Filament assembly induced by the recognition of target DNA activates the prokaryotic Argonaute SPARDA system

Domain: biology
Published: 2026-01-06
Editorial outcome: major_revision

Abstract
The short prokaryotic Argonaute (pAgo) proteins, in conjunction with their associated effector molecules, constitute a defence mechanism in prokaryotes that protects against phage infections. The SPARDA is characterized by its collateral nuclease activity, which, upon activation through target DNA recognition, non-specifically cleaves a wide range of nucleic acid substrates. Nevertheless, the structural underpinnings of its collateral activity have remained elusive. In this study, we investigate the NbaSPARDA system from Novosphingopyxis baekryungensis and reveal that RNA-guided DNA recognition triggers the assembly of a higher-order filamentous structure. This filamentation is essential for the tetramerization of the DREN nuclease domain, which in turn facilitates the accumulation and cleavage of substrate nucleic acids. Through the determination of the gRNA-bound and RNA-DNA duplex-bound cryo-EM structures, we delineate a sequential monomer-dimer-monomer-filament transition during SPARDA activation. These insights collectively elucidate a filament-dependent activation mechanism underpinning the short pAgo-mediated immune response, which is crucial for antiviral defence.

Introduction
Argonaute (Ago) proteins are a conserved family that is widely distributed across all domains of life. Eukaryotic Ago (eAgo) combines with 22-nt guide RNA (gRNA) to form the essential component of the RNA-induced silencing complex (RISC) in the RNA interference (RNAi) pathway, which silences the target RNA that is paired to the gRNA1,2. Extensive structural studies on Ago have demonstrated that canonical eAgo is composed of four structural domains (N, PAZ, MID, and PIWI), which are connected by two linkers (L1 and L2)3,4. The MID and PAZ domains are responsible for the binding of the 5’ and 3’ ends of gRNA and target DNA (tDNA), respectively5. The RNase H fold of PIWI possesses a DEDX catalytic triad to cleave the target. The architecture of eAgo displays a bilobed conformation in which the N and PAZ domains form one lobe, whereas the MID and PIWI domains form the other lobe6,7. The guide–target heteroduplex is bound in the central channel formed by the N-PAZ and MID-PIWI lobes8.

Unlike eAgo, the roles of their homologues in prokaryotes (pAgo) are not fully understood, despite pAgo being widely found in prokaryotes and archaea9,10. Compared with eAgo, pAgo is more diverse in terms of its domain composition and role in target cleavage5,10. Moreover, short pAgo is the largest group of pAgo, accounting for more than 60%10. However, short pAgo has only the MID domain and a catalytically inactive PIWI domain, which has led to the loss of its ability to cleave target nucleic acids10. Interestingly, recent studies revealed that short pAgo and neighbouring genes that encode fusion versions of proteins with a C-terminal APAZ (Analogue of PAZ) domain and diverse N-terminal effectors, including TIR, Sir2, and DUF3457, function together as antiviral systems in response to phage infection10,11. Furthermore, pAgo-related systems can be classified into four categories, in which the mechanisms used to defend against phage infection are distinct11–13. To date, the mechanisms underlying the majority of short pAgo-mediated antiphage have been elusive. Among the short pAgo systems, cellular NAD+ was rapidly depleted in the SPARTA14–18 and SPARSA19–22 systems because of the recognition of the tDNA of the gRNA in the positively charged channel formed by the APAZ and the pAgo. In the three-component pAgo of Sulfolobus islandicus (SiAgo), the activation of the associated transmembrane Aga2 effector can result in membrane depolarization and cell death23.

The SPARDA (Short Prokaryotic Argonaute and DNAse/RNase-Endonuclease) antiviral system is a newly identified antiphage system in which APAZ is fused with a nuclease domain24,25. Unlike eAgo proteins, in which the PIWI nuclease domain specifically cleaves paired targets6, SPARDA of Novosphingopyxis baekryungensis (NbaSPARDA) and Thermocrispum municipal (TmuSPARDA) unexpectedly exhibited collateral nuclease activity against various nucleic acid substrates dependent on the DREN nuclease domain upon target recognition24,25. However, the detailed mechanism of the assembly mechanism and the activation of the collateral indiscriminate nuclease activity of the DREN remains to be investigated. In this study, we demonstrate that fully paired tDNA can induce SPARDA filament formation; furthermore, we employ cryogenic electron microscopy (cryo-EM) to elucidate the tDNA-induced filament assembly of NbaSPARDA. A comparison of assembled and unassembled SPARDA structures reveals that filament assembly stabilizes the nuclease effector DREN and creates a binding channel for the substrate. Interestingly, the heteroduplex formed by gRNA and its complementary tDNA is located within the DREN domain, demonstrating the essential role of filament formation in activating SPARDA’s collateral nuclease activity.

Results
Overall structure of the gRNA-bound SPARDA
SPARDA comprises a short pAgo and the cognate DREN–APAZ (Fig. 1A), which can assemble into a heterodimer24. To investigate the SPARDA assembly mechanism, we first attempted to determine its apo structure using cryo-EM. Size-exclusion chromatography (SEC) and SDS-PAGE analyses confirmed the stable heterodimer formation between pAgo and its cognate DREN–APAZ (Supplementary Fig. 1A, B). Analytical ultracentrifugation (AUC) of the sample yielded a molecular weight of 114 kDa (Supplementary Fig. 1C), consistent with the SEC findings and indicative of a 1:1 stoichiometric complex similar to those of other pAgo homologues17–19,21,26. Unfortunately, preferential orientation during cryo-EM visualization hindered the structural resolution of SPARDA (Supplementary Fig. 2).

Based on an earlier study in which gRNA and complementary tDNA can induce the collateral nuclease activity of SPARDA24, we incubated a precise 20-nt gRNA with purified SPARDA to form the gRNA–SPARDA complex. Consistent with prior findings, the introduction of paired tDNA enabled the processing of a wide spectrum of substrates, including ssDNA, dsDNA and RNA (Fig. 1B). Cryo-EM analysis of the gRNA–SPARDA complex revealed a significant improvement in sample homogeneity with the addition of gRNA. Surprisingly, 2D classification indicated that gRNA-bound SPARDA adopted a dimeric conformation, distinct from the monomeric state of apo SPARDA (Supplementary Fig. 3). SEC analysis confirmed the formation of the gRNA–SPARDA complex (Supplementary Fig. 4A). Accordingly, the AUC results of gRNA–SPARDA yielded a calculated molecular weight of approximately 200 kDa (Supplementary Fig. 4B), which also reconfirmed that gRNA induces dimerization of SPARDA.

Finally, we resolved the cryo-EM structure of the gRNA–SPARDA complex at 2.77 Å resolution (Fig. 1C, D). The final map revealed that the N-terminal DREN nuclease domain was not visible, likely because of high flexibility, whereas only the first six nucleotides of the seed region of the gRNA could be traced (Fig. 1D), possibly because the remainder is solvent exposed, akin to the homologues in Ago proteins14,19. The structure contains two pAgo subunits and two copies of the cognate DREN–APAZ (Fig. 1D).

Like conventional Ago family homologues, each pAgo subunit adopts a canonical architecture, comprising an N-terminal MID domain (residues 1–250) and a C-terminal PIWI domain (residues 251–485). Although the DREN domain was not visible, the C-terminal APAZ domain (residues 161–442) presents a classical mixed α/β fold that resembles those in other short pAgo homologues7. The APAZ domain is positioned to the PIWI domain, forming a characteristic heterodimer with a buried interface of approximately 1,200 Å². Overall, the APAZ–pAgo heterodimer has an architecture similar to that of previous pAgo-related systems, generating a positively charged cavity to accumulate nucleic acids (Supplementary Fig. 5). Structural analysis revealed that the β1-strand, along with the C-terminal β12 and β11 strand protruding from the APAZ domain, forms a hand-like fold to lock the PIWI domain of pAgo (Fig. 1E). Hydrogen-bonding contacts were identified such that H410 from the last β-sheet of APAZ forms hydrogen bonds with T5 located in the nearby β-sheet of the MID domain. Residues L165 and N167 from the β1-β2 loop of APAZ establish hydrogen bonds with L376 and R374 located in the surrounding loop of the PIWI domain. In addition, R326 of the β11-sheet of APAZ forms a hydrogen bond with T372 in the PIWI domain (Fig. 1E). Complementary hydrophobic packing further reinforces the interface.

Like typical Ago family proteins, the gRNA is bound within the MID domain of both pAgo–APAZ heterodimers. In addition, Mg2+ was coordinated between the 5’-phosphate of the gRNA and the phosphate group of the third nucleotide (adenine) (Fig. 1F), providing a structural basis for the previously observed inability of SPARDA to cleave substrates in the absence of Mg2+24. The 5’-phosphate group is further recognized by the side chains of R183 and K240 in the MID domain (Fig. 1G), which is consistent with the conserved arginine–lysine motif in typical Ago proteins5,7. The first adenine of the gRNA is inserted into a hydrophobic pocket formed by aromatic amino acids W159, W201 and F182 in the MID domain and establishes a hydrogen bond with the side chain of N181 (Fig. 1G). The second nucleotide, U2, forms hydrogen bonds with the side chains of Q224 and N232, whereas A3 interacts with Q224. Notably, the 2’-OH groups of U2, A3, and C4 form hydrogen bonds with multiple residues in the MID domain (Fig. 1G), potentially explaining the preference of SPARDA for RNA over ssDNA as a guide24.

Structural basis for the dimerization assembly of SPARDA induced by gRNA
The addition of gRNA promoted the dimerization of the SPARDA heterodimer. We subsequently performed structural analysis on the interfaces between the individual SPARDA heterodimers. Cryo-EM structure analysis revealed that gRNA binding repositions the two APAZ domains in a head-to-tail orientation (Fig. 2A), facilitating the formation of hydrogen bonds. In the APAZ interfaces, Q353 forms a hydrogen bond with the side chain of R260, and F354 packs against F202 to form π‒π stacking interactions. Identical interactions occur at the symmetrical interface of the opposing APAZ domain (Fig. 2B). Similarly, the two pAgo subunits adopt a head-to-tail orientation mediated primarily by their PIWI domains. In particular, three loops protruding from the β-sheet core of each PIWI domain drive dimerization. Loop 1 (residues 166–181), Loop 2 (residues 265–275) and Loop 3 (residues 293–338) facilitate these interactions. Specifically, R301 in Loop 1 of pAgo1 forms hydrogen bonds with the side chains of E170 and T168 and the main chain of G176 in Loop 3 of pAgo2. Additionally, T298 and A296 in Loop 1 of pAgo1 establish hydrogen bonds with Q272 and Q273 in Loop 2 of pAgo2, with similar interactions observed between the Loop 1 of pAgo2 and the Loop 2 and Loop 3 in pAgo1 (Fig. 2C).

To investigate the role of gRNA-induced dimerization in regulating the collateral nuclease activity of SPARDA, we performed site-directed mutagenesis targeting key amino acids in SPARDA. Substitution of R183A/K187A/K240A, which are involved in the recognition of the 5’-phosphate group within the MID domain, significantly impaired SPARDA’s nuclease activity. Similarly, mutations in gRNA-binding sites critical for A1 recognition (F182A/N181A/W159A/W201A) or 2’-OH contacts (S447A/N451A) abolish the nuclease activity of SPARDA (Fig. 2D). To assess whether gRNA-triggered dimerization is necessary for SPARDA nuclease activity, we simultaneously mutated the residues involved in the APAZ–APAZ interfaces and those at the interface between pAgo–pAgo subunits. Nuclease activity assays demonstrated that simultaneous mutation of these critical residues nearly abolished the ability of SPARDA to cleave nucleic acid substrates (Fig. 2D). Collectively, these findings indicate that gRNA-induced dimerization is essential for SPARDA activation.

tDNA recognition induces the formation of SPARDA filaments
To further elucidate the mechanism of DREN activation upon tDNA recognition of the gRNA, we initially incubated purified SPARDA with gRNA and complementary tDNA. However, no density for the gRNA‒tDNA duplex was observed in the final map. We inferred that the gRNA–tDNA heteroduplex failed to be loaded in the SPARDA. Thus, we implemented an annealing step by heating the gRNA and tDNA to 95°C for 2 minutes and then slowly cooling it to room temperature. The annealed gRNA‒tDNA duplex was added to SPARDA at a ratio of 1:2 and subjected to cryo-EM. Unexpectedly, 2D classification (Supplementary Fig. 6) revealed that the micrographs exhibited regular helical filaments in all views, indicating that tDNA triggered the formation of a filamentous architecture in SPARDA (Fig. 3A). Analysis by size SEC and AUC demonstrated that gRNA–tDNA–SPARDA forms a supramolecular complex with a molecular mass of ~3.28 MDa (Supplementary Fig. 7A, B). Accordingly, in vitro nuclease activity assays established that collateral cleavage activity is exclusively observed in the high-molecular-weight fraction corresponding to the filamentous form (Supplementary Fig. 7C). In vivo BiFC assays further showed that SPARDA assembles into filamentous structures upon exogenous plasmid challenge (Supplementary Fig. 8). Collectively, these results suggested that tDNA recognition of the gRNA induces filament formation in SPARDA.

To better understand the link between SPARDA filamentation and collateral nuclease activity, we determined the structure of the complex of SPARDA bound to the gRNA and tDNA duplex through cryo-EM at 2.64 Å. The structure revealed that the activated SPARDA complex assembles into a regular helical filament with a helicase rise of approximately 130 Å and a diameter of approximately 180 Å (Fig. 3B). Each repeating unit of the SPARDA filament comprises four pAgo and four associated APAZ-DREN proteins. The four pAgo molecules are arranged as the backbone of the filament, whereas the cognate APAZ domains are pointed to the periphery of the filament (Fig. 3B). Notably, the N-terminal DREN domains, previously unresolved in inactive SPARDA, were unambiguously modelled within the SPARDA filament, where they were held together to further assemble into a tetrameric configuration (Fig. 3B). As observed in other Ago family proteins8,17,19, tDNA and gRNA are bound within a specific nucleic acid-binding lobe formed by pAgo and the APAZ domain.

In the gRNA–tDNA-bound filamentous SPARDA complex, the 20-nt gRNA and its complementary tDNA could be traced (Fig. 3B, C). Consistent with the recognition mode in the canonical Ago protein, the gRNA forms near-complete Watson‒Crick base pairing with the tDNA, except for the first two 5′-terminal nucleotides. The 5′ phosphate of gRNA inserts into a binding pocket within the MID domain, while the remaining nucleotides extend into solvent-exposed regions, accounting for the lack of discernible density beyond this region. Overall, the recognition pattern between gRNA and tDNA is similar to that of other Ago family proteins in which the 5’ end of gRNA binds within the nucleic acid binding pocket of the MID domain, while other nucleotides are solvent exposed. The 3’ terminus of tDNA is anchored between the MID of pAgo and the APAZ domain, whereas its 5’ end extends toward the solvent-facing direction of the APAZ domain. The recognition of gRNA depends on base-specific recognition, which primarily relies on the 5’-terminal nucleotides of gRNA and the residues located in the MID domain of pAgo, and the nonspecific backbone contacts between the phosphate groups of the gRNA/tDNA backbones and the resides across the nucleic acid binding tunnel (Fig. 3C).

Structural basis for the formation of SPARDA filaments
In the SPARDA filament, pAgo serves as the backbone, with the basic structural unit comprising four pAgo molecules. Two central pAgo molecules are oriented towards each other through their MID domains (Fig. 3B). However, the DREN–APAZ subunits exhibit two distinct states: an extended state and a compact state. In the extended conformation, the DREN and APAZ domains are separated by a distance of approximately 20 Å, resulting in no density for the connecting linker (Fig. 3D). Conversely, in the compact state, the linker folds back towards the DREN domain, enabling clear observation of its density (Fig. 3E). Superimposition of the dimeric SPARDA and the compact-state pAgo–DREN–APAZ of the SPARDA filament revealed obvious steric hindrance between the guide-target heteroduplex and the other pAgo molecule in the dimeric SPARDA complex (Fig. 3F), suggesting that the dimeric of gRNA-bound SPARDA transformed into a monomer, which then formed a filament during the propagation of guide-target duplex. In comparison with the pAgo in the gRNA-bound dimeric SPARDA and the core dimeric pAgo in the filamentous SPARDA reveals a substantial structural rearrangement. Superimposition of one pAgo molecule in the dimeric SPARDA and that within the SPARDA filament, it could be observed a rotation of approximately 90 degrees in the other molecule. In the gRNA-bound dimeric SPARDA complex, dimer formation is primarily mediated by interactions between the PIWI domains of adjacent pAgo molecules. In contrast, within the activated SPARDA filament, the dimer interface is largely reconfigured and stabilized predominantly by interactions between the MID domains of adjacent molecules, as well as through amino acid contacts formed between the PIWI domain of one subunit and the MID domain of its neighbouring subunit.

Despite the overall structural similarity of pAgo and DREN–APAZ in the gRNA-bound SPARDA and the SPARDA filament, subtle local differences were observed, primarily in regions responsible for nucleic acid binding in pAgo and the C-terminal portion in DREN–APAZ. In particular, the region comprising residues 160–178 in pAgo, which was invisible in the gRNA-bound SPARDA, was stabilized by the binding of tDNA (Supplementary Fig. 9A). In addition, the loop containing residues 269–302, which is crucial for nucleic acid recognition, undergoes significant reorientation (Supplementary Fig. 9A). In filamentous SPARDA, this loop was extended outwards in active SPARDA, which ultimately formed the PIWI–PIWI interface that is essential for filamentation. Local conformational changes were also observed in the region responsible for gRNA 3’-end binding of APAZ, enabling coordination with the extended tDNA strand during filament formation (Fig. 3F).

At the MID–MID interface, R244 forms hydrogen bonds with the backbone of G140 and the side chains of R142 and E97 (Fig. 3G). The central pAgo molecule interact with adjacent Ago molecules through the MID and the PIWI domains (Fig. 3G). The PIWI domain of pAgo engages with the MID domain of neighbouring pAgo, while the MID domain connects to the PIWI domain of another Ago. At the PIWI–MID interface, G140 and R142 form hydrogen bonds with E284 and D480 (Fig. 3G). In the PIWI–PIWI interaction interface, K324 participates in a charged interaction with E360, R287 forms another charged interaction with E253, and R285 establishes a hydrogen bond with G254; additionally, T329 and L330 form hydrogen bonds between the symmetric residues (Fig. 3H).

To investigate the functional significance of these interfaces, we introduced alanine substitutions at key residues within the MID–MID (E97A/G140A/R142A/R244A) and MID–PIWI (G140A/R142A/E284A/D480A) interfaces and evaluated the impact of these mutations on SPARDA using in vitro collateral cleavage assays. The results indicated that individual mutations had negligible effects, whereas simultaneously mutating key residues in both the MID–MID and MID–PIWI interfaces impaired SPARDA’s nuclease activity (Fig. 3I). Given that pAgo forms the backbone of the SPARDA filament, the conformational changes in the 269–302 region are likely to be critical for SPARDA filament assembly formation. Therefore, we introduced mutations into amino acids mediating PIWI–PIWI interactions within this region. Individual mutations did not markedly affect SPARDA function; however, simultaneous mutation of R285, R287, and K324 disrupted substrate cleavage and abolished the system’s ability to confer resistance against bacteriophage infection (Fig. 3I, J). Collectively, these results demonstrate that tDNA-induced SPARDA filament formation occurs in the active form, which is essential for its collateral nuclease activity.

Mechanism of the activation of the DERN nuclease effector
The DREN domain (residues 1–145) is connected to the C-terminal APAZ domain by a flexible loop (146–166). In contrast, in the activated SPARDA filament assembly, the DREN domain can be modelled. The DREN domain is composed of seven β-strands surrounded by three α-helices. In each repeating unit of SPARDA, four DREN protomers assemble into a tetramer with D2 symmetry (Fig. 4A, B). However, the nuclease catalytic sites (E16, D41, Q60, and K62) of the two extended DREN-2 and 3 protomers (displayed as translucent surfaces) are positioned to the pocket formed by the tetramer; in contrast, the catalytic sites of the other DREN-1 and 4 (displayed as cartoons) are buried and exposed to the other side of DREN (Fig. 4B). Additionally, two regions of extra density are observed in the DREN domain, which can be assigned as the gRNA–tDNA duplex (Fig. 4C). The RNA‒DNA duplex is stabilized by interactions between phosphate groups and residues within the activated DREN domain (Fig. 4D, E). Specifically, the binding of tDNA and gRNA to the DREN 2 and DREN 3 domains exhibits a mirror-image relationship. This structural arrangement allows tDNA to enter the catalytic amino acid pocket of DREN 3, which is composed of residues E16, D41, Q60, and K62, while the gRNA simultaneously occupies the corresponding catalytic pocket in DREN 2 (Fig. 4D, E). This configuration highlights the structural consistency underlying the recognition and cleavage of a broad spectrum of nucleic acid substrates.

To elucidate the molecular basis of DREN tetramer assembly, we characterized the intermolecular interactions among DREN protomers. DREN-1, DREN-2 and DREN-3 are utilized to analyse their interactions. Structural analysis revealed that these two regions predominantly contributed to their interactions. Specifically, the adjacent DREN protomers engage through their α1 helices, bringing residues E13 and R20 into proximity. Reciprocal hydrogen bonds form between E13 of one protomer and R20 of its neighbour, and vice versa, thereby stabilizing helix‒helix packing (Fig. 4F). Moreover, the β-sheet core of DREN-1 is oriented towards the β-sheet of DREN-3, forming interfacial interactions mediated by the residues in the β1–β2 strands of DREN-2. At the β1–β2 interface, residues located on β1 and β2 and the loop connecting them to the DREN molecule form a hydrogen bond network with the same regions in the other DREN. Specifically, D31 on β1 and E45 on β2 of DREN 2 form hydrogen bonds with R33 on the β1–β2 loop and Y88 on α2 of DREN 3, respectively. Additionally, R33 on the β1–β2 loop of DREN 2 form hydrogen bonds with D31 and E45 on β2 of DREN 3 (Fig. 4G). Structural homology analysis using Dali revealed that the endonuclease I-Ssp6803I is most similar to DREN. Notably, I-Ssp6803I also adopts a D2-symmetric tetrameric arrangement to cleave double-stranded DNA (dsDNA)27 (Supplementary Fig. 9B).

In the apo SPARDA, as predicted by AlphaFold 3, the N-terminal DREN was located outside the gRNA–tDNA binding channel. Similarly, in the gRNA–bound SPARDA complex, although the DREN domains themselves were unresolved in the structure, the orientations of their linkers suggested that the two DREN domains were spatially separated (Fig. 1C). In the activated SPARDA–gRNA–tDNA filament, the four DREN molecules exhibit nearly identical conformations to each other, but they are stabilized and further oligomerized within the SPARDA filament induced by tDNA recognition. Deletion of the DREN domain did not alter the filament formation induced by tDNA recognition and the gRNA according to the SEC elution file and the corresponding appearance on the micrographs (Supplementary Fig. 10). Based on these observations, we proposed that SPARDA filament-driven tetramerization of DREN is critical for its nuclease activity. We subsequently performed structure-guided mutagenesis of these interacting residues involved in DREN oligomerization. The results revealed that single-point mutations had a negligible impact, however, group alanine substitution on the resides involved in tetramerization (E13A/N17A/R20A/R33A/Q29A/E45A) significantly abolished both the nuclease activity against the substrate according to the in vitro nuclease assays and its capacity to defend against bacteriophages (Fig. 4H, I, Supplementary Fig. 11). Taken together, these findings demonstrate that the unique filament architecture of SPARDA drives DREN oligomerization, which is indispensable for substrate recognition and catalytic function (Fig. 5).

Methods
Plasmid construction
The NbaAgo gene (GenBank accession: WP_022673743.1) and the DREN-APAZ gene (WP_033317603.1) from the NCBI database were synthesized by Sangon Biotech (Shanghai, China). For expression of the NbaAgo/DREN-APAZ complex in E. coli, His-tagged NbaAgo and S-tagged DREN-APAZ were cloned into the first and second multiple cloning sites of pETDuet-1, respectively. Site-directed mutagenesis to generate mutant complexes was performed on the NbaAgo/DREN-APAZ protein-expression plasmid using whole-plasmid PCR amplification. The nucleic acid sequences used in this study are listed in Supplementary Data 1.

Protein expression and purification
The corresponding recombinant plasmids were transformed into E. coli BL21(DE3) cells. The transformants were inoculated in LB medium supplemented with 100 µM Amp and grown for approximately 6 h at 37 °C. When the OD600 reached 0.6–0.8, the cultures were induced with 0.4 mM IPTG and shifted to 16 °C for 16-18 h. The cells were collected by centrifugation and resuspended in buffer containing 20 mM Tris-HCl, pH 8.0, and 500 mM NaCl. After lysis by sonication on ice, the lysate was centrifuged at 17,000 × g for 30 min at 4 °C. The supernatant was then loaded onto a Ni2+-NTA column (Qiagen) pre-equilibrated with lysis buffer. The column was washed with 10 column volumes (CV) of equilibration buffer to remove unbound proteins. This was followed by a wash with 10 CV of equilibration buffer supplemented with 20 mM imidazole to remove weakly bound contaminants. Finally, the His-tagged protein complex was eluted with equilibration buffer supplemented with 300 mM imidazole. The eluate was then diluted fivefold with a buffer containing 20 mM Tris-HCl (pH 8.0) and 5% glycerol. The supernatant was loaded onto a 5 ml heparin-Sepharose column (Cytiva) pre-equilibrated with buffer containing 20 mM Tris-HCl (pH 8.0), 150 mM NaCl, and 2 mM DTT. The column was washed with the same buffer, and the protein complex was eluted using a linear NaCl gradient (from 150 mM to 1 M) in the identical buffer. Subsequently, the eluted protein was concentrated to 0.5 ml using a 30K-cutoff concentrator (Millipore), which was purified with a Superdex 200 increase column (Cytiva) equilibrated with buffer containing 20 mM Tris-HCl (pH 8.0), 150 mM NaCl and 2 mM DTT. The major chromatographic peak was fractionated in 0.5 ml aliquots, pooled, and concentrated for storage at −20 °C. Mutant complexes were purified following the same procedure as described above.

In vitro assembly of the SPARDA-gRNA-tDNA complex
To assemble SPARDA-gRNA complex, the purified SPARDA complex was incubated with guide RNA and target DNA at a molar ratio of 1:2:2. The mixture was then incubated at 4 °C for 30 min and loaded onto a Superdex 200 increase column (Cytiva). Fractions containing the SPARDA-gRNA-tDNA complex were pooled, flash frozen in liquid nitrogen, and stored at −80 °C.

Analytical ultracentrifugation (AUC)
Sedimentation velocity experiments were performed using an Optima analytical ultracentrifuge (Beckman Coulter, Inc, Brea, CA, USA) equipped with an absorbance optical system set to 280 nm. Samples were loaded into double-sector centerpieces and analyzed in an An-60 Ti rotor at 45,000 × g and 10 °C. All measurements were conducted in a buffer containing 20 mM Tris-HCl (pH 8.0) and 150 mM NaCl. The following complexes were characterized: (i) SPARDA complex, (ii) the gRNA-bound SPARDA complex, and (iii) the gRNA-tDNA SPARDA complex. Data were acquired at 280 nm with a radial spacing of 0.003 cm. Sedimentation coefficient distributions [c(s)] were determined using SEDFIT software (version 16.1c) with default parameters.

Cryo-EM sample preparation and data collection
For the SPARDA-gRNA complex dataset, aliquots (3 µL) of a 1.8 mg/mL mixture were loaded onto glow-discharged (80 s at 20 mA) holey carbon Quantifoil grids (AU 2/1, 300 mesh, Quantifoil Micro Tools) using a Vitrobot Mark IV (Thermo Fisher Scientific) at 100% humidity and 4 °C. For the SPARDA-gRNA-tDNA complex dataset, the sample was prepared and vitrified under identical conditions. All data were acquired using the EPU software on Titan Krios G4 transmission electron microscopy (Thermo Fisher Scientific) operated at 300 kV. Images were recorded using a Gatan K3 detector in a 48-frame movie mode at a nominal 130,000 × magnification with a pixel size of 0.325 Å, and the total electron dose was set to 48 e−·Å−2, with an exposure time of 1.35 s.

Image processing and 3D reconstruction
Drift and beam-induced motion correction were performed with MotionCor239 to produce a micrograph from each movie. Contrast transfer function (CTF) fitting and phase-shift estimation were conducted with Gctf40. Micrographs with astigmatism, obvious drift, or contamination were discarded before reconstruction. The following reconstruction procedures were performed using Cryosparc V341. In brief, particles were automatically picked using the “Blob picker” or “Filament Tracer”. Several rounds of reference-free 2D classifications were performed and the selected “good” particles were then subjected to ab initio reconstruction, heterogeneous refinement, and final non-uniform refinement. Localized refinement focusing of the local areas, were also performed if necessary. The resolutions of all density maps were determined by the gold-standard Fourier shell correlation curve, with a cutoff of 0.143442. Local resolutions of maps were estimated with ResMap43.

Atomic model building, refinement, and 3D visualization
The initial models of the NbaSPARDA complex were acquired from Alphafold344. We initially fitted the models into the corresponding final cryo-EM maps using Chimera45, and further corrected and adjusted them manually by real-space refinement in Coot46. The resulting models were then refined with phenix.real_space_refine in PHENIX47. These operations were executed iteratively until the problematic regions, Ramachandran outliers, and poor rotamers were either eliminated or moved to favoured regions. The final atomic models were validated with Molprobity48. All structural figures were generated with ChimeraX45.

Phage Efficiency of Plating (EOP) Assay
The efficiency of plating (EOP) assay was performed using E. coli BL21(DE3) cells harbouring the pET-Duet-1 vector expressing wild-type or mutant SPARDA. For the spot assay, overnight cultures of the expression strain were diluted 1:100 in MMB medium (LB supplemented with 0.1 mM MnCl₂, 5 mM MgCl₂, 5 mM CaCl₂) containing 0.5% agar and grown at 37 °C with shaking until OD₆₀₀ reached 0.3. Protein expression was induced with 0.3 mM IPTG, and incubation continued until OD₆₀₀ reached 0.6. Then, 500 µl of induced culture was mixed with 20 mL of molten MMB medium containing 0.2 mM IPTG and 100 µg/mL ampicillin to form bacterial lawns. Serially diluted phage T5 lysates (10-fold gradients) in MMB medium were spotted (4 µl per droplet) onto the lawns. Plates were incubated at room temperature for 24 hours before plaque counting and analysis.

Cleavage assays
Cleavage assays were performed to determine the activity of the SPARDA complex toward ssDNA, dsDNA and RNA substrates under various reaction conditions. Typically, 500 nM SPARDA complex was preincubated with 200 nM guide RNA for 15 min at 37 °C in a buffer containing 20 mM Tris-HCl (pH 8.0), 0.1 mg/ml BSA, 2 mM DTT and 10 mM Mg2+. Subsequently, 200 nM target DNA was added to the reaction mixture, followed by an additional 15 minute incubation. Substrate oligonucleotides (ssDNA, dsDNA and RNA) were then added to a final concentration of 100 nM. The reactions were incubated for 30 min, and then an equal volume of loading buffer containing 8 M urea, 20 mM EDTA and 2 × TBE prepared from a commercial 10 × TBE stock (Invitrogen). Reaction products were separated by 20% denaturing polyacrylamide gel (containing 8 M urea) and visualized by staining with SYBR Gold and scanning using an Amersham Typhoon imager (Cytiva).

Bimolecular fluorescence complementation assays
The superfolder green fluorescent protein (sfGFP) reporter was split into two distinct fragments: the large β1–β10 barrel (residues 1-214) and the small β11 strand (residues 215-239). The sfGFP β1–β10 fragment was fused to the DREN. Subsequently, an expression cassette encoding the sfGFP β1–β10-DREN-APAZ, and pAgo was cloned into the multiple cloning site of the pRSF-Duet-1 vector. Similarly, the sfGFP β11 fragment was fused to DREN, and the corresponding sfGFP β11-DREN-APAZ-pAgo cassette was assembled into the pET-Duet-1 vector. These two recombinant plasmids were then co-transformed with a third plasmid, pCDFDuet-1, into E. coli BL21(DE3) competent cells. Positive transformants were selected on LB agar plates containing ampicillin, kanamycin, and spectinomycin. A single colony was inoculated into LB liquid medium supplemented with the same antibiotics and grown at 37 °C with shaking until the OD₆₀₀ reached ~0.6. Protein expression was induced by adding 0.3 mM IPTG, followed by incubation at 16 °C for 16 h. Finally, cells were then harvested and resuspended in PBS buffer, and subjected to imaging using a Leica STELLARIS confocal microscope.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The formation of a filamentous structure by the Ago complex upon target recognition is an intriguing phenomenon. However, a study reporting a similar structure of the same protein was published five months ago (Cell Research (2025) 35: 510–519; Received: 26 November, Published online 24 March 2025). Judging from the workload in the current manuscript, the study could reasonably have been completed within this time frame. The only major difference is that, in the published study, the DREN nuclease domain is bound to tDNA, whereas in the current work it is bound to a gRNA–tDNA heteroduplex. This difference may simply result from sample preparation: in the published study, the Ago complex was first incubated with gRNA and then with excess tDNA, leading to tDNA occupying the DREN active site, while in the present work it was incubated directly with excess gRNA–tDNA heteroduplex. However, the novelty of this observation is insufficient to support publication in Nature Communications, particularly as the authors do not present data demonstrating the functional significance of this difference.\nThe biochemical assays and in vivo anti-phage experiments are also limited, and the manuscript does not clearly highlight findings that distinguish it from the previously published work. For functional validation, the authors employed combined mutants with substitutions at 4–6 amino acid residues, an approach that is overly coarse. This design does not allow the contribution of individual residues to be resolved—some may be critical, while others may not. Moreover, introducing multiple substitutions may have complex effects on activity, and at a minimum, the authors should exclude the possibility that these combined mutations disrupt secondary structure, thereby abolishing activity.\nFinally, the study appears to follow the anti-phage assay setup from the previous structural paper (Cell Research, 2025), in which T5 phage was used. However, in the first report identifying this system (Nature Microbiology, 2024, 9: 1368–1381), the system did not confer resistance to T5 phage but rather to λ phage. Whether this system provides resistance to T5 phage therefore requires careful verification.", "reviewer_2": "The manuscript presents a comprehensive structural and functional analysis of the prokaryotic Argonaute SPARDA system, revealing a filament-dependent activation mechanism. This work complements prior studies (NbaSPARDA, XauSPARDA, and SPARHA) and advances our understanding of prokaryotic Argonaute-associated anti-phage defense systems.\nAs the manuscript contains several grammatical, typographical, and formatting errors in the text and figures, A thorough, line-by-line revision is recommended prior to publication.\nBelow are specific suggestions:\n1. Lines 212–213: The statement “suggesting that the dimer of SPARDA was disrupted during the propagation of guide-target duplex, which is the prerequisite for SPARDA filamentation” appears inconsistent. The authors suggest that dimer disruption occurs spontaneously during filament formation, yet mutagenesis experiments disrupting the dimer interface abolish activity entirely. This apparent contradiction requires clarification.\n2. Lines 297–301: The comparison with SPARTA/SPARSA could be expanded to discuss the evolutionary implications of distinct effector mechanisms.\n3. Fig. 1G: Not cited in the main text.\n4. Supplementary Fig. 3A: Not cited.\n5. Supplementary Fig. 5: Not cited.\n6. Fig. S7B: Should be explicitly cited in lines 214–223.\n7. Line 121: Remove the redundant “Besides,” before “Additionally.”\n8. Fig. 1E: Residue R326 is missing from the labeling.\n9. Lines 213/397: Standardize nomenclature to consistently use “SPARDA” (instead of “SPARD” or “SPADA”).\n10. Fig. 1E: Add dashed lines for hydrogen bonds between residues H410 and T5.\n11. Fig. 1E: Label β1, β2, β11, and β12.\n12. Fig. 1E: Show the main chains for residues involved in backbone hydrogen bonds (e.g., L165–L376).\n13. Figs. 1G and 3C: Apply consistent domain-specific coloring for residues to reflect domain organization.\n14. Fig. S7 legend: Contains several errors. For example, correct “Ssp6301” to “Ssp6803I” and replace “heighted” with “highlighted.”\n15. Line 283: The phrase “according to the SEC elution file and the negative staining by cryo-EM” is unclear. Negative staining is a technique used in electron microscopy but not in cryo-EM. Please clarify the intended meaning.", "reviewer_3": "Review of “Filament assembly induced by the recognition of target DNA activates the prokaryotic Argonaute SPARDA system” by Zhang et al.\nThe manuscript by Zhang et al. presents the structural basis for filament assembly of NbaSPARDA upon recognition of guide RNA and target DNA. This filament formation is required to activate the DREN module of SPARDA, thereby initiating nuclease activity. The authors provide structural and biochemical data that shed important mechanistic insights into how SPARDA defends against phages and plasmids. Overall, this is a valuable contribution to the field. The structural data are of generally good quality and support the central claims, although several discrepancies and missing details need to be addressed.\nThe article would benefit from revisions as outlined below:\nMajor Concerns\n1. Incomplete or unclear Cryo-EM data presentation\nSeveral Cryo-EM datasets mentioned in the text are not shown or are insufficiently described, making it difficult to evaluate the claims. For instance:\n- Lines 91–92 mention the preferred orientation of the SPARDA monomer, but no supporting data are provided. The data processing workflow, including representative micrographs and 2D class averages, should be included. It would also be useful to provide a low-resolution 3D reconstruction of the monomer.\n- Lines 97–100 suggest that a separate dataset for SPARDA+gRNA was collected, but this is not clearly described in the Methods. It currently appears that all reconstructions were derived from a single dataset.\n- Supplementary Figures S2 and S5 show identical micrograph images, raising concerns. It appears that filament formation may also occur in the SPARDA+gRNA complex, which would contradict the authors’ claims. Micrographs for SPARDA alone should be provided to confirm whether filaments are also present.\n- Lines 166–167 imply that a 3D reconstruction of apo SPARDA was obtained from a different dataset, but this is not shown anywhere in the manuscript. Both the data processing workflow and the resulting 3D reconstruction should be included.\n2. The micrographs indicate that not all particles form filaments; a significant fraction of monomeric or dimeric particles or maybe tetramers appear to remain. It is unclear whether these contain nucleic acids, and if so, whether they include only gDNA or the duplex. The authors should clarify this. Importantly, it should be determined whether filament formation is essential for activity or whether only a subset of complexes engage in nuclease activity. And if so are these from the filament.\n3. A close-up view of the cryo-EM density for the gRNA should be shown at the same threshold used for the rest of the structure. This would help establish whether the reconstruction contains only gRNA or the gRNA+tDNA duplex. As currently described, the sample composition used for the reconstruction is unclear.\n4. The Introduction or Discussion would benefit from a more detailed description of what is known about DREN domains, including their structural conformations and oligomerization requirements in other systems. A comparative structural analysis of multiple active DREN domains, including SPARDA, would help readers appreciate the significance of tetramerization.\n5. It would considerably strengthen the manuscript if the authors could demonstrate filament formation in vivo during phage infection. For example, fluorescently labeled SPARDA expressed in cells could be used to visualize puncta corresponding to filamentous structures upon infection. While this experiment is not essential for publication, it would greatly enhance the physiological relevance of the findings if performed within a reasonable timeframe.\n6. The authors should provide more details on potential structural changes at the pAgo dimerization interface during filament assembly. Is this interface altered upon filament formation? Could dimerization play a regulatory role in nuclease activity by conferring specificity to the presence of tDNA? The current data chould be used to elaborate on this point.\n7. The SEC profile in Supplementary Figure S6 shows an additional peak at ~14 mL. Does this fraction also contain SPARDA bound to gRNA and tDNA? If duplex-bound SPARDA does not necessarily form filaments, is there evidence for tetramer formation in this fraction? Did the authors try a nuclease activity assay with this sample from the 14ml fraction compared to the 8ml fraction. This would be an important experiment.\n8. The cryoEM data processing workflow is not sufficiently detailed. For example, the FSC curves as well as local resolution maps are missing for the structures described. The authors should expand on the data processing showing ab initio reconstructions as well as further processing steps of possible 3d classification and refinemnt.\nMinor Comments\n1. The dimensions of the helical rise and diameter (lines 179–180) should be indicated in the figure.\n2. Line 213 contains a typo: “SPRADA” should be corrected to “SPARDA.”\n3. The sentence in lines 80–81 is unclear and should be rephrased for clarity."}, "author_response": "Response: We thank the reviewer for this valuable suggestion. As rightly pointed out, this represents the first instance where filament assembly and nuclease activation have been observed in short pAgo-associated defense systems, including NbaSPARDA and other homologous systems such as those from Enhydrobacter aerosaccus (EaeSPARDA) and Thermocrispum municipale (TmuSPARDA). These studies collectively demonstrate that, within this clade of short pAgos, the guide RNA-target DNA heteroduplex triggers filament formation of SPARDA proteins, thereby activating their collateral nuclease activity against a broad range of substrates.\nAnother notable feature of SPARDA is its broad substrate spectrum. As reported in the original study 1, it can cleave ssDNA, dsDNA, ssRNA, and RNA-DNA heteroduplexes. Coincidentally, while our manuscript was in preparation, a related study in Cell Research2 was reported that detailed SPARDA activation on ssDNA. A key distinction in our work is the successful capture of a pre-hydrolysis complex structure of SPARDA bound to an RNA-DNA heteroduplex, providing structural evidence for its hydrolytic mechanism for both DNA and RNA. Compared to other studies reporting different substrate interactions with the DREN nuclease, our research reveals a unique activation mode for SPARDA when binding to an RNA-DNA heteroduplex. This finding, along with our detailed elucidation of the common hydrolytic mechanism across various substrates, offers significant insights into the bacterial defense mechanisms within the short pAgo system. Additionally, after reevaluating the data based on feedback from other reviewers, we reprocessed the SPARDA monomer data from the gRNA-tDNA-SPARDA dataset, resulting in a relatively high-quality map suitable for rigid-body fitting. Unexpectedly, this map revealed the presence of a bound gRNA-tDNA heteroduplex, whereas the gRNA-SPARDA dimer complex clearly showed only gRNA binding. These results support our model in which SPARDA initially exists as a monomer, first assembles into a dimer upon gRNA binding, subsequently dissociates into monomers induced by target DNA, and ultimately reorganizes into active filaments.\nTogether, the two studies reveal a filament assembly-driven activation mechanism in bacterial defense SPARDA systems, which undergoes a monomer–dimer–monomer–filament transition during guide RNA loading, target DNA engagement, and propagation of the guide–target heteroduplex. We believe that the integration of our work with the published study will significantly advance the understanding of the filamentation-mediated mechanisms in short pAgo-based bacterial antiviral systems.\nThe biochemical assays and in vivo anti-phage experiments are also limited, and the manuscript does not clearly highlight findings that distinguish it from the previously published work. For functional validation, the authors employed combined mutants with substitutions at 4–6 amino acid residues, an approach that is overly coarse. This design does not allow the contribution of individual residues to be resolved—some may be critical, while others may not. Moreover, introducing multiple substitutions may have complex effects on activity, and at a minimum, the authors should exclude the possibility that these combined mutations disrupt secondary structure, thereby abolishing activity.\nResponse: We thank the reviewer for their valuable suggestion. To investigate the functional significance of key residues, we generated a series of point mutations. Our results demonstrate that the D41A mutation at the catalytic site of DREN completely abolishes both collateral cleavage activity and phage defense. To examine the role of amino acids involved in DREN tetramerization on SPARDA's collateral cleavage activity and phage defense, we conducted both single-point and combined mutations. Specifically, we generated single-point mutations at key residues (E13A, R20A, N17A, R33A, Q29A, E45A) implicated in DREN oligomerization. However, these individual mutations showed no significant effect. Consequently, we created a combined mutant by simultaneously altering all these residues. This multiple mutation significantly abolished both the nuclease activity in vitro and the system's ability to defend against bacteriophages. Additionally, we generated a series of individual point mutations at the MID-MID (E97A, G140A, R142A, R244A), MID-PIWI (G140A, R142A, E284A, D480A), and PIWI-PIWI (R285A, R287A, K324A) interfaces, however, all the mutants had no significant effect on its activity. Finally, a combined mutant on the interfacial residues abolished the related activity. The results of the single-point mutations have been included in the Supplementary figure 11.\nFurthermore, the reviewer expressed concern that mutating several amino acids simultaneously might affect the protein's structure. During the purification of these proteins, we found that the A260/A280 ratios of these mutants, as determined by size exclusion chromatography, were close to that of the WT. This indicates that these mutants retain the ability to bind gRNA and tDNA. Therefore, we speculate that these mutations do not adversely affect the protein itself. We have provided a representative size exclusion chromatography profile comparing a typical SPARDA multi-mutant as shown in the figure below (A260/A280 ~1). We appreciate the reviewer's question.\nFinally, the study appears to follow the anti-phage assay setup from the previous structural paper (Cell Research, 2025), in which T5 phage was used. However, in the first report identifying this system (Nature Microbiology, 2024, 9: 1368–1381), the system did not confer resistance to T5 phage but rather to λ phage. Whether this system provides resistance to T5 phage therefore requires careful verification.\nResponse: We are grateful to the reviewer for raising this point. In response, we would like to clarify our experimental approach regarding phage selection. We indeed followed the protocol from the original study (Nat Microbiol. 2024, PMID: 38622379) by testing NbaSPARDA against a panel of phages (λ, T1, T2, T4, T5). However, efficiency of plating (EOP) assays did not observe a reduction in lambda phage plating efficiency by NbaSPARDA. Instead, a pronounced reduction in EOP was specifically observed for phage T5. It is important to note that the λ phage infection data in that study (Fig. 4) were performed using the CmeSPARDA ortholog from Cupriavidus metallidurans. Conversely, the same study reported that NbaSPARDA itself exhibited significant antiphage activity against T5 phage, which directly supports and validates our choice of T5 phage for the experiments presented in our manuscript. We thank the reviewer for the opportunity to clarify this point.\nReviewer #2 (Remarks to the Author):\nThe manuscript presents a comprehensive structural and functional analysis of the prokaryotic Argonaute SPARDA system, revealing a filament-dependent activation mechanism. This work complements prior studies (NbaSPARDA, XauSPARDA, and SPARHA) and advances our understanding of prokaryotic Argonaute-associated anti-phage defense systems.\nAs the manuscript contains several grammatical, typographical, and formatting errors in the text and figures, A thorough, line-by-line revision is recommended prior to publication.\nResponse: We sincerely thank the reviewers for their positive and encouraging feedback on our work. Our findings on SPARDA, together with other recent reports 2-4, underscore the functional importance of filament assembly in this clade of pAgos-a mechanism that appears distinct from other recently characterized short pAgo systems such as SPARTA and SPARSA. We believe these collective advances deepen the understanding of pAgo protein diversity and their role in prokaryotic immunity. Furthermore, we have carefully addressed all the specific points raised by the reviewers, revised the manuscript accordingly, and corrected any typographical errors. We hope that the revised version now fully meets with the reviewers' approval.\nBelow are specific suggestions:\n1. Lines 212–213: The statement “suggesting that the dimer of SPARDA was disrupted during the propagation of guide-target duplex, which is the prerequisite for SPARDA filamentation” appears inconsistent. The authors suggest that dimer disruption occurs spontaneously during filament formation, yet mutagenesis experiments disrupting the dimer interface abolish activity entirely. This apparent contradiction requires clarification.\nResponse: We thank the reviewer for raising this important point. Our results indicate that SPARDA initially exists as a monomer, which then homodimerizes upon guide RNA recognition. This dimeric intermediate subsequently undergoes filamentation upon binding of target DNA and guide RNA. We therefore hypothesize that the guide RNA-bound homodimeric state represents a key intermediate prerequisite for filament assembly in the presence of a complementary target DNA.\nIn the revised manuscript, we revised this sentence to avoid any confusion to the authors as “suggesting that the dimeric of gRNA-bound SPARDA transformed into a monomer, which then formed a filament during the propagation of guide-target duplex”.\n2. Lines 297–301: The comparison with SPARTA/SPARSA could be expanded to discuss the evolutionary implications of distinct effector mechanisms.\nResponse: We thank the reviewer for the valuable comments to improvement of our manuscript. In the revised version of the manuscript, we discussed the similarity and the unique features of SPARDA and the prior short pAgo including SPARTA and SPARSA. Please see line 316 and 331: “Recent studies have demonstrated that pAgo family proteins, particularly short pAgos, play an essential role in combating phage infections 11,13,23. A featured characteristic of the pAgo systems is that the effector domains are fused to the APAZ domain via a long linker encoded by the pAgo-associated partner, whose enzymatic activities are obtained upon the recognition of guide and target nucleic acids. SPARTA, SPARSA and SPARDA belong to different phylogenetic clades. Both SPARTA and SPARSA utilize NADase-containing effectors that hydrolyse NAD+ upon RNA-guided recognition of target DNA, thereby leading to cell death to provide resistance against phage infections. Subsequent structural studies revealed that inactive SPARTA17 and SPARSA19 adopt a monomer in solution in the absence of target nucleic acids. The activation of their NADase activity depends on the tetramerization of the TIR domain in SPARTA and the Sir2 domain in SPARSA, which is induced by target ssDNA binding. In SPARTA and SPARSA, the active tetrameric architecture is formed by interfaces involving the pAgo subunit and the enzymatic domains located outside of the nucleic acid-binding channel. In SPARDA, the activation of the nuclease of DREN relies on its assembly into filaments upon the recognition of the guide RNA and target DNA. The tetramerization-dependent activation of DREN is similar to those of SPARTA and SPARSA, in which the Sir2 and TIR domains assemble into a tetramer upon nucleic acid binding.”\n3. Fig. 1G: Not cited in the main text.\nResponse: We thank the reviewer for pointing out this error. The citation for Fig. 1G has been added to the relevant results section (Lines 132–138).\n4. Supplementary Fig. 3A: Not cited.\nResponse: We thank the reviewer for pointing out this error. The Supplementary Figure 3A has been relocated to Supplementary Figure 4A, and this reference has been added to the relevant results section (Lines 103–104).\n5. Supplementary Fig. 5: Not cited.\nResponse: We thank the reviewer for pointing out this error. The Supplementary Fig. 5 has been relocated to Supplementary Figure 6, and this reference has been added to the relevant results section (Lines 173–174).\n6. Fig. S7B: Should be explicitly cited in lines 214–223.\nResponse: We thank the reviewer for pointing out this error. The Supplementary Fig. 7B has been relocated to Supplementary Figure 9A, and this reference has been added to the relevant results section (Lines 234–236).\n7. Line 121: Remove the redundant “Besides,” before “Additionally.”\nResponse: We thank the reviewer for pointing out this oversight. Now, they are deleted.\n8. Fig. 1E: Residue R326 is missing from the labeling.\nResponse: We appreciate the reviewer's comment. In the revised manuscript, we have now labeled residue R326 and clearly indicated the hydrogen bond between R326 and T372.\n9. Lines 213/397: Standardize nomenclature to consistently use “SPARDA” (instead of “SPARD” or “SPADA”).\nResponse: We thank the reviewer for pointing out this error. Now they are corrected in the revised manuscript.\n10. Fig. 1E: Add dashed lines for hydrogen bonds between residues H410 and T5.\nResponse: We thank the reviewer for this observation. In the original figure, the viewing angle obscured the hydrogen bond between H410 and T5. We have adjusted the figure orientation to better visualize this interaction.\n11. Fig. 1E: Label β1, β2, β11, and β12.\nResponse: We appreciate the reviewer's suggestion. In response, we have included β1, β2, β 11, and β12 in the figure and optimized the layout to improve the visibility of β11 and β12.\n12. Fig. 1E: Show the main chains for residues involved in backbone hydrogen bonds (e.g., L165–L376).\nResponse: Updated in the revised manuscript.\nRevised Fig. 1E\n13. Figs. 1G and 3C: Apply consistent domain-specific coloring for residues to reflect domain organization.\nResponse: Done.\nRevised Figs. 1G\nRevised Figs. 3C\n14. Fig. S7 legend: Contains several errors. For example, correct “Ssp6301” to “Ssp6803I” and replace “heighted” with “highlighted.”\nResponse: Corrected.\n15. Line 283: The phrase “according to the SEC elution file and the negative staining by cryo-EM” is unclear. Negative staining is a technique used in electron microscopy but not in cryo-EM. Please clarify the intended meaning.\nResponse: We thank the reviewer for the professional comment. We have revised the sentence in the manuscript according to your comment to more clearly and accurately describe our experimental results. We have modified the sentence to: “Deletion of the DREN domain did not alter the filament formation induced by tDNA recognition and the gRNA according to the SEC elution file and the corresponding appearance on the micrographs.”\nReviewer #3 (Remarks to the Author):\nReview of “Filament assembly induced by the recognition of target DNA activates the prokaryotic Argonaute SPARDA system” by Zhang et al.\nThe manuscript by Zhang et al. presents the structural basis for filament assembly of NbaSPARDA upon recognition of guide RNA and target DNA. This filament formation is required to activate the DREN module of SPARDA, thereby initiating nuclease activity. The authors provide structural and biochemical data that shed important mechanistic insights into how SPARDA defends against phages and plasmids. Overall, this is a valuable contribution to the field. The structural data are of generally good quality and support the central claims, although several discrepancies and missing details need to be addressed.\nResponse: We thank the reviewer for the positive comment and appreciate that the reviewer think our study is a valuable contribution to the field of pAgo-based antiphage defense system.\nThe article would benefit from revisions as outlined below:\nMajor Concerns\n1. Incomplete or unclear Cryo-EM data presentation\nSeveral Cryo-EM datasets mentioned in the text are not shown or are insufficiently described, making it difficult to evaluate the claims. For instance:\n- Lines 91–92 mention the preferred orientation of the SPARDA monomer, but no supporting data are provided. The data processing workflow, including representative micrographs and 2D class averages, should be included. It would also be useful to provide a low-resolution 3D reconstruction of the monomer.\nResponse: Thank you for highlighting this concern regarding the presentation of the SPARDA monomer data. We apologize for any lack of clarity in the original submission. In response to the reviewer's comments, we have included Supplementary Figure S2, which provides the 2D classification of the monomer and details of the data processing workflow. However, technical challenges including preferred orientation effects prevented us from obtaining a high-resolution structure.\n- Lines 97–100 suggest that a separate dataset for SPARDA+gRNA was collected, but this is not clearly described in the Methods. It currently appears that all reconstructions were derived from a single dataset.\nResponse: Thank you for highlighting the issue regarding the description of our datasets. We apologize for any confusion that the previous text may have caused. We confirm that a separate dataset for the SPARDA+gRNA complex was collected using the Titan Krios microscope. We have now amended the Methods section to include a detailed description of the data collection parameters for this specific dataset.\n- Supplementary Figures S2 and S5 show identical micrograph images, raising concerns. It appears that filament formation may also occur in the SPARDA+gRNA complex, which would contradict the authors’ claims. Micrographs for SPARDA alone should be provided to confirm whether filaments are also present.\nResponse: We sincerely thank the reviewer for their careful observation and for raising this important point. We sincerely apologize for the oversight in our supplementary figure preparation, where similar images were inadvertently reused in the original Supplementary Figures S2 and S5, which may have led to misunderstanding. To address this, we have made the following corrections:\nThe original Supplementary Figure S2 has been replaced with a new representative micrograph of the SPARDA+gRNA complex (now Supplementary Figure S3), which clearly shows no filamentous structures. Additionally, we have reorganized a new Supplementary Figure S2, which presents data from the SPARDA dataset collected using the Tecnai F30 microscope. This figure clearly demonstrates that SPARDA alone forms only as SPARDA monomers without any evidence of filament formation. Together, these supplementary results confirm that neither SPARDA alone nor the SPARDA+gRNA complex can autonomously assemble into filamentous structures. The formation of functional, long-range ordered filaments is strictly dependent on complete gRNA-tDNA target recognition.\n- Lines 166–167 imply that a 3D reconstruction of apo SPARDA was obtained from a different dataset, but this is not shown anywhere in the manuscript. Both the data processing workflow and the resulting 3D reconstruction should be included.\nResponse: Thank you for highlighting the need for further clarification regarding the 3D reconstruction of apo SPARDA. We apologize for any lack of clarity in our initial manuscript. To address this issue and enhance transparency, we have reorganized Supplementary Figures S2 in the supplementary materials. These figures clearly display the data processing workflow and the 3D reconstruction results of the SPARDA monomer. This addition provides a comprehensive understanding of the classification and reconstruction process of SPARDA.\n2. The micrographs indicate that not all particles form filaments; a significant fraction of monomeric or dimeric particles or maybe tetramers appear to remain. It is unclear whether these contain nucleic acids, and if so, whether they include only gDNA or the duplex. The authors should clarify this. Importantly, it should be determined whether filament formation is essential for activity or whether only a subset of complexes engage in nuclease activity. And if so are these from the filament.\nResponse: Thanks to the reviewers' insights, upon reprocessing the data for the small particles from the gRNA-tDNA-SPARDA dataset, we obtained a relatively high-quality map suitable for rigid-body fitting. Surprisingly, this map revealed the presence of a bound gRNA-tDNA heteroduplex, whereas the gRNA-SPARDA dimer complex clearly showed the presence of gRNA alone. The non-filamentous particles observed in the cryo-EM micrographs is likely due to that the SPARDA was incubated with annealed nucleic acids and then directly subjected to cryo-EM analysis.\nThis heterogeneity is consistent with our size exclusion chromatography results (Supplementary Figures S7A). In vitro nuclease activity assays established that collateral cleavage activity is exclusively associated with the high-molecular-weight fraction corresponding to the filamentous form, while the monomeric fraction shows no detectable activity (Supplementary Figures S7C). Similar to other oligomerization-dependent antiphage systems 9-11, when we mutated the amino acids affecting filament formation, SPARDA lost its collateral cleavage activity, indicating that filament formation is essential for SPARDA's activity.\nThe rigid body fitted model for the SPARDA monomer map from the gRNA-tDNA-SPARDA dataset.\n3. A close-up view of the cryo-EM density for the gRNA should be shown at the same threshold used for the rest of the structure. This would help establish whether the reconstruction contains only gRNA or the gRNA+tDNA duplex. As currently described, the sample composition used for the reconstruction is unclear.\nResponse: Thank you for this important observation. We have show a close-up view of the cryo-EM density for the gRNA, displayed at the same threshold used for the rest of the structure. This visualization confirms that the reconstruction contains only the gRNA, and not the gRNA+tDNA duplex. Consequently, we have appropriately named this structure the gRNA-bound SPARDA complex in our manuscript.\nA close-up view of the cryo-EM density and model for the gRNA-SPARDA dimer map.\n4. The Introduction or Discussion would benefit from a more detailed description of what is known about DREN domains, including their structural conformations and oligomerization requirements in other systems. A comparative structural analysis of multiple active DREN domains, including SPARDA, would help readers appreciate the significance of tetramerization.\nResponse: We appreciate the reviewer’s suggestion. We followed the reviewer’s suggestion and in the revised version of manuscript we enriched the Introduction and Discussion sections on the DREN domain, especially the oligomeric state of this domain for their functions. Please see line 331 and 342: “Dali search server reveals that the structure of DREN domain within SPARDA is similar to type II restriction endonuclease. Most of these restriction enzymes function as homodimers possessing two catalytic sites, a structural feature that appears to facilitate the binding and efficient cleavage of palindromic substrate sites. In addition, tetrameric assemblies are also common among type II restriction endonucleases, such as NgoMIV and SfiI. Structurally, these tetramers typically form through the head-to-head stacking of two dimers. In this manner, the two DNA-bound dimers in the restriction endonuclease places the PD-(D/E)-XK domains, their DNA-binding surfaces and the active sites close together in the individual protein. In contrast, the tetrameric I-SspI is able to accumulate a single long substrate and allows the catalytic PD-(D/E)-XK to far apart, thereby cleave a long target site. The tetramer assembly of the SPARDA DREN domain induced by ssDNA recognition can accumulate the substrate, thereby avoid the undesired activation under physiological conditions.”\n5. It would considerably strengthen the manuscript if the authors could demonstrate filament formation in vivo during phage infection. For example, fluorescently labeled SPARDA expressed in cells could be used to visualize puncta corresponding to filamentous structures upon infection. While this experiment is not essential for publication, it would greatly enhance the physiological relevance of the findings if performed within a reasonable timeframe.\nResponse: We thank the reviewer for this valuable suggestion. Accordingly, we applied the split GFP system to monitor SPARDA filament formation. In this assay, GFP is split into two non-fluorescent fragments, N-terminal β1-10 and C-terminal β11, which reconstitute fluorescence only when brought together by interacting proteins 12. Specifically, we fused GFP β1-10 to the N-terminus of DREN-APAZ and cloned it together with pAgo in separate reading frames of the pRSFDuet-1 vector. The GFP β11 fragment was fused to DREN-APAZ and co-cloned with pAgo into the pETDuet-1 vector. These two recombinant plasmids were co-transformed with a third plasmid, pCDFDuet-1, into E. coli BL21 (DE3) competent cells. Positive transformants were selected on LB agar plates containing ampicillin, kanamycin, and spectinomycin. A single colony was inoculated into LB liquid medium supplemented with the same antibiotics and grown at 37 °C with shaking until the OD₆₀₀ reached ~0.6. Protein expression was induced by adding 0.3 mM IPTG, followed by incubation at 16 °C for 16 h. Cells were then harvested and resuspended in PBS buffer.\nIn the absence of exogenous plasmid, no fluorescence was detected in bacteria co-expressing pAgo, GFP β1-10–DREN-APAZ, and GFP β11–DREN-APAZ. In contrast, upon introduction of the exogenous plasmid, clear fluorescence was observed in the co-expressing cells, indicating SPARDA filament formation. This indicates that pAgo and DREN-APAZ further assemble into a higher-order filamentous structure upon activation (Supplementary Figure 8).\nSupplementary Figure 8\n6. The authors should provide more details on potential structural changes at the pAgo dimerization interface during filament assembly. Is this interface altered upon filament formation? Could dimerization play a regulatory role in nuclease activity by conferring specificity to the presence of tDNA? The current data should be used to elaborate on this point.\nResponse: We thank the reviewer for raising this important point. In the revised manuscript, the structural changes on the dimeric pAgo are detailed described. Please see line 220 and 229, which now reads: “In comparison with the pAgo in the gRNA-bound dimeric SPARDA and the core dimeric pAgo in the filamentous SPARDA reveals a substantial structural rearrangement. Superimposition of one pAgo molecule in the dimeric SPARDA and that within SPARDA filament, it could be observed a rotation of approximately 90 degrees in the other molecule. In the gRNA-bound dimeric SPARDA complex, dimer formation is primarily mediated by interactions between the PIWI domains of adjacent pAgo molecules. In contrast, within the activated SPARDA filament, the dimer interface is largely reconfigured and stabilized predominantly by interactions between the MID domains of adjacent molecules, as well as through amino acid contacts formed between the PIWI domain of one subunit and the MID domain of its neighboring subunit.”\nThe activation of SPARDA involves a dynamic assembly process during gRNA loading, tDNA engagement, and propagation of the guide–target heteroduplex. In the apo state, SPARDA exists as a monomer. Upon gRNA binding, it transitions into a dimeric intermediate, which subsequently reverts to a monomeric form before assembling into a higher-order filament as the heteroduplex propagates. Therefore, we propose that the dimeric interfaces in the gRNA-bound SPARDA should be disrupted and further transition into the final filamentous assembly. The dimerization of SPARDA induced by gRNA is an intermediate state to form the functional filament, which we propose does not influence its nuclease activity.\n7. The SEC profile in Supplementary Figure S6 shows an additional peak at ~14 mL. Does this fraction also contain SPARDA bound to gRNA and tDNA? If duplex-bound SPARDA does not necessarily form filaments, is there evidence for tetramer formation in this fraction? Did the authors try a nuclease activity assay with this sample from the 14ml fraction compared to the 8ml fraction. This would be an important experiment.\nResponse: We thank the reviewer for raising this insightful question. Our size exclusion chromatography (SEC) data indicate that the second peak, eluting at approximately 14.34 mL, corresponds to monomeric SPARDA in complex with gRNA or tDNA. Ideally, the introduction of tDNA that triggers filament assembly should lead to the disappearance of this monomeric peak, as proteins would be expected to shift toward a higher-order oligomeric state. However, in all currently characterized prokaryotic Argonaute-based systems—such as SPARTA9,13 and SPARSA10 —the addition of gRNA and tDNA does not yield complete conversion to tetramers or filaments; a fraction of the sample typically remains in a monomeric or low-order form. At present, the precise mechanistic reason for this incomplete transition remains unclear.\nWe also performed the nuclease activity using SEC-fractionated samples from the two peaks. The results demonstrated that the filament-containing fraction (Peak1, eluting at ~8.13 mL) exhibited robust nuclease activity, whereas the monomeric complex fraction (Peak2, eluting at ~14.34 mL) showed no detectable nuclease activity. These results confirm that collateral cleavage activity is exclusively associated with the high-molecular-weight fraction corresponding to the filamentous form. We have added it in Supplementary Fig. 7C.\nSupplementary Fig. 7C.\n8. The cryoEM data processing workflow is not sufficiently detailed. For example, the FSC curves as well as local resolution maps are missing for the structures described. The authors should expand on the data processing showing ab initio reconstructions as well as further processing steps of possible 3d classification and refinement.\nResponse: We sincerely appreciate the reviewer's valuable suggestions. In response to these comments, we have systematically reorganized the cryo-EM data processing workflow (Supplementary Figure S3 and S6) to enhance both clarity and accuracy.\nMinor Comments\n1. The dimensions of the helical rise and diameter (lines 179–180) should be indicated in the figure.\nResponse: Done.\nRevised Fig. 3B\n2. Line 213 contains a typo: “SPRADA” should be corrected to “SPARDA.”\nResponse: Corrected.\n3. The sentence in lines 80–81 is unclear and should be rephrased for clarity.\nResponse: Thank you for pointing out the lack of clarity in this sentence. We have revised the sentence to enhance its clarity and precision. The revised sentence is as follows: “Interestingly, the heteroduplex formed by gRNA and its complementary tDNA is located within the DREN domain, demonstrating the essential role of filament formation in activating SPARDA's collateral nuclease activity.”"}, {"round": 2, "reviewer_comments": {"reviewer_1": "I have no further comments.", "reviewer_2": "The authors have addressed all my concerns.", "reviewer_3": "The authors have satisfactorily addressed all concerns raised in my previous review. The revised manuscript now includes the missing cryo-EM data workflows, corrected supplementary figures, clarified dataset descriptions, and additional analyses demonstrating the functional relevance of filament formation. These revisions substantially strengthen the mechanistic conclusions. I support publication of this manuscript in Nature Communications."}, "author_response": ""}]

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