of nucleotide metabolism, RNR uses multiple allosteric mechanisms to maintain the balanced deoxyribonucleoside triphosphate (dNTP) pools that are required for accurate DNA replication. First, allosteric activity regulation modulates the overall size of dNTP pools. ATP or dATP binding at an allosteric activity site, found at the N-terminus of α₂ (Figure 1D), leads to up-regulation or down-regulation of enzyme activity, respectively (Brown and Reichard, 1969). In E. coli class Ia RNR, this regulation is achieved by changes in the oligomeric arrangement of the α₂ and β₂ subunits (Brown and Reichard, 1969; Rofougaran et al., 2008; Ando et al., 2011). When ATP is bound at the activity site, an α₂β₂ complex is favored. Although no X-ray structure of the active complex has been determined, low resolution models have been generated using small-angle X-ray scattering (Ando et al., 2011), electron microscopy (Minnihan et al., 2013), and distance measurements made through spectroscopic analyses (Seyedsayamdost et al., 2007) (Figure 1D). This active α₂β₂ complex is capable of a long-range proton coupled electron transfer from β₂ to α₂, forming a transient thiyl radical on Cys439 to initiate catalysis (Licht et al., 1996). Alternatively, when concentrations of dATP become too high in the cell, dATP binds at the allosteric activity site and formation of an α₄β₄ complex is promoted. The structure of this complex was recently solved (Ando et al., 2011), revealing a ring of alternating α₂ and β₂ units that cannot form a productive electron transfer path, thus inhibiting the enzyme (Figure 1D).

Figure 1

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The second form of allosteric regulation is specificity regulation, which maintains the proper relative ratios of dNTPs in the cell. Briefly, the binding of (d)NTP effectors to an allosteric specificity site in α₂ influences the preference of RNR for its four nucleoside diphosphate (NDP) substrates. Whereas high levels of dATP inhibit class Ia RNR, at lower levels, dATP promotes CDP or UDP reduction. Likewise, TTP promotes GDP reduction, and dGTP promotes ADP reduction (Figure 1E) (Brown and Reichard, 1969; Rofougaran et al., 2008; von Döbeln and Reichard, 1976). Importantly, the affinity of the α₂ and β₂ subunits for each other is weak (~0.4 μM) in the absence of effectors, whereas the binding of a complementary substrate/specificity effector pair increases the affinity of the class Ia RNR subunits fivefold (Crona et al., 2010; Hassan et al., 2008). Previous structural work, which includes: X-ray structures of GDP and TTP bound to E. coli α₂ (Eriksson et al., 1997), structures of all four substrate/effector pairs bound to class Ia α₂ from Saccharomyces cerevisiae (Xu et al., 2006), and class II α₂ from Thermotoga maritima (Larsson et al., 2004), revealed the location of the allosteric specificity sites at the ends of a four helix bundle at the dimer interface (Figure 1B). These data, and accompanying in vitro and in vivo studies on S. cerevisiae (Ahmad et al., 2012; Kumar et al., 2010; Kumar et al., 2011), also implicated which residues (Gln294 and Arg298, E. coli numbering) and which regions of the structure are involved in the communication between the specificity site and the active site. A flexible loop, termed loop 2 (residues 292–301 in E. coli) (Figure 1B), bridges the two sites, which are approximately 15 Å apart, and becomes more ordered upon effector and substrate binding (Eriksson et al., 1997). An additional loop, termed loop 1 (residues 259–278), is also near the effector-binding site and is stabilized upon effector binding (Eriksson et al., 1997). Thus, previous work established the players involved in allosteric specificity regulation in RNR. This work supports those assignments, and goes on to provide a novel set of crystallographic snapshots that reveal how these residues in the prototypic RNR from E. coli are able to communicate and thereby regulate substrate preference.

We have utilized an α₄β₄ crystal form of the E. coli class Ia RNR (Figure 1D, right) (Ando et al., 2011; Zimanyi et al., 2012) to obtain structures of all four substrate/specificity effector pairs (CDP/dATP, UDP/dATP, ADP/dGTP or GDP/TTP) bound to the α₂ subunit (Table 2). Each model consists of two α₂ dimers, two β₂ dimers, and activity effector dATP in all four allosteric activity sites. The residues modeled for each chain of each structure are given in Table 3. Although the α₄β₄ form of the enzyme cannot turn over substrates due to abrogation of the radical transfer pathway, the substrate and effector binding sites are not affected (Video 1). In all structures, substrates and effectors were found ordered at their respective binding sites (loop 2 electron density shown in Figure 2).

Figure 2

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Video 1

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Substrate/effector binding induces a clamping of the (β/α)₁₀ barrel in the α₄β₄ crystal form

Each substrate/effector bound α₄β₄ structure reveals a closing of the (β/α)₁₀ barrel around the bound substrate that is not observed in the absence of substrate. An initial superposition of one α chain from the previously reported free α₂ soaked with AMP-PNP (PDB ID: 3R1R (Eriksson et al., 1997)) onto the α₄β₄ crystal form was performed using only residues 432–446, that comprise the conserved active site loop (‘finger loop’) found in the center of the (β/α)₁₀ barrel. In this structural alignment, one half of the active site barrel overlays exactly, whereas the second half of the barrel and the N-terminal portion of the structure has undergone a clamping movement (Figure 3). To better characterize this movement, a difference distance matrix (DDM) was calculated. This plot reveals relative movements in a reference-independent manner. Briefly, the distance between every possible pair of atoms in one chain of α from the α₄β₄ structure is subtracted from the same distance measured in the free α₂ structure, giving a representation of how much the distances between pairs of atoms change between the two structures. The pattern of movements seen in the plot indicates that the N-terminal 220 residues (region 1 of Figure 3B,C) shift substantially (>3Å), bringing them closer to half of the active site barrel (residues ~224–439, region 2 of Figure 3B,C). We also observe that loop 2 (region 3 of Figure 3B,C) and a flexible β hairpin that sits adjacent to the N-terminus (residues 646–651, region 5 of Figure 3B,C) undergo substantial motions.

Figure 3

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Closer inspection of the active site in the superposition (again, aligned by the active site finger loop) reveals that the movements described above directly affect substrate binding (Figure 4A). Residues Ser622, Ser625, and Thr209 form hydrogen bonds with the NDP substrate, whereas in the substrate-free state (pink ribbon in Figure 4A), these residues are shifted away from the center of the barrel. Thr209 is part of the N-terminal region that is shown to move in the DDM analysis. The structure superposition and DDM analysis also indicate that loop 2 shifts, bringing Arg298 closer to the active site. The Arg298 side chain now forms a cation-π interaction with the substrate base and a charge-charge interaction with the substrate β-phosphate. In the substrate-free structure, this side chain is pointing in the opposite direction. Protein contacts like Arg298 to the substrate diphosphate are expected to be particularly important for charge neutralization in enzymes, such as E. coli RNR, that do not employ metal cations such as Mg²⁺ for this purpose. The positions of residues that contact the ribose O3' (Glu441 and Asn437) are virtually unchanged between the NDP-free and NDP-bound structures (Figure 4A). All four of our substrate-bound structures show almost identical contacts between the enzyme and the substrate ribose and phosphates as described here and shown in Figure 4A,C and Figure 5 for GDP and CDP, respectively.

Figure 4

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Figure 5

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In contrast to the compact α₂ barrel observed in our substrate/effector-bound α₄β₄ structures, no clamping motion of the barrel is observed between substrate-free α₂ and a previously reported structure of α₂ with GDP substrate and TTP effector bound (Eriksson et al., 1997). In this structure, Ser622, Ser625, and Thr209 are all 3.0 Å or greater from the substrate diphosphate, and Arg298 points away from the active site (Figure 4B). These weaker (or missing) interactions between protein and substrate in this previously determined structure are consistent with the fact that substrate is observed in an altered orientation in the active site (Figure 4C,D) and is only present at half occupancy. We attribute the differences in substrate positioning in this structure to the inability of the active site barrel to clamp around substrate due to crystal lattice contacts. Symmetry-related molecules are closer to the active site in the α₂-only crystalline state than they are in the α₄β₄ crystal form. Importantly, differences in flexibility cannot be attributed to the fact that α₄β₄ is an inactive state of E. coli RNR because α₂ is also an inactive state; the former because β₂ is held at arm’s length from α₂, and the latter because β₂ is absent altogether. Additionally, other than contributing to an amenable crystal lattice, there is nothing to suggest that β₂ must be present for α₂ to clamp. When lattice contacts do not restrain α₂ movement, α₂ should be able to clamp in the presence of a cognate substrate/effector pair regardless of β_2. Thus, it seems that different crystal forms have given us two distinct snapshots of substrate-bound states for the E. coli class Ia enzyme: a high affinity state which is closed and ready for radical-based chemistry when β₂ becomes available (Figure 4C) and a lower affinity state in which substrate is still exposed to solvent and not ready to undergo catalysis, regardless of the availability of β₂ (Figure 4D).

Specificity effectors are anchored through conserved ribose and phosphate interactions

Regardless of the identity of the dNTP base, the ribose and phosphates of the three specificity effectors (dATP, dGTP, and TTP) form almost identical contacts with RNR (Figure 6). Despite the presence of the specificity effector site at the dimer interface, these common interactions are made with only one chain of the α₂ dimer. Asp232 and His275 are within hydrogen-bonding distance of O3' of the ribose. One Mg²⁺ ion appears to form an octahedral coordination complex with three phosphate oxygens of the specificity effector and three water molecules (Figure 6). There are also hydrogen bonds between the phosphates and the backbone amides of residues Arg269 and Leu234 and charge-charge interactions with the side chains of Arg269 and/or Arg262.

Figure 6

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Three different loop 2 conformations are responsible for substrate specificity

Loop 2 is ordered in all four structures as indicated by 2F_o–F_c composite omit electron density (Figure 2). As noted above, Arg298 of loop 2 forms a conserved interaction with the β-phosphate of all four substrates (Figures 7, 8). On the effector side of loop 2, Cys292 packs against the base of all three effectors (Figures 7, 8), consistent with binding studies that show the affinity for all specificity effectors is decreased when Cys292 is mutated to alanine (Ormö and Sjöberg, 1996). The remaining interactions with loop 2 are distinct for each substrate/effector pair (Figures 7, 8). dATP, which preferentially increases RNR activity for CDP and UDP substrates, makes two hydrogen bonds with loop 2. The backbone amide and carbonyl of Ser293 are in position to hydrogen bond to N1 and N6, respectively, of the adenine base of dATP. These hydrogen bonds hold Ser293 in place and position the adjacent Gln294 in to a conformation that points into the substrate-binding site (Figure 8A,B). In this conformation, Gln294 can form a hydrogen bond with O2 of CDP or UDP. There is no difference in substrate positioning between CDP and UDP, as the unique positions of the pyrimidine ring (N2 and N4/O4) are not involved in any contacts with the protein. Furthermore, the positioning of Gln294 by dATP selects for pyrimidine binding, as binding of a purine base would be disfavored due to steric occlusion by Gln294.

Figure 7

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Figure 8

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dGTP, which preferentially increases RNR activity for substrate ADP, forms one hydrogen bond with loop 1 residue Thr276 (Figure 6C) in addition to its loop 2 contacts. It is the only effector for which a hydrogen bond is formed between the base and loop 1. Because the position of hydrogen bond donors and acceptors at N1 and O6 in dGTP is opposite from that of N1 and N6 of dATP, the interaction with the backbone amide of Ser293 in loop 2 is unfavorable in this structure. Instead, Ser293 is shifted away from the base, and N1 and N2 – both hydrogen bond donors – form an interaction with the backbone carbonyl of Gln294, which has shifted away from the substrate (Figure 7A–C and 8A–C). Ser293 thus makes no hydrogen bond contacts to dGTP and instead provides hydrogen bonds to Gly300 on the other side of loop 2 through both its side chain and backbone. Stabilization of Gly300 in this position in turn allows for a hydrogen bond between the carbonyl of Gly299 with the N6 position of the preferred substrate, ADP. The movement of Gln294 away from the substrate-binding site allows room for the larger ADP to bind. Thus, Ser293 and Gln294 are responsible both for differentiating between the purine bases adenine and guanine at the effector-binding site and for differentiating pyrimidines from purines at the substrate site, either directly (in the case of Gln294 hydrogen bonding to CDP or UDP) or indirectly through stabilization of Gly299 and Gly300. With dATP bound, Ser293 plays a role in effector recognition and Gln294 in substrate recognition; these roles are reversed for ADP/dGTP.

TTP, which preferentially increases RNR activity for substrate GDP, binds to the specificity site such that only the carbonyl of Cys292 from loop 2 is positioned to form a hydrogen-bonding interaction with the thymine base (Figure 8D). As in the ADP/dGTP structure, Gln294 is swung away from the active site to make room for the binding of a purine substrate (Figure 8D). On the substrate side of loop 2, N1 and N2 of GDP are both within hydrogen-bonding distance of the backbone carbonyl of Gly299. N2 of GDP is also within hydrogen-bonding distance of the backbone carbonyl of Ala252 (not shown), giving GDP one more hydrogen bond than the other substrates

For the purine substrates ADP and GDP, preference appears to rely on a shift of the carbonyl of Gly299 in loop 2 forward or backward in the active site, enabled by the binding of dGTP or TTP, respectively, at the specificity site. With dGTP bound, Ser293 is positioned such that it pushes the carbonyl of Gly299 'forward' (away from the effector-side of loop 2, see Figure 8C), allowing a hydrogen bond to form between the carbonyl of Gly299 and N6 of adenine. With the smaller TTP in the effector site, Ser293 rearranges such that the carbonyl of Gly299 can relax ‘backward’ (toward the effector-side of loop 2, see Figure 8D), allowing for a hydrogen bond between the carbonyl of Gly299 and N1 and N2 of guanine. Specificity is imparted by the fact that the ‘forward’ position of the Gly299 carbonyl interacts with the 6-position of the purine, which is a hydrogen bond donor (NH₂) in adenine and a hydrogen bond acceptor (O) in guanine; and the ‘backward’ position of the Gly299 carbonyl interacts with the 1-position and 2-position of the purine, which are both hydrogen bond donors (NH and NH₂) in guanine but not in adenine (Figure 8C,D). To briefly summarize purine specificity, purines are favored over pyrimidines when Gln294 is positioned away from the active site creating a larger substrate-binding pocket, and ADP is favored over GDP when dGTP binding stabilizes Ser293 close to the Gly299 backbone pushing its carbonyl forward, and GDP is favored over ADP when TTP allows Ser293 to fall back.

Activity of mutant proteins Gln294Ala-α₂ and Arg298Ala-α₂ support structural observations

Our sets of structures implicate Arg298 and Gln294 as key residues in the specificity regulation of E. coli RNR. To confirm this prediction, we prepared Gln294Ala and Arg298Ala mutant enzymes and tested the activity for each substrate/effector pair. Because dATP at high concentrations inhibits RNR activity by binding to the allosteric activity site, whereas at lower concentrations it is a specificity effector, we first carried out control experiments to determine activity levels for CDP reduction in the absence of dATP and at an inhibitory dATP concentration (175 μM) (Figure 9). Results show that wild-type RNR is as active (CDP/dATP) or more active (UDP/dATP) at 1 μM dATP than it is in the positive control (CDP/ATP), indicating that at 1 μM, dATP is acting as a specificity effector and not as an allosteric inhibitor of activity, consistent with previous studies (Birgander et al., 2004). The negative control shows the substantial decrease in wild-type RNR activity on CDP when the concentration of dATP is high enough (175 μM) for dATP to bind to the allosteric activity site and inhibit the enzyme. With these controls in place, the activity of mutant RNRs with all four substrate/effector pairs can be established. Our structures predict that mutation of Arg298 to alanine should decrease activity for all substrates, and that is exactly what we observe (Figure 9). The effect is dramatic for this mutant, with negligible activity observed in all cases. In fact, the small amount of activity detected could be due to low levels of contamination of wild-type E. coli RNR instead of the Arg298Ala mutant protein. Thus, Arg298Ala-RNR is either mostly or completely inactive. In contrast, for Gln294Ala-RNR, we would expect purine and pyrimidine substrates to be differentially affected, with minimal or no loss of activity on ADP/GDP and measurable loss on CDP/UDP, and that is again what we observe. For Gln294Ala, activity is the same as wild-type for ADP/dGTP and perhaps even higher than wild-type for GDP/TTP, whereas for both CDP and UDP with 1 μM dATP, activity is decreased (Figure 9). These results are consistent with the removal of the Gln294 side chain from the active site to create room for the larger substrates (ADP/GDP) to bind and also with the positioning of Gln294 into the active site to stabilize CDP/UDP binding through hydrogen-bonding to the respective bases.

Figure 9

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In this work, we have sought to decipher the molecular rules of substrate/effector recognition in RNR and determine the basis for increased substrate affinity in the presence of a cognate effector. We observe that in the absence of substrates or effectors (for example, in the previously reported structure of α₂ (PDB ID: 3R1R (Eriksson et al., 1997)), the barrel is not clamped and loop 2 is not found in any of the conformations seen in our effector-bound structures (Figure 7E). However, in the presence of substrate/effector pairs, we find a clamped down barrel and a stabilized loop 2 that adopts three different conformations depending on the allosteric effector that is bound (Figure 7A–D) (Video 2). Loop 2 residues Cys292, Ser293, and Gln294 appear to be involved in specific interactions that read out the identity of the base and communicate that identity to the active site. Briefly, dATP, which enhances reduction of CDP/UDP (von Döbeln and Reichard, 1976), makes specific contacts with Ser293, orienting Gln294 toward the active site and stabilizing the binding of both CDP and UDP (Figure 8A–B). The lack of discrimination between CDP and UDP substrates by RNR is not problematic for the cell as cytidine deaminase provides another level of control for dCTP/TTP ratios (Wang and Weiss, 1992; O'Donovan et al., 1971). In contrast, dGTP binding stabilizes Gln294 away from the RNR active site, so that CDP/UDP binding is not stabilized (although it is not prohibited) and there is room for the larger purine substrates to bind and to hydrogen bond to the carbonyl of Gly299 (Figure 8C–D). The creation of a more expansive active site is consistent with the ability of RNR to reduce both purine substrates in the presence of dGTPγS, albeit with a much greater fold activity increase for preferred substrate ADP (von Döbeln and Reichard, 1976). Specificity for ADP versus GDP appears to be modulated by whether effector-loop contacts stabilize the carbonyl of Gly299 in a forward or a backward position, respectively (Figure 8C–D). Thus, our structural data suggest that movement of Gln294 in and out of the active site alternatively shrinks and expands the active site for CDP/UDP versus ADP/GDP, whereas a more modest shift of the Gly299 carbonyl is involved in ADP/GDP selectivity. Mutagenesis of Gln294 to Ala is consistent with this proposal, showing decreased activity on CDP/UDP and no change or increased activity on ADP/GDP (Figure 9).

Video 2

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When a cognate substrate/effector pair is bound and the barrel is clamped, Arg298 is able to reach across the active site, stack against the NDP base, and hydrogen bond to the NDP β-phosphate, thereby sequestering the substrate in the active site, and neutralizing the substrate negative charge. These interactions yield what appears to be a high affinity substrate-bound state that is sequestered by solvent and thus amenable to radical-based chemistry. To invoke an analogy, Arg298 is like a latch on a suitcase, locking the active site barrel in cases in which the active site is appropriately packed (Figure 10A). When a suitcase is packed with too many clothes, the latch cannot reach the lock, and the clothes are not secured. Similarly, a mismatched substrate/effector pair such as a purine nucleotide with dATP would not be expected to allow the barrel to clamp and loop 2 to rearrange such that Arg298 can reach the substrate phosphate and thus form a ‘latched’ complex (Figure 10B). In the latter case, one would expect the mismatched substrate to be released from the enzyme, so that a complementary one can bind. Chemical logic tells us that allosteric regulation of specificity would require both low and high affinity substrate/effector-enzyme states; low affinity states to sample substrate/effector pairs, and high affinity states to capture the correct pair, recruit β₂ (class I) or adenosylcobalamin (class II), and initiate catalysis (Figure 10). Although the molecular basis for the fivefold increase in binding affinity of β₂ for α₂ in E. coli class Ia RNR in the presence of bound substrate/effector pairs is not understood, the clamping of the barrel that we observe here may be part of the molecular explanation. The use of Arg298 as a molecular latch to secure the barrel in this closed state is beautiful in its simplicity. It is also consistent with both our structural and our biochemical data that show almost complete loss of activity in the Arg298Ala variant (Figure 9).

Figure 10

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Though it is generally accepted that the rules of specificity regulation are conserved among class I, II, and III RNRs, this hypothesis is based on a relatively small number of characterized RNRs. Loop 2 is clearly an important player in specificity regulation, and yet it is not highly conserved, even among characterized RNRs (Figure 11). This being said, our structures show that backbone atoms of loop 2 make the vast majority of contacts to the substrate base and the effector base, limiting the number of residues that need to be strictly conserved to allow for the same molecular mechanism. Also, despite the fairly low sequence identities between the E. coli and the T. maritima α subunits (20.5%) and the E. coli and the S. cerevisiae α subunits (29.3%), a structural comparison shows that all three enzymes interact with the phosphate groups and ribose moieties of their substrates and effectors in similar ways (Table 4).

Figure 11

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The only noteworthy difference is that Arg298, critical for reduction of all four NDPs in E. coli, directly contacts the β-phosphate of all four substrates. In contrast, none of the four S. cerevisiae RNR structures show a direct contact to the β-phosphate by the equivalent arginine, although one structure shows a through-water contact, and only one of the substrate/effector bound structures of T. maritima RNR (GDP/TTP) shows direct contact. This GDP/TTP-bound structure is the only T. maritima RNR structure in which loop 2 is fully ordered and it is remarkably similar to our E. coli structures (Figure 10H,I). All contacts made to GDP are the same: Arg side chain to β-phosphate, and loop 2 carbonyl to N1 and N2 of base (Figure 10H, 4C). The TTP contacts are not identical, but in both cases, the resulting loop 2 conformations allow the carbonyl of 299 (E. coli numbering) and 208 (T. maritima numbering) to ‘fall back’ to contact N1 and N2 of GDP.

Excitingly, the T. maritima class II RNR structures also reveal a movement of Gln294 in and out of the active site in response to effector binding. In particular, the GDP/TTP and CDP/dATP bound structures from T. maritima class II RNR, which have an ordered and semi-ordered loop 2, respectively, show that Gln203 (equivalent to Gln294 in E. coli) interacts directly with CDP when dATP is bound, and moves away from the active site when GDP/TTP are bound (Larsson et al., 2004). Unfortunately, many loop 2 residues are disordered and thus not modeled in the T. maritima structures with CDP/dATP, UDP/dATP, and ADP/dGTP, preventing further comparison to our E. coli structures.

In stark contrast to the behavior of Gln294 in E. coli and Gln203 in T. maritima, Gln288 in S. cerevisiae class Ia RNR is observed to point into the active site in all structures, regardless of which substrate/effector pairs are bound. Structural comparisons show that the protein does not pack as tightly around purine substrates when Gln points into the active site as it does when Gln is flipped away (Figure 10E,F compared with 10H,I). It is possible that this difference in Gln positioning in S. cerevisiae, and differences described above for Arg298 (E. coli numbering), indicate that the mechanism of allosteric regulation is not conserved among RNRs. However, it is also possible that these observed structural differences are a result of crystal packing variations or other deviations in how crystals were prepared. Additional structures of high-affinity RNR-substrate complexes will help to clarify the degree to which the roles of Gln294 and Arg298 are conserved across RNR species. Regardless of their exact roles in S. cerevisiae RNR, there are a number of studies that support the importance of these residues (Ahmad et al., 2012; Kumar et al., 2010; Kumar et al., 2011). In particular, when mutant RNR is the only RNR being expressed in S. cerevisiae, mutation of Arg293 (the Arg298 equivalent) to Ala is lethal, and mutation of Gln288 (the Gln294 equivalent) to Ala yields a severe S phase defect (Ahmad et al., 2012). Gln288Ala mutation in S. cerevisiae also leads to substantially elevated dGTP/dATP levels compared with dCTP/TTP levels when one compares wild-type S. cerevisiae expressing two wild-type RNRs with mutant S. cerevisiae expressing one wild-type RNR and one Gln288Ala mutant RNR (Kumar et al., 2010). Under the same experimental conditions, all four dNTPs are elevated by similar amounts (within 1–3%) for the Arg293Ala mutation in S. cerevisiae (Kumar et al., 2010). Although more studies are clearly needed to confirm or refute that the molecular basis of allosteric specificity regulation among RNRs is conserved, it is interesting to note that our in vitro data on E. coli (Figure 9) is consistent with the more severe phenotype in S. cerevisiae for Arg than Gln mutation and is consistent with the observation that mutation of Gln in S. cerevisiae elevates purines over pyrimidines whereas mutation of Arg does not show differential elevation.

In conclusion, the work presented here provides a unifying mechanism for substrate specificity regulation in the most studied RNR, the E. coli class Ia enzyme. Our structures show how each specificity effector is read out at a distal allosteric site and how that information is communicated to the active site where residues rearrange such that specific hydrogen bonds can be formed with the cognate substrate base. When an effector/substrate match is discovered, the barrel is clamped and latched in preparation for catalysis. Just as DNA replication and transcription take advantage of the unique hydrogen-bonding properties of each nucleotide base, enzymatic ribonucleotide reduction also employs these unique hydrogen-bonding properties for specificity regulation. Through an elegant set of protein rearrangements, E. coli RNR screens and selects its substrate from the four potential NDPs, ensuring appropriate pools of deoxynucleotides are available for DNA biosynthesis and repair.

1. 1. Chen P.Y.-T
   2. Zimanyi C.M.
   3. Drennan C.L.

   (2015) Crystal structure of the dATP inhibited E. coli class Ia ribonucleotide reductase complex bound to CDP and dATP at 2.97 Angstroms resolution

   5CNS.

   http://www.rcsb.org/pdb/explore.do?structureId=5CNS
2. 1. Chen P.Y.-T
   2. Zimanyi C.M.
   3. Drennan C.L.

   (2015) Crystal structure of the dATP inhibited E. coli class Ia ribonucleotide reductase complex bound to UDP and dATP at 3.25 Angstroms resolution

   5CNT.

   http://www.rcsb.org/pdb/explore/explore.do?structureId=5CNT
3. 1. Chen P.Y.-T
   2. Zimanyi C.M.
   3. Drennan C.L.

   (2015) Crystal structure of the dATP inhibited E. coli class Ia ribonucleotide reductase complex bound to ADP and dGTP at 3.40 Angstroms resolution

   5CNU.

   http://www.rcsb.org/pdb/explore/explore.do?structureId=5CNU
4. 1. Chen P.Y.-T
   2. Zimanyi C.M.
   3. Drennan C.L.

   (2015) Crystal structure of the dATP inhibited E. coli class Ia ribonucleotide reductase complex bound to GDP and TTP at 3.20 Angstroms resolution

   5CNV.

   http://www.rcsb.org/pdb/explore/explore.do?structureId=5CNV

1. 1. Eriksson M
   2. Uhlin U
   3. Ramaswamy S
   4. Ekberg M
   5. Regnstrom K
   6. Sjoberg BM
   7. Eklund H

   (1997) Ribonucleotide reductase R1 protein with substrate, GDP and effector dTTP fro Escherichia coli

   4R1R.

   http://www.rcsb.org/pdb/explore.do?structureId=4r1r
2. 1. Eriksson M
   2. Uhlin U
   3. Ramaswamy S
   4. Ekberg M
   5. Regnstrom K
   6. Sjoberg BM
   7. Eklund H

   (1997) Ribonucleotide reductase R1 protein with AMPPNP occupying the activity site from Echerichia coli

   3R1R.

   http://www.rcsb.org/pdb/explore/explore.do?structureId=3R1R
3. 1. Ando N
   2. Brignole EJ
   3. Christina M Zimanyi
   4. Michael A Funk
   5. Yokoyama K
   6. Asturias FJ
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Author details

1. Christina M Zimanyi

   Department of Chemistry, Massachusetts Institute of Technology, Cambridge, United States

   Present address

   Department of Molecular and Cellular Biology, Harvard University, Cambridge, United States

   Contribution

   CMZ, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Percival Yang-Ting Chen

   Department of Chemistry, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

   PYTC, The second and third authors contributed equally, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Contributed equally with

   Gyunghoon Kang

   Competing interests

   The authors declare that no competing interests exist.

3. Gyunghoon Kang

   Department of Chemistry, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

   GK, The second and third authors contributed equally, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Contributed equally with

   Percival Yang-Ting Chen

   Competing interests

   The authors declare that no competing interests exist.

4. Michael A Funk

   Department of Chemistry, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

   MAF, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

   Competing interests

   The authors declare that no competing interests exist.

5. Catherine L Drennan

   1. Department of Chemistry, Massachusetts Institute of Technology, Cambridge, United States
   2. Department of Biology, Howard Hughes Medical Institute, Massachusetts Institute of Technology, Cambridge, United States
   3. Center for Environmental Health Sciences, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

   CLD, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   cdrennan@mit.edu

   Competing interests

   The authors declare that no competing interests exist.

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