Tau condensation on DNA mediates microtubule attachment suggesting a mitotic role for centromere-localized tau

Domain: physics
Published: 2026-01-07
Editorial outcome: major_revision

Abstract
Tau protein, traditionally recognized for stabilizing microtubules and forming pathological aggregates, has recently been observed to form condensates in various contexts. While its condensation with RNA has been well studied, the interaction between tau and DNA, along with its biological significance, remains less explored. Here, using single-molecule experiments, we find that tau binds stably to naked DNA at nanomolar concentrations, leading to the local co-condensation of tau and DNA. These tau condensates on DNA can also interface with microtubules, leveraging tau’s known role in promoting microtubule growth and organization. The dynamic nature of these condensates facilitates the remodeling of the DNA–microtubule assembly. Interestingly, phosphorylated tau and nucleosomal DNA exhibit distinct capacities to form condensates and recruit microtubules. Furthermore, imaging of mitotic cells with endogenous or exogenous tau reveals its localization to centromeres, engaging mitotic spindles, whereas expression of phosphomimetic tau mutants (T231D/S235D and S262D) causes defects in chromosome alignment. Building on these observations, we speculate that tau may play a role in mitosis, where tau clusters facilitate the early registration of mitotic spindles to chromosomes before kinetochore-mediated attachment. We also discuss the implications of this model in conditions where abnormal cell cycle re-entry and tau activity may disrupt cell division.

Introduction
Tau protein has long been known for its primary role in stabilizing axonal microtubules in neurons1,2. More recently, its ability to facilitate the nucleation and growth of microtubules has gained attention3. This function relies on tau’s tendency to form condensates at high concentrations4–7, a mechanism increasingly recognized as an organizing principle in many areas of cell biology8,9. At the same time, tau has been a major focus of research in neurodegenerative diseases due to its critical involvement in forming pathological aggregates10. The abnormal aggregation of tau is a hallmark of tauopathies, including Alzheimer’s disease, and is strongly associated with neuronal dysfunction and cell death.

In addition to binding microtubules, tau’s multifaceted biophysical properties enable a variety of dynamic interactions11. As an intrinsically disordered protein with distinct domains12, tau adopts flexible conformations, allowing for weak multivalent interactions with both itself and other molecules. This ability is crucial for driving the co-condensation of other factors within tau condensates. One example is tubulin, which has been shown to be recruited and to locally nucleate microtubule growth3. Another is the complexation between tau and RNA, which has been explored in the context of electrostatic interactions4,13, participation in stress granules14, and the development of tau fibrils and aggregates5,15,16. As a corollary, the interactions between tau, tubulin, and RNA can also modulate tau’s ability to organize microtubules17. These examples suggest that tau condensates may act as hubs for various biomolecules sparking distinct cellular activities.

Unlike tau–RNA interactions, the interaction between tau and DNA has received less attention. Since tau primarily resides in the cytoplasm, its limited access to nuclear DNA may have reduced the perceived physiological relevance of tau–DNA binding. Nevertheless, interactions between tau and naked DNA molecules have been reported18–25, albeit infrequently. A recent biophysical investigation by Abasi et al. demonstrated that tau can co-condense with naked DNA and nucleosome arrays under low-salt conditions25, which can be disrupted by aberrant hyperphosphorylation of tau. While it is plausible that tau, having a net positive charge of ~6, may bind dsDNA via electrostatic interactions similar to its RNA binding, the intrinsic stiffness of dsDNA likely renders tau–DNA interactions distinct. The biophysical strength of this interaction, as well as its potential role in living cells, remains largely unexplored. One notable exception is tau’s role in the nucleus, where it has been proposed to protect the genome and influence gene expression26–28.

Here, we report that tau directly interacts with DNA, forming co-condensates that serve as dynamic platforms for microtubule organization. Our single-molecule observations reveal that 2N4R tau, the longest isoform of tau expressed in the adult human brain, forms mobile condensates on DNA strands at low concentrations. These tau–DNA co-condensates act as dynamic microtubule attachment sites, assembling microtubules around the DNA, with tau phosphorylation and nucleosomes adding further complexity to this process. Furthermore, by imaging tau-expressing cells during division, we observed tau clustering around the centromeres of mitotic chromosomes during prometaphase, poised to interface with nascent mitotic spindles. Expression of phosphomimetic tau variants in cells also resulted in the failure of some chromosomes to align at the metaphase plate. Based on these findings, we speculate a mitotic function of tau in tau-expressing cells such as neurons. We hypothesize that as the nuclear envelope breaks down and chromosomes become exposed to cytoplasmic tau, the dynamic rearrangement of tau on mitotic chromosomes facilitates the “search and capture” process29 of newly forming microtubules, a process traditionally considered random. This model further proposes that as tau–DNA co-condensates mature and migrate toward the centromeres, these transient attachment sites are gradually replaced by kinetochores for a more stable connection.

Results
Tau stably binds to naked DNA at nanomolar concentrations
To probe the interaction between tau protein (2N4R, tau-441) and naked DNA, we first examined the mobility of DNA in an agarose gel after mixing it with purified tau protein (Supplementary Fig. 1). The prepared dsDNA samples of varying lengths (24–5000 bp) all displayed a significant mobility shift when incubated with wild-type tau (500 nM or higher) (Supplementary Fig. 2), indicating a strong association between tau and DNA. This interaction was further enhanced when the NaCl concentration was lowered from 100 mM to 25 mM, suggesting an electrostatic nature of the binding, similar to tau–RNA complexation4. Given that the shortest DNA we prepared (24 bp) allowed tau binding, it is likely that the longer DNA constructs can accommodate proportionally more tau molecules along their length. A recent report on tau’s phase separation behavior with naked DNA25 is in quantitative agreement with our results.

While this bulk assay confirmed tau’s DNA-binding properties, it did not reveal the distribution of tau within and across DNA strands, as well as the impact of this binding on DNA conformation. To address these questions, we set up a series of single-molecule imaging experiments that directly visualize the binding of tau protein to DNA substrates. We first performed a classic flow-stretching experiment in a flow chamber, where 48.5-kb λ-DNA molecules were attached to a polyethyleneglycol (PEG)-coated glass coverslip by one end and then stretched upon buffer flow (7 μL min−1) (Fig. 1A). The stretched DNA strands were visualized using a fluorescent stain (SYTOX Orange) on a total internal reflection fluorescence (TIRF) microscope (Fig. 1B). The tethered λ-DNA molecules were gently stretched to 6.2 ± 0.6 μm (~40% of the contour length) (Fig. 1C, D), indicating ~0.1 pN of flow-induced stretching force. When 500 nM tau (10% labeled with Cy5) was added at the same flow rate, the λ-DNA molecules gradually shortened and collapsed onto the attachment point (Fig. 1B–D and Supplementary Movie 1). We checked that the fluorescently labeled tau also accumulated on the condensed DNA spots (Fig. 1B, E), indicating that tau binding is the direct cause of DNA compaction. Note that these observations were made without highly concentrated tau solutions or a crowding agent, both of which are typically needed to induce condensation behavior through liquid–liquid phase separation (LLPS)5,6,25,30. This indicates that a stable interaction between tau and naked DNA at nanomolar concentrations, which locally enriches tau, is sufficient to induce DNA compaction prior to the onset of LLPS-dependent tau–DNA co-condensation25.

Tau binding to DNA generates local co-condensates
Recent discoveries indicate that DNA-binding proteins can form condensates on DNA, thereby compacting and remodeling the underlying structure31–37. Motivated by these findings, we hypothesized that tau could exert similar effects on DNA. To test this, we prepared λ-DNA molecules tethered at both ends to a surface and introduced Cy5-labeled tau again (Fig. 2A), simultaneously imaging the fluorescence of both tau and DNA. As observed in single-tether experiments, there was significant fluorescence from tau binding to the DNA (Fig. 2B, C, and Supplementary Movie 2). The DNA backbone was diffusely coated with tau, interspersed with multiple bright foci of ~1 μm or smaller, as revealed by the intensity profile cross section (Fig. 2B, inset). As expected from the bulk gel shift assay, tau binding was both tau and salt concentration-dependent (Fig. 2C). Higher concentrations of tau resulted in more intense spots, even though we kept the concentration of the Cy5-labeled tau constant (50 nM) across all samples.

Although the DNA coverage by tau varied among DNA strands due to different levels of foci formation, the locations of these bright regions corresponded closely between DNA and tau signals (Fig. 2B inset and Fig. 2D), suggesting that clustered tau binding leads to local compaction of DNA. The level of local compaction was estimated by the coefficient of variation of the DNA fluorescence signal (the standard deviation of pixel intensities divided by their mean), which increased progressively as more tau was added (Fig. 2E). In contrast, DNA strands without tau showed uniform fluorescence along their length. Consistent with the local compaction effect, the binding of tau noticeably reduced the thermal fluctuation of DNA (Fig. 2F and Supplementary Movie 2), indicating that the DNA became shorter and tauter. The mean envelope width, or the amplitude of off-axis deviation due to fluctuation, gradually decreased as tau concentration increased from 50 nM to 5 μM (Fig. 2G), consistent with the increase in tau binding. Notably, even 50 nM tau was sufficient to induce strong condensation of DNA under low-salt conditions (25 mM NaCl), as demonstrated by both foci formation (Fig. 2B, E) and reduced fluctuation (Fig. 2G). Importantly, these findings were confirmed under 150 mM NaCl, which more closely approximates physiological salt conditions (Supplementary Fig. 3). These results demonstrate that wild-type tau protein at nano-to-micromolar concentrations can stably bind to naked DNA under physiological conditions, consistent with the behavior reported by Abasi et al25.

To ensure our findings on tau–DNA co-condensation extend beyond single-molecule observations, we checked bulk phase separation and subsequent co-condensation of high tau concentrations with dsDNA in the presence of 10% PEG, a standard crowding agent to drive LLPS. Under these conditions, tau and DNA readily formed co-condensates across various concentrations of tau, DNA, and salt (Supplementary Fig. 4), again consistent with the previous results25. Importantly, the co-condensation behavior was robust across a wide range of DNA lengths, from 24 bp to λ-DNA, indicating that DNA length did not significantly influence its partitioning into tau droplets. This tau-driven DNA condensation, reversing the above single-molecule observations of DNA-driven tau condensation, reinforces the strong molecular interaction between tau and DNA.

Tau condensates slide along DNA to compact local segments
Surprisingly, despite the fast accumulation of tau on DNA (Fig. 1B), many of the tau condensates remained mobile along the DNA substrates (Fig. 2H and Supplementary Movie 3). Moreover, the location of DNA foci also followed tau movement across all tau concentrations we tested (50 nM – 5 μM). This suggests that tau clusters can slide along DNA to dynamically remodel the underlying segments, reminiscent of other LLPS-driven protein–DNA co-condensates. However, not all condensates exhibited uniform mobility, likely due to heterogeneity in their mechanical properties (e.g., differential maturation to more solid-like states), friction from contact with the PEG-coated glass surface, or a combination of both. To probe the internal dynamics of tau clusters, we performed fluorescence recovery after photobleaching (FRAP) on tau clusters assembled with 5 μM tau along double-tethered λ-DNA (Supplementary Fig. 5). Bleaching and monitoring of tau fluorescence (Cy5) recovery yielded highly heterogeneous traces, reflecting frequent condensate movement, merging, and splitting, yet many traces exhibited substantial fluorescence return. A single-exponential fit of the mean recovery for representative traces gave a time constant of ~30 min and a mobile fraction of ~82%. However, these values represent a composite of fast- and slow-recovering populations and are affected by the loss of some condensates during monitoring. Overall, these results confirm that DNA-bound tau exchanges dynamically with the bulk pool in a liquid-like manner.

To alleviate the surface-imposed friction, we re-evaluated the mobility of tau–DNA condensates on the DNA skybridge platform (Fig. 2I)38. In this technique, λ-DNA is suspended between microfabricated quartz barriers, reducing surface interaction artifacts and enabling high-throughput visualization of condensate mobility. Indeed, by allowing sufficient spacing between the DNA strands and the substrate surface, we confirmed that the tau and DNA foci migrated together along the DNA strands (Fig. 2J and Supplementary Movie 4). Essentially all of the observed co-condensates (n=52) showed sub-diffusive behavior, with an anomalous exponent (α) centered around 0.5 and diffusion coefficients of 0.01–0.02 μm2 s−1 (Fig. 2K), indicating transient confinement and the need to reorganize or drag compacted DNA segments as they slide along the polymer. The skybridge platform further provided a convenient means to probe the sequence preference of tau condensate localization (Supplementary Fig. 6). Condensates were observed more frequently in the AT-rich half of λ-DNA, consistent with tau’s known preference for AT-rich sequences39, although the overall broad distribution reflects its predominantly nonspecific binding. In conclusion, these findings indicate that tau binding to DNA leads to dynamic co-condensation, which spatiotemporally modulates the local compaction of DNA.

Tau binding exerts piconewton condensation force on DNA
The flow-stretching and double-tether experiments showed that tau binding can induce DNA condensation, counteracting a force of at least ~0.1 pN from the buffer flow. We next employed magnetic tweezers to accurately measure the force generated by tau condensates on DNA and to determine if the condensation can be mechanically reversed (Fig. 3A). A 10-kb dsDNA fragment was prepared as a substrate, with one end attached to a PEG-coated glass surface and the other to a magnetic bead for manipulation. Changes in DNA extension were monitored by tracking the vertical movements of the bead. This setup was also equipped with a TIRF microscopy module40, which checked the fluorescence signal from tau molecules binding to DNA (Supplementary Fig. 7).

In the absence of tau, the classic worm-like-chain extension of dsDNA was reproduced by applying a force ramp between 0.1 and 10 pN (Fig. 3B, left). To ensure that the DNA constructs were free of single-stranded nicks, we used only those that collapsed under applied torque, exhibiting supercoiling behavior (Supplementary Fig. 7). However, when 500 nM tau was added to the buffer, the overall extension of DNA was notably reduced, especially in the low-force range between 0.1 and 3 pN (Fig. 3B, center), consistent with tau-induced condensation of DNA. Fluorescence verification confirmed the accumulation of tau molecules on the DNA strands (Supplementary Fig. 7, Supplementary Movie 5). At higher forces, in contrast, the extension returned fully to that of naked dsDNA, implying that the tau molecules were largely displaced by stretching the DNA. It was possible to drive this compaction and decompaction cycle by repeatedly stretching and relaxing the DNA. Although the exact force levels at which these transitions occurred varied across trials (Fig. 3C), slightly higher forces (1–3 pN) were required than those at which condensation first occurred (<1 pN), suggesting some mechanical hysteresis and energy dissipation in the tau–DNA interactions. Finally, increasing the concentration of tau (from 500 nM to 5 μM) raised both condensation and decondensation forces, consistent with an increased tendency to undergo condensation.

Tau–DNA co-condensation is cooperative
The force-extension curves in the presence of tau revealed both smooth and stepwise changes in DNA extension. We considered whether the stepwise changes might represent a specific unit length of DNA compacted by tau condensates of relatively uniform size. To better resolve these events, we applied force-jump cycles (Fig. 3D), where the force was abruptly switched between 0.5 and 3 pN to trigger compaction and decompaction events. During the stretching phase, we detected many stepwise length changes (Fig. 3E), with a median size of 72 nm (Fig. 3F). This stepwise unbinding reflects the cooperative nature of the tau–DNA condensates, as each step likely corresponds to the concerted rupture of a sub-domain held together by multivalent tau–tau and tau–DNA interactions.

To test this interpretation at the molecular level, we performed MD simulations of tau–DNA co-condensation under applied force (Supplementary Fig. 8). Here, ionic effects at physiological monovalent salt (~100 mM) were implicitly captured by setting the DNA persistence length to ~50 nm, which subsumes short-scale bending and salt-dependent flexibility (see Supplementary Note 1, “Molecular dynamics simulation for tau–DNA co-condensates” for more details). In silico, tau and DNA coalesce into droplets featuring looped DNA segments and characteristic bending, without strict periodicity (Fig. 3G). Simulated force–extension profiles recapitulated the experimentally observed stepwise jumps (Fig. 3H), and inspection of snapshots and local DNA densities before versus after individual jumps confirmed that each event corresponds to the rupture of a compacted DNA domain (Supplementary Fig. 8). Importantly, the distribution of simulated step sizes peaked near the DNA persistence length (Fig. 3I), demonstrating that DNA bending rigidity defines the natural rupture unit. In contrast, simulations lacking bending energy (i.e., a freely jointed chain model) produced much smaller, irregular steps (Supplementary Fig. 8).

During the condensation measurements under a constant 0.5 pN load, individual DNA tethers showed a binary response (Fig. 3J): they either collapsed rapidly into a condensed state (occasionally via discrete, slower steps) or remained fully extended, with large fluctuations over 1 min. Both the switch-like collapse and the slower fluctuations likely stem from cooperative tau condensation: rapid compaction follows cluster formation, while the larger, slower changes reflect cluster-mediated remodeling. To quantify this cooperativity, we measured the extent of DNA shortening as a function of tau concentration and observed a sharp transition near 500 nM (Fig. 3K). A Hill-plot analysis yielded a Hill coefficient of n=3.1±2.5 and an apparent Kd=0.55±0.14 μM (95% confidence interval), reinforcing the conclusion that multiple tau molecules bind DNA cooperatively with a moderate-to-high affinity.

Tau condensates facilitate microtubule attachment to DNA
The tau condensates formed on DNA substrates are reminiscent of those that facilitate microtubule growth and assembly3. Presumably, the DNA-bound tau condensates might function similarly to organize microtubules, possibly by cooperating with tau molecules already present on the microtubule surface. To test this idea, we introduced a mixture of tau and tubulin over the surface-bound tau–DNA assembly and evaluated whether the tau condensates maintain their potential to organize a microtubule network. We first checked that mixing tubulin with tau in the presence of GTP and 5% PEG induced rapid polymerization of microtubules (Supplementary Fig. 9). When this solution of nascent microtubules was introduced over surface-tethered DNA molecules prepared at a low density, tau–DNA co-condensates effectively captured them (Fig. 4), suggesting that tau retains its essential microtubule binding activity. Importantly, fluorescence from free tubulin also localized within the tau condensates (Fig. 4A), indicating that these spots might serve as seeds for nucleating new microtubules.

The microtubules engaged with the DNA strands either along their sides or via their ends, analogous to the lateral and end-on attachments posited for mitotic spindle coupling to chromosomes during mitosis41. Intriguingly, microtubules attached to both singly and doubly tethered DNA strands exhibited dynamic behaviors, including diffusive movements along DNA due to mobile condensates (Fig. 4A–C, Supplementary Movie 6) and pivoting around anchoring tau condensates (Fig. 4D, Supplementary Movie 7). This was particularly evident in the occasional diffusive translation of entire microtubule filaments along double-tethered DNA substrates. With a slight increase in DNA surface density, more microtubules were captured, and tau clusters on different DNA strands cooperatively bridged and bundled nearby microtubules, forming a tripartite network of DNA, tau, and microtubules (Supplementary Movie 8). These observations suggest that tau condensates may facilitate the repositioning and stabilization of microtubules relative to DNA, in processes such as mitotic spindle attachment to chromosomes.

To directly demonstrate the role of tau condensates in microtubule capture, we designed a microtubule capture assay in a more complex environment with higher concentrations of tau and DNA. In this assay, we densely packed the surface with DNA (Fig. 4E, Supplementary Movie 9), allowing multiple strands to collaboratively induce tau condensation. We increased the tau concentration to 5 μM and added 5% PEG to simulate molecular crowding. The formation of tau–DNA co-condensates appeared as discrete bright nodes interconnected by surface-tethered DNA, which manifested as thin, dimmer lines forming a reticular network (Fig. 4F, Supplementary Movie 9). The tau–DNA co-condensates appeared as distinct bright foci, while the surface-tethered DNA formed an intricate mesh connecting these nodes. This reticular structure suggests a complex and organized arrangement of tau–DNA interactions, highlighting tau’s ability to induce localized DNA condensates while maintaining a networked distribution across the nucleic acid surface, similar to the behavior of various DNA-compacting proteins. Under this condition, the overall structure became more stabilized, and the mobility of tau locations was significantly reduced (Supplementary Movie 9).

Under these conditions, tau–DNA clusters at distant sites were able to cooperatively recruit longer microtubules (Fig. 4G, Supplementary Movie 9), effectively bridging the gap in length scale. Since the DNA molecules were arranged on a 2D plane, the captured microtubules conformed to the substrate surface. Most importantly, when tau condensates were not preformed before the introduction of tubulin, only a few surface-bound microtubules were observed (Fig. 4H vs. 4I), despite the presence of numerous microtubules away from the DNA surface (Supplementary Movie 10), contributing to background fluorescence. By quantifying the number of microtubules attached to the DNA surface, we confirmed that the presence of preformed tau–DNA co-condensates is indeed critical for capturing microtubules from the solution (Fig. 4J). We propose that this conformal attachment of microtubules to a densely packed DNA surface via tau–DNA co-condensates may represent a mechanism for docking nascent microtubules to complex DNA structures, such as mitotic chromosomes.

Nucleosome arrays support tau binding and microtubule recruitment
To place our in vitro observations in a more physiologically relevant context, we replaced λ-DNA with a 2.3-kb dsDNA bearing 12 Widom-601 sites (Fig. 5G), reconstituted nucleosomes on this template, and repeated the tau-binding and microtubule-capture assays. A set of biochemical and biophysical assays indicated densely loaded, stable nucleosome arrays (see Supplementary Note 2, “Verification of Nucleosome Assembly on 601 DNA”, and Supplementary Figs. 10–11). For a controlled comparison to naked DNA, we also prepared naked 601 DNA (no nucleosome reconstitution) at similar surface density for imaging (Fig. 5H, I). Because the 601 construct is much shorter than λ-DNA (2.3 kb vs. 48.5 kb), individual DNA strands were not resolvable by TIRF. Upon adding 5 μM tau, discrete tau clusters formed and colocalized with H2A-labeled nucleosomes (Fig. 5J). Although overall tau binding was lower than on λ-DNA, likely reflecting the shorter template length, colocalization analysis showed a high correlation between nucleosome and tau signals for both nucleosomal and naked 601 DNA at near-comparable levels (Fig. 5K).

Finally, introducing a tau–tubulin mixture containing nascent microtubules yielded robust microtubule capture on surfaces bearing nucleosome-pinned tau clusters (Fig. 5L), comparable to λ-DNA-induced condensates (Fig. 4J). Microtubule counts were indistinguishable between naked DNA and nucleosome arrays (Fig. 5M), despite the reduced amount of free DNA segments in the latter. Together, these data suggest that nucleosomal DNA remains competent to recruit tau and interface with microtubules to a similar extent as its naked-DNA counterpart, consistent with recent bulk measurements of tau association with nucleosome arrays and linker DNA25.

Tau clusters localize on centromeres during prometaphase of mitosis
The tripartite assembly of DNA, tau, and microtubules we observed in vitro led us to hypothesize a role for tau in the division of tau-expressing cells, especially in how nascent mitotic spindles capture chromosomes. To test this in a neuronal setting, we conducted immunofluorescence imaging of SH-SY5Y cells, a neuroblastoma line that endogenously expresses tau. Immunostaining of thymidine-synchronized, dividing cells (identified by condensed chromosomes) revealed a consistent level of endogenous tau forming small puncta with distinct distributions across mitotic stages (Supplementary Fig. 12). We focused on cells in prometaphase, just before chromosome congression, which typically displayed the characteristic rosette structure formed by mitotic chromosomes surrounding the nascent spindle (Fig. 6A). These images revealed strong colocalization between tau and tubulin, indicating that tau remains closely associated with microtubules during prometaphase. Notably, tau formed distinct clusters on mitotic chromosomes, most of which colocalized with centromeres labeled by CREST anti-centromere antibody (Fig. 6A, magnified views in the bottom panels). To quantitatively assess the colocalization between tau and centromeres, we pooled images of 181 centromeres from 11 prometaphase cells and averaged them after precise alignment (Fig. 6B). Intensity profiles of centromeres and tau fluorescence in the averaged images indeed showed a clear overlap. Pearson correlation coefficients calculated for each SH-SY5Y cell image confirmed strong colocalization of tau with both tubulin and centromeres (Fig. 6C, D). Interestingly, prophase SH-SY5Y cells also revealed clear association of nuclear tau with centromeres (Supplementary Fig. 12), consistent with its known pericentromeric localization39,46.

To extend these findings beyond a neuronal context, we next turned to HEK-293 cells, which have low endogenous tau. We transiently expressed mCherry-tagged 2N4R tau and synchronized the cells in the same way. Confocal imaging of mCherry fluorescence for tau, alongside immunostaining for tubulin and centromeres (CREST antibody), again showed tau clusters on the prometaphase rosette of mitotic chromosomes and tight colocalization with spindle microtubules (Fig. 6E). Analysis of 460 centromeres from 18 prometaphase HEK-293 cells verified overlap between tau and centromere signals (Fig. 6F), mirroring our SH-SY5Y results and supporting tau’s role in chromosome–spindle attachment. Again, Pearson correlation across these cell images quantified strong colocalization of tau with both tubulin and centromeres (Fig. 6C, D).

To assess whether tau directly associates with mitotic chromosomes at higher resolution, we prepared chromosome spreads from these HEK-293 cells expressing mCherry–Tau (Fig. 6G). In these preparations, small punctate tau clusters decorated the surface of condensed chromosomes, demonstrating that tau retains its DNA-binding activity within a chromatin context. We did not observe centromere-specific enrichment in this case, likely because the more transient or weak interactions that mediate centromere recruitment are disrupted during the spreading.

While native HEK-293 cells showed no detectable tau, transient expression of 2N4R mCherry–Tau produced tau levels far exceeding those in SH-SY5Y (Fig. 6H). To probe whether tau expression alters cell-cycle behavior, we performed RNA-seq on these HEK-293 cells. Tau expression caused modest transcriptomic changes (Fig. 6I), including upregulation of stress-response genes CHAC1, ATF3, HMOX1, and DDIT3. We also observed downregulation of CENPE, involved in chromosome movement, and BRCA2, a key DNA-repair and cell-cycle regulator (Fig. 6J). These shifts mainly affected cell-division, microtubule-organization and DNA-repair pathways, consistent with a mitotic role for tau. Despite these transcriptomic shifts, overall proliferation was largely unchanged, likely because other mitotic factors compensate for tau’s function in its absence.

Collectively, these findings highlight the consistent involvement of tau in interfacing microtubules with centromeres during mitosis in tau-expressing cells, whether tau is present endogenously or introduced exogenously.

Phosphomimetic tau mutants condense DNA but differ in microtubule-interfacing potential
Phosphorylation of tau at multiple sites is tightly regulated by various kinases and phosphatases, influencing its affinity for microtubules47. To investigate how phosphorylation-like changes affect tau–DNA interactions and their role in recruiting microtubules, we generated two additional recombinant tau proteins with phosphomimetic mutations: T231D/S235D (a double mutant) and S262D. Given the critical association of these sites with microtubule binding and Alzheimer’s disease48–51, these mutants are often used to model pathological modifications seen in tauopathies52–55.

We first confirmed through single-molecule experiments that both T231D/S235D and S262D tau proteins successfully undergo co-condensation with DNA, similar to wild-type tau (Fig. 7A). Next, we examined whether these mutants could still support the assembly of the DNA–tau–microtubule network. When mixed with tubulin in the presence of GTP, the phosphomimetic mutants were able to nucleate microtubule growth, though to a much lesser extent than wild-type tau (Fig. 7B). Specifically, the microtubules formed in the presence of these mutants were shorter, less bundled (as indicated by a dimmer appearance), and depolymerized more rapidly compared to those formed with wild-type tau (Supplementary Fig. 9). Nonetheless, in this buffer condition, microtubule growth was entirely absent without tau, indicating that the tau mutants still retain some microtubule-binding activity. As a result, wild-type tau outperformed the phosphomimetic mutants in promoting microtubule growth, as reflected by the higher microtubule count (Fig. 7C). These findings are consistent with previous observations that phosphorylated tau has impaired microtubule binding, leading to a reduced stabilizing effect.

To compare the capacity of different tau mutants to couple microtubules to DNA, we performed a microtubule capture assay using tau–DNA co-condensates, as described above for wild-type tau (Fig. 4G). The images of microtubules on the DNA surface were largely indistinguishable between wild-type and the T231D/S235D mutant. However, in the case of the S262D mutant, almost no microtubules were observed; instead, clustered aggregates of tubulin formed (Fig. 7D). Quantification of the microtubule count revealed that, despite its impaired ability to promote microtubule formation in solution (Fig. 7B, C), the T231D/S235D tau mutant captured and interfaced with microtubules on the DNA surface at a level comparable to wild-type tau (Fig. 7D, E). This suggests that the T231D/S235D mutant retains a particularly strong capacity for mediating interactions between DNA and microtubules, even though it exhibits reduced microtubule nucleation and bundling overall. In contrast, the S262D mutant showed a marked decrease in microtubule capture on the DNA surface, presumably due to its impaired microtubule binding and stabilization49. Overall, these results suggest that tau’s ability to interface between DNA and microtubules may be regulated by site-specific phosphorylation.

Phosphomimetic tau localizes to centromeres but impairs chromosome alignment
To test whether the in vitro differences in DNA–microtubule interfacing by phosphomimetic tau mutants manifest in cells, we performed immunofluorescence on mitotic HEK-293 cells transiently expressing mCherry-tagged T231D/S235D or S262D tau, using the same approach as for wild-type tau (Fig. 6). Although both mutants largely retained microtubule binding during mitosis (Fig. 7F), Pearson correlation analysis revealed their reduced association with spindle microtubules (Fig. 7G), consistent with the impaired microtubule-stabilizing activity (Fig. 7B). Nevertheless, both mutants exhibited robust centromeric localization comparable to wild-type tau (Fig. 7H), reflecting their retained DNA-binding activity (Fig. 7A). Analysis of averaged centromere images further confirmed overlapping tau signal (Fig. 7I). Surprisingly, in metaphase cells after chromosomes have aligned at the metaphase plate, both phosphomimetic mutants showed prominent tau-rich chromosomal regions that failed to align (Fig. 7J). These tau–DNA clusters did not colocalize with the central tubulin signal, indicating a lack of spindle attachment, and were most frequent in S262D-expressing cells, implicating aberrant tau–DNA binding in defective microtubule capture. Quantitative analysis of DNA and tau amount (measured by Hoechst 33258 and mCherry fluorescence, respectively) confirmed these findings. In S262D cells, misaligned DNA foci comprised about 10% of total chromosomal DNA, significantly more than in wild-type or T231D/S235D cells (Fig. 7K; see Supplementary Fig. 13 and Supplementary Methods for analysis). Tau fluorescence at these foci accounted for nearly 30% of total cellular tau in S262D cells—over a five-fold increase versus wild type (Fig. 7L). These data demonstrate that phosphomimetic tau mutants not only lose their ability to engage spindle microtubules but also aberrantly accumulate on misaligned chromatin, thus compromising proper chromosome segregation.

Methods
Protein purification and labeling
The cDNA encoding the 6×His-tagged 2N4R isoform of human tau was cloned into the pRK172 vector (a gift from the Feinstein lab, UCSB) and expressed in LEMO21 cells (NEB #C2528J). For preparing phosphomimetic tau mutants, site-directed mutagenesis was performed on the wild-type plasmid via overlap-extension PCR (primer sequences provided in Supplementary Data 1), and the resulting products were assembled into full-length plasmids using Gibson assembly (NEB, #E2611S). For bacterial expression, cells were cultured in TB media at 37°C until the OD600 reached 0.6, followed by treatment with 10 mM betaine and 500 mM NaCl. After a 30-min incubation at 30°C, 0.4 mM IPTG was added, and the culture continued for an additional 3–4 h. The cells were homogenized and lysed in lysis buffer (20 mM MES (pH 6.0), 0.2 mM MgCl2, 1 mM EGTA, 1.5 mM PMSF, 1× protease inhibitor cocktail, 5 mM DTT). The lysate was sonicated at 30% amplitude in 30-s on-off cycles for 3 min, followed by the immediate addition of 500 mM NaCl. The lysate was boiled for 15 min and centrifuged at 15000 × g for 1 h. Supernatants were purified using Ni-NTA resin (Qiagen #30210). The bound resin was washed with wash buffer (50 mM phosphate buffer (pH 7.2), 200 mM NaCl, 20 mM imidazole) and eluted with elution buffer (50 mM phosphate buffer (pH 7.2), 200 mM NaCl, 500 mM imidazole). TEV protease purified in-house from a TEV expression plasmid obtained from Addgene (#135464) was then added to remove the His-tag, and the solution was incubated overnight at 4°C. To further purify the protein, ion exchange chromatography (Bio-Rad #1580030) was performed to remove dimers and other undesired proteins. The protein elutes were concentrated using a protein concentrator (Sartorius #VS15T01). Phosphorylation of wild-type tau was performed in vitro following Abasi et al.25. A total of 100 μM tau was incubated with 2 μL PKA (New England BioLabs #P6000S) in kinase buffer containing 20 mM HEPES (pH 7.2), 50 mM NaCl, 0.5 mM TCEP, 4 mM adenosine-5′-triphosphate (Sigma #A7699-5G), 1 mM PMSF and 10 mM MgCl2 at 30°C for 16 h. To inactivate the kinase, the mixture was incubated at 65°C for 20 min. Subsequently, buffer exchange was performed using a 30 kDa-cutoff Amicon centrifugal filter to replace the kinase buffer with 50 mM HEPES and 25 mM NaCl. Phosphorylation results were verified by SDS-PAGE mobility shift and LC–MS/MS analysis (methods in Supplementary Methods). For protein labeling, the purified protein was mixed with a 10-fold molar excess of Cy5-maleimide (Cytiva #25800674) and incubated for 2 h at room temperature (RT) in reaction buffer (50 mM phosphate buffer, 0.05 mM TCEP). Free dye was removed by gel filtration (Bio-Rad #1500738). The Cy5-labeled tau proteins were then concentrated using a protein concentrator (Sartorius #VS15T01).

Single-molecule imaging
To prepare DNA constructs for surface attachment, λ-DNA (Thermo #SD0021) was linearized by ligating 5′- and 3′-biotinylated oligonucleotides targeting the cohesive ends at a 100× molar excess in 1× NEB ligase buffer for 5 min at 65°C (the full list of oligonucleotides used in this study is provided in Supplementary Data 1). The mixture was then slowly cooled over 3 h. T4 DNA ligase (NEB #M0202L) and 1 mM ATP were added, followed by overnight ligation. The sample was purified using a 100 kDa cutoff buffer exchange filter (Merck, #UFC510024) to remove free primers and exchange the buffer for distilled water. A flow chamber for imaging was constructed using previously described methods98,99, In brief, two PEGylated glass coverslips were assembled with double-sided adhesive spacers to create a narrow flow channel. The coverslips were PEGylated through a first round using a 99:1 mixture of 5-kDa mPEG and biotin-PEG (Laysan Bio, #BIO-PEG-SVA-5K-100MG & MPEG-SVA-5K-1g), the coverslips were subjected to a second round of overnight PEGylation with a solution of 2-kDa mPEG (Laysan Bio #MPEG-SVA-2000-1g) without biotin-PEG to ensure optimal passivation. Before imaging, sample solutions were introduced into the flow chamber using a syringe pump. For capturing DNA samples, 8 nM NeutrAvidin (Thermo #31000) was introduced into the flow cell, incubated for 5 min, and then washed with PBS. The flow cell surface was further passivated by infusing 5% BSA (Sigma #A0100-010) for 10 min, followed by another PBS wash. Biotinylated λ-DNA was diluted to 10–20 pM, injected, and immobilized on the surface for 7 min. For imaging, the buffer was exchanged with 200 μL of imaging buffer (50 mM HEPES (pH 7.1), 100 mM NaCl, 2 mM Trolox, 59.5 nM protocatechuate 3,4-dioxygenase (PCD #P8279-25UN), and 2.5 mM protocatechuic acid (PCA #37580-25G-F)) containing 20 nM SYTOX Orange (Thermo #S11368) and incubated for 10 min. To examine tau–DNA co-condensation, tau protein was diluted in the imaging buffer to concentrations ranging from 50 nM to 5 μM and then injected into the flow cell. All images were taken on a home-built objective-type TIRF microscopy setup. Samples were illuminated with 488 nm, 532 nm, and 633 nm CW lasers (Cobolt) and imaged through a 60× oil-immersion lens (Olympus, NA 1.49 for TIRF microscopy) onto a high-sensitivity sCMOS camera (Photometrics Prime BSI Express). Images were acquired typically at 108-nm spatial resolution with 100-ms exposure time using custom LabVIEW software. When necessary, fluorescence in two channels was separated by a dual-view image splitter (Cairn OptoSplit II) and recorded simultaneously. Additional methods for characterizing tau–DNA co-condensation, image analysis, including mobility shift and bulk LLPS assays, FRAP, DNA skybridge experiments, and nucleosome reconstitution and verification (Supplementary Note 2) are described in the Supplementary Information.

Single-particle tracking
Tracking of tau condensates on DNA was performed by analyzing movies from DNA skybridge experiments using custom MATLAB scripts. After registering the tau and DNA fluorescence channels, tau particles were detected by intensity thresholding, and only those colocalizing with DNA foci were included in the diffusion analysis. Additionally, to avoid surface artifacts the skybridge substrates, particles within 7 pixels of the ridges or adhered to the bottom surface were excluded from analysis. Mean-squared displacements (MSD) were calculated as a function of time lag Δt and fit to MSD(Δt) = 2D(Δt)α to extract the diffusion coefficient D and anomalous exponent α.

Magnetic tweezers experiments
The preparation of the 10-kb dsDNA construct used in this work is detailed in our previous work40. Briefly, three DNA fragments were generated by PCR from λ-DNA: one 10 kb fragment and two 500 bp fragments (the list of primers is provided in Supplementary Data 1). During PCR, the 500 bp fragments were labeled with multiple biotin or digoxigenin molecules by incorporating biotin- or digoxigenin-labeled dUTP (Sigma, #11093070910, #11093088910). All three PCR products were digested with BsaI (NEB, #R3733S) for 4 h at 37°C, and after cleanup, were ligated overnight using T4 DNA ligase (NEB, #M0202L) at RT. The ligated product was then purified by agarose gel electrophoresis. Details of the flow cell construction, sample assembly, and the home-built magnetic tweezers setup are available in our previous work100,101. Briefly, the magnetic tweezers consisted of a pair of permanent magnets positioned above the flow cell on an inverted microscope, allowing controlled application of force to paramagnetic beads tethered to single DNA molecules. For measurements, the DNA samples were injected into the flow cell so that the biotinylated ends could attach to PEGylated glass surfaces via biotin-avidin interactions, while the digoxigenin-labeled ends were coupled to anti-digoxigenin-coated magnetic beads (Sigma, #11333089001, Invitrogen, #M-270), which were injected afterward. To apply force, an antiparallel pair of cubic neodymium magnets (N52, 3/8′′) was placed above the flow cell, with a 1-mm gap between the magnets. The height and rotation of the magnets were controlled using a translation stage (PI, #L-40620DD10) and a stepper motor. Magnetic beads were illuminated with a 680 nm super-luminescent diode and imaged using a 100× objective lens (Olympus, #UPLXAPO100XO) with a high-speed CMOS camera (Mikrotron, #EoSens 3CXP). Before measurements, the integrity of the dsDNA constructs (i.e., the absence of nicks) was checked by rotating the samples and observing supercoiling behavior (Supplementary Fig. 7). When necessary, the force-extension curves were corrected for the off-axis tilting of magnetic beads102. For fluorescence verification, DNA-bound tau protein labeled with Cy3 was imaged simultaneously with magnetic tweezing using a 532 nm laser aligned for TIRF illumination and an sCMOS camera (Photometrics, #Prime BSI Express)40. All measurements were conducted in a buffer containing 50 mM HEPES (pH 7.1), 100 mM NaCl, 59.5 nM PCD, 2.5 mM PCA, and 0.1% BSA.

Molecular dynamics simulation
Tau and DNA were coarse-grained as 3.4 nm beads and simulated via Langevin dynamics in HOOMD-blue (see Supplementary Note 1 for more details). A 10 kb DNA strand was represented by 1000 beads connected by harmonic springs and an angular potential set to yield a 150 bp persistence length; tau was modeled using identical beads with short-range attractive interactions to DNA and to itself. Interaction parameters were calibrated so that tau phase separation occurred only in the presence of DNA, and control simulations were conducted with DNA bending rigidity removed. Simulations comprised two phases: first, DNA ends were held at 50% extension while the tau–DNA co-condensate was allowed to assemble and equilibrate; second, a constant force was applied to one DNA terminus, with the opposite end fixed, to probe the mechanical response of the stabilized co-condensate.

Microtubule capture assay
Tau protein was diluted to 5 μM in imaging buffer (same as above) supplemented with 5% 10 kDa PEG, and then slowly injected into a sample with biotinylated λ-DNA to form condensates. After co-condensate formation, HiLyte 488-labeled tubulin (Cytoskeleton, #TL488M-A) was mixed with unlabeled tubulin (Cytoskeleton, #T240-B), making up 4% of the total tubulin concentration. The mixture was prepared in imaging buffer supplemented with 1 mM GTP and 5% 10-kDa PEG to achieve a final tubulin concentration of 5–10 μM, while maintaining the tau concentration from the co-condensation experiments. After microtubule polymerization, the solution was slowly injected (7 μL min−1) into the chamber, with or without tau–DNA co-condensates. The assay with 12× 601 DNA (naked and nucleosomal) was identical to the above protocol except for three changes: (1) the flow chamber was additionally blocked with 5% (w/v) BSA for 15 min; (2) after blocking, buffers were switched to HEPES/NaCl instead of PBS; and (3) the fluorophore set was changed to Alexa 488 (Sigma, #A10254 1 mg)–H2A (nucleosomes), Cy3 (Cytiva #25800673)–tau, and HiLyte 647–tubulin (Cytoskeleton, #TL670M-A).

Immunofluorescence imaging
For mammalian expression of mCherry-tagged tau variants, tau inserts were PCR-amplified from the bacterial plasmid and inserted into a mammalian expression vector (pmCherry-C1) by Gibson assembly. HEK-293 cells (Korean cell line bank, #21573) were then reverse-transfected with the appropriate tau plasmids using polyethyleneimine (Sigma, #P3143). A total of 2.5 × 105 HEK-293 cells were seeded onto 35 mm glass-bottom confocal dishes (SPL, #210350) with 2 μg mL−1 mCherry–Tau plasmids and 4 μg mL−1 polyethyleneimine (PEI, #P3143-100ML) in DMEM (Sigma #D6429) supplemented with 10% FBS (Welgene, #S101), and incubated for 36 h. For SH-SY5Y cells (Korean cell line bank, #22266), 1 × 106 cells were seeded onto the same dishes in RPMI-1640 (Sigma #R8758-500ML) supplemented with 10% FBS and were incubated for 24 h. To synchronize the cells, they were treated with 2 mM thymidine (Sigma, #T9250) for 16 h to induce the first synchronization block, then released into their respective media (with 10% FBS) for 8 h. This was followed by a second 16 h thymidine block. After the second block, the cells were washed with PBS and released into their specific media with 10% FBS. Cells in the M phase were collected 9 h after release. The cells were then fixed with ice-cold methanol for 10 min, permeabilized with 0.25% Triton X-100 (Sigma, #X100) in PBS for 15 min, and blocked with 5% BSA in PBS with 0.05% Tween-20 (Sigma, #P1379) (PBST) for 1 h at RT. The full list of antibodies for staining is provided in the Supplementary Materials. Primary antibodies were diluted in PBST and incubated for 1 h at RT or overnight at 4°C. After washing in PBST, cells were incubated with secondary antibodies diluted in PBST for 1 h at RT. After antibody staining, the cells were washed in PBST and incubated with 1 mg mL−1 Hoechst 33258 (Sigma, #B2883-25mg) diluted 1:1,000 in PBST for 10 min at RT. Following a final wash in PBS, coverslips were mounted on confocal dishes using mounting solution (Agilent, #S203280-2). Finally, the cells were imaged using a Zeiss LSM900 confocal microscope equipped with Airyscan 2. Details of image analysis procedures and western blot methods (including antibodies) are provided in the Supplementary Materials.

RNA sequencing and analysis
RNA was extracted from mCherry–Tau-expressing HEK-293 cells and untreated HEK-293 cells for control. The extracted RNA was purified and sequenced using the NovaSeqX platform at Macrogen (Seoul, Korea). Paired-end raw sequencing reads were pre-processed with FastQC and trimmed using Trimmomatic-0.39. The cDNA reads were aligned with HISAT2.1, and differential expression was defined and quantified using the DESeq program with criteria of log2 fold change >0.7 or <−0.7, along with p values <0.05. To identify biological pathways associated with the differentially expressed genes, ShinyGO 0.80 was used for analysis with the Gene Ontology (GO) database. For the heatmap, log2 fold changes of genes corresponding to specific pathways were plotted, and a Venn diagram was generated for pathways involving downregulated genes.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In the manuscript by Park, et.al., the authors report a condensate like interaction between Tau and DNA that can also interact with microtubules. Overexpressed Tau protein localizes to centromeres in mitosis, and staining in cells that express endogenous Tau shows that it also localizes to centromeres in mitosis. Extensive biophysical investigation of these interactions reveals that Tau exerts a 1-3 pN resistive force when it binds to and gathers DNA into a LLPS fashion. Tau has been recently studied as a nucleator of microtubules likely by a local concentration increase (LLPS?) of free dimers. The same is found here where Tau/DNA condensates bind to and seem to promote nucleation of tubulin. Mutational analysis suggests that Tau nucleating microtubule assembly is mediated by the cell cycle, however binding to DNA seems to obviate this effect.\nOverall this is an interesting report that carefully examines Tau interacting with DNA and it’s continued role microtubule biology. I am generally in favor of acceptance for publication, however I have some serious concerns listed below that should be addressed prior to publication.\nMajor Concerns:\n1) My major concern here is the function of these phenomenon in normal cell division. These can be broken into sub-concerns:\na. Given that not all cells express Tau, however all cells divide, this would lend these observations irrelevant to normal cell division.\nb. Many of the in vitro experiments use a wide range of Tau concentrations, ranging by 3 orders of magnitude or more in some cases. What is the concentration of endogenous tau in cells that express it? Are these concentrations in any way physiological?\nc. Both DNA and MTs are acidic in nature. In the world of MT biology, it has long been known/observed that proteins and stains that bind MTs often also bind DNA and vise versa in high enough concentrations. This is true even with things like antibodies etc. It is not so surprising, knowing that, that Tau can bind DNA. It being “specific” to centromeres is significant and should be highlighted.\n2) The force curves are very interesting. Given the apparent LLPS nature of the particles in question, is the binding cooperative? It is unclear to me, however, might be of interest.\n3) Is there a way to evaluate this phenomenon in an Alzheimer’s model system? How to the authors feel this could play a role in disease?\nMinor Concerns:\n1) The methods sections are lacking in details. More details on imaging, including microscope settings (lens, etc?), source and lot number on antibodies used, etc.\n2) The imaging of cells seems really limited. In the supplementary material, it would be nice to see a field of cells stained for Tau so the reader can get a better feeling for this localization.\n3) The fact that many cells do not express Tau (or it can not be detected) needs more focus out of full discloser. I do not think the authors should shy away from this elephant in the room as it does not necessarily diminish their work in my opinion.", "reviewer_2": "The manuscript investigates a less explored nuclear role of Tau and describes its role in linking DNA condensation with microtubule formation. The authors found that Tau interacts with DNA, resulting in condensation and compaction of the DNA. This interaction, facilitates the incorporation of tubulin within the condensates, supporting the nucleation and growth of microtubule. Based on these findings, the authors propose an interesting model in which Tau’s condensation on DNA promotes DNA compaction and mitotic spindle formation. Further, they propose a role in Tau in cell division: when the nuclear membrane is fragmented, Tau binds DNA promoting condensation and microtubule attachment. Overall, the manuscript presents compelling experiments, and the conclusions are well-supported by the data.\nMajor comments:\n1. Validation of the proposed model in cells should be strengthen. First, authors should test the impact of Tau depletion Tau and/or expression of Tau mutants on spindle formation and/or chromosome segregation in cells that naturally express Tau. Second, including a centromere marker, such as CENPA, is important to claim the presence of Tau at centromeres, rather than relying solely on DAPI-dense regions.\n2. Line 130 “This suggests that, in addition to the tightly bound tau-DNA complexes within the core, a more loosely associated fraction of tau remains dynamic and capable of modulating DNA architecture through rearrangement, reminiscent of the liquid-like behavior characteristic of LLPS-driven condensates”. While the experiments demonstrate mobility of the DNA-Tau condensates, it is unclear if this reflects diffusion of the stable DNA-Tau condensates or Tau sliding along the DNA. This distinction should be clarified, as the current phrasing is ambiguous and could be misleading. Furthermore, the fraction of Tua mobile vs not mobile is not quantified to support the above claim. If the authors intent to make this statement, FRAP experiments are necessary.\n3. I suggest choosing a title that more precisely summarizes the findings of the study rather than implying a potential link with cell division, as this aspect is not experimentally tested. For example, a title like: \"Tau and DNA Condensation Promote Microtubule Formation\" would be more accurate.\nMinor comments:\n- It would be helpful to include a “conventional” phase separation assay where condensates are imaged on a passivated glass surface, instead of using tethered DNA.\n- Line 127: the phrase “despite the strong affinity between Tau and DNA” should be reconsidered since affinity is not measured. Consider removing this sentence or replacing “strong” with an appropriate Kd range.\n- The authors employ 100mM salt, and it is important to test physiological 150mM salt.\n- The observation that Tau bridges DNA and microtubules only when DNA-Tau condensates are pre-formed is interesting and could indicate a conformational change in Tau upon DNA binding. The authors could comment on this possibility in the discussion.\n- What is the size range of Tau-DNA condensates?", "reviewer_3": "In this study, Park et al. investigated DNA-Tau co-condensation primarily using single-molecule fluorescence microscopy and force spectroscopy. The authors report intriguing findings, such as the ability of these co-condensates to generate mechanical forces and interact with microtubules. Through fluorescence imaging, they observed colocalization between overexpressed Tau protein and nuclear DNA structures, proposing a potential role for Tau in mitosis. While I find the concept of this study compelling, I am not convinced that the novelty of biological discoveries and the depth of the study at the current stage meet the standards of Nature Communications. I do not recommend a publication, unfortunately. Please see specific critiques below.\n1. The authors reported Tau-naked DNA interactions at nanomolar concentrations, as previously demonstrated in a study published earlier this year (Abasi et al., PMID: 38187700). Several other claims and observations in the current work are closely related to findings from the Abasi et al. paper. However, the authors have not adequately credited this prior work.\n2. The authors observed local Tau-DNA condensation using lambda DNA. Based on the foci locations, they could potentially estimate the sequences or regions to which the Tau protein preferentially binds.\n3. Line 106, a reference or additional explanation is needed to justify the use of RMS roughness analysis for assessing local compaction, although the approach appears reasonable.\n4. In line 115, it is noted that Tau protein binds to naked DNA molecules at low concentrations, a phenomenon that has been previously reported (Abasi et al., PMID: 38187700).\n5. In line 121, reference 35 should be introduced earlier.\n6. In line 130, it is unclear how the authors define Tau-DNA as being at the core or loosely associated. As a result, I could not interpret the data in Figure 2H in the same way the authors did.\n7. In line 142, please clarify how the diffusion coefficient and diffusion exponent were measured. I could not find any details in the Methods section about how single-particle tracking was performed. I assume the authors calculated the MSD as a function of time.\n8. Measuring the piconewton forces applied by Tau-DNA condensation using magnetic tweezers is an excellent experiment. Could the authors speculate on the 72 nm step size observed during the force jump assay?\n9. During mitosis, chromosomes are highly condensed, and DNA is fully chromatinized. Can Tau bind to such DNA and remain bound? Is there any literature supporting this? In this study, the authors reported that Tau-naked DNA co-condensates capture microtubules. However, it is unclear how biologically relevant this observation is.\n10. The investigation of Tau-DNA-microtubule assemblies and their roles in cells is underdeveloped. Proper controls are missing to confirm a direct interaction between Tau and the centromere, despite the colocalization images provided.\n11. In lines 287 and 296, why are only two centromeres analyzed per cell, given that each cell contains so many more centromeres? Maybe I am missing something.\n12. Similarly, additional experiments can help elucidate the role of phosphorylation in co-condensates interacting with microtubules.\n13. Statistical analysis is missing for Figure 7D. Why not perform a Pearson correlation analysis to assess the colocalization between centromeres and Tau foci?\n14. Long-term cell imaging of Tau-mCherry through the mitotic process may provide more convincing evidence of Tau interacting with condensed DNA.\n15. The authors could consider expressing phosphomimetic Tau-mCherry in cells and comparing its dynamic behavior to wild-type Tau to assess potential differences in centromere colocalization. This approach may provide insights supporting a role for Tau in regulating mitosis.", "reviewer_4": "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 Reviewer Comments on:\nPark et al., “Tau condensation on DNA mediates\nmicrotubule attachment: A potential mitotic role for\ncentromere-localized tau”\nReviewer #1 (Remarks to the Author):\nIn the manuscript by Park, et.al., the authors report a condensate like interaction between Tau\nand DNA that can also interact with microtubules. Overexpressed Tau protein localizes to\ncentromeres in mitosis, and staining in cells that express endogenous Tau shows that it also\nlocalizes to centromeres in mitosis. Extensive biophysical investigation of these interactions\nreveals that Tau exerts a 1-3 pN resistive force when it binds to and gathers DNA into a LLPS\nfashion. Tau has been recently studied as a nucleator of microtubules likely by a local\nconcentration increase (LLPS?) of free dimers. The same is found here where Tau/DNA\ncondensates bind to and seem to promote nucleation of tubulin. Mutational analysis suggests\nthat Tau nucleating microtubule assembly is mediated by the cell cycle, however binding to\nDNA seems to obviate this effect.\nOverall this is an interesting report that carefully examines Tau interacting with DNA and it’s\ncontinued role microtubule biology. I am generally in favor of acceptance for publication,\nhowever I have some serious concerns listed below that should be addressed prior to\npublication.\n[Reply] We appreciate the positive feedback and thoughtful suggestions, which have greatly\nenhanced our work in this revision.\nMajor Concerns:\n1) My major concern here is the function of these phenomenon in normal cell division. These\ncan be broken into sub-concerns:\n1-a) Given that not all cells express Tau, however all cells divide, this would lend these\nobservations irrelevant to normal cell division.\n[Reply] We thank the reviewer for highlighting this point, which we address alongside a related\nminor comment by the same referee:\n3) The fact that many cells do not express Tau (or it can not be detected) needs more focus\nout of full discloser. I do not think the authors should shy away from this elephant in the room\nas it does not necessarily diminish their work in my opinion.\nCertainly, we modeled high-tau conditions using SH-SY5Y cells with endogenous tau and\nHEK cells engineered to overexpress tau. Thus, our findings are most relevant to neuronal\ncells and other contexts with elevated tau levels (e.g., certain cancers). In cells with low tau,\nother mitotic regulators likely predominate in spindle capture. It is also possible that other\nMAPs compensate for tau’s role, but this definitely warrants further investigation.\nTo clarify the reach and limitations of our findings, we have revised the manuscript as follows:\n• [Title change] We have updated the manuscript title to: “Tau condensation on DNA\nmediates microtubule attachment: A potential mitotic role for centromere-localized tau.”\n• [Text revised] We have made changes to limit the scope of our finding to tau-expressing\ncells, including neurons. Some snippets are appended below:\n[Introduction, Line 79] “Based on these findings, we speculate a new mitotic function of tau in tau-\no\nexpressing cells such as neurons.”\n[Results, Line 336] “(…) to hypothesize a role for tau in the division of tau-expressing cells, (…)”\no\n[Results, Line 338] “To test this in a neuronal setting, we conducted immunofluorescence imaging\no\nof SH-SY5Y cells, (…)”\n[Discussion, Line 555] “Because tau is not constitutively expressed in most cell types, its mitotic\no\nfunction via tau–DNA co-condensation is likely restricted to contexts with elevated tau levels. (…)”\n1-b) Many of the in vitro experiments use a wide range of Tau concentrations, ranging by 3\norders of magnitude or more in some cases. What is the concentration of endogenous tau in\ncells that express it? Are these concentrations in any way physiological?\n[Reply] Endogenous tau levels in the brain range between 1 and 7 μM (Binder et al., Journal\nof Cell Biology 101, 1371 (1985); Kanaan & Grabinski, Front. Mol. Neurosci. 14, (2021)),\nalthough this range varies depending on cell type and developmental stage. About 16% of this\npool has been reported to localize to the nucleus (Greenwood & Johnson, Exp. Cell Res. 220,\n332 (1995)), yielding sub-micromolar nuclear tau. In our single-molecule and bulk assays, we\nspanned 50 nM to 5 μM tau: the lower end models initial binding events at limited tau levels,\nwhile the upper range targets physiological concentrations. We used 5 μM tau throughout the\nmicrotubule capture assay to ensure we probe mitotically relevant condensation and affinity.\n• [Text revised] We have updated the manuscript to include this information on tau levels:\n[Discussion, Line 467] “With total tau levels in the brain ranging from 1 to 7 μM54–56 and a sub-\no\nmicromolar nuclear fraction57, the concentrations we tested (50 nM to 5 μM) span early tau–DNA\nbinding events at the low to physiological and pathologically elevated tau levels at the high end.”\n1-c) Both DNA and MTs are acidic in nature. In the world of MT biology, it has long been\nknown/observed that proteins and stains that bind MTs often also bind DNA and vise versa in\nhigh enough concentrations. This is true even with things like antibodies etc. It is not so\nsurprising, knowing that, that Tau can bind DNA. It being “specific” to centromeres is significant\nand should be highlighted.\n[Reply] To reflect this comment, we have revised the manuscript in two ways:\n• [Title change] As noted above, we revised the title to “Tau condensation on DNA mediates\nmicrotubule attachment: a potential mitotic role for centromere-localized tau,” explicitly\nhighlighting the centromere specificity of tau–DNA interactions.\n• [Text revised] We have added remarks on both the nonspecific electrostatic attraction\nbetween tau and DNA and the centromere-specific localization of tau in cells. The changes\nare highlighted in yellow in the manuscript. Some snippets are appended below:\n[Discussion, Line 447] “The microtubule-binding region (MTBR) of tau is enriched in basic residues\no\nthat likely confer an overall affinity for polyanions such as DNA, consistent with the salt dependence\nof tau–DNA binding (Fig. 2). (…)”\n[Discussion, Line 502] “The unexpected enrichment of tau at centromeres during prometaphase in\no\nboth SH-SY5Y (endogenous tau) and HEK-293 (exogenous tau) cells (Fig. 6) occurs despite tau’s\nlargely nonspecific affinity for DNA. (…)”\n• [Results added] To highlight the tau clusters associating with centromeres, we quantified\ntheir colocalization in our confocal images by calculating the Pearson correlation coefficient\nbetween tau and centromere fluorescence signals. The results clearly show a good overlap\nbetween the two locations, indicating specificity in binding.\n[Fig. 6C, D] Pearson correlation coefficients for tau\no\ncolocalization with tubulin and centromeres, calculated\nfrom the cell images used in (B) and (F). Box plots show\nmedian (red line), interquartile range, and whiskers to the\nminimum/maximum; each point represents one cell.\n[Results, Line 351] “Pearson correlation coefficients\no\ncalculated for each SH-SY5Y cell image confirmed\nstrong colocalization of tau with both tubulin and\ncentromeres (Fig. 6C, D).”\n• [Text revised] During this revision, we also found that nucleosomes serve as docking sites\nfor both tau and microtubules, which helps explain tau’s centromere-specific localization in\ncells, since centromeric regions are highly nucleosome-dense. We added text regarding this\npoint in Discussion:\n[Discussion, Line 504] “(…) Formation of nucleosome-pinned tau condensates (Fig. 5A, B), which\no\nretain the ability to mediate microtubule assembly, partially explains how tau condensates form\npreferentially on nucleosome-rich centromeres in dividing cells. (…)”\n2) The force curves are very interesting. Given the apparent LLPS nature of the particles in\nquestion, is the binding cooperative? It is unclear to me, however, might be of interest.\n[Reply] Indeed, the sharp, threshold-like onset of DNA compaction above ~500 nM tau (blue\ncurve in Fig. 3B) and the ensuing all-or-none collapse (Fig. 3J) demonstrate that tau–DNA\nbinding is cooperative rather than incremental. The subsequent DNA re-extension also occurs\nin sudden, quantized steps (Fig. 3E), showing that condensates dissociate in multi-molecular\nbursts. A similarly abrupt transition around 500 nM tau in both bulk (Fig. S2) and single-\nmolecule assays (Fig. 2C, E) further supports this cooperativity.\nTo validate these claims, we added new data, analyses, and comments:\n• [Results added] By measuring tau–DNA co-condensation with magnetic tweezers at more\nclosely spaced tau concentrations, we determined its Hill coefficient. The results indeed\nindicate that tau–DNA binding is very cooperative with a Hill coefficient of ~3.\n[Fig. 3K] Compaction length of DNA under 0.5 pN force clamp as a\no\nfunction of tau concentration. Black dots represent individual\ncondensation events (n = 15 per concentration; mean ± s.d.). The solid\nline represents a Hill equation fit, yielding a Hill coefficient of 3.1 ± 2.5\nand an apparent Kd of 0.55 ± 0.14 µM (values ± 95% confidence\nintervals).\n[Results, Line 234] “During the condensation measurements under a\no\nconstant 0.5 pN load, individual DNA tethers showed a binary response\n(Fig. 3J): they either collapsed rapidly into a condensed state (occasionally via discrete, slower\nsteps) or remained fully extended, with large fluctuations over 1 min. Both the switch-like collapse\nand the slower fluctuations likely stem from cooperative tau condensation: rapid compaction follows\ncluster formation, while the larger, slower changes reflect cluster-mediated remodeling. To quantify\nthis cooperativity, we measured the extent of DNA shortening as a function of tau concentration\nand observed a sharp transition near 500 nM (Fig. 3K). A Hill-plot analysis yielded a Hill coefficient\nof n = 3.1 ± 2.5 and an apparent Kd = 0.55 ± 0.14 μM (95% confidence interval), reinforcing the\nconclusion that multiple tau molecules bind DNA cooperatively with a moderate-to-high affinity.”\n[Discussion, Line 471] “Additionally, our mechanical analysis using magnetic tweezers (Fig. 3)\no\nshowed that tau binding to DNA is strongly cooperative, thus driving condensation below csat.”\n• [Results added] To gain molecular-level insight into tau–DNA co-condensation and its\nforce-induced decondensation, we ran MD simulations mirroring our magnetic-tweezers\nassays. These simulations showed that tau–DNA and tau–tau attractions alone drive the\nformation of local tau–DNA clusters, and that under force these clusters disassemble in\ndiscrete steps, consistent with a cooperatively assembled network that breaks apart\nmodularly.\n[Fig. 3G–I] Coarse-grained simulation of stretching tau–DNA co-condensates. (G) Representative\no\nsnapshot of a tau–DNA co-condensate captured before force application. The right image shows\nDNA alone in the same configuration for clarity. (H) DNA extension during force-induced\ndecondensation. Right, zoomed-in views of the boxed region, with snapshots of the DNA\nconformation before and after each stepwise jump. (I) Histograms of step sizes for extension events.\nThe persistence length of dsDNA (Lp) is indicated by the red dashed line for reference.\n[Supplementary Materials, Page 9] Supplementary Notes: Molecular dynamics simulation for\no\ntau–DNA co-condensates\n[Fig. S8] Molecular dynamics simulation of stretching tau–DNA co-condensates\no\n[Results, Line 224] “To test this interpretation at the molecular level, we performed MD simulations\no\nof tau–DNA co-condensation under applied force (Fig. S8). In silico, tau and DNA coalesce into\ndroplets featuring looped DNA segments and characteristic bending, without strict periodicity\n(Fig. 3G). Simulated force–extension profiles recapitulated the experimentally observed stepwise\njumps (Fig. 3H), and inspection of snapshots and local DNA densities before versus after individual\njumps confirmed that each event corresponds to the rupture of a compacted DNA domain (Fig. S8).\nImportantly, the distribution of simulated step sizes peaked near the DNA persistence length\n(Fig. 3I), demonstrating that DNA bending rigidity defines the natural rupture unit. In contrast,\nsimulations lacking bending energy (i.e., a freely jointed chain model) produced much smaller,\nirregular steps (Fig. S8).”\n[Discussion, Line 472] “Given the contour length of dsDNA (~0.34 nm/bp), the 50–70-nm\no\nincrements observed during the stretching of tau–DNA co-condensates (Fig. 3F) correspond to\n150–200 base pairs. This is intriguingly close to the persistence length of dsDNA (~150 bp),\nsuggesting that tau condensates compact DNA into domains of this characteristic size, set by the\nhelix’s bendability. Indeed, our simulations reproduce these steps of similar sizes (Fig. 3I), each\nreflecting the force-induced rupture of an individual tau–DNA cluster. (…) The all-or-none behavior\nduring the condensation phase also supports the cooperative nature of tau condensation on DNA,\nrapidly drawing nearby DNA segments into maturing tau condensates.”\n3) Is there a way to evaluate this phenomenon in an Alzheimer’s model system? How to the\nauthors feel this could play a role in disease?\n[Reply] It is indeed tempting to verify the implications of our findings for Alzheimer’s disease\n(AD). Although in vivo testing in models such as 5xFAD mice is beyond this study’s scope,\nwe\nexamined the effects of AD-associated phosphorylation by employing site-specific\nphosphomimetic variants.\n• [Results added] In our original submission, we demonstrated that the phosphomimetic\nmutants T231D/S235D and S262D differentially bridged microtubules and DNA. In this\nrevision, we extended our investigation to their effects on cell division by expressing these\nmutants in HEK-293 cells and comparing them to wild-type tau. The results are informative:\nthe phosphomimetic variants retained centromere localization (Fig. 7F–I) but impaired\nchromosome congression, yielding tau-associated misaligned chromosomes (Fig. 7J–L).\nWe speculate that altered partitioning of these mutants between microtubules and DNA\nunderlies this defect.\n[Fig. 7F–L] (F) Immunofluorescence images of prometaphase HEK-293 cells transiently expressing\no\nphosphomimetic (T231D/S235D and S262D) mCherry–Tau, showing the distribution of tubulin, tau,\nand centromeres. (G, H) Pearson correlation coefficients for tau colocalization with tubulin and\ncentromeres (n = 9 cells for T231D/S235D, n = 8 cells for S262D). Box plots show median (red\nline), interquartile range, and whiskers to the minimum/maximum; each point represents one cell.\nFor comparison, the wild-type tau data from Fig. 6C and 6D are reproduced here. (I) Intensity\nprofiles of centromere (blue) and tau (green) measured across aligned centromere regions,\nobtained from the cell images used in (G) and (H) (n = 429 centromeres for T231D/S235D, n = 288\ncentromeres for S262D). (J) Immunofluorescence of metaphase HEK-293 cells expressing WT or\nphosphomimetic mCherry–Tau. DNA image (Hoechst 33258-stained) is overlaid in gray for\nreference. Misaligned DNA foci away from the metaphase plate is outlined in yellow. (K, L)\nQuantification of misaligned DNA (Hoechst 33258) and tau (mCherry) intensities, normalized to\neach cell’s total signal (n = 6 images each; mean ± s.d.).\n[Results, Line 419] Phosphomimetic tau localizes to centromeres but impairs chromosome\no\nalignment\n“To test whether the in vitro differences in DNA–microtubule interfacing by phosphomimetic tau\nmutants manifest in cells, we performed immunofluorescence on mitotic HEK-293 cells transiently\nexpressing mCherry-tagged T231D/S235D or S262D tau, using the same approach as for wild-type\ntau (Fig. 6). Although both mutants largely retained microtubule binding during mitosis (Fig. 7F),\nPearson correlation analysis revealed their reduced association with spindle microtubules (Fig. 7G),\nconsistent with the impaired microtubule-stabilizing activity (Fig. 7B). Nevertheless, both mutants\nexhibited robust centromeric localization comparable to wild-type tau (Fig. 7H), reflecting their\nretained DNA-binding activity (Fig. 7A). Analysis of averaged centromere images further confirmed\noverlapping tau signal (Fig. 7I).\nSurprisingly, in metaphase cells after chromosomes have aligned at the metaphase plate, both\nphosphomimetic mutants showed prominent tau-rich chromosomal regions that failed to align\n(Fig. 7J). These tau–DNA clusters did not colocalize with the central tubulin signal, indicating a lack\nof spindle attachment, and were most frequent in S262D-expressing cells, implicating aberrant tau–\nDNA binding in defective microtubule capture. Quantitative analysis of DNA and tau amount\n(measured by Hoechst 33258 and mCherry fluorescence, respectively) confirmed these findings.\nIn S262D cells, misaligned DNA foci comprised about 10% of total chromosomal DNA, significantly\nmore than in wild-type or T231D/S235D cells (Fig. 7K; see Fig. S12 and Supplementary Methods\nfor analysis). Tau fluorescence at these foci accounted for nearly 30% of total cellular tau in S262D\ncells—over a five-fold increase versus wild type (Fig. 7L). These data demonstrate that\nphosphomimetic tau mutants not only lose their ability to engage spindle microtubules but also\naberrantly accumulate on misaligned chromatin, thus compromising proper chromosome\nsegregation.”\n[Discussion, Line 542] “The failure of some chromosomes to align at the metaphase plate in cells\no\nexpressing Alzheimer’s-associated phosphomimetic tau mutants is informative in this regard\n(Fig. 7J–L). It suggests that altered phosphorylation could disrupt tau’s dynamic partitioning\nbetween DNA and microtubules, remain abnormally coated on chromatin, and escape timely\nspindle capture, potentially contributing to chromosome missegregation and aneuploidy. These\ncell-based findings not only validate the biophysical function of tau observed in vitro but also reflects\nthe loss-of-function effect seen in microtubule capture assays. This bidirectional correspondence\nstrongly reinforces the idea that site-specific phosphorylation of tau disrupts its mitotic function.”\nMinor Concerns:\n1) The methods sections are lacking in details. More details on imaging, including microscope\nsettings (lens, etc?), source and lot number on antibodies used, etc.\n[Reply] We have updated the Materials and Methods section, including Supplementary\nMaterials, with additional details:\n• [Text revised] More details on imaging setup and materials have been added:\n[Materials and Methods, Line 633] “Single-molecule imaging. (…) All images were taken on a\no\nhome-built objective-type TIRF microscopy setup. Samples were illuminated with 488 nm, 532 nm,\nand 633 nm CW lasers (Cobolt) and imaged through a 60× oil-immersion lens (Olympus, NA 1.49\nfor TIRF microscopy) onto a high-sensitivity sCMOS camera (Photometrics Prime BSI Express).\nImages were acquired typically at 108-nm spatial resolution with 100-ms exposure time using\ncustom LabVIEW software. When necessary, fluorescence in two channels was separated by a\ndual-view image splitter (Cairn OptoSplit II) and recorded simultaneously. Additional methods for\ncharacterizing tau–DNA co-condensation, including mobility shift and bulk LLPS assays, FRAP,\nand sequence-dependence measurements, are described in the Supplementary Methods.”\n[Materials and Methods, Line 708] “Immunofluorescence imaging. (…) The full list of antibodies\no\nfor staining is provided in the Supplementary Materials. Primary antibodies were diluted in PBST\nand incubated for 1 h at RT or overnight at 4 °C. After washing in PBST, cells were incubated with\nsecondary antibodies diluted in PBST for 1 h at RT. After antibody staining, the cells were washed\nin PBST and incubated with 1 mg/mL Hoechst 33258 (Sigma) diluted 1:1000 in PBST for 10 min at\nRT. Following a final wash in PBS, coverslips were mounted on confocal dishes using mounting\nsolution (Agilent, S203280-2). Finally, the cells were imaged using a Zeiss LSM 900 confocal\nmicroscope equipped with Airyscan 2. Details of image analysis procedures and western blot\nmethods (including antibodies) are provided in the Supplementary Materials.”\n[Supplementary Materials, Page 2]\no\n List of PCR primers and oligonucleotides\n List of antibodies for immunofluorescence imaging\n List of antibodies for western blot\n2) The imaging of cells seems really limited. In the supplementary material, it would be nice to\nsee a field of cells stained for Tau so the reader can get a better feeling for this localization.\n[Reply] To reflect this comment, we have added a large, zoomed-out field of cell images:\n[Fig. S11A] (A) Overview of immunofluorescence images of native SH-SY5Y cells and HEK-293\no\ncells expressing wild-type 2N4R mCherry–Tau. Cells were synchronized using a double-thymidine\nblock protocol. Distributions of tubulin (red, Alexa 647), tau (green, Alexa 568 for SH-SY5Y and\nmCherry for HEK-293), centromeres (blue, Alexa 488), and DNA (gray, Hoechst 33258) are shown.\nScale, 10 μm.\n[Results, Line 340] “Immunostaining of thymidine-synchronized, dividing cells (identified by\no\ncondensed chromosomes) revealed a consistent level of endogenous tau forming small puncta with\ndistinct distributions across mitotic stages (Fig. S11).”\n3) The fact that many cells do not express Tau (or it can not be detected) needs more focus\nout of full discloser. I do not think the authors should shy away from this elephant in the room\nas it does not necessarily diminish their work in my opinion.\n[Reply] This comment has been addressed with the Major Concern 1-a) above.\nReviewer #2 (Remarks to the Author):\nThe manuscript investigates a less explored nuclear role of Tau and describes its role in linking\nDNA condensation with microtubule formation. The authors found that Tau interacts with DNA,\nresulting in condensation and compaction of the DNA. This interaction, facilitates the\nincorporation of tubulin within the condensates, supporting the nucleation and growth of\nmicrotubule. Based on these findings, the authors propose an interesting model in which Tau’s\ncondensation on DNA promotes DNA compaction and mitotic spindle formation. Further, they\npropose a role in Tau in cell division: when the nuclear membrane is fragmented, Tau binds\nDNA promoting condensation and microtubule attachment. Overall, the manuscript presents\ncompelling experiments, and the conclusions are well-supported by the data.\n[Reply] We appreciate the positive feedback and thoughtful suggestions, which have greatly\nenhanced our work in this revision.\nMajor comments:\n1. Validation of the proposed model in cells should be strengthen. First, authors should test\nthe impact of Tau depletion Tau and/or expression of Tau mutants on spindle formation and/or\nchromosome segregation in cells that naturally express Tau. Second, including a centromere\nmarker, such as CENPA, is important to claim the presence of Tau at centromeres, rather than\nrelying solely on DAPI-dense regions.\n[Reply] We thank the reviewer for this suggestion. To further validate our model in cells, we\nfirst attempted to knock down endogenous tau in SH-SY5Y cells but were unable to obtain\nreliable results. This may be because tau’s critical role in stabilizing the microtubule network\ncomplicates assessment of its proposed DNA-binding function. Instead, we tested the effects\nof phosphomimetic tau mutants (T231D/S235D and S262D) in HEK cells (a tau-free\nbackground to minimize confounding effects):\n• [Results added] To assess the cellular effects of phosphomimetic tau mutants, we\nexpressed them in HEK 293 cells and compared them with wild-type tau. The results are\ninformative: the phosphomimetic variants retained centromere localization (Fig. 7F–I) but\nimpaired chromosome congression, yielding tau-associated misaligned chromosomes\n(Fig. 7J–L). We speculate that altered partitioning of these mutants between microtubules\nand DNA underlies this defect.\n[Fig. 7F–L] (F) Immunofluorescence images of prometaphase HEK-293 cells transiently expressing\no\nphosphomimetic (T231D/S235D and S262D) mCherry–Tau, showing the distribution of tubulin, tau,\nand centromeres. (G, H) Pearson correlation coefficients for tau colocalization with tubulin and\ncentromeres (n = 9 cells for T231D/S235D, n = 8 cells for S262D). Box plots show median (red\nline), interquartile range, and whiskers to the minimum/maximum; each point represents one cell.\nFor comparison, the wild-type tau data from Fig. 6C and 6D are reproduced here. (I) Intensity\nprofiles of centromere (blue) and tau (green) measured across aligned centromere regions,\nobtained from the cell images used in (G) and (H) (n = 429 centromeres for T231D/S235D, n = 288\ncentromeres for S262D). (J) Immunofluorescence of metaphase HEK-293 cells expressing WT or\nphosphomimetic mCherry–Tau. DNA image (Hoechst 33258-stained) is overlaid in gray for\nreference. Misaligned DNA foci away from the metaphase plate is outlined in yellow. (K, L)\nQuantification of misaligned DNA (Hoechst 33258) and tau (mCherry) intensities, normalized to\neach cell’s total signal (n = 6 images each; mean ± s.d.).\n[Results, Line 419] Phosphomimetic tau localizes to centromeres but impairs chromosome\no\nalignment\n“To test whether the in vitro differences in DNA–microtubule interfacing by phosphomimetic tau\nmutants manifest in cells, we performed immunofluorescence on mitotic HEK-293 cells transiently\nexpressing mCherry-tagged T231D/S235D or S262D tau, using the same approach as for wild-type\ntau (Fig. 6). Although both mutants largely retained microtubule binding during mitosis (Fig. 7F),\nPearson correlation analysis revealed their reduced association with spindle microtubules (Fig. 7G),\nconsistent with the impaired microtubule-stabilizing activity (Fig. 7B). Nevertheless, both mutants\nexhibited robust centromeric localization comparable to wild-type tau (Fig. 7H), reflecting their\nretained DNA-binding activity (Fig. 7A). Analysis of averaged centromere images further confirmed\noverlapping tau signal (Fig. 7I).\nSurprisingly, in metaphase cells after chromosomes have aligned at the metaphase plate, both\nphosphomimetic mutants showed prominent tau-rich chromosomal regions that failed to align\n(Fig. 7J). These tau–DNA clusters did not colocalize with the central tubulin signal, indicating a lack\nof spindle attachment, and were most frequent in S262D-expressing cells, implicating aberrant tau–\nDNA binding in defective microtubule capture. Quantitative analysis of DNA and tau amount\n(measured by Hoechst 33258 and mCherry fluorescence, respectively) confirmed these findings.\nIn S262D cells, misaligned DNA foci comprised about 10% of total chromosomal DNA, significantly\nmore than in wild-type or T231D/S235D cells (Fig. 7K; see Fig. S12 and Supplementary Methods\nfor analysis). Tau fluorescence at these foci accounted for nearly 30% of total cellular tau in S262D\ncells—over a five-fold increase versus wild type (Fig. 7L). These data demonstrate that\nphosphomimetic tau mutants not only lose their ability to engage spindle microtubules but also\naberrantly accumulate on misaligned chromatin, thus compromising proper chromosome\nsegregation.”\n[Discussion, Line 542] “The failure of some chromosomes to align at the metaphase plate in cells\no\nexpressing Alzheimer’s-associated phosphomimetic tau mutants is informative in this regard\n(Fig. 7J–L). It suggests that altered phosphorylation could disrupt tau’s dynamic partitioning\nbetween DNA and microtubules, remain abnormally coated on chromatin, and escape timely\nspindle capture, potentially contributing to chromosome missegregation and aneuploidy. These\ncell-based findings not only validate the biophysical function of tau observed in vitro but also reflects\nthe loss-of-function effect seen in microtubule capture assays. This bidirectional correspondence\nstrongly reinforces the idea that site-specific phosphorylation of tau disrupts its mitotic function.”\nRegarding the reviewer’s second point on centromere labeling, we appreciate the\nopportunity to clarify. In all relevant figures (Figs. 6A–D, 7F, 7I, and 7J), centromeres were\nspecifically labeled with anti-centromere (CREST) antibody. The DNA stain (Hoechst 33258)\nwas not shown in the original manuscript but included in this revision for Fig. 7J, only to\nshow overall chromosome distribution during mitosis and may have led to confusion. We\napologize and have clarified the text accordingly:\n• [Text revised] We clarified centromere versus DNA\nimages:\n[Fig. 6] (A) Immunofluorescence images of prometaphase\no\nSH-SY5Y cells endogenously expressing tau showing the\ndistribution of tubulin, tau, and centromeres. (…) In (A)–(D),\ncentromeres were specifically labeled using an anti-\ncentromere (CREST) antibody.\n[Fig. 7] (F) Immunofluorescence images of\no\nprometaphase HEK-293 cells transiently expressing\nphosphomimetic (T231D/S235D and S262D) mCherry–\nTau, showing the distribution of tubulin, tau, and\ncentromeres. (…) (J) Immunofluorescence of\nmetaphase HEK-293 cells expressing WT or\nphosphomimetic mCherry–Tau. DNA image (Hoechst\n33258-stained) is overlaid in gray for reference.\nMisaligned DNA foci away from the metaphase plate is\noutlined in yellow. In (F), (I), and (J), centromeres were\nspecifically labeled using an anti-centromere (CREST) antibody.\n[Supplementary Materials, Page 2]\no\n List of antibodies for immunofluorescence imaging:\nFor HEK-293 cells expressing mCherry–Tau:\nHuman anti-CREST (#15-234, Antibodies\nIncorporated, 1:200 dilution)\n2. Line 130 “This suggests that, in addition to the tightly bound tau-tau-DNA complexes within the\ncore, a more loosely associated fraction of tau remains dynamic and capable of modulating\nDNA architecture through rearrangement, reminiscent of the liquid-like behavior characteristic\nof LLPS-driven condensates”. While the experiments demonstrate mobility of the DNA-Tau\ncondensates, it is unclear if this reflects diffusion of the stable DNA-Tau condensates or Tau\nsliding along the DNA. This distinction should be clarified, as the current phrasing is\nambiguous and could be misleading. Furthermore, the fraction of Tua mobile vs not mobile is\nnot quantified to support the above claim. If the authors intent to make this statement, FRAP\nexperiments are necessary.\n[Reply] The previous wording mistakenly implied that stable tau–DNA complexes themselves\nrelocate. As the reviewer noted, it is actually mobile tau clusters that slide along DNA,\ncompacting underlying segments so that the site of co-condensation appears to migrate.\n• [Text revised] We clarified the observation:\n[Section title, Line 155] Now reads: “Tau condensates slide along DNA to compact local\no\nsegments”\n[Results, Line 157] “Moreover, the location of DNA foci also followed tau movement across all tau\no\nconcentrations we tested (50 nM – 5 μM). This suggests that tau clusters can slide along DNA to\ndynamically remodel the underlying segments, reminiscent of other LLPS-driven protein–DNA co-\ncondensates.”\nThe reviewer additionally recommended FRAP experiments to estimate the mobile fraction of\nDNA-bound tau, which we have now performed and included.\n• [Results added] Although the level of recovery was rather ill-defined (these\ncondensates frequently moved, merged, and split) and thus heterogeneous across\ncondensates, we observed consistent fluorescence recovery, indicating that a substantial\nfraction of DNA-bound tau is liquid-like and dynamically exchanges with bulk tau.\n[Supplementary Materials, Page 3] Fluorescence recovery after photobleaching (FRAP)\no\n“To probe the internal dynamics of tau–DNA condensates, we performed FRAP on individual tau\nclusters on double-tethered λ-DNA, assembled with 5 μM tau. (…)”\n[Fig. S5] FRAP analysis of DNA-bound tau condensates\no\n(A) Representative tau\nfluorescence along λ-DNA\nsubstrates (green dashed\noutline) before bleaching\nand after recovery for 0, 5,\n10 and 20 min. All tau\nfluorescence on the DNA\nwas fully photobleached\nat t = 0 min. (B) Normalized fluorescence intensity of individual tau condensates (colored traces, n\n= 5) showing heterogeneous recovery kinetics. (C) Normalized fluorescence recovery over time.\nPink shading represents the mean normalized intensity ± SD (individual data not shown), and the\nblue curve is a single-exponential fit to the recovery (τ ~ 30 min). Horizontal dashed lines indicate\nthe post-bleach intensity (Ibleach = 0.13) and the plateau intensity (I∞ = 0.84). Assuming a normalized\npre-bleach intensity (Ipre = 1), the average mobile fraction was calculated (I∞ − Ibleach )/(Ipre − Ibleach )\n= 0.82 (82%).\n[Results, Line 163] “To probe the internal dynamics of tau clusters, we performed fluorescence\no\nrecovery after photobleaching (FRAP) on tau clusters assembled with 5 μM tau along double-\ntethered λ-DNA (Fig. S5). Bleaching and monitoring of tau fluorescence (Cy5) recovery yielded\nhighly heterogeneous traces, reflecting frequent condensate movement, merging, and splitting, yet\nmany traces exhibited substantial fluorescence return. A single-exponential fit of the mean recovery\nfor representative traces gave a time constant of ~30 min and a mobile fraction of ~82%. However,\nthese values represent a composite of fast- and slow-recovering populations and are affected by\nthe loss of some condensates during monitoring. Overall, these results confirm that DNA-bound tau\nexchanges dynamically with the bulk pool in a liquid-like manner.”\n3. I suggest choosing a title that more precisely summarizes the findings of the study rather\nthan implying a potential link with cell division, as this aspect is not experimentally tested.\nFor example, a title like: \"Tau and DNA Condensation Promote Microtubule Formation\" would be\nmore accurate.\n[Reply] We respect the suggestion to adopt a more neutral title, and addressed it in two ways:\n• [Title change] To narrow the scope and focus on our findings, we have revised the title to:\n“Tau condensation on DNA mediates microtubule attachment: A potential mitotic role for\ncentromere-localized tau.”\n• [Results added] As we have attached above in response to Comment #1, we have now\ndirectly tested the proposed mitotic function of tau by expressing phosphomimetic mutants\nin cells. As shown in Figs. 7F–I, these variants still localize to centromeres, yet they produce\na clear defect in chromosome congression, yielding tau-associated misaligned\nchromosomes (Figs. 7J–L). These findings provide the first experimental evidence that\ncentromere-localized tau can influence mitotic chromosome behavior. Accordingly, we have\nrefined the title as above to retain the minimal mitotic reference supported by our data.\nMinor comments:\n1. It would be helpful to include a “conventional” phase separation assay where condensates\nare imaged on a passivated glass surface, instead of using tethered DNA.\n[Reply] Although these bulk phase separation data were already included (currently Fig. S4),\nthey may not have received the attention they deserve.\n• [Text revised] We have now added an explicit citation to Fig. S4 in the main Results text,\nso that readers immediately see our LLPS assay alongside the single-molecule\nmeasurements:\n[Fig. S4] Co-condensation of tau and DNA in bulk via LLPS\no\n(A) Co-condensation of tau (5 μM) and λ-DNA (10 ng/μL) observed with 10% PEG. (B) Merging of\ntwo co-condensates, characteristic of liquid droplets. (C-E) Effects of tau, DNA, and salt\nconcentrations, with λ-DNA and tau kept at 10 ng/μL and 5 μM respectively. (F) Effect of DNA length,\nwith 1 ng/μL DNA and 5 μM tau. Scale: 2 μm in (B), 5 μm for the rest.\n[Results, Line 145] “To ensure our findings on tau–DNA co-condensation extend beyond single-\no\nmolecule observations, we checked bulk phase separation and subsequent co-condensation of\nhigh tau concentrations with dsDNA in the presence of 10% PEG, a standard crowding agent to\ndrive LLPS. Under these conditions, tau and DNA readily formed co-condensates across various\nconcentrations of tau, DNA, and salt (Fig. S4), again consistent with the previous results25.”\n2. Line 127: the phrase “despite the strong affinity between Tau and DNA” should be\nreconsidered since affinity is not measured. Consider removing this sentence or replacing\n“strong” with an appropriate Kd range.\n[Reply] To obtain a ballpark estimate of tau’s affinity for DNA, we titrated tau in our magnetic\ntweezers assay and obtained an apparent K of 0.55 ± 0.14 μM with a high Hill coefficient,\nd\nindicating moderate-to-high affinity. We call this value “apparent” because it reflects tau-driven\ncondensation rather than a simple two-body binding equilibrium, and because the high Hill\ncoefficient complicates precise K determination. Nonetheless, this estimate aligns with the\nd\ntau concentrations at which single-molecule imaging shows a sharp increase in binding and\ngel assays exhibit corresponding DNA mobility shifts.\n• [Results added] K and cooperativity measurements by magnetic tweezers have been\nd\nincluded.\n[Fig. 3K] (K) Compaction length of DNA under 0.5 pN force clamp as a\no\nfunction of tau concentration. Black dots represent individual\ncondensation events (n = 15 per concentration; mean ± s.d.). The solid\nline represents a Hill equation fit, yielding a Hill coefficient of 3.1 ± 2.5\nand an apparent Kd of 0.55 ± 0.14 µM (values ± 95% confidence\nintervals).\n[Results, Line 234] “During the condensation measurements under a\no\nconstant 0.5 pN load, individual DNA tethers showed a binary response\n(Fig. 3J): they either collapsed rapidly into a condensed state (occasionally via discrete, slower\nsteps) or remained fully extended, with large fluctuations over 1 min. Both the switch-like collapse\nand the slower fluctuations likely stem from cooperative tau condensation: rapid compaction follows\ncluster formation, while the larger, slower changes reflect cluster-mediated remodeling. To quantify\nthis cooperativity, we measured the extent of DNA shortening as a function of tau concentration\nand observed a sharp transition near 500 nM (Fig. 3K). A Hill-plot analysis yielded a Hill coefficient\nof n = 3.1 ± 2.5 and an apparent Kd = 0.55 ± 0.14 μM (95% confidence interval), reinforcing the\nconclusion that multiple tau molecules bind DNA cooperatively with a moderate-to-high affinity.”\n• [Text revised] We removed the phrase “despite the strong affinity” that still appears before\nour K measurements. Instead, we now emphasize the rapid kinetics of tau–DNA association\nd\nobserved in single-molecule binding assays (Fig. 1):\n[Results, Line 156] “Surprisingly, despite the fast accumulation of tau on DNA (Fig. 1B), many of\no\nthe tau condensates remained mobile along the DNA substrates (Fig. 2H and Movie S3). (…)”\n3. The authors employ 100mM salt, and it is important to test physiological 150mM salt.\n[Reply] As suggested, we repeated the single-molecule experiments under 150 mM NaCl\nconditions and largely recapitulated the previously observed tau–DNA co-condensation\nbehavior.\n• [Results added] Because Fig. 2 is already crowded, we have included the results obtained\nunder 150 mM NaCl conditions in the supplement (Fig. S3). These data confirm tau binding\nto DNA, colocalization of tau with DNA clusters, and a reduction in DNA fluctuations:\n[Fig. S3] Verification of tau–DNA co-condensation at 150 mM NaCl\no\n(A) TIRF images of doubly tethered λ-DNA in the absence and presence of 5 μM tau at 150 mM\nNaCl. DNA is labeled with SYTOX Orange (green) and tau with Cy5 (magenta). (B) Intensity profiles\nalong λ-DNA at 5 μM tau, demonstrating colocalization between DNA and tau signals. (C)\nQuantification of tau fluorescence intensity for conditions shown in (A). (D) Pearson correlation\ncoefficients between DNA and tau fluorescence intensities, indicating their spatial colocalization.\n(E) Coefficient of variation of DNA fluorescence images, reflecting the extent of DNA puncta\nformation induced by tau. (F) Maximum intensity projections of DNA fluctuations over 5 s, illustrating\ntau's effect on DNA flexibility. Magenta lines indicate measured envelope widths. (G) Distribution of\nDNA envelope width with and without tau, quantifying the reduction in DNA fluctuations due to tau\nbinding at 150 mM NaCl.\n[Results, Line 141] “Importantly, these findings were confirmed under 150 mM NaCl, which more\no\nclosely approximates physiological salt conditions (Fig. S3). These results demonstrate that wild-\ntype tau protein at nano- to micromolar concentrations can stably bind to naked DNA under\nphysiological conditions, consistent with the behavior reported by Abasi et al.25”\n4. The observation that Tau bridges DNA and microtubules only when DNA-Tau condensates\nare pre-formed is interesting and could indicate a conformational change in Tau upon DNA\nbinding. The authors could comment on this possibility in the discussion.\n[Reply] We appreciate the reviewer’s suggestion regarding DNA-induced conformational\nchanges in tau. Previous NMR studies have identified key DNA-binding residues in the P2 and\nR2 regions, which are adjacent to or overlap with the microtubule-binding region (MTBR). This\nrationalizes the possibility that DNA binding could help reorganize MTBR to promote\nmicrotubule interactions. At the same time, our work points to a larger-scale mechanism in\nwhich many tau molecules in a condensate collectively present MTBRs to the microtubule\nlattice. These two models are certainly not mutually exclusive.\n• [Text revised] We have revised the Discussion to incorporate both: local conformational\npriming of tau by DNA and cooperative recruitment via condensate assembly:\n[Discussion, Line 493] “This indicates that tau condensates on DNA are fundamentally distinct from\no\nthose formed with tubulin, which may be explained by conformational changes in tau upon DNA\nbinding that enhance microtubule interaction. Indeed, the DNA-binding region of tau is proposed to\npartially overlap with MTBR24, supporting the conformational priming model. Beyond this monomer-\nlevel mechanism, we favor a larger-scale, multivalent mode of affinity arising from clustered tau\nmolecules, potentially involving a composite tau–DNA interface. These two mechanisms\n(conformational priming and condensate-mediated multivalent recruitment) are not mutually\nexclusive and likely act together to promote robust microtubule bridging.”\n5. What is the size range of Tau-DNA condensates?\n[Reply] The condensates are highly heterogeneous in size: some fuse into larger assemblies,\nwhile the smallest clusters fall below our diffraction-limited resolution and cannot be\nindividually resolved. Nevertheless, intensity cross sections show that the majority of\nresolvable condensates are on the order of 1 μm in diameter.\n• [Text revised] We have included this information:\n[Results, Line 124] “The DNA backbone was diffusely coated with tau, interspersed with multiple\no\nbright foci of approximately 1 μm or smaller, as revealed by the intensity profile cross section\n(Fig. 2B, inset).”\nReviewer #3 (Remarks to the Author):\nIn this study, Park et al. investigated DNA-Tau co-condensation primarily using single-\nmolecule fluorescence microscopy and force spectroscopy. The authors report intriguing\nfindings, such as the ability of these co-condensates to generate mechanical forces and\ninteract with microtubules. Through fluorescence imaging, they observed colocalization\nbetween overexpressed Tau protein and nuclear DNA structures, proposing a potential role\nfor Tau in mitosis. While I find the concept of this study compelling, I am not convinced that\nthe novelty of biological discoveries and the depth of the study at the current stage meet the\nstandards of Nature Communications. I do not recommend a publication, unfortunately.\nPlease see specific critiques below.\n[Reply] We appreciate this evaluation and have substantially revised the manuscript,\nadding\nmultiple new data sets and analyses that, hopefully, address the concerns and meet the\njournal’s standards.\n1. The authors reported Tau-naked DNA interactions at nanomolar concentrations, as\npreviously demonstrated in a study published earlier this year (Abasi et al., PMID: 38187700).\nSeveral other claims and observations in the current work are closely related to findings from\nthe Abasi et al. paper. However, the authors have not adequately credited this prior work.\n[Reply] We address this comment alongside a related note by the same referee:\n4. In line 115, it is noted that Tau protein binds to naked DNA molecules at low concentrations,\na phenomenon that has been previously reported (Abasi et al., PMID: 38187700).\nThe paper by Abasi et al. appeared during the final stages of our study and indeed motivated\nseveral of our experiments. Although we had cited it, we regret not emphasizing its influence\nmore clearly. We have now integrated this reference prominently in the Introduction and\nDiscussion and hope this clarifies its contribution to our work.\n• [Text revised] Manuscript updated to include additional references to Abasi et al. (currently\nRef. 25):\n[Introduction, Line 61] “A recent biophysical investigation by Abasi et al. demonstrated that tau can\no\nco-condense with naked DNA and nucleosome arrays under low-salt conditions25, which can be\ndisrupted by aberrant hyperphosphorylation of tau.”\n[Results, Line 96] “A recent report on tau’s phase separation behavior with naked DNA25 is in\no\nquantitative agreement with our results.”\n[Results, Line 113] “This indicates that a stable interaction between tau and naked DNA at\no\nnanomolar concentrations, which locally enriches tau, is sufficient to induce DNA compaction prior\nto the onset of LLPS-dependent tau–DNA co-condensation25.”\n[Results, Line 141] “Importantly, these findings were confirmed under 150 mM NaCl, which more\no\nclosely approximates physiological salt conditions (Fig. S3). These results demonstrate that wild-\ntype tau protein at nano- to micromolar concentrations can stably bind to naked DNA under\nphysiological conditions, consistent with the behavior reported by Abasi et al.25”\n[Results, Line 302] “This high degree of colocalization suggests that tau–DNA co-condensation\no\nresponds to nucleosome landmarks, consistent with recent bulk studies showing tau co-\ncondensation with nucleosome arrays25, and implies that mobile tau clusters are pinned at\nnucleosome sites.”\n[Discussion, Line 449] “A previous NMR study mapped the DNA-binding region of tau to the proline-\no\nrich domain and R2 region24, and a recent biophysical study showed a strong salt dependence of\ntau–DNA and tau–chromatin associations25, consistent with an electrostatic picture.”\n[Discussion, Line 461] “The liquid-like properties of tau clusters on DNA, including their mobility\no\n(Fig. 2) and content exchange (Fig. S5), are reminiscent of phase-separated condensates in free\nsolution, as systematically investigated by Abasi et al.25”\n[Materials and Methods, Line 602] “Phosphorylation of wild-type tau was performed in vitro following\no\nAbasi et al.25 (…)”\n2. The authors observed local Tau-DNA condensation using lambda DNA. Based on the foci\nlocations, they could potentially estimate the sequences or regions to which the Tau protein\npreferentially binds.\n[Reply] Following the reviewer’s suggestion, we examined the distribution of tau–DNA\ncondensates on λ-DNA using the DNA skybridge platform:\n• [Results added] To determine condensate sequence preferences, we modified the\nskybridge flow-stretch assay by integrating the DNA alignment method of Kopūstas et al.\n(Langmuir 2021, doi: 10.1021/acs.langmuir.1c00066), ensuring that all λ-DNA molecules\nwere oriented in one direction.\n[Supplementary Methods, Page 4] “Sample preparation and imaging for DNA skybridge\no\nexperiments. (…) To orient λ-DNA in one direction for sequence-preference measurements, a\ndigoxigenin(dig)-tagged oligonucleotide targeting the cosR end was used in place of the biotin-\ntagged one, following Kopu̅stas et al.2 (…)”\n[Fig. S6] Sequence preference of tau condensate location\no\n(A) Schematic of the skybridge assay, in which λ-DNA molecules are aligned in one direction. (B)\nTop: GC content (%) along λ-DNA plotted against genomic coordinates (kbp, top axis, from cosL to\ncosR end) and normalized position (bottom axis); red shading marks the inaccessible regions\nblocked by quartz ridges. Bottom: Histogram of condensate positions relative to the tether center,\nshowing a positional bias toward the GC-rich region on the left (green) compared to the right (blue).\nInset pie chart indicates the fraction of condensates in each half.\n[Results, Line 182] “The skybridge platform further provided a convenient means to probe the\no\nsequence preference of tau condensate localization (Fig. S6). Condensates were observed more\nfrequently in the AT-rich half of λ-DNA, consistent with tau’s known preference for AT-rich\nsequences38, although the overall broad distribution reflects its predominantly nonspecific binding.”\n[Discussion, Line 454] “Interestingly, despite the largely nonspecific nature of tau–DNA interactions,\no\nwe observed a slight preference for AT-rich regions in condensate positioning (Fig. S6), in\nagreement with gel assays and tau’s association with α-satellite DNA sequences38. Because tau-\ninduced DNA compaction involves bending of DNA segments (see below for mechanical\ndiscussion), the observed sequence dependence would combinedly reflect both biophysical\ndynamics and biochemical affinity. Importantly, this in vitro result does not account for the influence\nof additional cellular components.”\n3. Line 106, a reference or additional explanation is needed to justify the use of RMS\nroughness analysis for assessing local compaction, although the approach appears\nreasonable.\n[Reply] We recognized that RMS roughness is not a widely used metric for assessing\ncondensate formation. In this revision, we quantified fluorescence heterogeneity using the\ncoefficient of variation of pixel intensities (standard deviation divided by mean), a more\ncommon measure in imaging and LLPS studies (for example, Zhang et al. J. Cell Biol. (2020);\ndoi:10.1083/jcb.202006054). This approach recapitulated the same salt- and tau\nconcentration-dependent behavior.\n• [Figure updated] We have revised our quantification of DNA condensation, replacing RMS\nroughness with the coefficient of variation of fluorescence intensity (SYTOX Orange DNA\nstrain):\n[Fig. 2E] Coefficient of variation of DNA fluorescence images, reflecting\no\nthe extent of DNA puncta formation induced by tau.\n[Results, Line 132] “The level of local compaction was estimated by the\no\ncoefficient of variation of the DNA fluorescence signal (the standard\ndeviation of pixel intensities divided by their mean), which increased\nprogressively as more tau was added (Fig. 2E).”\n4. In line 115, it is noted that Tau protein binds to naked DNA molecules at low concentrations,\na phenomenon that has been previously reported (Abasi et al., PMID: 38187700).\n[Reply] This comment has been addressed with the Comment #1 as stated above. Specifically,\nwe added:\n[Results, Line 141] “Importantly, these findings were confirmed under 150 mM NaCl, which more\no\nclosely approximates physiological salt conditions (Fig. S3). These results demonstrate that wild-\ntype tau protein at nano- to micromolar concentrations can stably bind to naked DNA under\nphysiological conditions, consistent with the behavior reported by Abasi et al.25”\n5. In line 130, it is unclear how the authors define Tau-DNA as being at the core or loosely\nassociated. As a result, I could not interpret the data in Figure 2H in the same way the authors\ndid.\n[Reply] The original statement was speculative, so we have revised the part in two ways:\n• [Text revised] We have rewritten it to focus on the dynamic nature of the condensates:\n[Results, Line 160] “Surprisingly, despite the fast accumulation of tau on DNA (Fig. 1B), many of\no\nthe tau condensates remained mobile along the DNA substrates (Fig. 2H and Movie S3). Moreover,\nthe location of DNA foci also followed tau movement across all tau concentrations we tested (50 nM\n– 5 μM). This suggests that tau clusters can slide along DNA to dynamically remodel the underlying\nsegments, reminiscent of other LLPS-driven protein–DNA co-condensates.”\n• [Results added] We have included FRAP results characterizing the mobile and immobile\nfractions of tau on DNA. Although the level of recovery was rather ill-defined (these\ncondensates frequently moved, merged, and split) and thus heterogeneous across\ncondensates, we observed consistent fluorescence recovery, indicating that a substantial\nfraction of DNA-bound tau is liquid-like and dynamically exchanges with bulk tau.\n[Supplementary Materials, Page 3] Fluorescence recovery after photobleaching (FRAP)\no\n“To probe the internal dynamics of tau–DNA condensates, we performed FRAP on individual tau\nclusters on double-tethered λ-DNA, assembled with 5 μM tau. (…)”\n[Fig. S5] FRAP analysis of DNA-bound tau condensates\no\n(A) Representative tau\nfluorescence along λ-DNA\nsubstrates (green dashed\noutline) before bleaching\nand after recovery for 0, 5,\n10 and 20 min. All tau\nfluorescence on the DNA\nwas fully photobleached\nat t = 0 min. (B) Normalized fluorescence intensity of individual tau condensates (colored traces, n\n= 5) showing heterogeneous recovery kinetics. (C) Normalized fluorescence recovery over time.\nPink shading represents the mean normalized intensity ± SD (individual data not shown), and the\nblue curve is a single-exponential fit to the recovery (τ ~ 30 min). Horizontal dashed lines indicate\nthe post-bleach intensity (Ibleach = 0.13) and the plateau intensity (I∞ = 0.84). Assuming a normalized\npre-bleach intensity (Ipre = 1), the average mobile fraction was calculated (I∞ − Ibleach )/(Ipre − Ibleach )\n= 0.82 (82%).\n[Results, Line 163] “To probe the internal dynamics of tau clusters, we performed fluorescence\no\nrecovery after photobleaching (FRAP) on tau clusters assembled with 5 μM tau along double-\ntethered λ-DNA (Fig. S5). Bleaching and monitoring of tau fluorescence (Cy5) recovery yielded\nhighly heterogeneous traces, reflecting frequent condensate movement, merging, and splitting, yet\nmany traces exhibited substantial fluorescence return. A single-exponential fit of the mean recovery\nfor representative traces gave a time constant of ~30 min and a mobile fraction of ~82%. However,\nthese values represent a composite of fast- and slow-recovering populations and are affected by\nthe loss of some condensates during monitoring. Overall, these results confirm that DNA-bound tau\nexchanges dynamically with the bulk pool in a liquid-like manner.”\n6. In line 142, please clarify how the diffusion coefficient and diffusion exponent were\nmeasured. I could not find any details in the Methods section about how single-particle tracking\nwas performed. I assume the authors calculated the MSD as a function of time.\n[Reply] We apologize for this oversight. Yes, The diffusion parameters were obtained by fitting\nthe experimental MSD to the generalized power-law model for 1D diffusion, MSD(Δt) = 2D(Δt)α.\n• [Methods added] We have now added the details for the single-particle tracking in the\nMethods section:\n[Materials and Methods, Line 642] “Single-particle tracking. Tracking of tau condensates on\no\nDNA was performed by analyzing movies from DNA skybridge experiments using custom MATLAB\nscripts. After registering the tau and DNA fluorescence channels, tau particles were detected by\nintensity thresholding, and only those colocalizing with DNA foci were included in the diffusion\nanalysis. Additionally, to avoid surface artifacts the skybridge substrates, particles within 7 pixels of\nthe ridges or adhered to the bottom surface were excluded from analysis. Mean-squared\ndisplacements (MSD) were calculated as a function of time lag Δt and fit to MSD(Δt) = 2D(Δt)α to\nextract the diffusion coefficient D and anomalous exponent α.”\n7. Measuring the piconewton forces applied by Tau-DNA condensation using magnetic\ntweezers is an excellent experiment. Could the authors speculate on the 72 nm step size\nobserved during the force jump assay?\n[Reply] We thank the reviewer for highlighting this point. The step size distribution peaks\nbetween 50 and 70 nm (with a median of 72 nm including larger steps likely arising from two\nunresolved events), which is comparable to the intrinsic persistence length (~50 nm) of dsDNA\nin physiological conditions, drawing attention to the role of DNA bending energy. We suggest\nthat tau condensation supplies just enough energy to compact segments roughly one to two\npersistence lengths long, which under applied force then rupture in quantized steps rather\nthan unwind continuously.\n• [Results added] To test the above hypothesis, we added coarse-grained molecular\ndynamics simulations that mimic our force-jump protocol. These simulations showed that\ntau–DNA and tau–tau attractions alone drive the formation of local tau-DNA clusters, and\nreproduced discrete rupture events of sizes similar to the L of DNA, supporting the notion\np\nthat DNA bending rigidity dictates the step size.\n[Fig. 3G–I] Coarse-grained simulation of stretching tau–DNA co-condensates. (G) Representative\no\nsnapshot of a tau–DNA co-condensate captured before force application. The right image shows\nDNA alone in the same configuration for clarity. (H) DNA extension during force-induced\ndecondensation. Right, zoomed-in views of the boxed region, with snapshots of the DNA\nconformation before and after each stepwise jump. (I) Histograms of step sizes for extension events.\nThe persistence length of dsDNA (Lp) is indicated by the red dashed line for reference.\n[Supplementary Materials, Page 9] Supplementary Notes: Molecular dynamics simulation for\no\ntau–DNA co-condensates\n[Fig. S8] Molecular dynamics simulation of stretching tau–DNA co-condensates\no\n[Results, Line 224] “To test this interpretation at the molecular level, we performed MD simulations\no\nof tau–DNA co-condensation under applied force (Fig. S8). In silico, tau and DNA coalesce into\ndroplets featuring looped DNA segments and characteristic bending, without strict periodicity\n(Fig. 3G). Simulated force–extension profiles recapitulated the experimentally observed stepwise\njumps (Fig. 3H), and inspection of snapshots and local DNA densities before versus after individual\njumps confirmed that each event corresponds to the rupture of a compacted DNA domain (Fig. S8).\nImportantly, the distribution of simulated step sizes peaked near the DNA persistence length\n(Fig. 3I), demonstrating that DNA bending rigidity defines the natural rupture unit. In contrast,\nsimulations lacking bending energy (i.e., a freely jointed chain model) produced much smaller,\nirregular steps (Fig. S8).”\n[Discussion, Line 472] “Given the contour length of dsDNA (~0.34 nm/bp), the 50–70-nm\no\nincrements observed during the stretching of tau–DNA co-condensates (Fig. 3F) correspond to\n150–200 base pairs. This is intriguingly close to the persistence length of dsDNA (~150 bp),\nsuggesting that tau condensates compact DNA into domains of this characteristic size, set by the\nhelix’s bendability. Indeed, our simulations reproduce these steps of similar sizes (Fig. 3I), each\nreflecting the force-induced rupture of an individual tau–DNA cluster. (…) The all-or-none behavior\nduring the condensation phase also supports the cooperative nature of tau condensation on DNA,\nrapidly drawing nearby DNA segments into maturing tau condensates.”\n8. During mitosis, chromosomes are highly condensed, and DNA is fully chromatinized. Can\nTau bind to such DNA and remain bound? Is there any literature supporting this? In this study,\nthe authors reported that Tau-naked DNA co-condensates capture microtubules. However, it\nis unclear how biologically relevant this observation is.\n[Reply] We agree that enhancing the biological relevance of our tau-DNA interaction findings\nis important. As the reviewer suggested, testing whether these interactions persist on\nchromatin is essential. In this revision, we include new experiments demonstrating that our\nobservations extend to chromatin substrates. Alongside the tau phosphorylation and mutant\nanalyses described below, these data further support the cellular relevance of our model.\n• [Results added] We repeated our investigations after preparing nucleosomes-reconstituted\nDNA and found that it supports both tau binding and subsequent microtubule capture.\n[Fig. 5A, B] Microtubule capture by tau–DNA co-\no\ncondensates with nucleosomes and phosphorylated tau\n(A) TIRF images of nucleosome-reconstituted λ-DNA\nshowing DNA (green), tau (magenta), and Alexa 488–H2A\nnucleosomes (cyan). Left panels: addition of 5 μM tau yields\ndiscrete tau–DNA co-condensates colocalized with\nnucleosome positions (cyan circles). Right panels: upon\nintroduction of a tau/tubulin mixture, nascent microtubules\n(cyan) attach to these condensates. (B) Representative\nlarger fields of view showing tau (magenta) and microtubules\n(cyan) on nucleosome-decorated DNA strands (green).\n[Results, Line 294] Tau condensates form on\no\nnucleosomes to mediate microtubule attachment\n“To place our in vitro observations in a more physiologically\nrelevant context, we reconstituted nucleosomes on λ-DNA\nand examined how tau condensation and subsequent\nmicrotubule binding were affected. After purifying histones,\nassembling octamers, and loading them onto λ-DNA,\nnucleosome formation was confirmed by visualizing Alexa\n488-labeled H2A on the DNA skybridge, with each λ-DNA\nmolecule exhibiting the characteristic beads-on-a-string configuration (Fig. S10 and Movie S11).\nThese nucleosome-containing λ-DNA molecules were tethered to a TIRF imaging surface,\nproducing a mixture of singly and doubly tethered substrates. Addition of 5 μM tau to this surface\nyielded discrete foci of tau–DNA co-condensates, 85 ± 5% of which colocalized with nucleosomes\n(Fig. 5A; cyan circles). This high degree of colocalization suggests that tau–DNA co-condensation\nresponds to nucleosome landmarks, consistent with recent bulk studies showing tau co-\ncondensation with nucleosome arrays25, and implies that mobile tau clusters are pinned at\nnucleosome sites. However, on singly tethered DNA adopting a ball-like conformation, nucleosome\nlocations were not laterally resolved, precluding precise colocalization analysis. Importantly, when\nmixtures of tau and tubulin containing nascent microtubules were introduced, the microtubules\nattached to this surface to an extent comparable to that observed for tau-bare DNA condensates\n(Fig. 5B). These results suggest that tau-nucleosome clusters can also serve as effective\nmicrotubule-interfacing platforms within a chromatin context.”\n[Discussion, Line 504] “(…) Formation of nucleosome-pinned tau condensates (Fig. 5A, B), which\no\nretain the ability to mediate microtubule assembly, partially explains how tau condensates form\npreferentially on nucleosome-rich centromeres in dividing cells. (…)”\n• [Results added] To verify tau binding to mitotic chromosomes at a higher resolution, we\nextracted chromosomes from mitotic HEK-293 cells expressing mCherry–Tau and revealed\na strong tau signal on the surface of these chromosomes, indicating that tau remains bound\nto chromatin.\n[Fig. 6G] Confocal images of chromosome spreads\no\nfrom mitotic HEK-293 cells expressing mCherry–Tau\n(green) and stained with Hoechst 33258 (blue); the\nright-hand panel shows a magnified view of the white-\nboxed region.\n[Results, Line 364] “To assess whether tau directly\no\nassociates with mitotic chromosomes at higher\nresolution, we prepared chromosome spreads from\nthese HEK-293 cells expressing mCherry–Tau\n(Fig. 6G). In these preparations, small punctate tau clusters decorated the surface of condensed\nchromosomes, demonstrating that tau retains its DNA-binding activity within a chromatin context.\nWe did not observe centromere-specific enrichment in this case, likely because the more transient\nor weak interactions that mediate centromere recruitment are disrupted during the spreading.”\n9. The investigation of Tau-DNA-microtubule assemblies and their roles in cells is\nunderdeveloped. Proper controls are missing to confirm a direct interaction between Tau and\nthe centromere, despite the colocalization images provided.\n[Reply] We hope that our response above (to Comment #8) on the cellular relevance of tau–\nDNA assembly partially addresses this point. Below, we expand specifically on tau’s\ncentromere localization:\n(a) We previously verified centromere localization of tau in two cellular contexts: in\nSH-SY5Y cells, which endogenously express tau, and in HEK-293 cells, where we\nexogenously expressed wild-type tau. In HEK-293 cells, the localization of exogenous\ntau to centromeres in an otherwise tau-free background, now further validated by\nPearson correlation analysis of tau and centromere signals (see our response to\nComment #12 below), strongly indicates a centromere preference. This observation is\nnot totally unexpected given previous reports of “pericentromeric nuclear tau” (for\nexample, Sjöberg et al. J. of Cell Science (2006); doi: 10.1242/jcs.02907), and we\nextend this concept to the mitotic context.\n(b) For this revision, we further assessed the cellular role of tau’s centromere localization\nby expressing phosphomimetic tau variants in tau-free HEK-293 cells (see our\nresponse to Comment #14 below). In contrast to wild-type tau, expression of these\nmutants consistently impaired metaphase chromosome alignment. Notably,\nmisaligned chromosomes were broadly coated with tau without centromere preference,\nsupporting that selective tau localization at centromeres is required for microtubule\ncapture and chromosome segregation.\n(c) Finally, we do not presume that, in cells, tau’s localization to centromeres depends\nsolely on direct tau–DNA interactions. Our data show that tau binds naked DNA and,\nin this revision, nucleosomes, suggesting that its inherent affinity for densely\nnucleosomal regions may partly account for centromere enrichment, consistent with\nAbasi et al. However, additional factors such as HP1α likely contribute to the\nmaturation of tau assemblies at centromeres, which may even pre-form in the nucleus\nbefore nuclear envelope breakdown (our new prophase images attached below\nsupport this notion, too). This mechanism likely acts alongside tau condensates’\nbroader DNA-binding at more randomly distributed sites on mitotic chromosomes,\nsubsequently refining into centromere-focused structures that funnel tau to\ncentromeres. This model is illustrated in Fig. 8. We agree that probing direct\ninteractions between tau and DNA in dividing cells with appropriate controls would be\nhighly informative. However, technical challenges associated with tau’s multifaceted\nroles in cytoskeletal organization currently constrain such targeted perturbations. We\nplan to address this challenge by undertaking a molecular level dissection of tau–DNA\nbinding in a follow up study.\n‐\n• [Text revised] In addi‐tion to the new experimental results mentioned above, we have\nupdated the manuscript to incorporate the conceptual refinements:\n[Discussion, Line 502] “The unexpected enrichment of tau at centromeres during prometaphase in\no\nboth SH-SY5Y (endogenous tau) and HEK-293 (exogenous tau) cells (Fig. 6) occurs despite tau’s\nlargely nonspecific affinity for DNA. Formation of nucleosome-pinned tau condensates (Fig. 5A, B),\nwhich retain the ability to mediate microtubule assembly, partially explains how tau condensates\nform preferentially on nucleosome-rich centromeres in dividing cells. This model is supported by\ntau’s strong complexation with nucleosome arrays25, its nuclear colocalization with HP1α-marked\npericentromeric heterochromatin38,42, and its chromatin-related functions60,61. Although the tripartite\nassembly of tau–DNA–microtubule itself does not necessarily invoke the nuclear function of tau28,\nwe speculate that centromeric tau condensates that assemble in the nucleus during prophase\n(Fig. S11) may behave analogously to our in vitro preformed condensates, guide the migration of\ntau on other chromosomal sites toward the centromeres, and then merge with cytosolic tau clusters\non microtubules after nuclear envelope breakdown. Future experiments should employ centromere-\nspecific CENP-A nucleosomes and prolonged live-cell imaging to validate this idea.”\n10. In lines 287 and 296, why are only two centromeres analyzed per cell, given that each cell\ncontains so many more centromeres? Maybe I am missing something.\n[Reply] We thank the reviewer for identifying this error. The numbers were simply miswritten,\nand we have corrected the text.\n[Results, Line 348] “To quantitatively assess the colocalization between tau and centromeres, we\no\npooled images of 181 centromeres from 11 prometaphase cells and averaged them after precise\nalignment (Fig. 6B).”\n[Results, Line 360] “Analysis of 460 centromeres from 18 prometaphase HEK-293 cells verified\no\noverlap between tau and centromere signals (Fig. 6F), (…)”\n11. Similarly, additional experiments can help elucidate the role of phosphorylation in co-\ncondensates interacting with microtubules.\n[Reply] Following the reviewer’s advice, we prepared PKA-phosphorylated tau (pTau) as\ndescribed in Abasi et al. and repeated the microtubule capture assays with pTau:\n• [Results added] pTau retained DNA binding and condensate formation but showed\nmodestly reduced microtubule recruitment compared with WT tau. More interestingly,\nmicrotubule fragments produced with pTau still bound efficiently to WT-derived condensates\non DNA, suggesting that tau phosphorylation during mitosis may sustain DNA-microtubule\nassembly in cooperation with unmodified tau.\n[Fig. 5C–H] (C) Schematic of the microtubule capture assay using two tau conditions: pre-formed\no\ntau–DNA condensates or polymerizing microtubules. (D) Comparison of tau–DNA co-condensate\nformation with wild-type (WT) and PKA-phosphorylated tau (pTau). (E) Quantification of tau\ncondensate density for WT and pTau on DNA substrates (n = 5 images; mean ± s.d.). (F) TIRF\nimages of tau (magenta) and microtubules (cyan) observed with different combinations of WT and\npTau. DNA (green) is shown in the merged images. (G) Fluorescence intensity of WT tau-\npolymerized microtubules recruited by WT versus pTau condensates (n = 9 images; mean ± s.d.).\n(H) Fluorescence intensity of tubulin captured by tau–DNA condensates (n = 8 images; mean ±\ns.d.).\n[Results, Line 312] Phosphorylated tau retains DNA binding and enables microtubule capture\no\nwith wild-type tau\n“Microtubule dynamics are tightly regulated by tau and its phosphorylation state, particularly during\nmitosis when increased phosphorylation correlates with enhanced microtubule turnover. To\ndetermine how phosphorylation alters tau’s DNA condensation and microtubule-recruiting capacity,\nrecombinant human 2N4R tau was phosphorylated in vitro with cAMP-dependent protein kinase\n(PKA), which primes tau for subsequent modification by GSK-3β41. Efficient phosphorylation was\nconfirmed by the characteristic upward shift on SDS–PAGE (Fig. S1). Phosphorylated tau (pTau)\nwas then introduced either onto λ-DNA tethers to assess condensation or into the tubulin mix during\nnascent microtubule polymerization (Fig. 5C). Although pTau retained DNA-binding and\ncondensate-forming potential, both activities were modestly reduced compared with wild-type (WT)\ntau (Fig. 5D, E).\nIn microtubule capture assays, pTau–DNA condensates recruited fewer microtubules when\nexposed to filaments formed with WT tau and tubulin (Fig. 5F, G). Unexpectedly, when microtubules\nwere polymerized with pTau, the shorter, less stable filaments still attached efficiently to WT tau–\nDNA condensates, as indicated by intense fluorescence from bound segments (Fig. 5F, H).\nThis\nresult suggests that dynamic pTau-associated microtubule fragments can engage tau–DNA\ncondensates in the presence of WT tau. By contrast, assays using pTau for both DNA condensation\nand microtubule assembly produced only tubulin aggregates on the DNA surface, without\nfilamentous structures, demonstrating that pTau alone lacks robust microtubule-interfacing\ncapability. Together, these findings indicate that although phosphorylation diminishes tau’s affinity\nfor DNA and microtubules, pTau-associated microtubule fragments during mitotic progression can\nstill interact with tau–DNA co-condensates when WT tau is available.”\n[Discussion, Line 520] “(…) Based on the differential effects of enzymatic phosphorylation and\no\nphosphomimetic variants (Figs. 5 and 7), we propose that multisite phosphorylation fine-tunes tau’s\naffinity for DNA and microtubules and regulates their dynamic interplay. Because our microtubule\ncapture assay was specifically configured to distinguish DNA-preformed from tubulin-associated\ncondensates, the distinct activities of unphosphorylated versus phosphorylated tau in these two\ncontexts (Fig. 5C–H) provide a support for our two-stage model of nuclear-origin and spindle-\nassociated condensates, in line with the high phosphorylation state of cytosolic tau during mitosis.”\n12. Statistical analysis is missing for Figure 7D. Why not perform a Pearson correlation\nanalysis to assess the colocalization between centromeres and Tau foci?\n[Reply] We have reflected this suggestion by calculating Pearson correlation coefficients\nbetween the tau and centromere images, and between the tubulin and tau images. We also\nperformed this analysis for the newly added HEK cell images expressing phosphomimetic tau.\n[Fig. 6C, D] Pearson correlation coefficients for tau\no\ncolocalization with tubulin and centromeres, calculated\nfrom the cell images used in (B) and (F). Box plots show\nmedian (red line), interquartile range, and whiskers to the\nminimum/maximum; each point represents one cell.\n[Results, Line 351] “Pearson correlation coefficients\no\ncalculated for each SH-SY5Y cell image confirmed\nstrong colocalization of tau with both tubulin and\ncentromeres (Fig. 6C, D).”\n[Fig. 7G, H] Pearson correlation\no\ncoefficients for tau colocalization with\ntubulin and centromeres (n = 9 cells for\nT231D/S235D, n = 8 cells for S262D).\nBox plots show median (red line),\ninterquartile range, and whiskers to the\nminimum/maximum; each point\nrepresents one cell. For comparison, the\nwild-type tau data from Fig. 6C and 6D are reproduced here.\n[Results, Line 424] “Pearson correlation analysis revealed their reduced association with spindle\no\nmicrotubules (Fig. 7G), consistent with the impaired microtubule-stabilizing activity (Fig. 7B).”\n13. Long-term cell imaging of Tau-mCherry through the mitotic process may provide more\nconvincing evidence of Tau interacting with condensed DNA.\n[Reply] Although long-term live-cell imaging of mCherry-tagged tau would provide valuable\ndynamic insights, we relied on fixed cell preparations because co expressing multiple\nfluorescent markers (to track chromosomes, microtubules, and tau) and imaging them over\nextended periods imposed too great a b‐urden on the cells and proved un‐successful. Instead,\nto reconstruct the temporal sequence of tau localization, we revisited endogenous tau in\nSH-SY5Y cells synchronized at different cell-cycle stages (Fig. S11).\n• [Results added] During prophase, abundant tau clusters colocalized with centromeric\nmarkers, consistent with previous reports of pericentromeric nuclear tau and supporting the\nnotion that tau condensates assemble prior to nuclear envelope breakdown. By metaphase\nalignment, however, centromere-associated tau was largely lost, underscoring the transient,\ncell cycle-specific nature of this localization.\n[Fig. S11B] Images of SH-SY5Y across mitotic stages. Scale, 10 μm. In addition to tau’s centromere\no\nlocalization in prometaphase and metaphase, nuclear tau associated with centromeres in prophase.\n[Results, Line 338] “To test this in a neuronal setting, we conducted immunofluorescence imaging\no\nof SH-SY5Y cells, a neuroblastoma line that endogenously expresses tau. Immunostaining of\nthymidine-synchronized, dividing cells (identified by condensed chromosomes) revealed a\nconsistent level of endogenous tau forming small puncta with distinct distributions across mitotic\nstages (Fig. S11). (…) Interestingly, prophase SH-SY5Y cells also revealed clear association of\nnuclear tau with centromeres (Fig. S11), consistent with its known pericentromeric localization38,42”\n[Discussion, Line 508] “(…) Although the tripartite assembly of tau–DNA–microtubule itself does\no\nnot necessarily invoke the nuclear function of tau28, we speculate that centromeric tau condensates\nthat assemble in the nucleus during prophase (Fig. S11) may behave analogously to our in vitro\npreformed condensates, guide the migration of tau on other chromosomal sites toward the\ncentromeres, and then merge with cytosolic tau clusters on microtubules after nuclear envelope\nbreakdown. Future experiments should employ centromere-specific CENP-A nucleosomes and\nprolonged live-cell imaging to validate this idea.”\n14. The authors could consider expressing phosphomimetic Tau-mCherry in cells and\ncomparing its dynamic behavior to wild-type Tau to assess potential differences in centromere\ncolocalization. This approach may provide insights supporting a role for Tau in regulating\nmitosis.\n[Reply] We appreciate this suggestion. Indeed, expression of phosphomimetic mCherry–tau\nin HEK-293 cells led to clear defects in metaphase chromosome alignment.\n• [Results added] Both S262D and T231D/S235D mutants increased the fraction of\nmisaligned chromosomes, with S262D showing a more pronounced effect. We interpret the\ntau coating of misaligned chromosomes as indicating that, although the mutant tau retains\nDNA-binding affinity, its altered partitioning between microtubule and chromosome binding\nleads to aberrant coverage that disrupts spindle engagement.\n[Fig. 7F–L] (F) Immunofluorescence images of prometaphase HEK-293 cells transiently expressing\no\nphosphomimetic (T231D/S235D and S262D) mCherry–Tau, showing the distribution of tubulin, tau,\nand centromeres. (G, H) Pearson correlation coefficients for tau colocalization with tubulin and\ncentromeres (n = 9 cells for T231D/S235D, n = 8 cells for S262D). Box plots show median (red\nline), interquartile range, and whiskers to the minimum/maximum; each point represents one cell.\nFor comparison, the wild-type tau data from Fig. 6C and 6D are reproduced here. (I) Intensity\nprofiles of centromere (blue) and tau (green) measured across aligned centromere regions,\nobtained from the cell images used in (G) and (H) (n = 429 centromeres for T231D/S235D, n = 288\ncentromeres for S262D). (J) Immunofluorescence of metaphase HEK-293 cells expressing WT or\nphosphomimetic mCherry–Tau. DNA image (Hoechst 33258-stained) is overlaid in gray for\nreference. Misaligned DNA foci away from the metaphase plate is outlined in yellow. (K, L)\nQuantification of misaligned DNA (Hoechst 33258) and tau (mCherry) intensities, normalized to\neach cell’s total signal (n = 6 images each; mean ± s.d.).\n[Results, Line 419] Phosphomimetic tau localizes to centromeres but impairs chromosome\no\nalignment\n“To test whether the in vitro differences in DNA–microtubule interfacing by phosphomimetic tau\nmutants manifest in cells, we performed immunofluorescence on mitotic HEK-293 cells transiently\nexpressing mCherry-tagged T231D/S235D or S262D tau, using the same approach as for wild-type\ntau (Fig. 6). Although both mutants largely retained microtubule binding during mitosis (Fig. 7F),\nPearson correlation analysis revealed their reduced association with spindle microtubules (Fig. 7G),\nconsistent with the impaired microtubule-stabilizing activity (Fig. 7B). Nevertheless, both mutants\nexhibited robust centromeric localization comparable to wild-type tau (Fig. 7H), reflecting their\nretained DNA-binding activity (Fig. 7A). Analysis of averaged centromere images further confirmed\noverlapping tau signal (Fig. 7I).\nSurprisingly, in metaphase cells after chromosomes have aligned at the metaphase plate, both\nphosphomimetic mutants showed prominent tau-rich chromosomal regions that failed to align\n(Fig. 7J). These tau–DNA clusters did not colocalize with the central tubulin signal, indicating a lack\nof spindle attachment, and were most frequent in S262D-expressing cells, implicating aberrant tau–\nDNA binding in defective microtubule capture. Quantitative analysis of DNA and tau amount\n(measured by Hoechst 33258 and mCherry fluorescence, respectively) confirmed these findings.\nIn S262D cells, misaligned DNA foci comprised about 10% of total chromosomal DNA, significantly\nmore than in wild-type or T231D/S235D cells (Fig. 7K; see Fig. S12 and Supplementary Methods\nfor analysis). Tau fluorescence at these foci accounted for nearly 30% of total cellular tau in S262D\ncells—over a five-fold increase versus wild type (Fig. 7L). These data demonstrate that\nphosphomimetic tau mutants not only lose their ability to engage spindle microtubules but also\naberrantly accumulate on misaligned chromatin, thus compromising proper chromosome\nsegregation.”\n[Discussion, Line 542] “The failure of some chromosomes to align at the metaphase plate in cells\no\nexpressing Alzheimer’s-associated phosphomimetic tau mutants is informative in this regard\n(Fig. 7J–L). It suggests that altered phosphorylation could disrupt tau’s dynamic partitioning\nbetween DNA and microtubules, remain abnormally coated on chromatin, and escape timely\nspindle capture, potentially contributing to chromosome missegregation and aneuploidy. These\ncell-based findings not only validate the biophysical function of tau observed in vitro but also reflects\nthe loss-of-function effect seen in microtubule capture assays. This bidirectional correspondence\nstrongly reinforces the idea that site-specific phosphorylation of tau disrupts its mitotic function.”\nReviewer #3 (Remarks on code availability):\nI didn't review all the codes in depth. However, after a quick check, I couldn't find the README\nfile for most codes (if not all).\n[Reply] We apologize for the oversight and have now added README files to all code\ndirectories to ensure proper documentation and ease of use. For example:\n• At https://github.com/ShonLab/tau-dna-co-condensation/tree/main/TIRF:\nreadme_tirf.txt\n• At https://github.com/ShonLab/tau-dna-co-condensation/tree/main/MT:\nreadme_MT.txt\n• At https://github.com/ShonLab/tau-dna-co-condensation/tree/main/confocal:\nreadme_confocal.txt\nResponse to Reviewer Comments on:\nPark et al., “Tau condensation on DNA mediates\nmicrotubule attachment: A potential mitotic role for\ncentromere-localized tau”\nReviewer #1 (Remarks to the Author):\nIn the revised manuscript the authors admirably updated the text as I suggested. I think this is\nan important piece of science that needs to be available. I still have concerns that people might\nmistakenly believe that this mechanism is fundamental to mitosis. However I believe that is for\nothers to decide. I accept the corrected manuscript.\n[Reply] We sincerely thank the reviewer for the positive and thoughtful evaluation, the\nencouraging recommendation, and the constructive concerns and insights shared alongside.\nReviewer #2 (Remarks to the Author):\nThe authors have improved the manuscript and addressed or clarified most of the major\nconcerns. Tau DNA binding and localization on microtubules is well supported by experimental\ndata. However, the functional implications for such interactions in cells are limited. The authors\nprovide additional data using phosphomimetic Tau in 293T cells, but the lack of a biologically\nrelevant model—such as modulation of Tau levels or phosphorylation in SH-SY5Y cells—\nlimits the biological significance of their findings. They have addressed this in part by\ntempering the interpretation in the revised text. While the conclusions are supported by the\ndata, the absence of physiological validation reduces the impact and excitement of the work.\nSome major concerns remain with the new experiments presented:\n1. The extent of nucleosome assembly is not shown, which is particularly important for the\nnon-601 DNA templates. Additional quality control data must be shown. This is critical for\ninterpreting Tau-nucleosome interactions: without efficient assembly, Tau could be binding\npredominantly to free or poorly packaged DNA (Fig. S10E show the degree of assembly od\nlambda- DNA).\n[Reply] We thank the reviewer for this suggestion. In this revision, we have quantified\nnucleosome assembly by counting clusters on λ-DNA at the single-molecule level from images\nsuch as Fig. S10E (DNA skybridge platform). Each λ-DNA molecule typically displayed three\nto five clusters, with some cluster fluorescence intensities (from Alexa 488-labeled H2A)\nseveral-fold higher than those of single nucleosomes (Fig. S10F, G), indicating multiple\nhistone octamers per cluster. Some nucleosome signals appeared to overlap, partially\nsacrificing the precision of our cluster estimate. But more importantly, these clusters remained\nstably positioned on DNA for tens of minutes (observation time) without movement or\ndisassembly, consistent with their survival through rigorous purification. Moreover, assembled\nnucleosome arrays retained integrity when stored at 4 °C for several days, further\ndemonstrating the stability of the complexes. Although the nucleosome coverage does not\nfully saturate the DNA, it confirms that a substantial fraction of λ-DNA is properly packaged,\nsupporting our interpretation of colocalization between nucleosome clusters and tau\nassemblies (Fig. 5A).\nAt the same time, we acknowledge that these data cannot exclude a model in which tau\npredominantly associates with free DNA regions between clusters, so we have added this\nreservation in the revised text, too. We are currently investigating tau interactions at the single-\nnucleosome level in a separate study.\n• [Results added] We added the above results to the main text and as a supplementary figure.\n[Results, Line 299] “After purifying histones, assembling octamers, and loading them onto λ-DNA,\no\nnucleosome formation was confirmed by imaging Alexa 488-labeled H2A on stretched DNA\nmolecules, revealing the characteristic beads-on-a-string pattern (Fig. S10 and Movie S11).\nEach\nλ-DNA molecule typically exhibited 3–5 nucleosome clusters (Fig. S10), with some clusters\ncontaining multiple octamers based on fluorescence intensity, although individual nucleosomes\ncould not be resolved due to the diffraction limit. Importantly, these nucleosome arrays remained\nintact and stable in solution for several days, indicating successful and robust assembly. (…) \nHowever, on singly tethered DNA adopting a ball-like conformation, nucleosome locations were not\nlaterally resolved, ding precise colocalization analysis and necessitating further investigation at the\nsingle-nucleosome level. Nevertheless, when tau–tubulin mixtures containing nascent microtubules\nwere introduced, microtubules readily attached to the surface, at levels comparable to those\nobserved with tau–bare DNA condensates (Fig. 5B). These results suggest that tau–nucleosome\ncomplexes can also serve as effective microtubule-interfacing platforms, even within a partially\nchromatinized context.”\n[Fig. S10] (F) Representative fluorescence images of individual nucleosome-loaded λ-DNA, with\no\ncorresponding Alexa 488 intensity profiles. Arrows indicate nucleosome clusters identified from\nlocal intensity maxima in line profiles. (G) Histogram of nucleosome cluster counts per λ-DNA\nmolecule (n = 89).\n2. Tau phosphorylation sites are not clearly described. How many are found in cells? Are all\nsites equally phosphorylated? What is the extent of phosphorylation achieved in the in vitro\nsystem using PKA? Mass spectrometry is needed to accurately assess phosphorylation status.\n[Reply] We thank the reviewer for the comment, which we had previously overlooked. We\nhave now revised the manuscript to address the questions and suggestions:\n• [Text revised] Additional details on tau phosphorylation have been included:\n[Results, Line 322] “Although tau contains up to 85 potential phosphorylation sites, only a small\no\nsubset is modified under physiological conditions41. In healthy brains, tau carries an average of 2–\n3 phosphate groups, while aberrant forms found in Alzheimer’s disease exhibit 3–4 times higher\nlevels, contributing to tau aggregation42.”\n41. Buée, L., Bussière, T., Buée-Scherrer, V., Delacourte, A. & Hof, P. R. Tau protein isoforms,\nphosphorylation and role in neurodegenerative disorders1. Brain Res. Rev. 33, 95–130 (2000).\n42. Köpke, E. et al. Microtubule-associated protein tau. Abnormal phosphorylation of a non-paired\nhelical filament pool in Alzheimer disease. J. Biol. Chem. 268, 24374–24384 (1993).\n• [Results added] We have conducted mass spec analysis of PKA-phosphorylated tau and\nincluded the results:\n[Results, Line 330] “Site-specific verification by LC–MS/MS confirmed strong phosphorylation at\no\nknown PKA target sites (Fig. S1), including S214, S324, S352, S356, S409, and S416, consistent\nwith previous reports25,44. These modifications collectively yield an overall phosphorylation\noccupancy of ~4 sites per tau molecule.”\n[Fig. S1D and E] (D) Phosphosite occupancy across the full length of 2N4R tau as measured by\no\nLC–MS/MS. Schematic showing phosphorylation sites with >20% occupancy are indicated by\narrows on top.\nMinor comments:\n1. The use of CREST as a centromere marker should be explicitly mentioned in the main text.\n[Reply] We appreciate this suggestion and updated the text accordingly:\n[Result, Line 360] “Notably, tau formed distinct clusters on mitotic chromosomes, most of which\no\ncolocalized with centromeres labeled by CREST anti-centromere antibody (Fig. 6A, magnified\nviews in the bottom panels).”\n[Result, Line 372] “Confocal imaging of mCherry fluorescence for tau, alongside immunostaining\no\nfor tubulin and centromeres (CREST antibody), (…)”\n2. Several sections/figures of the manuscript offer only marginal additions over one another,\nand some experiments are redundant. Consolidation of figures and movement of some panels\nto supplementary materials would improve clarity and focus.\n[Reply] We sincerely appreciate this point and understand the concern. However, we hope to\nretain the current structure, as we have made a concerted effort to include only essential data\nin the main figures. We are concerned that moving seemingly similar but mechanistically\ndistinct results to the supplement may risk misleading readers, obscuring important contextual\ndifferences, and/or underrepresenting key results.\nIn summary, the in vitro characterization of Tau interactions with DNA and microtubules is\nsolid. Although the biological implications are limited, the study proposes an intriguing and\nnovel model for Tau’s unexpected role in microtubule attachment.\n[Reply] We appreciate the positive evaluation again.\nReviewer #3 (Remarks to the Author):\nThe claims in the manuscript have been greatly strengthened through revision. Therefore, I\nrecommend publication of this manuscript.\n[Reply] We sincerely thank the reviewer for the thoughtful evaluation and greatly appreciate\nthe constructive feedback throughout the review process.\nA few minor comments:\n1. I appreciate that the authors conducted several technically challenging but critical\nexperiments, including (1) FRAP to confirm the dynamic exchange of Tau proteins within\ncondensates, (2) multi-color imaging to demonstrate Tau interaction with nucleosome-bound\nDNA, and (3) imaging assays showing that phosphomimetic Tau results in mitotic defects.\nThese additions significantly strengthen the story.\n[Reply] The reviewer’s suggestions helped us improve the clarity, rigor, and overall quality of\nthe manuscript. We are pleased that our efforts were seen as strengthening the overall story\nof the manuscript.\n2. The manuscript lacks a description of how the images were quantified and the coefficient\nof variation was calculated (line 132). It would be helpful to include this information in the\nSupplementary Information.\n[Reply] Following the reviewer’s suggestion, we have included more details on single-\nmolecule image analysis:\n[Supplementary Methods, Page 3] Single-molecule image analysis of surface-tethered DNA\no\n“Single-molecule fluorescence images of DNA and tau were analyzed using custom MATLAB\nscripts (available at https://github.com/ShonLab). For each doubly tethered DNA molecule, the\nattachment points were identified from minimum-intensity projections over time, as these points\nremain stable while the DNA contour fluctuates and fades. Fluorescence intensity profiles of DNA\nand tau were then extracted along the contour between these points by averaging 5–7 rows\nperpendicular to the DNA axis. From these line profiles (e.g., Fig. 1B, C and S3B), tau–DNA\ncolocalization was quantified using Pearson correlation, calculated as the covariance of DNA and\ntau intensities divided by the product of their standard deviations (Fig. 1D and S3D). Local intensity\nvariation was quantified using the coefficient of variation, the standard deviation divided by the\nmean of pixel intensities (Fig. 1E and S3E). Additionally, DNA fluctuation was measured from time-\nlapse maximum-projection images by calculating the envelope width at the midpoint of the DNA\ncontour, defined as the spatial extent of DNA fluorescence orthogonal to the axis (Fig. 1G and S3G).\nMolecules showing abnormal fluctuation behavior or end-to-end lengths outside a defined range\n(5–9 μm) were excluded from analysis.”\n3. What salt concentrations were used in the molecular dynamics simulations of DNA\nstretching? dsDNA shorter than 150 bp can bend under high-salt conditions, which may affect\nthe results.\n[Reply] Our simulations operate without explicit ions, but we implicitly capture ionic screening\nby tuning the DNA bending rigidity to a persistence length of 50 nm, matching experimental\nvalues at 100 mM monovalent salt (Hagerman, Ann. Rev. Biophys. (1988); doi:\n10.1146/annurev.bb.17.060188.001405). This effectively pla~ces our model in the 100–200\nmM regime,∼ where salt-dependent changes in persistence length are <10% (1–2 beads)\n(Baumann et al. PNAS (1997); doi: 10.1073/pnas.94.12.6185). Given that our force-extension\ntransitions occur over long segments of DNA (spanning tens of beads), this small change\ncorresponding to salt-dependent persistence length is effectively negligible at this scale.\nAdditionally, any short-length-scale bending (<150 bp) is subsumed into the single\npersistence-length parameter and does not manifest as a distinct fluctuation mode, further\nminimizing its impact on our long-segment stretching analysis. We therefore consider our\neffective salt condition to lie within the physiological, high-salt regime.\n• [Text revised] To clarify, we have updated the text:\n[Results, Line 225] “Here, ionic effects at physiological monovalent salt (~100 mM) were implicitly\no\ncaptured by setting the DNA persistence length to ~50 nm, which subsumes short-scale bending\nand salt-dependent flexibility (see Supplementary Notes for more details).”\nReviewer #3 (Remarks on code availability):\nThe authors included the correct documentation. I have no further comments.\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This\nis part of the Nature Communications initiative to facilitate training in peer review and to\nprovide appropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse to Reviewer Comments on:\nPark et al., “Tau condensation on DNA mediates\nmicrotubule attachment: A potential mitotic role for\ncentromere-localized tau”\nReviewer #2 (Remarks to the Author):\nReviewer #2 (Remarks to the Author):\nI appreciate the additional data provided. However, major concerns regarding nucleosome\narray quality remain unresolved. As currently presented, the nucleosome-related data do not\nsupport the claim that Tau condensates form on nucleosomes to mediate microtubule\nattachment. Without rigorous controls and proper validation of nucleosome assembly, the\nconclusions drawn from these experiments are not sufficiently robust. The observed results\ncould be equally explained by octamer aggregation, misassembled nucleosomes, or\nnonspecific Tau co-aggregation with free DNA or histones.\nQuality controls are required (detailed below). Alternatively, the nucleosome-specific claims\nshould be removed, and the manuscript should focus solely on Tau–DNA co-condensation,\nwhich is better supported by the current data.\n[Reply] We thank the reviewer for the constructive comments and the chance to better support\nour nucleosome-related claims. In this revision, we have substantially updated the\nnucleosome experiments, refined the analyses, and clarified the data presentation. We\naddress each specific point below. In summary:\n• We replaced λ-DNA with a 2.3-kb DNA\nharboring a 12× array of the Widom-601\nsequence (right, Fig. 5G), a canonical choice\nfor robust nucleosome assembly.\n• To confirm the integrity of the nucleosome assemblies, we used multiple biochemical and\nbiophysical assays.\n• We compared nucleosomal versus naked DNA of the same size and sequence for their\nability to bind tau and recruit microtubules.\n• We found that nucleosome-bound DNA showed\ntau and microtubule binding largely comparable\nto naked DNA (right, Fig. 5J–M), despite the\nreduced stretches of free DNA\n• We note that residual free DNA regions may\nunderlie part of the effect, citing prior work, and\nadjust conclusions to avoid over-interpretation.\nWe believe these additions and clarifications fully\naddress the reviewer’s concerns about nucleosome\nintegrity and strengthen the overall conclusions of\nthe study.\nDetailed point-by-point responses are provided below.\n• The manuscript does not provide details on the octamer preparation, assembly conditions,\npurification steps. The claim of “rigorous purification” after mixing DNA and octamers is not\nexplained in the Methods or Results. Usually this entails a sucrose gradient.\n[Reply] Since the previous λ-DNA data did not adequately address nucleosome quality, we\ncompletely replaced λ-DNA with a 2.3-kb dsDNA bearing 12 consecutive Widom 601\nsequences for all nucleosome experiments (quality control results are appended below for the\nspecific concerns). In addition, we now provide more detailed methods for histone-octamer\npreparation, nucleosome assembly on the 601 DNA, and their use in single-molecule imaging.\nThe octamer preparation was already present in the previous Supplementary Information but\nmay have been overlooked because it was bundled with nucleosome reconstitution; we have\nnow separated it into its own section (Supplementary Information: page 8 and Fig. S10B, C).\nBriefly, individual purified histones were denatured, H2A–H2B and H3–H4 were combined with\na slight H2A–H2B excess, refolded by dialysis into high-salt buffer, and purified by size-\nexclusion chromatography. Fractions containing equimolar amounts of all four histones\n(verified by SDS-PAGE) were pooled and stored. Octamer formation was validated by\nreconstitution of mononucleosomes on a single stretch of 601 DNA sequence.\n[Supplementary Methods, Page 7] Preparation of histone octamers\no\nHistone octamers were assembled by refolding denatured histones3,4. Each histone was dissolved\nin unfolding buffer (20 mM Tris-HCl, pH 7.5; 7 M guanidinium-HCl; 10 mM DTT) at room\ntemperature for 1 h. Protein concentrations were determined by A280. H2A and H2B were mixed\nat a 1:1 molar ratio, as were H3 and H4, with the H2A–H2B dimers at 10% molar excess relative to\nthe H3–H4 tetramers. The final protein concentration was adjusted to 1 mg/mL in unfolding buffer,\nand the mixture was dialyzed in a 3.5 kDa MWCO cassette (Thermo Scientific 87722) against\nrefolding buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA; 2 M NaCl; 5 mM β-mercaptoethanol) at 4 °C\nfor 48 h with periodic buffer changes. The dialysate was concentrated to 1 mL using a 5 kDa MWCO\ncentrifugal filter (Millipore), then applied to a HiLoad 16/600 Superdex 200 column (Cytiva\n28989335). Fractions containing equimolar amounts of all four histones (confirmed by SDS-PAGE;\nFig. S10B) were pooled, adjusted to 30% glycerol (v/v), and stored at −20 °C until use. Successful\nhistone octamer assembly was confirmed by reconstitution of mononucleosomes on a test template\nbearing a single Widom 601 nucleosome positioning sequence (Fig. S10C).\nUsing these octamers, nucleosomes were assembled by salt-gradient dialysis, a standard\nmethod that has been extensively validated (Dyer et al., Methods in Enzymology (2003); doi:\n10.1016/S0076-6879(03)75002-2) (Kim et al., Nucleic Acids Research (2019); doi:\n10.1093/nar/gky1244). Although we did not perform optional post-assembly purification, such\nas a sucrose-gradient step, note that our surface-tethered single-molecule assay inherently\nselects for the targeted species: only the particles containing biotinylated DNA molecules were\ncaptured by surface-anchored NeutrAvidin, washing away unbound or misassembled histones\nand their aggregates from the imaging surface.\n[Supplementary Methods, Page 8] Nucleosome reconstitution with 12×601 DNA\no\nNucleosome arrays were reconstituted using the 2.3-kb 601 DNA template (following a previously\nestablished method3,4) with a single biotin modification at one DNA terminus. Purified recombinant\nhuman histone octamers were mixed with the DNA at a molar ratio of 1:12 (based on the twelve\nnucleosome binding sites) in high-salt buffer containing 10 mM Tris-HCl (pH 8.0), 1 mM EDTA,\n1 mM DTT, and 2 M KCl. All buffers were filtered through a 0.22-μm-pore membrane before use\n(CA Syringe filter 0.2 μm, Advantec). The mixture was incubated at 4 °C for 2 h and then subjected\nto stepwise salt-gradient dialysis (2 M → 1.5 M → 1 M → 0.8 M → 0.6 M → 0.4 M → 0.2 M →\n0.1 M), allowing 2-h equilibration at each step followed by an overnight dialysis at 0.1 M to complete\nnucleosome assembly. Dialysis was performed using Slide-A-Lyzer MINI Dialysis Devices (20 kDa\nMWCO, Thermo Fisher Scientific) at 4 °C under gentle stirring. For the characterization of the\nobtained nucleosome assembly, see Supplementary Note 2, “Verification of Nucleosome Assembly\non 601 DNA”.\n• No standard QC performed e.g. Native gel electrophoresis (free DNA vs. nucleosome vs.\nsub-nucleosomal particle) or Sucrose gradient sedimentation, AFM or EM imaging of\nnucleosome arrays, Stoichiometry verification (e.g., SDS-PAGE/Coomassie of assembled\nnucleosomes, fiber-seq). H2A fluorescence alone is not sufficient to confirm full nucleosome\nassembly, since signal could arise from free histones, sub-nucleosomal particles, or aggregates.\n[Reply] As recommended, we performed comprehensive quality-control experiments to\ndemonstrate the integrity and stability of our nucleosome samples with 12×601 DNA,\ncombining standard gel mobility assays, electron microscopy, nucleosome occupancy\nanalysis, an ultimate mechanical-stability test using magnetic tweezers, and, finally, in-situ\ncross-correlation of fluorescence signals from histone (H2A) and DNA (SYTOX) for more\nquantitative verification. The full details are provided in Supplementary Note 2, “Verification\nof Nucleosome Assembly on 601 DNA” (pp.14–16 in Supplementary Information, with\nresults in Figs. S10–S11), with a summary given below:\n• [Native gel and nuclease digestion assay] We ran TBE–PAGE with identical aliquots\nfrom a single nucleosome preparation across lanes, applying different treatments\n(Fig. S10E). In the native lane (without detergent, nuclease, or proteinase), nucleosome-\nloaded 2.3-kb 601 DNA migrated much more slowly than naked DNA (largely retained in\nthe well with a faint smear below) consistent with densely loaded arrays. (Rather than\ndissect native mobility further, we quantified nucleosome occupancy with complementary\nbiophysical assays below.) In the lanes for DNA protection assay, samples were partially\ndigested with micrococcal nuclease (MNase) and then\ndeproteinized with proteinase K before loading; as\nexpected, MNase preferentially cleaved linker DNA,\nproducing a nucleosomal ladder (mono/di/tri/…) whose\nband spacing reports the nucleosome repeat length.\nOver the 0–180 s time course, the pattern resolved\nfrom high-MW material (0 s) into a clear ladder (1–4+)\nand, at longer digestions, into a predominant ~150 bp\nmononucleosome band. A minor smear just below the\n~150 bp band appeared only after extensive digestion,\nlikely reflecting over-digestion to sub-nucleosomal\nfragments rather than pre-existing species.\n[Fig. S10E] Native gel electrophoresis (6% TBE–PAGE, SYBR Gold) with MNase protection for\no\nnucleosome-reconstituted 12×601 DNA. Lane 1: Native sample (without detergent, nuclease, or\nproteinase); Lane 2–5: MNase digestion time course followed by proteinase K, displaying the\ncharacteristic nucleosomal ladder (see Supplementary Note 2).\n• [Electron microscopy] Negative-stain TEM revealed clear beads-on-a-string arrays, very\nfrequently showing ~12 nucleosomes per DNA molecule with uniform spacing (Fig. S11A),\nas expected from the construct design.\n[Fig. S11A] (A) Negative-stain TEM of 12×601 nucleosome arrays; magnified field (inset) shows\no\nbeads-on-a-string morphology.\n• [Colocalization of histone and DNA] After\ncapturing biotinylated nucleosomal DNA on the\nimaging surface, we confirmed nucleosome\npresence in situ by correlating the histone\n(Alexa 488–H2A) and DNA (SYTOX Orange)\nfluorescence signals (Fig. S11B). The histone\nchannel was relatively dim (due to FRET-\nmediated quenching of Alexa 488 by nearby\nSYTOX) but still showed a high correlation with\nDNA fluorescence. A separate imaging without\nDNA fluorescence also estimated the\nnucleosome occupancy per DNA molecule\n(see below).\n[Fig. S11B] (B) Fluorescence images of surface-\no\ncaptured nucleosome arrays: DNA (SYTOX\nOrange) and H2A (Alexa 488) signals with\nPearson correlation quantified for matched versus\nunmatched ROIs (right) (n = 4 pairs).\n• [Nucleosome occupancy from single-molecule photobleaching] The TEM images\nabove provided a qualitative check for the nucleosome occupancy, though throughput was\nlimited. For more direct and quantitative statistics, we performed single-molecule\nphotobleaching analysis of surface-captured 601 nucleosomes labeled on H2A with Alexa\n488 dyes (Fig. S11C–E). During the full-field photobleaching, discrete bleaching steps of\nindividual spots were resolved. The Alexa 488 label counts were broadly distributed\n(reflecting some heterogeneity) but clustered most densely around 8–14 labeled H2A\n(median ~10.3). Given a labeling efficiency of\n~0.6 and two H2A per octamer, this corresponds\nto ~7–12 octamers per DNA, consistent with the\n12-site array and confirming high nucleosome\noccupancy.\n[Fig. S11C–E] (C) Single-molecule photobleaching\no\nstep analysis for nucleosome occupancy estimation.\nRepresentative single-spot photobleaching trace of\nAlexa 488–H2A exhibiting stepwise intensity\ndecreases is shown. Fluorophore count (left axis)\nwas estimated from the uniform bleaching step size.\nInsets: full field of view before and after\nphotobleaching. (D) Distribution of Alexa 488–H2A\ncounts per spot (n = 69 particles). (E) Estimated\noctamer counts per DNA converted from (D) using\nH2A labeling efficiency (~0.6) and 2 H2A per\noctamer.\n• [Magnetic Tweezers assay] Finally, we probed mechanical stability of the nucleosomes\nby force–extension ramps (0–40 pN) on nucleosome arrays assembled on 601 DNA.\nBeyond the low-force fiber-extension regime, we consistently detected ~10–13 discrete\nunwrapping steps per tether above 10 pN; fits to a standard four-state chromatin model\nreproduced these traces, confirming robust nucleosome loading.\n[Fig. S11F] (F) Force–extension curves from magnetic tweezers measurements (red) for two\no\nrepresentative nucleosome arrays, with reference curves overlaid (chromatin fiber, single-\nwrapped state, extended 12-mer, unwrapping transitions, and DNA WLC model).\n• The half-life or observation that arrays are “intact and stable in solution for several days” (for\nwhich no experimental data are shown) cannot be used as a measurement of nucleosome\nquality, as it does not demonstrate correct nucleosome positioning or occupancy.\n[Reply] We agree with this point and have removed the statement. Instead, we hope the\nquality-control results above better demonstrate the stability of our nucleosome samples.\n• The manuscript claims that nucleosome arrays were verified on the “skybridge,” revealing\nbeads-on-a-string (Fig. S10E, Movie S11). However, the presented images do not\nconvincingly show beads-on-a-string and a comparison side by side with canonical 601\nreference nucleosome arrays should be provided to validate morphology.\n[Reply] We believe the beads-on-a-string morphology is now better supported by the new\nWidom-array samples and the TEM imaging shown above (Fig. S11A). Accordingly, we have\nremoved the nucleosome images on λ-DNA that were obtained on the skybridge platform.\n• In Fig. 4A-D, Tau and DNA co-localize convincingly, and Tau+tubulin condensates track DNA\nfilaments. In contrast, Fig. 5 lacks comparable resolution — the DNA filaments are not clearly\nvisible (only large foci), and the Tau signal appears punctate. This could represent Tau\naggregation seeded by misassembled octamers rather than true nucleosome-specific co-\ncondensation.\n[Reply] Using the newly prepared 601 DNA nucleosomes described above, we repeated all\ntau and microtubule assays (Fig. 5G–M). As shown in the verification results above, the new\nDNA template produced densely occupied, well-resolved nucleosome arrays, reducing\nconcerns that misassembled or aggregated histones contributed to the observed signals. Note\nthat in TIRF images (Fig. 5H), DNA filaments were not visible in this sample because its\ncontour length (~0.7 µm) was far shorter than λ-DNA (~16 µm), so we could not resolve how\nindividual nucleosomes interacted with tau condensates. Instead, we compared whether\nnucleosomal DNA exhibited tau and microtubule binding comparable to naked DNA. As\nappended below, nucleosomal particles were sufficient to bind tau and microtubules at levels\ncomparable to naked DNA prepared at an equivalent surface density.\nWe added the new results with 12×601 DNA to the main text:\n[Results, Line 327] Nucleosome-pinned tau clusters support microtubule attachment\no\n“To place our in vitro observations in a more\nphysiologically relevant context, we replaced λ-\nDNA with a 2.3-kb dsDNA bearing 12 Widom-\n601 sites (Fig. 5G), reconstituted nucleosomes\non this template, and repeated the tau-binding\nand microtubule-capture assays. A set of\nbiochemical and biophysical assays indicated\ndensely loaded, stable nucleosome arrays (see\nSupplementary Note 2, “Verification of\nNucleosome Assembly on 601 DNA”, and\nFigs. S10–S11). For a controlled comparison\nto naked DNA, we also prepared naked 601\nDNA (no nucleosome reconstitution) at similar\nsurface density for imaging (Fig. 5H, I).\nBecause the 601 construct is much shorter\nthan λ-DNA (2.3 kb vs. 48.5 kb), individual\nDNA strands were not resolvable by TIRF.\nUpon adding 5 μM tau, discrete tau clusters\nformed and colocalized with H2A-labeled\nnucleosomes (Fig. 5J). Although overall tau\nbinding was lower than on λ-DNA, likely\nreflecting the shorter template length,\ncolocalization analysis showed a high\ncorrelation between nucleosome and tau\nsignals for both nucleosomal and naked 601\nDNA at near-comparable levels (Fig. 5K).\nFinally, introducing a tau–tubulin mixture containing nascent microtubules yielded robust\nmicrotubule capture on surfaces bearing nucleosome-pinned tau clusters (Fig. 5L),\ncomparable\nto λ-DNA-induced condensates (Fig. 4J). Microtubule counts were indistinguishable between\nnaked DNA and nucleosome arrays (Fig. 5M), despite the reduced amount of free DNA segments\nin the latter. Together, these data suggest that nucleosomal DNA remains competent to recruit\ntau and interface with microtubules to a similar extent as its naked-DNA counterpart, consistent\nwith recent bulk measurements of tau association with nucleosome arrays and linker DNA25.”\n[Fig. 5G–M] (G) Schematic of the 2.3-kb DNA construct bearing 12 repeats of the Widom-601\no\nnucleosome positioning sequence. The construct was used either as naked 12×601 DNA or\nassembled into nucleosomes (Nuc) with purified histone octamers. Negative-stain TEM of the\nassembled sample (inset) showed beads-on-a-string arrays with ~12 nucleosomes per DNA and\nuniform spacing (for more images, see Fig. S11). (H) TIRF fields of view for naked and\nnucleosomal conditions at matched surface density. (I) 601 DNA particle densities for the\nconditions shown in (H) (n = 5 images; mean ± s.d.). (J) Fluorescence images of nucleosomes\n(Alexa 488–H2A) and Cy3-labeled tau; arrows mark H2A puncta that colocalize with tau. (K)\nPearson correlation between nucleosome and tau signals for the images in (J) (n = 9 images).\nBox plot shows median (red line), interquartile range, and whiskers to the minimum/maximum.\n(L)\nFluorescence images of nucleosomes (Alexa 488–H2A), tau (Cy3), and microtubules (HiLyte\n647–labeled tubulin). (M) Microtubule counts per field for the conditions shown in (L) (n = 11\nimages; mean ± s.d.).\n[Discussion, Line 526] “In vitro formation of nucleosome-pinned tau condensates, which retain\no\nthe ability to mediate microtubule assembly (Fig. 5), may partially explain why tau condensates\nform preferentially on nucleosome-rich centromeres in dividing cells. Across assays, nucleosomal\ncounterpart. Since fully loaded 601 DNA leaves only short stretches of free DNA, this suggests\nthat nucleosome-wrapped DNA segments still provide effective binding sites for tau. This\ninterpretation is also consistent with a recent report showing strong complexation of tau with\nnucleosome arrays and linker DNA25. ”\n• The authors themselves state in the rebuttal: “We acknowledge that these data cannot\nexclude a model in which Tau predominantly associates with free DNA regions between\nclusters.”\nHowever, the main manuscript and title still claim nucleosome-specific interactions, which is\nmisleading.\n[Reply] We appreciate the feedback and have revised the text and title to clarify that tau may\ninteract with nucleosome arrays via both nucleosome-wrapped DNA and free linker DNA\nbetween nucleosome cores, and that this distinction cannot be resolved with our TIRF imaging.\nWe believe this revision more accurately reflects our experimental evidence.\n• [Section title changed] We changed the section title to “Nucleosome arrays support tau\nbinding and microtubule recruitment” to better reflect what the data show.\n• [Discussion revised] We updated the Discussion to incorporate the new results and to\nnote the above reservations:\n[Discussion Line 525] “In vitro formation of tau clusters on nucleosome arrays, which retain the\no\nability to mediate microtubule assembly (Fig. 5), may partially explain how tau condensates form\non nucleosome-rich centromeres in dividing cells. Across assays, nucleosomal 601 DNA bound\ntau and supported microtubule binding at levels comparable to its naked-DNA counterpart. Since\nfully loaded 601 DNA leaves only short stretches of free DNA, this suggests that nucleosome-\nwrapped DNA segments still provide effective binding sites for tau. Although our TIRF imaging\ncould not resolve whether tau predominantly associates with free linker DNA or with nucleosome\ncores, this interpretation is consistent with a recent bulk study reporting strong complexation of\ntau with nucleosome arrays and comparable nanomolar affinities for both nucleosomal and naked\nDNA25. It will be informative to track tau’s motions on longer DNA substrates (e.g., λ-DNA) to test\nwhether tau drives nucleosome rearrangements with potential epigenetic consequences. Given\ntau’s centromeric enrichment, it will be important to examine tau–nucleosome interactions with\nCENP-A nucleosomes, whose more flexible DNA ends may afford greater access to free DNA\nsegments65.\n• The nucleosome occupancy and clustering analysis are insufficiently supported. The claim\nof “3–4 nucleosomes per cluster” is unclear, how was this number obtained? No calibration\ncurve or reference standard is provided.\n[Reply] As noted above, we have now updated analysis of nucleosome occupancy from\nsingle-molecule fluorescence imaging of the new 601 DNA construct (Fig. S11C−E). The\nresults show ~7–12 nucleosomes per DNA strand (median = 8.6), indicating high occupancy.\nBecause this template is too short to resolve internal clustering, we have removed all claims\nabout nucleosome clusters in the revised manuscript.\n• Electrophoretic mobility shift assays (EMSA) should also be performed for Tau binding to\nnucleosomes, in addition to the DNA, using the same molar ratios side by side. This would\nclarify whether Tau binding or condensation is nucleosome specific.\n[Reply] We appreciate the suggestion to perform bulk EMSA for tau–nucleosome binding.\nWhile we did not include our own EMSA in this revision, Abasi et al. (Comm. Biology, 2024)\nhave already performed the relevant side-by-side assays systematically, which we now cite\nmore prominently. Briefly, they ran EMSAs on free DNA and on 147-bp (linker-free)\nand\n177-bp (linker-containing) mononucleosomes, observing clear band shifts and similar\nnanomolar affinities for DNA and for both mononucleosome substrates. They therefore\nconcluded that tau binds both nucleosomal and linker DNA (i.e., binding is not restricted to\nlinker regions), with linker DNA facilitating compaction/condensation.\nWe added the above citation to Abasi et al. (Ref. 25):\no [Discussion Line 531] “(…) this interpretation is consistent with a recent bulk study reporting\nstrong complexation of tau with nucleosome arrays and comparable nanomolar affinities for both\nnucleosomal and naked DNA25.”\nReviewer #3 (Remarks to the Author):\nThe claims in the manuscript have been greatly strengthened through revision. Therefore, I\nrecommend publication of this manuscript.\n[Reply] We sincerely thank the reviewer for the positive and thoughtful evaluation, the\nencouraging recommendation, and the constructive concerns and insights shared alongside.\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This\nis part of the Nature Communications initiative to facilitate training in peer review and to\nprovide appropriate recognition for Early Career Researchers who co-review manuscripts."}, {"round": 2, "reviewer_comments": {"reviewer_2": "I appreciate the additional data provided. However, major concerns regarding nucleosome array quality remain unresolved.\nAs currently presented, the nucleosome-related data do not support the claim that Tau condensates form on nucleosomes to mediate microtubule attachment. Without rigorous controls and proper validation of nucleosome assembly, the conclusions drawn from these experiments are not sufficiently robust. The observed results could be equally explained by octamer aggregation, misassembled nucleosomes, or nonspecific Tau co-aggregation with free DNA or histones.\nQuality controls are required (detailed below). Alternatively, the nucleosome-specific claims should be removed, and the manuscript should focus solely on Tau–DNA co-condensation, which is better supported by the current data.\n• The manuscript does not provide details on the octamer preparation, assembly conditions, purification steps. The claim of “rigorous purification” after mixing DNA and octamers is not explained in the Methods or Results. Usually this entails a sucrose gradient.\n• No standard QC performed e.g. Native gel electrophoresis (free DNA vs. nucleosome vs. sub-nucleosomal particle) or Sucrose gradient sedimentation, AFM or EM imaging of nucleosome arrays, Stoichiometry verification (e.g., SDS-PAGE/Coomassie of assembled nucleosomes, fiber-seq). H2A fluorescence alone is not sufficient to confirm full nucleosome assembly, since signal could arise from free histones, sub-nucleosomal particles, or aggregates.\n• The half-life or observation that arrays are “intact and stable in solution for several days” (for which no experimental data are shown) cannot be used as a measurement of nucleosome quality, as it does not demonstrate correct nucleosome positioning or occupancy.\n• The manuscript claims that nucleosome arrays were verified on the “skybridge,” revealing beads-on-a-string (Fig. S10E, Movie S11). However, the presented images do not convincingly show beads-on-a-string and a comparison side by side with canonical 601 reference nucleosome arrays should be provided to validate morphology.\n• In Fig. 4A-D, Tau and DNA co-localize convincingly, and Tau+tubulin condensates track DNA filaments. In contrast, Fig. 5 lacks comparable resolution — the DNA filaments are not clearly visible (only large foci), and the Tau signal appears punctate. This could represent Tau aggregation seeded by misassembled octamers rather than true nucleosome-specific co-condensation.\n• The authors themselves state in the rebuttal: “We acknowledge that these data cannot exclude a model in which Tau predominantly associates with free DNA regions between clusters.”\nHowever, the main manuscript and title still claim nucleosome-specific interactions, which is misleading.\n• The nucleosome occupancy and clustering analysis are insufficiently supported. The claim of “3–4 nucleosomes per cluster” is unclear, how was this number obtained? No calibration curve or reference standard is provided.\n• Electrophoretic mobility shift assays (EMSA) should also be performed for Tau binding to nucleosomes, in addition to the DNA, using the same molar ratios side by side. This would clarify whether Tau binding or condensation is nucleosome specific.", "reviewer_3": "The claims in the manuscript have been greatly strengthened through revision. Therefore, I recommend publication of this manuscript.", "reviewer_4": "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 Reviewer Comments on:\nPark et al., “Tau condensation on DNA mediates\nmicrotubule attachment: A potential mitotic role for\ncentromere-localized tau”\nReviewer #2 (Remarks to the Author):\nReviewer #2 (Remarks to the Author):\nI appreciate the additional data provided. However, major concerns regarding nucleosome\narray quality remain unresolved. As currently presented, the nucleosome-related data do not\nsupport the claim that Tau condensates form on nucleosomes to mediate microtubule\nattachment. Without rigorous controls and proper validation of nucleosome assembly, the\nconclusions drawn from these experiments are not sufficiently robust. The observed results\ncould be equally explained by octamer aggregation, misassembled nucleosomes, or\nnonspecific Tau co-aggregation with free DNA or histones.\nQuality controls are required (detailed below). Alternatively, the nucleosome-specific claims\nshould be removed, and the manuscript should focus solely on Tau–DNA co-condensation,\nwhich is better supported by the current data.\n[Reply] We thank the reviewer for the constructive comments and the chance to better support\nour nucleosome-related claims. In this revision, we have substantially updated the\nnucleosome experiments, refined the analyses, and clarified the data presentation. We\naddress each specific point below. In summary:\n• We replaced λ-DNA with a 2.3-kb DNA\nharboring a 12× array of the Widom-601\nsequence (right, Fig. 5G), a canonical choice\nfor robust nucleosome assembly.\n• To confirm the integrity of the nucleosome assemblies, we used multiple biochemical and\nbiophysical assays.\n• We compared nucleosomal versus naked DNA of the same size and sequence for their\nability to bind tau and recruit microtubules.\n• We found that nucleosome-bound DNA showed\ntau and microtubule binding largely comparable\nto naked DNA (right, Fig. 5J–M), despite the\nreduced stretches of free DNA\n• We note that residual free DNA regions may\nunderlie part of the effect, citing prior work, and\nadjust conclusions to avoid over-interpretation.\nWe believe these additions and clarifications fully\naddress the reviewer’s concerns about nucleosome\nintegrity and strengthen the overall conclusions of\nthe study.\nDetailed point-by-point responses are provided below.\n• The manuscript does not provide details on the octamer preparation, assembly conditions,\npurification steps. The claim of “rigorous purification” after mixing DNA and octamers is not\nexplained in the Methods or Results. Usually this entails a sucrose gradient.\n[Reply] Since the previous λ-DNA data did not adequately address nucleosome quality, we\ncompletely replaced λ-DNA with a 2.3-kb dsDNA bearing 12 consecutive Widom 601\nsequences for all nucleosome experiments (quality control results are appended below for the\nspecific concerns). In addition, we now provide more detailed methods for histone-octamer\npreparation, nucleosome assembly on the 601 DNA, and their use in single-molecule imaging.\nThe octamer preparation was already present in the previous Supplementary Information but\nmay have been overlooked because it was bundled with nucleosome reconstitution; we have\nnow separated it into its own section (Supplementary Information: page 8 and Fig. S10B, C).\nBriefly, individual purified histones were denatured, H2A–H2B and H3–H4 were combined with\na slight H2A–H2B excess, refolded by dialysis into high-salt buffer, and purified by size-\nexclusion chromatography. Fractions containing equimolar amounts of all four histones\n(verified by SDS-PAGE) were pooled and stored. Octamer formation was validated by\nreconstitution of mononucleosomes on a single stretch of 601 DNA sequence.\n[Supplementary Methods, Page 7] Preparation of histone octamers\no\nHistone octamers were assembled by refolding denatured histones3,4. Each histone was dissolved\nin unfolding buffer (20 mM Tris-HCl, pH 7.5; 7 M guanidinium-HCl; 10 mM DTT) at room\ntemperature for 1 h. Protein concentrations were determined by A280. H2A and H2B were mixed\nat a 1:1 molar ratio, as were H3 and H4, with the H2A–H2B dimers at 10% molar excess relative to\nthe H3–H4 tetramers. The final protein concentration was adjusted to 1 mg/mL in unfolding buffer,\nand the mixture was dialyzed in a 3.5 kDa MWCO cassette (Thermo Scientific 87722) against\nrefolding buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA; 2 M NaCl; 5 mM β-mercaptoethanol) at 4 °C\nfor 48 h with periodic buffer changes. The dialysate was concentrated to 1 mL using a 5 kDa MWCO\ncentrifugal filter (Millipore), then applied to a HiLoad 16/600 Superdex 200 column (Cytiva\n28989335). Fractions containing equimolar amounts of all four histones (confirmed by SDS-PAGE;\nFig. S10B) were pooled, adjusted to 30% glycerol (v/v), and stored at −20 °C until use. Successful\nhistone octamer assembly was confirmed by reconstitution of mononucleosomes on a test template\nbearing a single Widom 601 nucleosome positioning sequence (Fig. S10C).\nUsing these octamers, nucleosomes were assembled by salt-gradient dialysis, a standard\nmethod that has been extensively validated (Dyer et al., Methods in Enzymology (2003); doi:\n10.1016/S0076-6879(03)75002-2) (Kim et al., Nucleic Acids Research (2019); doi:\n10.1093/nar/gky1244). Although we did not perform optional post-assembly purification, such\nas a sucrose-gradient step, note that our surface-tethered single-molecule assay inherently\nselects for the targeted species: only the particles containing biotinylated DNA molecules were\ncaptured by surface-anchored NeutrAvidin, washing away unbound or misassembled histones\nand their aggregates from the imaging surface.\n[Supplementary Methods, Page 8] Nucleosome reconstitution with 12×601 DNA\no\nNucleosome arrays were reconstituted using the 2.3-kb 601 DNA template (following a previously\nestablished method3,4) with a single biotin modification at one DNA terminus. Purified recombinant\nhuman histone octamers were mixed with the DNA at a molar ratio of 1:12 (based on the twelve\nnucleosome binding sites) in high-salt buffer containing 10 mM Tris-HCl (pH 8.0), 1 mM EDTA,\n1 mM DTT, and 2 M KCl. All buffers were filtered through a 0.22-μm-pore membrane before use\n(CA Syringe filter 0.2 μm, Advantec). The mixture was incubated at 4 °C for 2 h and then subjected\nto stepwise salt-gradient dialysis (2 M → 1.5 M → 1 M → 0.8 M → 0.6 M → 0.4 M → 0.2 M →\n0.1 M), allowing 2-h equilibration at each step followed by an overnight dialysis at 0.1 M to complete\nnucleosome assembly. Dialysis was performed using Slide-A-Lyzer MINI Dialysis Devices (20 kDa\nMWCO, Thermo Fisher Scientific) at 4 °C under gentle stirring. For the characterization of the\nobtained nucleosome assembly, see Supplementary Note 2, “Verification of Nucleosome Assembly\non 601 DNA”.\n• No standard QC performed e.g. Native gel electrophoresis (free DNA vs. nucleosome vs.\nsub-nucleosomal particle) or Sucrose gradient sedimentation, AFM or EM imaging of\nnucleosome arrays, Stoichiometry verification (e.g., SDS-PAGE/Coomassie of assembled\nnucleosomes, fiber-seq). H2A fluorescence alone is not sufficient to confirm full nucleosome\nassembly, since signal could arise from free histones, sub-nucleosomal particles, or aggregates.\n[Reply] As recommended, we performed comprehensive quality-control experiments to\ndemonstrate the integrity and stability of our nucleosome samples with 12×601 DNA,\ncombining standard gel mobility assays, electron microscopy, nucleosome occupancy\nanalysis, an ultimate mechanical-stability test using magnetic tweezers, and, finally, in-situ\ncross-correlation of fluorescence signals from histone (H2A) and DNA (SYTOX) for more\nquantitative verification. The full details are provided in Supplementary Note 2, “Verification\nof Nucleosome Assembly on 601 DNA” (pp.14–16 in Supplementary Information, with\nresults in Figs. S10–S11), with a summary given below:\n• [Native gel and nuclease digestion assay] We ran TBE–PAGE with identical aliquots\nfrom a single nucleosome preparation across lanes, applying different treatments\n(Fig. S10E). In the native lane (without detergent, nuclease, or proteinase), nucleosome-\nloaded 2.3-kb 601 DNA migrated much more slowly than naked DNA (largely retained in\nthe well with a faint smear below) consistent with densely loaded arrays. (Rather than\ndissect native mobility further, we quantified nucleosome occupancy with complementary\nbiophysical assays below.) In the lanes for DNA protection assay, samples were partially\ndigested with micrococcal nuclease (MNase) and then\ndeproteinized with proteinase K before loading; as\nexpected, MNase preferentially cleaved linker DNA,\nproducing a nucleosomal ladder (mono/di/tri/…) whose\nband spacing reports the nucleosome repeat length.\nOver the 0–180 s time course, the pattern resolved\nfrom high-MW material (0 s) into a clear ladder (1–4+)\nand, at longer digestions, into a predominant ~150 bp\nmononucleosome band. A minor smear just below the\n~150 bp band appeared only after extensive digestion,\nlikely reflecting over-digestion to sub-nucleosomal\nfragments rather than pre-existing species.\n[Fig. S10E] Native gel electrophoresis (6% TBE–PAGE, SYBR Gold) with MNase protection for\no\nnucleosome-reconstituted 12×601 DNA. Lane 1: Native sample (without detergent, nuclease, or\nproteinase); Lane 2–5: MNase digestion time course followed by proteinase K, displaying the\ncharacteristic nucleosomal ladder (see Supplementary Note 2).\n• [Electron microscopy] Negative-stain TEM revealed clear beads-on-a-string arrays, very\nfrequently showing ~12 nucleosomes per DNA molecule with uniform spacing (Fig. S11A),\nas expected from the construct design.\n[Fig. S11A] (A) Negative-stain TEM of 12×601 nucleosome arrays; magnified field (inset) shows\no\nbeads-on-a-string morphology.\n• [Colocalization of histone and DNA] After\ncapturing biotinylated nucleosomal DNA on the\nimaging surface, we confirmed nucleosome\npresence in situ by correlating the histone\n(Alexa 488–H2A) and DNA (SYTOX Orange)\nfluorescence signals (Fig. S11B). The histone\nchannel was relatively dim (due to FRET-\nmediated quenching of Alexa 488 by nearby\nSYTOX) but still showed a high correlation with\nDNA fluorescence. A separate imaging without\nDNA fluorescence also estimated the\nnucleosome occupancy per DNA molecule\n(see below).\n[Fig. S11B] (B) Fluorescence images of surface-\no\ncaptured nucleosome arrays: DNA (SYTOX\nOrange) and H2A (Alexa 488) signals with\nPearson correlation quantified for matched versus\nunmatched ROIs (right) (n = 4 pairs).\n• [Nucleosome occupancy from single-molecule photobleaching] The TEM images\nabove provided a qualitative check for the nucleosome occupancy, though throughput was\nlimited. For more direct and quantitative statistics, we performed single-molecule\nphotobleaching analysis of surface-captured 601 nucleosomes labeled on H2A with Alexa\n488 dyes (Fig. S11C–E). During the full-field photobleaching, discrete bleaching steps of\nindividual spots were resolved. The Alexa 488 label counts were broadly distributed\n(reflecting some heterogeneity) but clustered most densely around 8–14 labeled H2A\n(median ~10.3). Given a labeling efficiency of\n~0.6 and two H2A per octamer, this corresponds\nto ~7–12 octamers per DNA, consistent with the\n12-site array and confirming high nucleosome\noccupancy.\n[Fig. S11C–E] (C) Single-molecule photobleaching\no\nstep analysis for nucleosome occupancy estimation.\nRepresentative single-spot photobleaching trace of\nAlexa 488–H2A exhibiting stepwise intensity\ndecreases is shown. Fluorophore count (left axis)\nwas estimated from the uniform bleaching step size.\nInsets: full field of view before and after\nphotobleaching. (D) Distribution of Alexa 488–H2A\ncounts per spot (n = 69 particles). (E) Estimated\noctamer counts per DNA converted from (D) using\nH2A labeling efficiency (~0.6) and 2 H2A per\noctamer.\n• [Magnetic Tweezers assay] Finally, we probed mechanical stability of the nucleosomes\nby force–extension ramps (0–40 pN) on nucleosome arrays assembled on 601 DNA.\nBeyond the low-force fiber-extension regime, we consistently detected ~10–13 discrete\nunwrapping steps per tether above 10 pN; fits to a standard four-state chromatin model\nreproduced these traces, confirming robust nucleosome loading.\n[Fig. S11F] (F) Force–extension curves from magnetic tweezers measurements (red) for two\no\nrepresentative nucleosome arrays, with reference curves overlaid (chromatin fiber, single-\nwrapped state, extended 12-mer, unwrapping transitions, and DNA WLC model).\n• The half-life or observation that arrays are “intact and stable in solution for several days” (for\nwhich no experimental data are shown) cannot be used as a measurement of nucleosome\nquality, as it does not demonstrate correct nucleosome positioning or occupancy.\n[Reply] We agree with this point and have removed the statement. Instead, we hope the\nquality-control results above better demonstrate the stability of our nucleosome samples.\n• The manuscript claims that nucleosome arrays were verified on the “skybridge,” revealing\nbeads-on-a-string (Fig. S10E, Movie S11). However, the presented images do not\nconvincingly show beads-on-a-string and a comparison side by side with canonical 601\nreference nucleosome arrays should be provided to validate morphology.\n[Reply] We believe the beads-on-a-string morphology is now better supported by the new\nWidom-array samples and the TEM imaging shown above (Fig. S11A). Accordingly, we have\nremoved the nucleosome images on λ-DNA that were obtained on the skybridge platform.\n• In Fig. 4A-D, Tau and DNA co-localize convincingly, and Tau+tubulin condensates track DNA\nfilaments. In contrast, Fig. 5 lacks comparable resolution — the DNA filaments are not clearly\nvisible (only large foci), and the Tau signal appears punctate. This could represent Tau\naggregation seeded by misassembled octamers rather than true nucleosome-specific co-\ncondensation.\n[Reply] Using the newly prepared 601 DNA nucleosomes described above, we repeated all\ntau and microtubule assays (Fig. 5G–M). As shown in the verification results above, the new\nDNA template produced densely occupied, well-resolved nucleosome arrays, reducing\nconcerns that misassembled or aggregated histones contributed to the observed signals. Note\nthat in TIRF images (Fig. 5H), DNA filaments were not visible in this sample because its\ncontour length (~0.7 µm) was far shorter than λ-DNA (~16 µm), so we could not resolve how\nindividual nucleosomes interacted with tau condensates. Instead, we compared whether\nnucleosomal DNA exhibited tau and microtubule binding comparable to naked DNA. As\nappended below, nucleosomal particles were sufficient to bind tau and microtubules at levels\ncomparable to naked DNA prepared at an equivalent surface density.\nWe added the new results with 12×601 DNA to the main text:\n[Results, Line 327] Nucleosome-pinned tau clusters support microtubule attachment\no\n“To place our in vitro observations in a more\nphysiologically relevant context, we replaced λ-\nDNA with a 2.3-kb dsDNA bearing 12 Widom-\n601 sites (Fig. 5G), reconstituted nucleosomes\non this template, and repeated the tau-binding\nand microtubule-capture assays. A set of\nbiochemical and biophysical assays indicated\ndensely loaded, stable nucleosome arrays (see\nSupplementary Note 2, “Verification of\nNucleosome Assembly on 601 DNA”, and\nFigs. S10–S11). For a controlled comparison\nto naked DNA, we also prepared naked 601\nDNA (no nucleosome reconstitution) at similar\nsurface density for imaging (Fig. 5H, I).\nBecause the 601 construct is much shorter\nthan λ-DNA (2.3 kb vs. 48.5 kb), individual\nDNA strands were not resolvable by TIRF.\nUpon adding 5 μM tau, discrete tau clusters\nformed and colocalized with H2A-labeled\nnucleosomes (Fig. 5J). Although overall tau\nbinding was lower than on λ-DNA, likely\nreflecting the shorter template length,\ncolocalization analysis showed a high\ncorrelation between nucleosome and tau\nsignals for both nucleosomal and naked 601\nDNA at near-comparable levels (Fig. 5K).\nFinally, introducing a tau–tubulin mixture containing nascent microtubules yielded robust\nmicrotubule capture on surfaces bearing nucleosome-pinned tau clusters (Fig. 5L),\ncomparable\nto λ-DNA-induced condensates (Fig. 4J). Microtubule counts were indistinguishable between\nnaked DNA and nucleosome arrays (Fig. 5M), despite the reduced amount of free DNA segments\nin the latter. Together, these data suggest that nucleosomal DNA remains competent to recruit\ntau and interface with microtubules to a similar extent as its naked-DNA counterpart, consistent\nwith recent bulk measurements of tau association with nucleosome arrays and linker DNA25.”\n[Fig. 5G–M] (G) Schematic of the 2.3-kb DNA construct bearing 12 repeats of the Widom-601\no\nnucleosome positioning sequence. The construct was used either as naked 12×601 DNA or\nassembled into nucleosomes (Nuc) with purified histone octamers. Negative-stain TEM of the\nassembled sample (inset) showed beads-on-a-string arrays with ~12 nucleosomes per DNA and\nuniform spacing (for more images, see Fig. S11). (H) TIRF fields of view for naked and\nnucleosomal conditions at matched surface density. (I) 601 DNA particle densities for the\nconditions shown in (H) (n = 5 images; mean ± s.d.). (J) Fluorescence images of nucleosomes\n(Alexa 488–H2A) and Cy3-labeled tau; arrows mark H2A puncta that colocalize with tau. (K)\nPearson correlation between nucleosome and tau signals for the images in (J) (n = 9 images).\nBox plot shows median (red line), interquartile range, and whiskers to the minimum/maximum.\n(L)\nFluorescence images of nucleosomes (Alexa 488–H2A), tau (Cy3), and microtubules (HiLyte\n647–labeled tubulin). (M) Microtubule counts per field for the conditions shown in (L) (n = 11\nimages; mean ± s.d.).\n[Discussion, Line 526] “In vitro formation of nucleosome-pinned tau condensates, which retain\no\nthe ability to mediate microtubule assembly (Fig. 5), may partially explain why tau condensates\nform preferentially on nucleosome-rich centromeres in dividing cells. Across assays, nucleosomal\ncounterpart. Since fully loaded 601 DNA leaves only short stretches of free DNA, this suggests\nthat nucleosome-wrapped DNA segments still provide effective binding sites for tau. This\ninterpretation is also consistent with a recent report showing strong complexation of tau with\nnucleosome arrays and linker DNA25. ”\n• The authors themselves state in the rebuttal: “We acknowledge that these data cannot\nexclude a model in which Tau predominantly associates with free DNA regions between\nclusters.”\nHowever, the main manuscript and title still claim nucleosome-specific interactions, which is\nmisleading.\n[Reply] We appreciate the feedback and have revised the text and title to clarify that tau may\ninteract with nucleosome arrays via both nucleosome-wrapped DNA and free linker DNA\nbetween nucleosome cores, and that this distinction cannot be resolved with our TIRF imaging.\nWe believe this revision more accurately reflects our experimental evidence.\n• [Section title changed] We changed the section title to “Nucleosome arrays support tau\nbinding and microtubule recruitment” to better reflect what the data show.\n• [Discussion revised] We updated the Discussion to incorporate the new results and to\nnote the above reservations:\n[Discussion Line 525] “In vitro formation of tau clusters on nucleosome arrays, which retain the\no\nability to mediate microtubule assembly (Fig. 5), may partially explain how tau condensates form\non nucleosome-rich centromeres in dividing cells. Across assays, nucleosomal 601 DNA bound\ntau and supported microtubule binding at levels comparable to its naked-DNA counterpart. Since\nfully loaded 601 DNA leaves only short stretches of free DNA, this suggests that nucleosome-\nwrapped DNA segments still provide effective binding sites for tau. Although our TIRF imaging\ncould not resolve whether tau predominantly associates with free linker DNA or with nucleosome\ncores, this interpretation is consistent with a recent bulk study reporting strong complexation of\ntau with nucleosome arrays and comparable nanomolar affinities for both nucleosomal and naked\nDNA25. It will be informative to track tau’s motions on longer DNA substrates (e.g., λ-DNA) to test\nwhether tau drives nucleosome rearrangements with potential epigenetic consequences. Given\ntau’s centromeric enrichment, it will be important to examine tau–nucleosome interactions with\nCENP-A nucleosomes, whose more flexible DNA ends may afford greater access to free DNA\nsegments65.\n• The nucleosome occupancy and clustering analysis are insufficiently supported. The claim\nof “3–4 nucleosomes per cluster” is unclear, how was this number obtained? No calibration\ncurve or reference standard is provided.\n[Reply] As noted above, we have now updated analysis of nucleosome occupancy from\nsingle-molecule fluorescence imaging of the new 601 DNA construct (Fig. S11C−E). The\nresults show ~7–12 nucleosomes per DNA strand (median = 8.6), indicating high occupancy.\nBecause this template is too short to resolve internal clustering, we have removed all claims\nabout nucleosome clusters in the revised manuscript.\n• Electrophoretic mobility shift assays (EMSA) should also be performed for Tau binding to\nnucleosomes, in addition to the DNA, using the same molar ratios side by side. This would\nclarify whether Tau binding or condensation is nucleosome specific.\n[Reply] We appreciate the suggestion to perform bulk EMSA for tau–nucleosome binding.\nWhile we did not include our own EMSA in this revision, Abasi et al. (Comm. Biology, 2024)\nhave already performed the relevant side-by-side assays systematically, which we now cite\nmore prominently. Briefly, they ran EMSAs on free DNA and on 147-bp (linker-free)\nand\n177-bp (linker-containing) mononucleosomes, observing clear band shifts and similar\nnanomolar affinities for DNA and for both mononucleosome substrates. They therefore\nconcluded that tau binds both nucleosomal and linker DNA (i.e., binding is not restricted to\nlinker regions), with linker DNA facilitating compaction/condensation.\nWe added the above citation to Abasi et al. (Ref. 25):\no [Discussion Line 531] “(…) this interpretation is consistent with a recent bulk study reporting\nstrong complexation of tau with nucleosome arrays and comparable nanomolar affinities for both\nnucleosomal and naked DNA25.”\nReviewer #3 (Remarks to the Author):\nThe claims in the manuscript have been greatly strengthened through revision. Therefore, I\nrecommend publication of this manuscript.\n[Reply] We sincerely thank the reviewer for the positive and thoughtful evaluation, the\nencouraging recommendation, and the constructive concerns and insights shared alongside.\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This\nis part of the Nature Communications initiative to facilitate training in peer review and to\nprovide appropriate recognition for Early Career Researchers who co-review manuscripts."}, {"round": 3, "reviewer_comments": {"reviewer_2": "The newly added experiments adequately substantiate the manuscript’s claims. I recommend publication.", "reviewer_3": "The claims in the manuscript have been greatly strengthened through revision. Therefore, I recommend publication of this manuscript.", "reviewer_4": "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": ""}]

Decision letter