Directed evolution of a TNA polymerase identifies independent paths to fidelity and catalysis

Domain: chemistry
Published: 2025-12-19
Editorial outcome: major_revision

Abstract
Directed evolution facilitates functional adaptation through stepwise changes in sequence that alter protein structure. While most campaigns yield solutions that maintain the framework of a rigid protein architecture, a few have produced enzymes with more notable structural differences. One example is a polymerase that was evolved to synthesize threose nucleic acid (TNA) with near-natural activity. Understanding how this enzyme arose provides a model for studying pathways that guide enzymes toward more productive regions of the fitness landscape. Here, we trace the evolutionary trajectory of an unnatural polymerase by solving crystal structures of key intermediates along the pathway and evaluating their biochemical activity. Contrary to the view that fidelity is a product of increased catalytic efficiency, we find that accuracy and catalysis are decoupled activities guided by separate ground-state and transition-state discrimination events. Together, these results offer a glimpse into the forces responsible for shaping the emergence of new enzyme functions.

Introduction
Directed evolution is a powerful approach for generating enzymes that are needed to drive future applications in biotechnology and medicine1–3. Through recursive cycles of diversification and selection, populations of molecules adapt to novel biochemical functions that lie outside the sphere of activities found in nature4. Though effective, most protein evolution studies are unable to match the catalytic efficiency of natural enzymes5. The central challenge lies in understanding how to reshape the active site pocket to accommodate noncognate substrates while preserving the catalytic activity of the native enzyme fold. Although aspects of this problem have been studied by examining the products of directed evolution campaigns6–9, understanding the molecular basis of functional adaptation requires structural information about key intermediates along the pathway, each representing a critical step toward the enzyme’s final, functional form10. By studying the evolutionary journey from a low-activity variant to a highly functional enzyme, it may be possible to learn basic design principles that could accelerate future selections or expand the predictive power of AI algorithms to include dynamic enzymes with complicated reaction mechanisms.

In recent work, we demonstrated the directed evolution of a polymerase, called 10–92, that can synthesize α-L-threofuranosyl nucleic acid (TNA) with near-natural activity (Fig. 1)11. Biochemical studies reveal that 10–92 achieves a catalytic rate of ~1 nt s−1 and >99% fidelity by sequentially adding TNA triphosphates (tNTPs) to the 2′-hydroxyl group of the growing primer, yielding an unnatural 3′ → 2′ linked TNA product (Fig. 1a). The enzyme was discovered using the massively parallel screening technique of droplet-based optical polymerase sorting (DrOPS)12,13 to query a large combinatorial library prepared by recombining the sequences of four hyperthermophilic archaeal B-family DNA polymerases (DNAP) (Thermococcus kodakarensis, Kod; Thermococcus gorgonarius, Tgo; Pyrococcus Deep Vent, DV; and Thermococcus 9°N, 9°N) (Fig. 1b, c). Each polymerase carried the exonuclease silencing mutations (D141A and D143A) and four beneficial mutations (A485R, N491S, R606G, and T723A) found in Kod-RSGA, our previous best TNA polymerase (TNAP)14.

A crystal structure of the 10–92 TNAP uncovered large movements in the protein architecture that improved substrate binding, including positional rearrangements of α-helical secondary structures (up to 7 Å) that bring the thumb subdomain closer to the paired region of the DNA duplex and other large movements that realign the catalytic triad in the palm subdomain (Supplementary Figs. 1 and 2)11. Together, these structural changes enabled the transition from an enzyme capable of highly efficient DNA synthesis to one that is now specialized for TNA synthesis (Supplementary Fig. 3). Whether these structural changes arose early or late in the evolutionary process remains to be established, as do the underlying mechanisms responsible for achieving enhanced catalytic activity.

Here, we study the evolutionary trajectory of the 10–92 TNAP by solving crystal structures of key intermediates along the pathway and evaluating their biochemical activity. Crystal structures were resolved as binary and closed ternary complexes, and biochemical data were collected for fidelity, catalytic efficiency, thermal stability, and substrate specificity. Among the more notable trends is the observation that fidelity and catalysis appear to occur as decoupled activities, possibly guided by separate ground- and transition-state discrimination events. Molecular insights into these processes were obtained from crystal structures that were used to map the process of active site refinement across successive generations of the enzyme. Together, these findings provide a detailed mechanistic framework for understanding how directed evolution reshapes enzyme function to achieve accurate and efficient catalysis.

Results
Biochemical analysis of the evolutionary intermediates
To follow the biochemical changes that occurred along the evolutionary path, we evaluated the polymerases that most influenced the evolution of the 10–92 TNAP for fidelity, activity, thermal stability, and substrate specificity. This analysis included Kod-RSGA as well as intermediates 5–270, 7–47, and 8–64 (Fig. 1c and Supplementary Fig. 4), the most active variants identified in rounds 5, 7, and 8, respectively, and the parent sequences used for further diversification11.

Our analysis of fidelity and catalysis reveals a general trend in which steep gains in fidelity occur in earlier generations of the enzyme, whereas analogous improvements in activity are delayed until later stages of the evolutionary trajectory (Fig. 1d; Supplementary Figs. 5–7 and Supplementary Table 1). However, it should be noted that we do observe a concomitant improvement in fidelity as catalysis improves, presumably due to fine-tuning of the active site near the end of the evolutionary trajectory. Nevertheless, the overall trend of early improvements in fidelity and later improvements in catalysis caught our attention as it appears to counter the prevailing view that a selection for improved catalytic activity through the use of progressively shorter incubation times, as was the case for 10–9211, should indirectly enrich for enzymes with higher fidelity15,16. This is because less faithful polymerases are more likely to stall at misincorporation sites, thereby failing to complete the extension within the allotted time17–22. Thus, variants that maintain rapid synthesis under stringent incubation times are expected to exhibit the combined properties of enhanced catalytic efficiency with increased fidelity. While this interpretation is consistent with the selection results overall, our biochemical analysis of the evolutionary intermediates indicates that variants can emerge that exhibit enhanced fidelity without a proportional gain in catalytic activity. This observation implies that fidelity and catalysis may have determinants that can be separately tuned during the early stages of evolution.

We also found that all of the polymerases tested retain their extreme thermostability, showing no detectable loss in TNA synthesis activity after prolonged exposure to 90°C for 6 h (Fig. 1d and Supplementary Fig. 8). This result highlights the exceptional structural integrity of the variants and challenges the view that directed evolution inevitably compromises protein folding stability5. Such trade-offs are particularly relevant in enzyme engineering campaigns, where enhanced activity often comes at the expense of reduced thermostability23. Remarkably, the 10–92 TNAP, with its 51 amino acid mutations relative to its closest natural homolog, Kod DNAP (Fig. 1c), exhibits no measurable loss of thermostability (Supplementary Fig. 8). This observation supports the view that protein stability promotes evolvability24, which may be a result of the thermal lysis step of the DrOPS selection protocol25.

Finally, the 10–92 TNAP also exhibits a striking shift in substrate specificity, defined here as the ability for the polymerase to distinguish TNA versus DNA substrates based on their catalytic rates of incorporation. Kinetic analysis across generations of the polymerase reveals a consistent reluctance of the enzymes to relinquish their ancestral DNA synthesis activity until the final stages of evolution, at which point an inversion (13-fold) in substrate specificity occurs, favoring TNA synthesis over DNA synthesis (Supplementary Fig. 7). This inversion of substrate preference marks the transition of the enzyme from a broadly acting generalist to one that is increasingly specialized for TNA synthesis on DNA templates. Such a shift is a hallmark of functional divergence, arising from the accumulation of mutations that reconfigure the active site toward a new function.

A binary complex capturing an unexpected preorganized state
As an initial step towards solving the puzzle of how 10–92 emerged as a highly active enzyme from a diverse pool of synthetic homologs, crystal structures of major intermediates identified along the evolutionary pathway were first solved as binary structures of the post-catalytic complex with a bound DNA primer-template (P/T) duplex that was enzymatically extended with two TNA residues (3′-tTtA-2′) at the 3′ terminus. Four structures (5–270, 7–47, 8–64, and 10–92) spanning a resolution limit of 2.17–2.56 Å were determined by molecular replacement (Supplementary Table 2). Consistent with all known archaeal B-family DNAPs, the enzymes adopt a disk-shaped architecture (Supplementary Figs. 9 and 10) defined by an N-terminal domain (NTD), an exonuclease domain, and a catalytic domain comprised of the finger, palm, and thumb subdomains26. In all cases, the duplex is well resolved and tightly bound between the palm and thumb subdomains (Supplementary Fig. 9).

Surprisingly, comparative structural analysis reveals an unexpected, closed conformation for the finger subdomain in all four of the TNAP structures. In this conformation, helices α14 and α15 are shifted by ~22° relative to their position in the open binary structure of wild-type Kod DNAP (Fig. 2a–e)27. This finding contrasts with all known binary structures of replicative DNAPs, which typically exhibit an open finger conformation ready to accept the incoming triphosphate28. Until now, the closed state has only been observed in structures of ternary complexes trapped in a catalytically competent state of the reaction cycle (Fig. 2b, c)29,30.

While it is possible that one could consider this observation a crystallization artifact, several lines of evidence argue against this option. First, the TNAP variants (5–270, 7–47, 8–64, and 10–92) crystallized under two distinct precipitation conditions, suggesting that the conformation reflects a bona fide structural feature rather than a condition-specific artifact. Second, the closed binary conformation is absent in previous binary structures of Kod-RI31 and Kod-RSGA32 (Supplementary Fig. 11), further strengthening the view that the conformational difference arises from sequence changes rather than crystallization chemistry. Finally, the closed binary conformation emerged in variants that also exhibit changes in their biochemical activity, providing a functional correlation that supports its structural relevance.

To understand the molecular basis of this distinct conformation, we analyzed the interdomain interactions stabilizing the closed state. In the binary form of natural Kod DNAP, only limited contacts exist between the exonuclease domain and finger subdomain (Fig. 2f)27, whereas noticeably more contacts are observed as Kod transitions to a closed ternary conformation, supporting a catalytically competent state with the dNTP poised for chemical bond formation (Fig. 2g)26. Contrasting the natural system, the closed binary conformation of the TNAP intermediate observed in 5–270, 7–47, 8–64, and 10–92, exhibits a robust network of electrostatic and hydrophobic interactions that preorganize the enzyme in a catalytically active conformation despite the absence of a bound tNTP (Fig. 2h).

The structural similarity between the binary TNAP complexes and the closed ternary structure of 10–92 previously solved by X-ray crystallography is striking, particularly in the geometry of the active site, which differs only by the absence of substate-specific contacts (Supplementary Fig. 12)11. Because the closed binary motif first appears in the 5–270 intermediate and is absent in the binary structure of Kod-RSGA32 (Supplementary Fig. 11), we infer that this conformational switch arose from recombination events that facilitated TNA synthesis by stabilizing the enzyme in a catalytically productive state. In this scenario, the finger subdomain exists in a dynamic equilibrium between the open and closed conformations, reminiscent of natural DNAPs20. However, upon correct substrate binding and the ensuing conformational change19, the preorganized state would provide a selective advantage by biasing the enzyme toward the closed conformation, thus offering more time for catalysis to occur prior to substrate release.

The catalytically active state of the enzyme
To visualize how catalytic activity evolved within the TNAP lineage, we determined the crystal structures of key intermediates in a catalytically competent state with a closed finger conformation and a P/T duplex and tATP substrate bound in the enzyme active site. A key aspect of this effort was the chemical synthesis of a chain-terminating 2′-deoxy-α-L-threofuranosyl thymidine-3′-triphosphate (dtTTP) analog, prepared in 8 synthetic steps from a 3′-protected α-L-threofuranosyl thymidine (tT) precursor using a Barton deoxygenation strategy to remove the 2′ hydroxyl group (Supplementary Figs. 13–16)33,34. This substrate enabled us to capture the reaction state immediately following tNTP binding, but before chemical bond formation. Using this approach, we successfully obtained high-resolution crystal structures for 5–270 and 8–84, resolving to 3.03 and 2.38 Å, respectively (Supplementary Table 3).

By combining this data with previously solved structures for Kod-RI31 and 10–9211, the least and most optimized versions of the enzyme, respectively, a structural framework was established to study the biochemical changes that occurred along the evolutionary path. Superposition of the four TNAP structures (Kod-RI, 5–270, 8–64, and 10–92) reveals that a global divergence in structure occurred following recombination (Fig. 3a, e and Supplementary Fig. 17). The backbone Cα root mean square deviation (RMSD) relative to wild-type Kod DNAP increases from ~0.5 Å for Kod-RI to ≥1.0 Å for 5–270, 8–64, and 10–92, with movements of secondary structural elements observed in the thumb, palm, and NTDs. These changes correspond to a progressive remodeling of the active site pocket (Fig. 3b–f), which aligns with observed biochemical gains in TNA synthesis activity (Fig. 1d). Surprisingly, most of the 51 acquired mutations found in 10–92 are located >20 Å from the catalytic core (Fig. 3a and Supplementary Fig. 18), emphasizing the importance of long-range structural interactions in fine-tuning the active site geometry35.

Importantly, deep learning models failed to predict these transitions. Structure predictions by AlphaFold3 yielded only canonical B-family DNAP structures, with open binary conformations and no evidence of the experimentally observed rearrangements (Supplementary Figs. 19–23). This observation underscores a key limitation of current AI-based modeling, which is the inability to accurately capture long-range mutations and evolution-driven conformational shifts that underlie new protein functions. Future improvements in predictive accuracy will require experimental benchmarks and better mechanistic understanding of conformational plasticity during enzyme evolution.

Reshaping the active site pocket
Our previous crystal structure of the 10–92 TNAP trapped in a closed ternary complex revealed how recombination and molecular evolution enabled substantial active site remodeling to support efficient TNA synthesis11. The selected mutations produced local backbone shifts, including high-energy rearrangements, that restructured the active site pocket to promote optimal hydrogen bonding, electrostatic stabilization, and van der Waals complementarity with the bound tNTP. In this conformation, the catalytic aspartate triad (D405, D541, and D543) is precisely arranged to coordinate metal ions and facilitate robust nucleotide transfer chemistry.

By integrating structural data from intermediates along the evolutionary path, we now provide a high-resolution, atomic-level view of how these changes unfolded. A central theme to emerge is the stepwise refinement of the active site geometry, beginning with a disruptive expansion of the active site volume followed by precise, substrate-specific remodeling. For example, the active site volume increases ~250% from wild-type Kod DNAP to Kod-RI, before contracting through 5–270 and 8–64 to a more compact volume of 692 Å3 observed in 10–92, which is approximately 10% smaller than the starting enzyme (Fig. 3c, d and Supplementary Fig. 24). This process of active site refinement allows the enzyme to accommodate the smaller size of the tNTP substrate, noting that TNA derives from a four-carbon threose sugar, as compared to the five-carbon sugar found in DNA36.

At the local level, analysis of the catalytic triad reveals a clear stepwise refinement across the lineage. Overlaying the active site from successive TNAP generations shows that the catalytic residues undergo a series of positional and rotational movements, supported by Polder maps, that ultimately align the aspartate residues into a catalytic configuration closely mimicking the arrangement found in wild-type Kod DNAP (Fig. 3e, f; Supplementary Figs. 25 and 26). These refinements enable TNA substrate recognition while preserving the structural features essential for metal ion coordination and catalysis.

Our structural analysis reveals that base pair recognition and reaction center geometry emerged as two distinct evolutionary pressures shaping the evolutionary trajectory. The former is reflected in the conformation of the nascent Watson–Crick base pair formed between the incoming tATP and the templating thymine base. Following the active site distortion observed in Kod-RI, base pair geometry was largely restored in 5–270, as measured for the parameters of opening, propeller, and buckle (Fig. 4 and Supplementary Table 4). By comparison, the geometry of the reaction center improved more gradually across generations of the enzyme, with the distance from the C2′ group of the primer to the α-phosphate of the bound tATP steadily decreasing from 5.8–6.7 Å in 5–270 (Supplementary Fig. 27) to eventually 3.9 Å in 10–92. This final value closely approximates the ideal distance of 3.7 Å observed in wild-type Kod DNAP (Fig. 4b). Together, these findings underscore the evolutionary mechanism underpinning TNAP optimization; namely, early gains in fidelity correspond to improved base pair recognition, while later gains in catalytic activity emerge from a fine-tuning of the active site geometry. This structural and functional convergence culminates in a highly specialized enzyme whose performance reflects both macro-scale architectural adaptation and micro-scale atomic-level precision.

Mutational landscape underlying the activity of 10–92
The mutational landscape underlying the evolution of the 10–92 TNAPs reveals a clear shift from broadly distributed, largely conserved substitutions found in 5–270 to a set of later-arising, non-conserved mutations that cluster in the thumb subdomain and strongly impact TNA synthesis activity (Fig. 5 and Supplementary Table 5). This distribution is unsurprising as the mutations found in 5–270 are the result of homologous recombination, while those in 10–92 derive from focused mutagenesis11. Structural analyses show that mutations occur across both surface exposed and buried positions, with most mutations residing far from the catalytic center (Fig. 5), again underscoring the importance of long-range interactions in shaping new protein functions35. Computational free-energy predictions indicate that while early substitutions are generally neutral with respect to stability, most of the function-driving mutations obtained from directed evolution are destabilizing (Fig. 5), highlighting the trade-off between structural integrity and catalytic enhancement23. Limited reversion analysis of later-stage mutations demonstrates that at least one function-driving mutation is present in each of the key intermediates (Fig. 5)11. Together, these data suggest that polymerase specialization for TNA synthesis emerged through a process involving the accumulation of large numbers of mostly neutral mutations via homologous recombination that change the protein architecture, followed by a smaller set of adaptive mutations that refine catalytic activity at the cost of local stability.

Identifying the precise set of mutations responsible for improved fidelity is challenging because 5–270 carries 35 mutations, all derived from natural diversity. These substitutions collectively remodel the enzyme architecture, produce major structural alterations that reshape the active site, improve the geometry of the nascent Watson–Crick base pair of the incoming tNTP, and facilitate formation of the preorganized conformation observed in the binary structures. While this extensive remodeling likely underlies the fidelity gains observed in 5–270, the large number of changes makes it difficult to determine which mutations are adaptive drivers versus neutral passengers that simply accumulate through the process of directed evolution. Nevertheless, some mutations do appear to contribute disproportionately to these global changes, including the 381L insertion and substitutions within the thumb subdomain (G602D and K672R). Future studies aimed at disentangling the functional role of these and other mutations will be critical for elucidating the mechanistic basis of the fidelity improvement observed early in the evolutionary path.

Interpreting the mutations responsible for improved catalytic activity is aided by a single-point reversion analysis performed on the ten amino acid mutations observed between 5–270 and 10–9211. This analysis revealed that the top four most important mutations in terms of their contribution to catalytic improvement (D615G, A741P, T548P, and S493G) involve the gain of either a glycine or proline residue, while the fifth most important mutation (P550H) transitioned a proline to histidine (Figs. 5 and 6). Together, these changes highlight the importance of flexibility and rigidity as a mechanism for fine-tuning the activity of the catalytic center. While these mutations are located ~20 Å (except S493G, which is ~13 Å) from the active site, defined as the 2′ terminus of the primer, they have a profound impact on enzyme function. The D615G and A741P mutations, first observed in 7–47 and 8–64, respectively, directly contact the primer and template strands, respectively (Fig. 6; Supplementary Figs. 28 and 29). These interactions balance the need for the enzyme to recognize the duplex tight enough to allow for catalysis yet weaken enough to facilitate efficient translocation. The T548P and P550H mutations, both observed in 10–92, induce structural changes in a loop that optimizes the position of the catalytic triad for efficient coordination to divalent metals (Fig. 6 and Supplementary Figs. 28–30), allowing for enhanced catalytic efficiency. Finally, the S493G mutation observed in 10–92 is located in the active site pocket (Fig. 6; Supplementary Figs. 28 and 29). This mutation appears to reduce suboptimal interactions between the enzyme and templating base, resulting in better substrate binding and catalysis.

Methods
Reagents
DNA oligonucleotides were purchased from Integrated DNA Technologies (IDT, Coralville, Iowa). TNA triphosphates were obtained by chemical synthesis previously34,37. ThermoPol buffer, Q5 DNA polymerase, Xl1 Blue competent E. coli, BL21(DE3) competent E. coli, Taq DNA polymerase, NdeI and NotI restriction enzymes were purchased from New England Biolabs (Ipswich, MA). Qiaprep plasmid minikit was purchased from Qiagen (San Diego, CA). Amicon centrifugal filter, hen egg white lysozyme, and chemical reagents including dNTPs, sodium chloride, manganese chloride, magnesium chloride, Tris-HCl, isopropyl β-D-thiogalactoside (IPTG), 1,4-dithiothreitol (DTT), ampicillin, and ammonium persulfate (APS) were purchased from Sigma Aldrich (St. Louis, Missouri). TOPO TA cloning kit, ethylenediaminetetraacetic acid (EDTA), and Dynabeads M270 were purchased from Thermofisher Scientific (Waltham, Massachusetts). SequalGel UreaGel 29:1 Denaturing Gel System was purchased from National Diagnostics (Atlanta, GA). Tetramethyl-ethylenediamine (TEMED) was purchased from Bio-Rad (Hercules, California). Chromatography columns were purchased from Cytiva (Little Chalfont, United Kingdom). EvaGreen® dye was purchased from Biotium (Fremont, CA). Clear V-bottom 96-well plates were purchased from Greiner (Monroe, NC). Plastic 1.5 mL microcentrifuge tubes were purchased from Sigma-Aldrich (St. Louis, MO). Crystallization screens and individual crystallization reagents were purchased from Hampton Research (Aliso Viejo, CA), NeXtal Biosciences (Holland, OH), and Molecular Dimensions (Holland, OH). The Mosquito crystallization robot was purchased from SPT LabTech (Covina, CA). pET-21b(+) plasmid was purchased from Novagen Technology (Glendale, CA). Whole plasmid and Sanger sequencing were performed using Plasmidsaurus (Los Angeles, CA) and Genewiz, respectively (San Diego, CA).

TNAP expression and purification for biochemical assays
Plasmids for TNAP expression and purification were generated previously38. Briefly, a pGDR11 vector containing the TNAP of interest was transformed into XL1 Blue E. coli, grown, induced for expression, and harvested. Cells were resuspended and sonicated in TNAP buffer (10 mM Tris-HCl, pH 8.0, 500 mM NaCl, 10% glycerol). The cell lysate was heat-treated at 70°C for 1 h, cooled on ice for 1 h, and centrifuged (23,708 × g, 4°C, 20 min). The clarified supernatant was treated with a final concentration of 0.5% (v/v) PEI for 15 min and centrifuged (23,708 × g, 4°C, 20 min) to remove nucleic acids. Ammonium sulfate precipitation was performed (final concentration: 60% (w/v)), centrifuged (23,708 × g, 4°C, 20 min), and the protein pellet was resuspended in equilibration buffer (10 mM Tris-HCl, pH 8.0, 50 mM NaCl, 10% glycerol). The TNAP was purified by hand on a 5 mL heparin affinity column and eluted by stepwise addition of buffers containing: 10 mM Tris-HCl, pH 8.0, 10% glycerol and increasing concentrations of NaCl (50, 100, 250, 500, and 750 mM). Eluted TNAP at 500 mM NaCl was pooled and concentrated to 100 μM for biochemical assays.

TNAP kinetics
TNAP kinetic measurements were performed by monitoring the reaction progress as the DNA primer-template duplex (PBS8: GTCCCCTTGGGGATACCACC and EM619: CCCACACCCTCCTATCGCTAAACACACACTTAATAAAGTTGGTGGTATCCCCAAGGGGAC, Supplementary Table 1) was extended with dNTPs or tNTPs to measure DNA or TNA synthesis, respectively, over time39. For each time point, 2 individual reaction replicates (15 μL) from a single master mix poised under single-turnover conditions with equimolar concentrations of TNAP (0.2 μM) and pre-annealed duplex (0.2 μM each, heated at 90°C for 5 min, cooled on ice) in 1x ThermoPol buffer [20 mM Tris-HCl, 10 mM (NH4)2SO4, 10 mM KCl, 2 mM MgSO4, 0.1% Triton X-100, (pH 8.8)] were pre-equilibrated for 5 min at 55°C. The reactions were then initiated by adding a preheated triphosphates mixture (200 μM of each). The reactions were stopped at designated time points by plunging the reaction vessel into powdered dry ice. Once all the time points were collected, the frozen reactions were thawed on a cold plate by adding 15 μL (6 μM) EvaGreen® dye, previously identified as the optimal intercalating dye to monitor the synthesis on a DNA template39. Each reaction (25 μL) was transferred to a clear V-bottom 96-well plate, and fluorescence intensity was measured (ex: 487/20 nm, em: 528/20 nm) using a Synergy Neo2 plate reader with the Gen5 v3.14.03 software (BioTek).

For both DNA and TNA synthesis, baseline fluorescence was measured using Kod (exo-), the parent polymerase with negligible TNA synthesis ability, which was used to collect a 0-s time point by freezing the reaction immediately after adding tNTPs. The maximum fluorescence value was obtained from a 15-min reaction with each polymerase. The raw fluorescence data from the plate reader were normalized by subtracting the baseline and dividing by the difference between the maximum (15-min reaction) and minimum (baseline) fluorescence values. The fluorescence data were then converted to nucleotides per polymerase39 by multiplying the data with the conversion factor F = [Template]*L/[Polymerase], where L is the length of extension in bases, then plotted over time. Rates in nucleotides per minute are extracted as the slope of the linear range of each curve. Data were processed using Microsoft Excel v16.66.

TNAP fidelity
Fidelity analysis was performed in hydrogel particles40. In brief, an aliquot of ~12 million hydrogel particles displaying the acrydite PBS8 primer (GTCCCCTTGGGGATACCACC, Supplementary Table 1) was placed into a 1.5 mL and washed twice with 100 μL 1x ThermoPol buffer. The hydrogels were then resuspended in 100 μL 1x ThermoPol buffer containing 2 μM 4NT.9G fidelity template (Supplementary Table 1) and 1% KF-6012 and heated at 95°C for 3 min. The reaction mixture was then snap-cooled on ice for 5 min to promote primer-template annealing. To initiate TNA synthesis, 2 μL of tNTPs (5 mM stock) and 20 μL of TNAP (10 μM stock) were added, mixed, and then incubated at 55°C on a ThermoMixer C for 1 h. After the reaction is complete, hydrogels were washed with 100 μL breaking buffer (10 mM Tris-HCl pH 7.5, 100 mM NaCl, 1 mM EDTA, 1% (v/v) SDS, 1% (v/v) Triton X-100) twice, incubated with 100 μL 100 mM NaOH (37°C, 3 min) to strip the template, and neutralized with 100 μL breaking buffer. The hydrogels were resuspended in 100 μL 1x ThermoPol buffer containing 2 μM PBS7.PBS9 overhang primer (Supplementary Table 1) with a double nucleotide mismatch (TT-TT), 0.01% KF-6012, 3 mM magnesium sulfate and then heated at 95°C for 3 min. The reaction mixture was then snap-cooled on ice for 5 min to promote primer-template annealing. To initiate the reverse transcription reaction, 10 μL of dNTPs (5 mM stock) and 20 μL of Bst-LF (10 μM stock) were added, mixed, and then incubated at 50°C on a ThermoMixer C for 4 h. The cDNA was PCR amplified with Taq DNA polymerase using PBS8 and PBS9: CTTTTAAGAACCGGACGAAC as primers (Supplementary Table 1). PCR material was ligated into a TOPO vector using a TOPO-TA kit and cloned into DH5α competent E. coli cells. Individual clones were picked, grown in liquid LB media, and sequenced. DNA sequences were aligned and analyzed with the template using CLC Main software. Error rates were determined as μ exp→obs = (#observed/#expected) × 1000. The total error rate was determined by summing the error rate for each substitution.

TNAP thermal challenge
5 × 1.5 mL Eppendorf tubes, each containing 100 μL Kod or TNAP (10 μM), were placed into an Eppendorf ThermoMixer set at 90°C with a thermal lid to prevent evaporation. After 6 h, the tubes were spun at 16,3000 × g for 5 min at 4°C and UV quantified using a NanoDrop instrument. TNA synthesis was assayed with a primer-template duplex (PBS8_short: /5IRD680/GTCCCCTTGG and EM619: CCCACACCCTCCTATCGCTAAACACACACTTAATAAAGTTGGTGGTATCCCCAAGGGGAC, Supplementary Table 1). Polymerase activity screen was performed with the following final concentrations: 1 μM primer-template duplex, 100 μM tNTPs, 1 μM heat-treated polymerase in a 20 μL reaction. After 30 min of incubation, a 1 μL aliquot of the reaction was quenched by the addition of 39 μL of quenching buffer and visualized by denaturing polyacrylamide gel electrophoresis with fluorescent imaging on a Li-Cor Odyssey CLx Imager, Image Studio Lite v5.2.

TNAP cloning, expression, and purification for crystallography
Full-length tnap genes in pGDR11 were PCR amplified using the TNAP-Fwd (ATCCATATGATCCTCGACACTGACTAC) and TNAP-Rvs primers (ACGCATGCGGCCGCTCAAGTTCCTTTCGGCGTCAG), Supplementary Table 1, containing NdeI and NotI restriction enzyme sites, respectively. C-terminal truncation constructs for all TNAPs, except for 8–64, were PCR amplified by substituting the TNAP-Rvs primer with TNAP-Rvs_760 (ACGCATGCGGCCGCTCACTTCTGGTAGCGCAGGTC, Supplementary Table 1); 8–64-Rvs_760: ACGCATGCGGCCGCTCAAGTT CCTTTCGGCGTCAG was used to amplify 8–64, which harbored acquired mutations at the C-terminal. All reverse primers contain the NotI restriction enzyme site. Purified PCR product of each gene construct and pET-21b(+) were digested with NdeI and NotI restriction enzymes and ligated, resulting in pET21–5–270, pET21–5-270_760 pET21–7–47, pET21–7–47_760, pET21–8–64, pET21–8–64_760, and pET21–10–92. Post verification by whole plasmid sequencing, all TNAP constructs were expressed and purified31. Briefly, BL21(DE3) cells harboring pET21-tnap were grown aerobically at 37°C in LB medium containing 100 μg mL−1 ampicillin. At an OD600 of 0.8, expression of a tagless TNAP was induced with 0.8 mM isopropyl β-D-thiogalactoside at 18°C for 20 h. Cells were harvested by centrifugation for 20 min at 3315 × g at 4°C and lysed in 40 mL lysis buffer (10 mM Tris-HCl, pH 7/6.5-full-length/truncated TNAP, 100 mM NaCl, 0.1 mM EDTA, 1 mM DTT, 10% glycerol, 5 mg hen egg white lysozyme) by sonication. The cell lysate was centrifuged at 23,708 × g for 30 min, and the clarified supernatant was heat-treated for 20 min at 70°C and centrifuged again at 23,708 × g for 30 min. The supernatant was loaded onto 5 mL HiTrap Q HP and heparin HP columns assembled in series with the efflux of the Q column loaded in front of the heparin column. After washing with lysis buffer, the Q column was removed, and TNAP was eluted from the heparin column with a high salt buffer (10 mM Tris-HCl, pH 7/6.5-full-length/truncated TNAP, 1 M NaCl, 0.1 mM EDTA, 1 mM DTT, 10% glycerol) using a linear gradient. Eluted fractions containing TNAP were visualized by SDS-PAGE, pooled, and concentrated using a 30 kDa cutoff Amicon centrifugal filter. Further purification was achieved by size exclusion chromatography (Superdex 200 HiLoad 16/600) pre-equilibrated with TNAP buffer (50 mM Tris-HCl, pH 8.5, 200 mM NaCl, 0.1 mM EDTA, 1 mM DTT). Purified TNAP was concentrated to 20 mg mL−1 for crystallization trials.

TNAP crystallography
The binary DNA template, T1: /5Cy5/AAACGTACGCAGTTCGCG, was HPLC purified sample, while the remaining oligonucleotides, P: CGCGAACTGCG and T2: TATGCACGTACGCAGTTCGCG, were ordered with standard desalting (Supplementary Table 1). The primer-template duplexes, P/T1 and P/T2 for the binary and ternary complexes, respectively, were prepared by combining equal parts of the respective primer and template strands in TNAP buffer and supplemented with 20 mM MgCl2, heating at 95°C for 5 min and slow cooling to 10°C over 10 min.

All constructs were screened against ~900 conditions in a hanging-drop format using a Mosquito crystallization robot (SPT LabTech). Positive crystal hits were further optimized in 24-well hanging drop trays, with each drop consisting of 1 μL of sample mixed with 1 μL of mother liquor over 500 μL mother liquor in every well. The specific crystallization conditions for each TNAP complex are described below. Six diffraction datasets were collected at synchrotron sources (Advanced Light Source, Stanford Synchrotron Radiation Lightsource, and National Synchrotron Light Source II) from single crystals. Unless specified, images were indexed, integrated, and scaled using XDS41. Data collection statistics are summarized in Tables S2 and S3. Initial models were determined by molecular replacement using Phaser42; for the binary complexes, the Kod-RSGA binary complex (7RSU) was initially used as the search. However, the 10–92 closed ternary complex (8T3X) was subsequently used due to the closed conformation of the binary structures. 8T3x was used as the search model for the ternary complexes. All final models were determined using iterative rounds of manual building through Coot43 and refinement with phenix44. The stereochemistry and geometry of all structures were validated with Molprobity45 with the final refinement parameters summarized in Tables S2 and S3. Final coordinates and structure factors have been deposited in the Protein Data Bank.

Binary complexes were prepared by incubating full-length TNAPs (6 mg mL−1) with 1.2 molar equivalents of the annealed P/T1 at 37°C for 30 min. 5 M excess of tTTP and tATP were added and incubated at 37°C for 30 min separately. The resulting 5–270 binary complex crystallized in 0.1 M calcium acetate, 0.1 M 2-(N-morpholino)ethanesulfonic acid pH 6, and 15% polyethylene glycol 400 while the remaining binary complexes crystallized in 0.06 M magnesium chloride hexahydrate, 0.06 M calcium chloride dihydrate, 0.1 M imidazole, 0.1 M 2-(N-morpholino)ethanesulfonic acid pH 6, 17.5–19% glycerol, and 7.5–9% polyethylene glycol 4000. The images for the 7–47 binary complex were indexed, integrated, and scaled using iMOSFLM.

Initial binary complexes were prepared by incubating truncated TNAPs (6 mg mL−1) with 1.2 M equivalents of the annealed P/T2 duplex at 37°C for 30 min. 3 M excess of dtTTP was added and incubated at 37°C for 30 min. The resulting n+1 binary complex was further incubated with 10 M excess tATP at 37°C for another 30 min to yield final ternary complexes. 5–270 ternary complex crystals grew in 0.2 M sodium iodide, 20% polyethylene glycol 3350, and 0.1 M calcium chloride dihydrate, and 8–84 ternary complex crystals grew in 0.2 M sodium iodide, 20% polyethylene glycol 3350, and 0.1 M taurine.

TNAP structural analysis
Molecular visualizations, RMSD, and distance and torsion angle measurements were performed using licensed PyMOL (version 2.5.2; https://www.pymol.org)46 and open-source PyMOL (version 3.1.0; https://github.com/schrodinger/pymol-open-source). Structural alignments for visualization and analysis in Figs. 3 and 4 were carried out using the “align” function in PyMOL, applied to the entire protein–nucleic acid complex. Polder omit maps were generated in Phenix using phenix.polder and visualized in PyMOL47. Local base-pair and base-pair step parameters are computed using Web3DNA48. 2D interaction maps were assembled with data from Mapiya and DNA-proDB. Pocket volumes were calculated using PyVOL.

Integrated mutational analysis
Sequence alignments used to generate the fitness landscape (Supplementary Fig. 3) were performed using the EMBOSS Needle algorithm49 on the EMBL–EBI platform50. Conservation was quantified with BLOSUM62 substitution scores51. Functional effects of reversions to 5–270 were quantified from the 45-s primer extension assay as previously reported11. Band intensities were measured by densitometry in ImageJ52 as the fraction of full-length product (top band) relative to the total pixel density of each lane. Activities were normalized to the performance of 10–92. Mutational effects on folding stability were estimated using FoldX553 with the BuildModel command applied to a repaired Kod exo- closed ternary structure (PDB ID: 5OMF). Reported ΔΔG values are in kcal mol−1. Distances between each mutated alpha carbon atom and the active site, defined by the sugar moiety of the dideoxy primer terminus (C2′ in TNA or C3′ in DNA), were calculated in PyMOL on ternary complexes of Kod, 5–270, and 10–92. Relative solvent-accessible surface area (SASA) was computed on ternary structures using FreeSASA54 with the default parameters. Values were reported as relative exposure (%) in ternary complexes of Kod, 5–270, and 10–92.

AlphaFold3 predictions
AlphaFold355 structural predictions were performed on the online server provided by Google DeepMind (https://alphafoldserver.com/) on 2024-08-16, before the initial public release of the 10–92 ternary structure (PDB ID: 8T3X) on 2024-08-28. Two AlphaFold3 runs (denoted AF3-1 and AF3-2) were performed for every evaluated structure, producing 5 models per run (models 0–4, denoted by subscripts). The binary structural predictions were obtained by inputting the sequence of residues 1–776 in the presence of a DNA primer (CGCGAACTGC), DNA template (AAACGTACGCAGTTCGCG) and 2 Mg2+ ions. Ternary structural predictions were obtained by inputting the sequence of residues 1–760 in the presence of a DNA primer (CGCGAACTGCGT), DNA template (TATGCACGTACGCAGTTCGCG), an ATP, and 2 Mg2+ ions.

2′-deoxy-α-L-threofuranosylthymidine-3′-triphosphate (dtTTP) synthesis
Synthetic procedures. All moisture sensitive reactions were performed in anhydrous solvents under an argon or nitrogen atmosphere. All commercial reagents, solvents and anhydrous solvents were used without further purification. Reaction progresses were monitored by thin-layer chromatography using glass-backed analytical Silica Plate with UV-active F254 indicator. Flash column chromatography was performed with Silica Flash® P60 silica gel (40–63 μm particle size) for most of the crude reaction mixture. Yields are reported as isolated yields of pure compounds. 1H, 13C, and 31P NMR spectra were analysed on 400 MHz NMR spectrometer (Bruker Avance NEO). 1H values are reported in parts per million relative to Me4Si or corresponding deuterium solvents used as an internal standard. 13C values are reported in parts per million relative to corresponding deuterium solvents used as an internal standards. 31P NMR values are reported in parts per million relative to an external standard of 85% H3PO4. Splitting patterns are designated as follows: s, singlet; d, doublet; dd, doublet of doublets; t, triplet; m, multiplet. High-resolution mass spectrometry data were acquired using the electrospray ionization time-of-flight method at the University of California, Irvine Mass Spectrometry Core Facility.

Chemical synthesis, step 1. To a mixture containing the activated nucleoside monophosphate133 (240 mg, 0.6087 mmol, 1 equiv) and 1.2 equiv of 1-(2-(pyrenesulfonyl)ethyl)pyrophosphate (343 mg, 0.7305 mmol) was added, along with 8 equiv of a premade solution of ZnCl2 (664 mg, 4.8670 mmol) in 3.2 mL anhydrous DMF under a nitrogen atmosphere. The mixture was stirred at room temperature for 5 h, and the reaction progress was monitored by HPLC (MeCN/0.05 M TEAB buffer, from 0% to 70% over 42 min). After consumption of the starting material, the product was precipitated by adding the reaction mixture dropwise to stirred diethyl ether (170 mL). The precipitate was collected by centrifuging at 20,000 × g for 5 min at room temperature. The pellet was resuspended in 30 mL 20% H2O in MeCN containing 2% N,N-diisopropylethylamine (DIPEA) and centrifuged at 20,000 × g for 5 min, and the supernatant was collected. The above process was repeated two times, and the combined supernatants were evaporated under diminished pressure. The crude was loaded on a silica gel column (packed with 2% H2O/isopropanol containing 1% DIPEA) by liquid loading (2 mL of 2% H2O/isopropanol containing 1% DIPEA) with eluents 5%–10% H2O/isopropanol containing 1% DIPEA and then 5% H2O in (isopropanol/MeCN 1:1) containing 1% DIPEA. The fractions containing the product were collected and evaporated under diminished pressure at 30–40°C to afford pure 240 mg (0.2758 mmol) fully protected nucleoside triphosphate 2. 31P NMR (162 MHz, D2O) δ −11.93, −12.85 (d, J = 15.6 Hz), −23.10.

Chemical synthesis, step 2. A solution of fully protected nucleoside triphosphate 240 mg (0.2758 mmol) in 30–33% aqueous NH4OH was stirred for 18 h at 37°C in a sealed tube. After the reaction, the solvent was evaporated under diminished pressure. The solid was resuspended with MilliQ water (5 mL) and the aqueous solution was washed with DCM (10 mL) and EtOAc (10 mL). The organic portion was discarded, and the aqueous extract was collected and evaporated under diminished pressure. The crude solid was resuspended with 2 mL of RNAse free water, filtrated by 0.22 μm syringe filter and dropwise added to the forty times volume of acetone (80 mL) at room temperature containing NaClO4 (390 mg, 3.1815 mmol, 15 equiv). The resulting suspension was centrifuged at 20,000 × g for 5 min at room temperature. The supernatant was discarded, and the pellet was washed with organic solution (acetone/DCM, 10:1, 2 × 30 mL) to afford the 1a (85 mg, 0.1577 mmol, 26% two-step yield). 31P NMR (162 MHz, D2O) δ −8.09, −12.42 (d, J = 19.5 Hz), −22.52 (t, J = 19.5 Hz, 1H).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript presents a comprehensive structural and biochemical characterization of the evolutionary trajectory of an engineered polymerase capable of efficiently synthesizing TNA, offering valuable insights into how mutagenesis and selection reshape enzyme architecture and function. These findings not only enhance our understanding of polymerase adaptation but also provide a framework for future protein engineering efforts. By resolving crystal structures of key intermediate variants and correlating them with biochemical data, the authors propose that catalytic efficiency and fidelity can evolve through distinct pathways. While the results are promising, additional clarification and discussion are needed to strengthen some of the conclusions, particularly regarding the mechanistic link between structural changes and functional decoupling. I recommend the manuscript for publication only after the following points have been fully addressed.\nMajor points\n1. In lines 86–87, the manuscript states that “crystal structures were solved as binary and closed ternary complexes, and biochemical data were collected for fidelity, catalytic efficiency, thermal stability, and substrate specificity.” However, limited information is provided about changes in substrate specificity during evolution. Please clarify whether substrate specificity correlates with fidelity or catalytic efficiency.\n2. The manuscript emphasizes significant conformational changes in the finger subdomain and active site during evolution. However, it remains unclear how the accumulation of mutations induces these structural rearrangements. Additionally, electron density maps for key residues in the active site should be provided alongside Figures 3 and 4 to support the structural interpretations. In particular, in Figure 4b, the increased distance from the primer to the α-phosphate in the 5-270 variant, accompanied by the divergent conformation of tATP, requires further explanation.\n3. In lines 104–109 and 281–282, the authors state: “our analysis shows that fidelity increased substantially in earlier generations of the enzyme, whereas strong improvements in catalytic turnover emerge only during the later stages of evolution. Although not widely discussed in the literature, this observation implies that mutations acquired by directed evolution can differentially affect fidelity and catalysis, challenging the conventional view that these traits evolve in parallel as coupled events.” However, Figure 1 shows that in the later stages of evolution both activity and fidelity increase substantially from variant 8-64 to 10-92, which seems inconsistent with complete decoupling. Could the authors comment on this apparent discrepancy? Additionally, beyond reference 16, are there other studies supporting the “conventional view” of coupled evolution?\n4. Variant 10-92, which carries five additional mutations compared to 8-64 (Extended Data Figure 2), shows a sharp increase in activity. How do these mutations lead to local or global structural rearrangements that enhance catalytic efficiency? Are these changes reflected in the active site conformations presented in Figures 3 and 4?\n5. Structural alignments are presented in Figures 3 and 4. Please specify whether these alignments were performed using the overall structure, individual protein domains, or DNA/TNA components. Since the alignment strategy can substantially influence the interpretation of structural differences, a clear description of the methodology is necessary.\n6. The authors state that “the closed binary motif first appears in the 5-270 intermediate and is absent in the binary structure of Kod-RSGA” (lines 184–185). This suggests that the closed binary motif arises during the transition from Kod-RSGA to 5-270. Please elaborate on how these structural changes contribute to the marked improvement in fidelity observed in the early evolutionary stages. A clearer comparison between the structural features of Kod-RSGA and 5-270 would help clarify the mechanistic basis for this enhancement.\nMinor points\n1. In Figure 1b, the term “sequence space” should be clearly defined in the legend or the main text.\n2. In Supplementary Figure S6, please clarify whether the structure corresponds to wild-type KOD, the 10-92 variant, or another form. If a previously published structure was used, the relevant PDB code should be provided. In addition, the description in lines 150–152 is currently unclear and should be revised for clarity.\n3. In Figures 2f, 2g, 2h, and S8, all interactions are depicted using dashed lines. However, some represent hydrogen bonds and others hydrophobic interactions. These should be distinguished visually to avoid confusion. For example, hydrogen bonds could be shown as dashed lines, while hydrophobic interactions should be depicted differently.\n4. In Figure 2g, the residue labeled T267 appears to actually be I268. Please verify and correct.", "reviewer_2": "Hajjar and colleagues present an extensive crystallographic analysis of directed evolution intermediates found in the development of an efficient TNA synthetase. I put below my comments roughly in the order they appear in the manuscript, with more important comments highlighted (*).\n*1. Difference between n=1 and generalizations. The authors repeatedly claim they are demonstrating evolutionary principles based on the single evolutionary path they analysed, without discussing how the selection pressure may have influenced the evolutionary path. I don’t think that is needed and anchoring the path to polymerase biochemistry will not detract from the novelty of the paper.\n*2. The differential impact on fidelity and catalysis for polymerases is widely discussed in the literature, but rarely in B-family thermophilic proteins. There is extensive literature available on Pol beta, which I tried to summarise (and integrate with DE efforts) here: 10.1016/j.copbio.2018.11.008.\n3. Impact of 51 mutations. While I agree that the screening methodology actively excludes polymerases that are no longer thermostable, I think the nature and location of the mutations can also be taken into consideration. Surface residues (therefore not part of significant contact networks) and conservative mutations that are present in similar thermostable proteins, would not be expected to significantly affect protein stability.\n4. TNAP kinetics. It would be useful to add the reaction buffer composition rather than rely on reference to previous work. I also wondered if the authors ever tried to optimise the reaction buffer (beyond supplementing reactions with Mn2+). Some of the enzymes I have worked with had shifted optima Mg concentration, which is often ignored by some in the field.\n*5. TNAP Fidelity. The substrate used to determine fidelity has biases (no GG, known to affect fidelity of TNA polymerases), therefore the values obtained do not sample all possible dinucleotide transitions and may be overestimating fidelity. In addition, based on our own calculations, 1000 bases are not enough data to claim differences between 99.1% and 99.9% incorporation accuracy. What are the confidence intervals for the determined fidelities?\n*6. Crystallization poses. While I find very interesting that the evolutionary intermediates crystallized in a “closed” conformation, can this not simply be a crystallization artifact? How do the crystallization conditions of these variants compare to the conditions the starting variant (Kod-RI/Kod-RSGA) was crystallized? Have the authors ever explored how these proteins behave in solution? For the avoidance of doubt, I think the authors need to acknowledge and discuss this point in the manuscript. I am not asking for single-molecule FRET experiments which would be the only way to demonstrate that.\n*7. Link between fidelity and catalysis. Given the title, I looked forward to a more extended discussion on these differences for polymerases. If the crystallization pose is relevant in solution, pre-organising binary complex could suggest that the initial gains in TNA incorporation are coming from stabilising the closed conformation of the enzyme. That would be expected to reduce the fidelity of the polymerase, unless that step is also affecting the enzyme’s processivity (measured as bound time rather than number of incorporations per binding). In line with that comment, I disagree that the prevailing assumption in the field is that fidelity and catalysis go hand-in-hand for polymerases.\n8. Figure 3 (but also applicable to other single amino acid superposition figures). Presenting the relative positioning of the residues without making reference to the electron density and B-factor of the residue in those positions in the crystal reduces the quality of the comparison. Including those directly in the figure may make it too busy, but that could be included in SI.\n*9 “…importance of long-range structural… geometry”. Unfortunately, ref. 29 does not support that statement. Ref. 29 simply lists how residues are in non-sequential networks and the size and distance may depend on the local structure.\n10. AlphaFold. It may be worth including in the main text that AlphaFold3 was used and tempering the statement. AlphaFold2 was trained differently and would have been far more susceptible to homology matches – justifying the side chain positioning as per training data. AlphaFold3 should be less susceptible to homology matches but it provides a static pose, much like crystallography. Relaxing the poses can have a significant impact on the argument that AI cannot deliver accurate structures (or at least meaningful structural insights).\nVitor Pinheiro", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response: We thank the reviewer for their positive assessment of the study.\nMajor points\n1. In lines 86–87, the manuscript states that “crystal structures were solved as binary and closed ternary complexes, and biochemical data were collected for fidelity, catalytic efficiency, thermal stability, and substrate specificity.” However, limited information is provided about changes in substrate specificity during evolution. Please clarify whether substrate specificity correlates with fidelity or catalytic efficiency.\nResponse: In the context of this study, substrate specificity refers to the ability of the polymerase to distinguish TNA versus DNA substrates based on their catalytic rates of incorporation. The main text was updated to clarify this point and Supplementary Figure 7 was revised with an additional panel showing the rates of synthesis along with a matching bar graph of substrate specificity.\n2. The manuscript emphasizes significant conformational changes in the finger subdomain and active site during evolution. However, it remains unclear how the accumulation of mutations induces these structural rearrangements. Additionally, electron density maps for key residues in the active site should be provided alongside Figures 3 and 4 to support the structural interpretations. In particular, in Figure 4b, the increased distance from the primer to the α-phosphate in the 5-270 variant, accompanied by the divergent conformation of tATP, requires further explanation.\nResponse: Supplementary Figures 1 and 2 were added as new images showing global changes in the protein architecture between 10-92 and its closest natural homolog, Kod DNA polymerase. The structural differences first appeared in 5-270 and are retained in 10-92. This pattern suggests that the changes arose through recombination and represent a defining structural signature of the engineered polymerase.\nAdditionally, the manuscript was revised to include a discussion of the mutational landscape underlying the activity of 10-92. This new section, and accompanying figures, describe the mutations and/or structural changes responsible for improved fidelity and catalytic activity.\nLast, the addition of polder maps to Figures 3 and 4 yielded a change to the image that was too small to visualize at 100% magnification. Recognizing the importance of this information, we prepared supplementary figures showing the electron density of key residues in the active site, as well as electron density of the nascent base pair (dT:tATP) and adjacent template-primer region. In the case of 5-270, we analyzed a second crystal structure with an alternate tATP conformation, which provided a primer to a-phosphate distance of 5.7 Å. The main text was updated with this new information, which we attribute to flexibility within the increased size of the active site pocket.\n1\n3. In lines 104–109 and 281–282, the authors state: “our analysis shows that fidelity increased substantially in earlier generations of the enzyme, whereas strong improvements in catalytic turnover emerge only during the later stages of evolution. Although not widely discussed in the literature, this observation implies that mutations acquired by directed evolution can differentially affect fidelity and catalysis, challenging the conventional view that these traits evolve in parallel as coupled events.” However, Figure 1 shows that in the later stages of evolution both activity and fidelity increase substantially from variant 8-64 to 10-92, which seems inconsistent with complete decoupling. Could the authors comment on this apparent discrepancy? Additionally, beyond reference 16, are there other studies supporting the “conventional view” of coupled evolution?\nResponse: We did not intend to suggest that fidelity and catalysis are fully decoupled, but rather that substantial improvements in each trait arise at different stages of the directed evolution pathway. As the reviewer correctly notes, the strong gain in catalytic activity coincides with a second increase in fidelity. To clarify this point, we have re-written this section of the results to temper our language and better frame the relationship between fidelity and catalysis. Finally, additional references were added to support the claim that fidelity is expected to improve with faster catalysis.\n4. Variant 10-92, which carries five additional mutations compared to 8-64 (Extended Data Figure 2), shows a sharp increase in activity. How do these mutations lead to local or global structural rearrangements that enhance catalytic efficiency? Are these changes reflected in the active site conformations presented in Figures 3 and 4?\nResponse: A single-point reversion analysis described previously implicates the T548P mutation and to a lesser extent the P550H mutation as sequence changes that are likely responsible for improved catalytic activity. Since global architectural changes were largely fixed at 5-270, these mutations appear to fine tune the active site through their proximity to the catalytic aspartates (D405, D541, and D543). The manuscript and supplementary material were updated to clarify this point.\n5. Structural alignments are presented in Figures 3 and 4. Please specify whether these alignments were performed using the overall structure, individual protein domains, or DNA/TNA components. Since the alignment strategy can substantially influence the interpretation of structural differences, a clear description of the methodology is necessary.\nResponse: All structural alignments were performed by global alignment of the C-alpha residues. The figure legends and Methods were updated to clarify this point.\n6. The authors state that “the closed binary motif first appears in the 5-270 intermediate and is absent in the binary structure of Kod-RSGA” (lines 184–185). This suggests that the closed binary motif arises during the transition from Kod-RSGA to 5-270. Please elaborate on how these structural changes contribute to the marked improvement in fidelity observed in the early evolutionary stages. A clearer comparison between the structural features of Kod-RSGA and 5-270 would help clarify the mechanistic basis for this enhancement.\nResponse: The manuscript was revised to include a new supplementary figure (Supplementary Figure 12), comparing the structures of Kod-RSGA and 5-270. Additionally, the results section was modified to include a discussion of the mutational landscape underlying the activity of 10-92, which delineates mutational and/or structural changes impacting fidelity and activity.\nMinor points\n1. In Figure 1b, the term “sequence space” should be clearly defined in the legend or the main text.\nResponse: The text was revised to define the term ‘sequence space’.\n2\n2. In Supplementary Figure S6, please clarify whether the structure corresponds to wild-type KOD, the 10-92 variant, or another form. If a previously published structure was used, the relevant PDB code should be provided. In addition, the description in lines 150–152 is currently unclear and should be revised for clarity.\nResponse: The legend for Supplementary Figure 8 was revised to clarify that the structure shown corresponds to that of wildtype Kod DNA polymerase (PDB ID: 5OMF).\n3. In Figures 2f, 2g, 2h, and S8, all interactions are depicted using dashed lines. However, some represent hydrogen bonds and others hydrophobic interactions. These should be distinguished visually to avoid confusion. For example, hydrogen bonds could be shown as dashed lines, while hydrophobic interactions should be depicted differently.\nResponse: Figure 2 (panels f, g, and h) and Supplementary Figure 12 were updated to delineate the different types of interactions observed in the active site.\n4. In Figure 2g, the residue labeled T267 appears to actually be I268. Please verify and correct.\nResponse: The figure was updated to correct the mistake.\nREVIEWER 2\nHajjar and colleagues present an extensive crystallographic analysis of directed evolution intermediates found in the development of an efficient TNA synthetase. I put below my comments roughly in the order they appear in the manuscript, with more important comments highlighted (*).\nResponse: We thank the reviewer for their positive assessment of the study.\n*1. Difference between n=1 and generalizations. The authors repeatedly claim they are demonstrating evolutionary principles based on the single evolutionary path they analysed, without discussing how the selection pressure may have influenced the evolutionary path. I don’t think that is needed and anchoring the path to polymerase biochemistry will not detract from the novelty of the paper.\nResponse: We agree that our analysis is based on a single evolutionary trajectory, and we were careful to acknowledge this limitation in the revised manuscript. Our goal was to use this trajectory as an example of how evolutionary pressures can shape the emergence of specific biochemical traits in a polymerase scaffold. The manuscript was revised to clarify this point by discussing the imposed selection pressure, which in this case, involved progressively reducing the incubation time to favor improved catalysis. We agree that future studies analyzing alternate trajectories will be important to assess the generality of our observations.\n*2. The differential impact on fidelity and catalysis for polymerases is widely discussed in the literature, but rarely in B-family thermophilic proteins. There is extensive literature available on Pol beta, which I tried to summarise (and integrate with DE efforts) here: 10.1016/j.copbio.2018.11.008.\nResponse: The manuscript was revised to include a discussion of the differential impact of mutations on polymerase fidelity and catalytic activity. This included a new section on the mutational landscape underlying the activity of 10-92 as well as accompanying figures, describing the mutations and/or structural changes responsible for improved fidelity and catalytic activity.\n3. Impact of 51 mutations. While I agree that the screening methodology actively excludes polymerases that are no longer thermostable, I think the nature and location of the mutations can also be taken into consideration. Surface residues (therefore not part of significant contact networks) and conservative mutations that are present in similar thermostable proteins, would not be expected to significantly affect protein stability.\nResponse: The manuscript was revised to include a deeper analysis of the mutations responsible for improved fidelity versus those responsible for improved catalysis. In addition to textural changes, this includes a new main text figure summarizing an integrated analysis of each mutation.\n4. TNAP kinetics. It would be useful to add the reaction buffer composition rather than rely on reference to previous work. I also wondered if the authors ever tried to optimise the reaction buffer (beyond supplementing reactions with Mn2+). Some of the enzymes I have worked with had shifted optima Mg concentration, which is often ignored by some in the field.\nResponse: The methods section was revised to include the composition of ThermoPol buffer from NEB, which was used for all polymerase reaction described in the study. We have not attempted to optimize the reaction buffer further, but could consider this option in the future.\n*5. TNAP Fidelity. The substrate used to determine fidelity has biases (no GG, known to affect fidelity of TNA polymerases), therefore the values obtained do not sample all possible dinucleotide transitions and may be overestimating fidelity. In addition, based on our own calculations, 1000 bases are not enough data to claim differences between 99.1% and 99.9% incorporation accuracy. What are the confidence intervals for the determined fidelities?\nResponse: The template used for the fidelity assay was chosen to allow comparison with our past work. The sequencing results indicate a clear improvement in fidelity as the polymerase progresses along the evolutionary trajectory. While future studies may explore this question in more detail, for example, using NGS approaches to evaluate the fidelity of TNA replication from thousands of different templates, as was the case for our earlier study on information storage, we believe that the current experiment adequately addresses the fidelity of 10-92. We also wish to point out that it is common practice in the field to perform this assay on a single template with data deriving from 1000 nucleotide incorporation events.\n*6. Crystallization poses. While I find very interesting that the evolutionary intermediates crystallized in a “closed” conformation, can this not simply be a crystallization artifact? How do the crystallization conditions of these variants compare to the conditions the starting variant (Kod-RI/Kod-RSGA) was crystallized? Have the authors ever explored how these proteins behave in solution? For the avoidance of doubt, I think the authors need to acknowledge and discuss this point in the manuscript. I am not asking for single-molecule FRET experiments which would be the only way to demonstrate that.\nResponse: We agree that crystallographic artifacts should always be considered when evaluating new protein conformations by X-ray crystallography. However, in this case, there are multiple lines of evidence arguing against the closed binary conformation being an artifact of crystallization. First, binary structures of the TNAP variants in question (5-270, 7-47, 8-64, and 10-92) crystallized under two distinct precipitation conditions. The reoccurrence of this conformation under two different crystallization conditions suggests that the conformation reflects a bone fide structural feature rather than a condition-specific artifact. Second, the closed binary conformation is absent in Kod-RI and 4\nKod-RSGA, further strengthening the view that the conformational difference arises from sequence changes rather than crystallization chemistry. Finally, the closed binary conformation emerges in variants that also exhibit changes in their biochemical activity, providing a functional correlation that supports its structural relevance. The manuscript was updated to clarify this point. No solution-based experiments have been undertaken to further explore this point, but could be worthwhile exercises in the future.\n*7. Link between fidelity and catalysis. Given the title, I looked forward to a more extended discussion on these differences for polymerases. If the crystallization pose is relevant in solution, pre-organising binary complex could suggest that the initial gains in TNA incorporation are coming from stabilising the closed conformation of the enzyme. That would be expected to reduce the fidelity of the polymerase, unless that step is also affecting the enzyme’s processivity (measured as bound time rather than number of incorporations per binding). In line with that comment, I disagree that the prevailing assumption in the field is that fidelity and catalysis go hand-in-hand for polymerases.\nResponse: The manuscript was revised to clarify the relationship between fidelity and catalysis. The prevailing view is that a selection for improved catalytic activity through the use of progressively shorter incubation times should, indirectly, enrich for enzymes with higher fidelity. This is because less faithful polymerases are more likely to stall at misincorporation sites, thereby failing to complete the extension within the allotted time. Thus, variants that maintain rapid synthesis under stringent incubation times are expected to combine enhanced catalytic efficiency with increased fidelity.\n8. Figure 3 (but also applicable to other single amino acid superposition figures). Presenting the relative positioning of the residues without making reference to the electron density and B-factor of the residue in those positions in the crystal reduces the quality of the comparison. Including those directly in the figure may make it too busy, but that could be included in SI.\nResponse: The manuscript was updated to include polder maps of key residues and nucleotide conformations in the active site. These data provide confidence that the observed structural changes correspond to regions of high experimental confidence.\n*9 “…importance of long-range structural… geometry”. Unfortunately, ref. 29 does not support that statement. Ref. 29 simply lists how residues are in non-sequential networks and the size and distance may depend on the local structure.\nResponse: Reference 29 (now ref 35 in the revised manuscript) is a review on the importance of long-range interactions in protein folding.\n10. AlphaFold. It may be worth including in the main text that AlphaFold3 was used and tempering the statement. AlphaFold2 was trained differently and would have been far more susceptible to homology matches – justifying the side chain positioning as per training data. AlphaFold3 should be less susceptible to homology matches but it provides a static pose, much like crystallography. Relaxing the poses can have a significant impact on the argument that AI cannot deliver accurate structures (or at least meaningful structural insights).\nResponse: The Results and Methods sections of the manuscript were revised to clarify that Alpha Fold3 was used for the analysis. Additionally, we redesigned Supplementary Figure 15 and added Supplementary Figure 17 to visualize AlphaFold3 predictions.\nVitor Pinheiro\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse: We thank the reviewer for their positive assessment of the study. Their comments were addressed in one of the reports provided above.\n6"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have thoroughly addressed most of the concerns raised in the initial review, and the manuscript has been significantly strengthened. The study presents valuable insights, and I recommend acceptance after final revision. The following are two remaining minor suggestions to further improve the manuscript:\n1. In Figure 2, short dashed lines denote electrostatic interactions, while long dashed lines show hydrophobic interactions, which helps distinguish between different types of interactions. This notation should be applied consistently in Figure 6 and Supplementary Figure S12.\n2. In the response, the authors state that \"substrate specificity refers to the ability of the polymerase to distinguish TNA versus DNA substrates based on their catalytic rates of incorporation.\" Supplementary Figure S7 shows a sharp rise in TNA catalytic activity between variants 8-64 and 10-92, which coincides with a notable enhancement in substrate specificity. To elucidate the structural basis for the evolutionary changes in fidelity and catalytic activity, the manuscript offers a thorough structural analysis of the catalytic core, including active site organization, interactions and so on, while lack of structural comparison regarding how the enzyme's interactions with dA and tA change during the evolution from the dA to tA substrate. Including such an analysis is essential to fully clarify the mechanism.", "reviewer_2": "I thank the authors for the changes introduced in the manuscript.\nPoints raised here (where possible) use the same numbering as in the original comments.\n#5 - TNAP fidelity: This point remains unaddressed. The field's common practice has shifted significantly in the last decade. If the authors persist in applying this approach, it would still benefit from a statistical test to measure the significance of the findings. A test to compare 2 Poisson distributions (using individual reads as the estimate of an error rate) would be the bare minimum.\n#7 - fidelity vs. catalysis. Based on my own experience with Tgo, KOD and Phi29 evolution campaigns, gains in processivity dominate early detection in selection campaigns. Here, increases in fidelity delivered processivity faster than improved catalysis as it would be expected from the search pattern, which started from homologous recombinations. Nevertheless, this point is academic and requires no change to the manuscript.\n#9 - While I agree that the review is of importance, it does not support the statement being made by the authors that \"long-range interactions support the fine-tuning of active site geometry\". My particular disagreement here is focused on the use of \"fine-tuning\". The review supports that long-range interactions can be implicated in catalysis and functional specificity.\nI don't expect further changes to the manuscript, but I leave it to the editors whether the points raised here would need to be addressed before publication.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "RESPONSE TO THE REVIEWER’S COMMENTS\nReviewer #1:\nThe authors have thoroughly addressed most of the concerns raised in the initial review, and the manuscript has been significantly strengthened. The study presents valuable insights, and I recommend acceptance after final revision. The following are two remaining minor suggestions to further improve the manuscript:\n1. In Figure 2, short dashed lines denote electrostatic interactions, while long dashed lines show hydrophobic interactions, which helps distinguish between different types of interactions. This notation should be applied consistently in Figure 6 and Supplementary Figure S12.\nResponse: Figure S12 was updated to include long dashes for the hydrophobic interactions. However, we decided not to change Figure 6, because some of the lines, especially those between amino acid side chains and DNA bases, contain both hydrophobic and electrostatic interactions.\n2. In the response, the authors state that \"substrate specificity refers to the ability of the polymerase to distinguish TNA versus DNA substrates based on their catalytic rates of incorporation.\" Supplementary Figure S7 shows a sharp rise in TNA catalytic activity between variants 8-64 and 10-92, which coincides with a notable enhancement in substrate specificity. To elucidate the structural basis for the evolutionary changes in fidelity and catalytic activity, the manuscript offers a thorough structural analysis of the catalytic core, including active site organization, interactions and so on, while lack of structural comparison regarding how the enzyme's interactions with dA and tA change during the evolution from the dA to tA substrate. Including such an analysis is essential to fully clarify the mechanism.\nResponse: We agree that it would be interesting to study how the enzyme changed its substrate specificity from DNA to TNA. However, such an analysis would require crystal structures of the evolved enzymes with the natural substrate, which is beyond the scope of the current study. Nevertheless, we do hope to provide such information in a future study.\nReviewer #2:\nI thank the authors for the changes introduced in the manuscript.\nPoints raised here (where possible) use the same numbering as in the original comments.\n#5 - TNAP fidelity: This point remains unaddressed. The field's common practice has shifted significantly in the last decade. If the authors persist in applying this approach, it would still benefit from a statistical test to measure the significance of the findings. A test to compare 2 Poisson distributions (using individual reads as the estimate of an error rate) would be the bare minimum.\nResponse: We updated Figure S6 to include statistical data with and without insertions and deletions (indel). The comparison between Kod-RSGA and 5-270 with and without indels was significant in both cases. This result is consistent with the interpretation that fidelity improved during the early stages of evolution, while gains in catalysis arose later in the selection. For consistency, the axis in Figure 1d was also updated to include the fidelity value with indels.\n1\n#7 - fidelity vs. catalysis. Based on my own experience with Tgo, KOD and Phi29 evolution campaigns, gains in processivity dominate early detection in selection campaigns. Here, increases in fidelity delivered processivity faster than improved catalysis as it would be expected from the search pattern, which started from homologous recombinations. Nevertheless, this point is academic and requires no change to the manuscript.\nResponse: We are not aware of any prior work in the field where kinetic rates and fidelity have been examined in detail across an evolutionary trajectory for an engineered polymerase, and as such, have no basis of comparison for a competing model.\n#9 - While I agree that the review is of importance, it does not support the statement being made by the authors that \"long-range interactions support the fine-tuning of active site geometry\". My particular disagreement here is focused on the use of \"fine-tuning\". The review supports that long-range interactions can be implicated in catalysis and functional specificity.\nResponse: We do not feel that this is a valid concern, as the mutations are nearly all located at long-distance positions and their accumulation in the protein sequence resulted in changes in the enzyme active site that were visualized by protein X-ray crystallography. Thus, our use of the term “fine-tuning” is consistent with other examples in the literature, where mutations accumulated by directed evolution leads to improvements in substrate recognition.\nI don't expect further changes to the manuscript, but I leave it to the editors whether the points raised here would need to be addressed before publication.\nReviewer #3:\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse: We thank the reviewer for their positive assessment of the manuscript.\n2"}, {"round": 3, "reviewer_comments": {"reviewer_2": "I thank the authors for adding the statistical test. It would have been good to include that result also in Figure 1d. Other than that, I think the research is ready for publication.\nBest wishes,\nVitor Pinheiro"}, "author_response": ""}]

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