Synthetic aptamer mechanoreceptors enable cell-specific force sensing and temporal control via DNA circuits

Domain: biology
Published: 2026-03-15
Editorial outcome: major_revision

Abstract
Cells interpret mechanical cues from their microenvironment with spatio-temporal precision to guide adaptive behaviors. However, engineering synthetic mechanosensing systems with both cell-specificity and programmability remains challenging, especially when targeting ubiquitous classical mechanoreceptors. Here, we introduce an all-DNA mechanosensing platform based on aptamers that transmit force through noncanonical surface receptors. Aptamer–receptor recognition acts as a molecular gate for force transduction, enabling the design of mechanoprobes with cell-type selectivity. These probes interpret diverse mechanical inputs via distinct mechanisms, including actomyosin-driven contractility and membrane ruffling during macropinocytosis. By integrating aptamer mechanoprobes with upstream DNA reaction networks, we achieve reversible and temporally programmable mechanoresponses. This modular, all-nucleic-acid system offers a general framework for constructing tunable mechanotransduction circuits. It expands the design space for synthetic mechanobiology and provides opportunities for autonomous, multi-layered mechanical–biochemical regulation in tissue engineering, morphogenesis, and dynamic cell programming.

Introduction
Mechanical forces are key regulators of cellular behavior, shaping processes from morphogenesis to disease progression by guiding how cells sense, adapt, and respond to their microenvironment1–4. These responses are typically mediated by mechanoreceptors, which transduce physical stimuli into biochemical signals controlling cell fate, tissue structure, and homeostasis. Understanding and customizing the mechanotransduction process is pivotal for advancing mechanotargeting-based therapies, tissue engineering, and regenerative medicine5,6. However, orchestrating orthogonal cell-specific processes and implementing autonomous control mechanisms during complex morphogenesis is still challenged by the limited specificity, versatility, and dynamic tunability of the current mechanosensing toolbox7–9.

Selectivity and differential force sensing at the cellular level are key design principles for engineering synthetic mechanosensing materials. Cells endogenously express a variety of mechanoreceptors, including integrins, cadherins, T cell receptors, and Piezo channels to accommodate diverse mechanical demands10–13. To engage with those, integrin-targeting mechanoprobes (MPs) have been most widely explored14. Pioneering studies have employed DNA-based MPs with Arg-Gly-Asp (RGD) motifs to measure piconewton (pN) forces transmitted through integrins. Depending on the DNA architecture, such probes can provide either irreversible (historical) readouts via duplex rupture or reversible (real-time) readouts via hairpin unfolding15. Recent innovations have even enabled quantification of single-molecule force loading rates16,17. However, the broad and often non-specific expression of integrins across cell types undermines terminal specificity10,18, limiting their applicability in mixed or heterogeneous cell populations. Greater selectivity can be achieved using protein- or antibody-based ligands conjugated to DNA-MPs, which have proven valuable for interrogating T cell signaling and for constructing mechanosensitive cell-cell junctions12,19. In addition to sensing forces, previous DNA MPs have made use of force-induced strand separation to reveal cryptic domains amenable for downstream reactivity, such as in situ dimerization or for ex situ amplification20,21. Fully protein-based sensors that mimic Notch signaling offer additional strategies to encode both receptor specificity and force responsiveness, yet these approaches typically require labor-intensive protein engineering, including insertion-site optimization to maintain receptor localization and biological function—thereby limiting accessibility, modularity, and scalability22,23.

We hypothesized that aptamers—short single-stranded DNA sequences (ssDNA) selected for high-affinity binding to specific cell surface receptors—could serve as a versatile and new family of synthetic mechanosensors. Unlike classical mechanotransduction targets such as integrins or cadherins, many aptamer-accessible receptors are not canonically associated with force transmission. However, given their roles in signaling, trafficking, and cell migration, and since cells continuously recycle surface proteins for quality control, these receptors may transiently interact with the cytoskeleton or become indirectly coupled to intracellular force-generating machinery—potentially enabling mechanical engagement through previously unrecognized mechanisms. Indeed, a recent proof-of-concept using the CI-M6PR-targeting aptamer demonstrated that an aptamer can function as a force-reception module to sense weak endocytic forces (~4 pN)24 and then translate this force cue into intracellular signals via DNA-mediated receptor dimerization20. However, that system required soft DNA hairpins suffering from dynamic breathing behavior, which limits force resolution and complicates integration with DNA reaction circuits. More generally, the design space for aptamer-based force sensors remains largely unclear—particularly in terms of cell-type specificity, force thresholding, and mechanistic understanding of how force is generated at the aptamer-receptor interface.

Importantly, because aptamers are nucleic acids, they are inherently compatible with DNA-based strand displacement reactions and programmable logic circuits25–27. This opens opportunities for dynamic and autonomous regulation of mechanosensing—features that are difficult to achieve with conventional peptide- or protein-based systems like RGD-integrin probes or SynNotch receptors.

Here, we present a modular and fully programmable mechanosensing platform built entirely from nucleic acids, in which DNA aptamers function as synthetic receptor ligands embedded within force-responsive DNA MPs. This approach combines the receptor specificity of aptamers with the tunable force thresholds of DNA mechanics, enabling the systematic exploration of non-canonical and previously unrecognized force-generating surface receptors across diverse cell types. Using this platform, we elucidate the magnitude of noncanonically transmitted forces through DNA MP geometry, and robust cell-type selectivity dictated by aptamer-receptor binding. In contrast to conventional integrin-targeting RGD probes, our aptamer MPs display distinct force transmission behaviors, revealing mechanistic insights into how specific receptor contexts modulate mechanical engagement. Going further, leveraging the nucleic acid nature of aptamers, we integrate these probes with upstream DNA-based reaction networks to achieve temporal gating and switchable mechanosensitivity. This platform establishes a general framework for constructing adaptive, programmable mechano-interfaces and expands the synthetic biology toolkit for force-guided cell control.

Results
General concept for aptamer-based mechanosensing platform
Our aptamer-based MPs are composed of three modular elements: a terminal aptamer that selectively binds a target receptor, a DNA duplex that acts as a tunable force-sensitive element, and a fluorophore–quencher pair that converts mechanical unfolding into a fluorescence signal (Fig. 1). The aptamer functions as a programmable ligand, enabling receptor-specific engagement and ensuring that only receptor-expressing cells generate detectable force signals. To demonstrate the versatility of this platform, we employ five distinct and frequently used aptamers targeting diverse cell-surface receptors across five different cell lines. Moreover, the nucleic acid-based architecture of the aptamer permits dynamic control of receptor recognition via complementary blocker strands, which can be regulated through DNA-based reaction networks to achieve multi-layered regulatory complexity, such as temporally gated mechanosensing. For surface functionalization, we immobilized biotinylated MPs onto streptavidin-coated glass slides and co-immobilized biotinylated RGD peptides to support baseline integrin-mediated cell adhesion. Supported lipid bilayer-based surface density calibration assays reveal that the densities of different aptamer MPs are in the range of 3500–4500 molecules/µm², and the overall stoichiometric ratio of MPs to RGD is ~3:1 (Supplementary Fig. 1, Supplementary Table 2).

Force transmission of aptamer MPs requires specific aptamer-receptor recognition
To investigate the role of aptamer–receptor recognition in mechanosensing, we designed a MP incorporating the AS1411 aptamer, which binds nucleolin (dissociation constant Kd = 69.1 nM)28—a membrane-associated receptor overexpressed in many cancer cells29. We selected nucleolin-rich HeLa cells as the test system and used an AS1411 MP with a zipper-type duplex designed to rupture under a low force tolerance (Ttol) of 12 pN. Subsequently, we seeded cells onto the MP-functionalized surface and imaged them by total internal reflection fluorescence (TIRF) microscopy after 4 h incubation at 37 °C in 1% FBS/DMEM medium. Two controls highlight the successful specific activation: One control bears a randomized sequence without affinity to nucleolin, and one lacks any ligand (Fig. 2a). As expected, only the AS1411 MP generates significant fluorescence, confirming that specific aptamer-receptor binding enables force transmission (Fig. 2a, b). Neither control produces signal, confirming the absence of any non-specific interactions, such as from dye-membrane interactions (Fig. 2b). Interestingly, cell spreading area and cell aspect ratio are similar for the three surfaces (AS1411 MP, random MP and negative control) within the 4 h observation period, indicating that the AS1411-nucleolin couple does not promote cell adhesion (Fig. 2c, Supplementary Fig. 2).

Next, we validate that the AS1411 MP specifically engages the nucleolin receptor by performing colocalization and competitive binding assays. To this end, we employed the TB-5 aptamer30, which binds to a distinct epitope on nucleolin compared to AS1411, and co-immobilized TB-5 and AS1411 MP on the surface at a 1:1 ratio. The mechanosignals from both MPs colocalize, confirming that they both engage nucleolin (Fig. 2d). To further confirm specificity, we blocked the AS1411–nucleolin interaction using endostatin, a natural ligand for nucleolin29, which reduces the AS1411-derived mechanosignals by 21.7% (Fig. 2e). Flow cytometry corroborates this finding, showing a 23.2% decrease in AS1411 binding to HeLa cells in the presence of endostatin (Supplementary Fig. 3). Together, these results demonstrate that force transmission through the AS1411 MP originates from specific aptamer–receptor recognition.

Aptamer MPs reveal force magnitudes across noncanonical mechanoreceptors
To compare the force magnitude of aptamer-targeted receptors with common RGD-targeted integrin21, we designed each MP with two distinct force geometries. In the unzipping mode, the duplex opens base-by-base, resulting in a low Ttol of 12 pN. In contrast, the shearing mode applies force across multiple base pairs near the duplex termini, yielding a higher Ttol of 54 pN based on sequence design (Fig. 3a)15. Comparing first both RGD MPs, the RGD-unzipping MP produces markedly stronger fluorescence signals than the RGD-shearing MP, particularly at peripheral focal adhesions (FAs) and at central regions indicative of early-stage adhesion events (Fig. 3b). This pattern reflects the lower Ttol of the unzipping mode, which enables detection of nascent, low-force adhesion events that fail to activate RGD-shearing MP. Although altering the force geometry modulates the average mechanoresponse pattern per cell (Fig. 3c), the overall response based on the integrated intensity remains indistinguishable and independent of the MP geometry (Fig. 3d). The integrated intensity reflects the cumulative mechano-signal across the entire cell footprint and is thus not only regulated by intrinsic Ttol of the probe but inherently dependent on cell area19. This apparent convergence arises because RGD-integrin linkages are capable of transmitting single-molecule forces that exceed both Ttol values (>54 pN), and in some cases can even rupture the streptavidin-biotin linkage (>100 pN)31. However, the generation of such high forces is contingent on adequate mechanical resistance from the substrate—typically requiring 33-44 pN counterforce to sustain cytoskeletal maturation and stable adhesion32. As a result, shearing-mode MPs promote cell spreading and increase overall force generation, thereby blurring the difference in integrated mechanoresponse between both geometries (Fig. 3e)19.

In contrast, AS1411 MPs show a pronounced distinction between unzipping and shearing geometry. The unzipping mode consistently produces stronger signals in both mean and integrated intensity (Fig. 3b–d). Two factors account for this behavior: First, the AS1411-nucleolin couple transmits substantially lower force (consistently below 54 pN) than RGD-integrin engagement. Second, unlike integrins, the AS1411-nucleolin pair alone does not promote cell spreading (Figs. 2c, 3e). This property makes the shearing mode of aptamer MPs particularly well-suited for tandem integration with other DNA nanodevices33,34, where mechanical stability is essential to avoid unintended force-induced detachment.

Since the above results suggest that aptamers cannot promote cell adhesion, it raises the question to what extent baseline adhesion is required for aptamer MPs to function. To address this, we prepared surfaces functionalized solely with aptamer MPs without co-immobilized RGD to investigate mechanosensing under minimal adhesion conditions. HeLa cells are largely unable to spread on surfaces presenting AS1411 MPs alone, regardless of whether unzipping or shearing MPs are used (Fig. 3f). Compared with RGD co-immobilized conditions, AS1411 unzipping-mediated mechanosignals are significantly reduced to below 24.4% (Fig. 3g), implying that baseline cell adhesion is a prerequisite for AS1411-based force sensing.

Considering that we aim to develop receptor-orthogonal aptamer MPs, we further investigated the Sgc8-aptamer as a potential MP for the protein tyrosine kinase 7 (PTK7), using HepG2 cells that overexpress this receptor. Interestingly, Sgc8 MPs-functionalized surfaces exhibit a somewhat different behavior (Fig. 3h). Even though Sgc8 MPs alone also fail to promote HepG2 cell spreading, robust mechanoactivation occurs (Fig. 3i). Even in a non-spread, quasi-suspended state (see brightfield images), HepG2 cells effectively activate Sgc8 unzipping MPs, yielding ring-like mechanosignals. This indicates that for Sgc8-PTK7, baseline cell adhesion is not a prerequisite for mechanoactivation. The differential adhesion dependence of these two aptamer MPs suggests that they transmit forces through fundamentally distinct mechanisms, which we will further discuss below.

Source of forces associated with noncanonical mechanoreceptors
Although nucleolin is a well-established molecular target35,36, it has not been previously associated with the exertion of mechanical force on the extracellular environment. Our data reveal distinct differences to integrin-targeting MPs, prompting us to investigate the source of nucleolin-associated forces. To test whether aptamer-receptor internalization contributes to force generation, we treated HeLa cells with pathway-specific inhibitors. Neither chlorpromazine (CPZ, 28 µM), which inhibits clathrin-mediated internalization37, nor methyl-β-cyclodextrin (MβCD, 1.9 mM), which inhibits caveolin- and lipid-raft-mediated uptake38, alter the mechanosignal intensity (Fig. 4a, b). In contrast, treatment with 5-(N-ethyl-N-isopropyl) amiloride (EIPA, 100 µM), an inhibitor of macropinocytosis39, reduces mechanosignals by 29.8% (Fig. 4b). This implicates macropinocytosis in the mechanoactivation of the AS1411 MP, consistent with previous findings that cancer cells take up free AS1411 aptamers predominantly via macropinocytosis40. Flow cytometry furthers support this conclusion, showing a 76.6% reduction in the uptake of free AS1411 aptamers by HeLa cells in the presence of EIPA (Supplementary Fig. 4). Notably, compared to the free state in solution, the inhibitory effect of EIPA decreases by 46% when AS1411 is immobilized as an MP on a surface. These results suggest that while macropinocytosis contributes to force generation, it may not be the main source of nucleolin-mediated force. Interestingly, previous studies reported molecular associations between nucleolin and multiple focal adhesion-related or cytoskeletal components, including integrins αvβ3, αvβ5, β1, focal adhesion protein talin44, and myosin heavy chain 945. To determine whether nucleolin-mediated forces spatially correlate with FA structures, we transfected HeLa cells with a plasmid for constitutive expression of monomeric enhanced green fluorescent protein (mEGFP)-paxillin and visualized the actin cytoskeleton with phalloidin staining. Notably, nucleolin-generated forces are clearly localized at peripheral FA sites marked by paxillin, suggesting a spatial and functional association between nucleolin and FAs (Fig. 4c, d, Supplementary Fig. 5). Based on this, we hypothesized nucleolin may indirectly couple to the actomyosin cytoskeleton to transmit mechanical force (Fig. 4e). To identify the cytoskeletal components responsible for this transmission, we treated cells with specific inhibitors after allowing 30 min of initial adhesion16,46. Inhibiting actin polymerization with cytochalasin D (10 µM) moderately reduces mechanosignals by 38.9% (Fig. 4f, Supplementary Fig. 6). In contrast, blocking myosin II ATPase activity with blebbistatin (25 µM) causes a pronounced 77.5% decrease. Together, these results demonstrate that nucleolin-mediated forces primarily depend on myosin-driven contractility, which may be relayed through FAs or other physical linkages independently coupled to the actomyosin network before reaching nucleolin.

To understand the differential dependence of Sgc8 aptamer MPs on baseline adhesion (see above, Fig. 3f, h), we investigated the underlying source of Sgc8-PTK7 forces. Unlike the FA-like patterns observed for nucleolin, PTK7-activated mechanosignals display spatially distinct, ring-like force patterns with submicron size (Fig. 4g). Those are consistent with a macropinocytic origin, as clathrin- and caveolin-mediated internalization typically generates vesicles in the nanometer range47. To test this hypothesis, we inhibited macropinocytosis using EIPA (100 µM), which substantially reduces mechanoactivation by 51.9 %, whereas CPZ (28 µM) and MβCD (1.9 mM) inhibiting clathrin- and caveolin-mediated endocytosis, have no effect (Fig. 4g, h, Supplementary Fig. 7). To further examine the spatial organization of these signals, we stained the plasma membrane and performed TIRF microscopy. As expected, the ring-like mechanofluorescence predominantly localizes to the peripheries of dark membrane invaginations, consistent with force generation during membrane ruffling in early-stage macropinocytosis (Fig. 4i, j, Supplementary Fig. 8)48.

To understand the cytoskeletal origin of this force generation, we next examined the roles of actin polymerization and myosin action during different macropinocytic phases. Actin polymerization is critical for driving membrane ruffling and protrusion formation during the early-stage macropinocytosis, whereas myosins contribute to actin network contraction at the base of ruffles as the membrane closes to form macropinosomes49. Treatment with cytochalasin D (10 µM, actin polymerization inhibitor) and blebbistatin (25 µM, myosin II inhibitor) reduces mechanosignals by 56.1 % and 32.6%, respectively, suggesting that force generation is dominated by actin polymerization rather than myosin-driven contraction (Fig. 4k, Supplementary Fig. 9). Phosphoinositide 3-kinase (PI3K) catalyzes membrane sealing by generating PIP3 at the ruffle bases, thereby recruiting effectors for actin remodeling and vesicle closure50. While PI3K activity is essential for membrane sealing, it is dispensable for ruffle formation51. Consistent with this, inhibition of PI3K using LY294002 (50 µM) does not affect mechanosignals, further confirming that PTK7-mediated forces originate primarily from early-stage membrane ruffling during macropinocytosis with minimal contribution from the sealing phase (Fig. 4l, m).

To further distinguish between two force transmission mechanisms, we knocked down the orthogonal regulator neuronal Wiskott-Aldrich syndrome protein (N-WASP). As a critical cytoskeletal regulator, N-WASP interacts with the Arp2/3 complex to drive branched actin polymerization52, thereby regulating essential cellular processes such as membrane ruffling, endocytosis, and cytoskeletal remodeling. Inhibition of the Arp2/3 complex has been shown to inhibit Arp2/3-mediated actin polymerization in phagocytic adhesion rings of macrophages but has little effect on integrin-mediated tensions in FAs53. The latter depends on myosin II-loaded stress fibers which lack the Arp2/3 complex. For the AS1411-nucleolin pair, compared with cells transfected with scrambled siRNA, N-WASP knockdown HeLa cells still exhibit a stripe-like force pattern, although the signal amplitude is reduced by 27.9% (Fig. 4n, Supplementary Fig. 10a). This result indicates that force transmission through the AS1411-nucleolin pair is weakly dependent on N-WASP. In contrast, for the Sgc8-PTK7 pair, N-WASP knockdown in HepG2 cells leads to an almost complete loss of the ring-like pattern, with a 57.8% reduction in signal intensity relative to scrambled controls, suggesting a strong dependence on N-WASP-mediated actin dynamics (Fig. 4o, Supplementary Fig. 10b). Moreover, the different effects of N-WASP knockdown in both cell lines closely mirror the results observed upon EIPA treatment (Fig. 4b, h). This is consistent with the fact that EIPA inhibits Rac1/Cdc42 activity, and Cdc42 is a key upstream activator of N-WASP54. Together, these results demonstrate that the AS1411-nucleolin and Sgc8-PTK7 pairs exhibit distinct dependencies on N-WASP and downstream Arp2/3-mediated branched actin polymerization, reflecting fundamentally different force transmission pathways.

Versatility of aptamer MPs to customize cell-type-specific mechanoresponses
The specificity of aptamer-receptor recognition enables the modular customization of MPs for cell-type-specific mechanosensing. We demonstrate this principle using three aptamer MPs—Sgc8, MUC1 S2.2, and SYL3c—which target PTK7 (Kd = 0.85 nM)55, mucin-1 (Kd = 0.12 nM)56, and epithelial cell adhesion molecule (EpCAM, Kd = 38 nM)57, respectively. We tested each MP on both receptor-high-expressing and receptor-low-expressing cell lines, selected based on differential receptor expression profiles confirmed by flow cytometry (Supplementary Fig. 11). Mucin-1-high-expressing HeLa cells strongly activate the MUC1 S2.2 MP, whereas HepG2 cells with substantially lower levels of mucin-1 show minimal response (Fig. 5a, b). Conversely, HeLa cells fail to activate the SYL3c MP due to low EpCAM receptor levels. This contrasts with EpCAM-high-expressing MDA-MB-231 cells (Fig. 5c, d). Similarly, HepG2 cells generate stronger activation of the Sgc8 MP than A549 cells, consistent with their higher PTK7 expression (Fig. 5e, f). Interestingly, beyond expression level-dependent signal magnitude, different cell types expressing the same receptor can exhibit distinct, cell-type-dependent force patterns. HeLa cells express PTK7 at an intermediate level (Supplementary Fig. 11e, f). Mechanosensing using Sgc8 MPs reveals a stripe-like force pattern in contrast to the characteristic ring-like mechano-signals observed in HepG2 cells (Fig. 5e, f). These results indicate that different cell types may transmit forces through the same receptor via distinct pathways, a phenomenon that cannot be inferred from receptor expression levels alone. Collectively, the high differential selectivity of aptamer-receptor MP pairs highlights the broad adaptability of our platform for sensing mechanoresponses across diverse cell types and receptor classes.

Motivated by the need for selective and precise mechanoresponse in complex multicellular environments, we extend our validation beyond simple single-cell-type systems. To this end, we labeled receptor-high-expressing and receptor-low-expressing cells with CellTrace Deep Red and Violet dyes, respectively, and co-seeded them in wells containing the corresponding aptamer MPs to assess cell-type-specific mechanoresponse (Fig. 5g). Indeed, mechanoactivation of the Sgc8 MP occurs exclusively in the Deep Red-labeled PTK7-high-expressing HepG2 cells, while the Violet-labeled PTK7-low-expressing A549 cells exhibit much lower signals (Fig. 5h, i). Switching to the MUC1 S2.2 MP leads to strong mechano-signals in the Deep Red-labeled mucin-1-high-expressing HeLa cells, whereas Violet-labeled mucin-1-low-expressing HepG2 cells show minimal signals (Fig. 5j, k). These results underscore the utility of aptamer MPs as customizable mechanical elements for precise, cell-type-specific mechanochemical transduction in heterogeneous cell populations.

Aptamer MPs enable programmable mechanoresponses via integration with DNA-based reaction networks (DRNs)
Natural mechanotransduction involves temporally orchestrated biochemical signals that guide adaptive cellular responses. Aptamers, unlike protein or peptide ligands, offer the unique advantage that their DNA structure can be seamlessly integrated with DRNs to emulate this dynamic regulation. Importantly, aptamers can be manipulated directly at their binding site with DRNs without detaching them from the underlying surface by for instance breaking the force-sensitive duplex, thus ensuring their correct configuration and integrity.

To introduce temporal control over mechanoactivation, we regulate aptamer accessibility using designer blocker and activator strands that can be designed with tunable affinities based on base-pairing principles (Fig. 6a). We first use a blocker fully complementary to AS1411 to completely prevent binding to nucleolin. This blocker also contains a 5-nucleotide (nt) toehold, enabling its subsequent removal via a toehold-mediated strand displacement reaction (SDR) triggered by an activator strand. In this design, mechanoactivation becomes conditional on the possibility for aptamer-receptor engagement, which can be dynamically regulated through SDR. Cell-free surface assays confirm robust and reversible aptamer blocking (Supplementary Fig. 12). The blocker quenches >90% of a fluorescently labeled, surface-tethered AS1411 aptamer within 30 min, indicating effective blocking. Subsequent medium exchange with the equimolar DNA activator restores ~50% of the signal within 30 min with a reaction rate of 2.14×10−2 min−1, confirming aptamer reactivation (see Methods, Supplementary Fig. 12a–d, Supplementary Table 3). We next validate this mechanism under live-cell conditions for two consecutive suppression-recovery cycles. In the first cycle, adding 200 nM blocker 30 min after seeding suppresses AS1411-nucleolin mechanoactivation by 75.1 %. Subsequent medium exchange with 200 nM activator restores 82.5 % of the original signal, consistent with aptamer re-engagement and functional recovery of mechanosensing (Fig. 6b). The second blocker-activator cycle closely mirrors the suppression-recovery behavior observed in the initial cycle (Supplementary Fig. 12e). The overall higher signal level can be attributed to cumulative mechanoactivation accumulated during the additional activation period.

Going a step beyond external switching, we next introduce more advanced enzyme-catalyzed DRNs upstream of mechanoactivation to autonomously program the temporal accessibility of aptamer MPs26,27. To this end, we replace the DNA blocker with an RNA analogue. This RNA blocker can be selectively degraded by RNase H only when bonded as RNA-DNA duplexes at the MP. This degradation releases the aptamer and restores mechanoactivation within programmable time windows (Fig. 6c). In this context, RNase H acts as a biochemical gatekeeper. Additional temporal control can be gained by introducing freely diffusing RNA-DNA duplexes as decoy substrates, which compete with the RNA-blocked MP for RNase H activity.

The kinetics of the timer circuit for MP activation depend on the DRN parameters. First, the RNase H concentration can be used to control the time for mechanoactivation to occur. Monitoring the aptamer reconfiguration kinetics with simple fluorescence assays reveals RNase H concentration-dependent reaction rates of 0.71×10−2 min−1 at 10 U/mL and 5.26×10−2 min−1 at 100 U/mL (see Methods, Supplementary Fig. 13a, Supplementary Table 3). When it comes to force sensing, low RNase H concentrations (≤1 U/ml) fail to initiate measurable mechanoresponse within 4 h. At 10 U/mL, mechanoactivation is restored to 78.2%, while 100 U/mL yields near-complete recovery, consistent with efficient RNA blocker degradation (Fig. 6d, e). To further tune the system, we then fixed RNase H at 10 U/mL and introduced the RNA-DNA decoy (Fig. 6f). The introduction of the decoy strand exerts a clear, dose-dependent inhibitory effect on the reaction rate of RNA blocker degradation (Supplementary Fig. 13b, Supplementary Table 3). Consistent with this trend, 10 nM decoy is sufficient to slightly delay the mechanoresponse, whereas 100 nM nearly abolishes the response within 4 h and a significant response is observed only at 7 h (Fig. 6g, h, Supplementary Fig. 13c).

Because RNA-RNase H modules can be extended into more complex signaling DRNs25, our aptamer manipulation strategy opens doors for more complex temporal programs in the future. We note that similar programmable mechanoactivation could also be achieved using antibody- or nanobody-based probes that block receptor access via engineered protein blockers and activators with progressively tuned affinities. However, such protein-based systems would require extensive efforts, including site-directed mutagenesis, affinity optimization, and computational modeling to fine-tune the underlying protein–protein interactions. In contrast, aptamer systems offer a far more accessible and modular alternative, leveraging predictable base-pairing rules to design dynamic, multistage mechanical response circuits in a straightforward manner.

Methods
Materials
Hydrogen peroxide (35%), sulfuric acid (95%), (3-Aminopropyl)triethoxysilane (APTES), phosphate buffered saline (PBS), and ethanol were purchased from Carl Roth. Biotin-PEG-NHS 5K (HE041023-5K) was purchased from Biopharma. 5(6)-(Biotinamidohexanoylamido) pentylthioureidylfluorescein was purchased from BLDpharm. Cyclo[Arg-Gly-Asp-d-Phe-Lys(PEG-PEG-azide)] (RGD-3759-PI), and Cyclo[Arg-Gly-Asp-D-Phe-Lys(Biotin)](cRGD-Biotin, PCI-3895-PI) were purchased from Biosynth. 25 mm × 75 mm glass coverslips (10812) and sticky slide 8 well (80828) were purchased from ibidi. All oligonucleotides were purchased from Biomers. 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC), 5-(N-ethyl-N-isopropyl)amiloride, chlorpromazine hydrochloride, Methyl-β-cyclodextrin, LY294002, Cytochalasin D, and Blebbistatin were purchased from Sigma-Aldrich. Streptavidin (434302), Alexa Fluor Plus 405 Phalloidin (A30104), CellTrace violet cell profileration kit(C34571), CellTrace far red cell proliferation kit(C34564), bovine albumin fraction V (BSA, 7.5% solution), paraformaldehyde solution (4% in PBS), anti-PTK7 monoclonal antibody (MA5-25774, OTI2E7), Alexa Fluor 488 anti-CD227 (Mucin-1) monoclonal antibody (53-9893-82, SM3), Alexa Fluor 488 anti-CD326 (EpCAM) monoclonal antibody (53-8326-42, MH99), Alexa Fluor 488 goat anti-mouse IgG (H+L) cross-adsorbed secondary antibody (A-11001), Alexa Fluor™ 647 goat anti-mouse IgG (H+L) highly cross-adsorbed secondary antibody (A-21236), Texas Red™ 1,2-Dihexadecanoyl-sn-Glycerin-3-Phosphoethanolamin, Triethylammonium salz (Texas Red™ DHPE), and Oregon Green™ 488 1,2-Dihexadecanoyl-sn-Glycerin-3-Phosphoethanolamin (Oregon Green™ 488 DHPE) were purchased from Thermo Fisher Scientific. N-WASP antibody (C-1, sc-271484) was purchased from Santa Cruz. RNase H was purchased from New England Biolabs. Actin protein (>95% pure, rabbit skeletal muscle) was purchased from Cytoskeleton, Inc. All buffers were prepared with nuclease free water. All chemicals and enzymes were used as such without any further purification.

DNA Hybridization
DNA concentrations were determined using a DeNovix-S-06873 (DeNovix OS 0.8.1 v4.1.5) spectrophotometer. DNA oligonucleotides were hybridized at 10 µM in a TE buffer additionally containing 12.5 mM MgCl2 in 200 µL PCR tubes and subsequently diluted to 125 nM. DNA oligonucleotides were heated to 95 °C and then cooled at a rate of 1.3 °C/min to 25 °C.

Surface preparation
Surface preparation method was modified from previously published protocols21. Briefly, glass coverslips (25 × 75 mm) were rinsed with ultrapure water and sequentially sonicated in water and ethanol for 20 min each. The coverslips were then immersed in freshly prepared piranha solution (3:1 v/v concentrated sulfuric acid (95%) to hydrogen peroxide (30%)) for 1 h. Caution: Piranha solution becomes very hot upon mixing, and is highly oxidizing and may explode upon contact with organic solvents. Then, the coverslips were washed 6 times with ultrapure water, followed by 4 successive ethanol washes. For amine functionalization, the coverslips were incubated in 3% v/v APTES at room temperature for 1 h. The coverslips were then washed 6 times with ethanol, and baked at 80 °C for 20 min. Slides were then reacted with Biotin-PEG-NHS (1% w/v) for 1 h in ultrapure water. Next, the coverslips were washed 3 times with ultrapure water, dried under N2 gas, and stored at −80 °C until use.

The biotinylated coverslips were adhered to sticky-slide 8-well chambers. Wells were incubated with 50 µg/mL of streptavidin for 1 h at room temperature, followed by three washes with nuclease-free water. Wells were then incubated with 200 µL of 250 nM probe solution (125 nM cRGD-biotin and 125 nM DNA probes) for 1 h at room temperature. After washing with nuclease free water, cell imaging media (1 % FBS, 1 % P/S, DMEM; see below) was added to the well followed by the addition of cells.

Probe density calibration
Lipid vesicles. Lipid vesicles were prepared with 100% 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC) or with 99.5 mol% DOPC and 0.5 mol% Texas Red DHPE (TR-DHPE) or Oregon Green™ 488 DHPE (OG-DHPE). Lipid was dissolved in chloroform in a round-bottom glass tube and then carefully dried with a nitrogen stream while rotating the tube. The films were then placed in vacuum for 2 h for complete evaporation of the solvent and stored at −20 °C until further use. Small unilamellar vesicles (SUVs) were formed by sonicating the lipids in nanopure water, with a lipid concentration of 2 mg/mL. The dried lipids were resuspended in Nanopure water at 2 mg/ml by sonicating. To generate small unilamellar vesicles (SUVs), lipids were then extruded 20 times through a mini-extruder (Avanti Research 610000-1EA) assembled with a 0.1 µm polycarbonate membrane.

Solution phase standard curve preparation. To relate yebrightness, the brightness of known concentrations of labeled oligonucleotides and SUVs was measured in solution72. A glass bottom 96-well plate was first washed with ethanol and water and passivated with 1% BSA for 1 h to prevent any surface adsorption. Different concentrations of labeled oligonucleotides and SUVs (0-800 nM) were then added to each well, and their bulk fluorescence intensity in solution was measured with a fluorescence microscope (20x objective lens) to create a standard curve. The scaling factor Ffactor was defined as:

F = S labeledDNA / S labeledSUVs (1)

where S labeledDNA and S labeledSUVs represent the slopes of the plots of fluorescence intensity versus concentration for the labeled DNA and SUVs, respectively.

Supported lipid bilayer preparation. A glass-bottom 96-well plate was used for preparing supported lipid bilayers21,34. Each well was filled and soaked with ethanol for 15 min and rinsed with nuclease free water. Subsequently, 200 µL of 6 M NaOH solution was added to each well for base etching at room temperature for 1 h. After washing each well with Nanopure water, lipid mixtures containing different percentage of labeled SUVs (0, 0.01, 0.05, 0.1, 0.25, 0.5 mol.%) were added at 0.5 mg/mL in PBS with 2.5 mM calcium ions (100 µL) to coat the glass surfaces for 20 min. After the lipid vesicles fused to the surfaces, the wells were rinsed with PBS and imaged with a fluorescence microscope to create a standard curve for probe density. Note that the substrate should never come in contact with air, as this will damage the supported membrane.

Cell culture and transfection
HeLa (ACC 57) and HepG2 (ACC 180) were purchased from DSMZ. A549 (DSMZ, ACC 107) and MDA-MB-231 (DSMZ, ACC 732) were gifts from Tanja Weil laboratory at Max Planck Institute for Polymer Research. HeLa cells were cultured in Minimum Essential Medium (MEM) (10% fetal bovine serum (FBS), 1% penicillin–streptomycin (P/S)) at 37 °C with 5% CO2. A549 and MDA-MB-231 cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) (10% FBS, 1% P/S) at 37 °C with 5% CO2. HepG2 cells were cultured in RPMI 1640 Medium (10% FBS, 1% P/S) at 37 °C with 5% CO2. mEGFP-paxillin WT plasmid was a gift from Michael Rosen (Addgene plasmid #186152)73. Transfection was carried out using the Lipofectamine 3000 Transfection Reagent (Invitrogen, L3000015) following the manufacturer’s instructions. N-WASP knockdown was carried out using human N-WASP siRNA (Santa Cruz, sc-36006) or control siRNA-A (Santa Cruz, sc-37007) with Lipofectamine RNAiMAX Transfection Reagent (Invitrogen, 13778030) following the manufacturer’s instructions.

Force imaging, image acquisition and analysis
All cells were seeded onto the MP surfaces in cell imaging medium (1% FBS, 1% P/S, DMEM) and incubated at 37 °C with 5% CO2 for 4 h, except for Fig. 6h–i, where incubation times were 2, 4, and 7 h. TIRF and brightfield microscopy were performed on a Zeiss Elyra 7 Imaging System equipped with 405 nm, 488 nm, 561 nm, and 642 nm excitation lasers using alpha Plan-Apochromat 63x, N.A. 1.46. oil immersion, TIRF objective, pco.edge 4.2 CLHS water-cooled sCMOS cameras. Live cell imaging was conducted at 37 °C in humidified air with 5 % CO2. Microscopy experiments were typically conducted using the following parameters: 405 nm laser: 0.4% output (~0.01 mW), 488 nm laser: 0.8 % output (~0.7 mW), 561 nm laser: 0.8 % output (~0.8 mW), and 642 nm laser: 0.4% output (~0.4 mW). Beamsplitters (primary, secondary): LBF 405/488/561/642, SBS BP 490-560+LP 640. Exposure time: 50 ms (except for Fig. 2d, RRX-AS1411 200 ms, Atto488-TB-5 200 ms; Fig. 5h, A549-Violet 200 ms; Fig. 5j, HeLa-Deep Red 100 ms, HepG2-Violet 200 ms). TIRF angle: 66° (noil: 1.518). All images were contrasted equally.

Background subtraction methods were adapted from previously published protocols68. Briefly, raw images were exported to FIJI (software) followed by selecting 3 ROIs in the fluorescent channel, next the background intensity was measured and subtracted from the images. Immunofluorescence staining of N-WASP was imaged by confocal laser scanning microscopy (CLSM) using a Leica Stellaris 5 microscope (LasX v4.3.0.24308) with four laser lines and three HyDS detectors using plan-apochromat objectives (63×, 1.40 numerical aperture, oil immersion).

Inhibition experiments
100 µM 5-(N-ethyl-N-isopropyl) amiloride (EIPA), 1.9 mM Methyl-β-cyclodextrin (MβCD), 28 µM chlorpromazine hydrochloride (CPZ) were used to inhibit the macropinocytosis, caveolae, and clathrin-mediated internalization, respectively. 50 µM LY294002 was used to inhibit the PI3K. 10 µM cytochalasin D and 25 µM Blebbistatin were used to inhibit the actin polymerization and myosin ATPase, respectively.

Immunostaining
Cells were fixed with 4% (w/v) paraformaldehyde in PBS for 15 min at room temperature and permeabilized with 0.25% (v/v) Triton X-100 in PBS for 10 min, followed by washes in PBS twice. Unspecific binding was blocked using 10% normal goat serum in PBS for 30 min at room temperature. After removing blocking solution, cells were incubated with anti N-WASP primary antibody (1:25 dilution) at 4 °C overnight. After rinsing three times with PBS, cells were incubated in secondary antibody (1:500 dilution) and Hoechst 34580 (1:200 dilution) in 1% BSA for 1 h at room temperature. For actin staining, fixed cells were incubated with Alexa Fluor Plus 405 Phalloidin (1:200 dilution) in 2.5% (w/v) BSA/PBS at room temperature for 1 h. Samples were then washed three times in PBS for 5 min each before microscopic observations.

Flow cytometry
To assess the competitive binding of endostatin to nucleolin, 5 × 105 cells were incubated with 25 µg/mL endostatin and 2.5 µM FAM-AS1411 in 3% (w/v) BSA/PBS at 4 °C for 60 min. Cells were washed and then resuspended in PBS before running through flow cytometer. To quantify the receptor expression levels, 5 × 105 cells were labelled with fluorescent primary antibodies (Alexa Fluor™ 488-anti-MUC1: 5 µg/mL, Alexa Fluor™ 488-anti-EpCAM: 5 µg/mL) in 3% (w/v) BSA/PBS at 4 °C for 30 min. For labelling PTK7, cells were incubated with primary antibody anti-PTK7 (2 µg/mL) in 3% (w/v) BSA/PBS at 4 °C for 30 min. Cells were washed three times with PBS and then incubated in Alexa Fluor™ 488-secondary antibody (5 µg/mL) in 3% (w/v) BSA/PBS at 37 °C for 30 min. After labelling, cells were washed and resuspended in PBS. To assess the effect of EIPA on AS1411 internalization, 2 × 105 cells were seeded in standard cell medium in 12-well plates and incubated overnight at 37 °C. Cells were then incubated with 100 µM EIPA and 500 nM FAM-AS1411 in 1% FBS/DMEM at 37 °C for 4 h. Cells were washed and then resuspended in PBS before running through flow cytometer.

Flow cytometry experiments were performed on a Novocyte Quanteon (Agilent, NovoExpress v.1.6.0) with 4 excitation lasers (violet 405 nm, blue 488 nm, yellow-green 561 nm, and red 640 nm) and 16 fluorescence detectors. Gating strategy is shown in Supplementary Fig. 3a. A total of 10,000 events were collected per sample. Flow cytometry data were analyzed using FLOWJO (v10.8.1) software.

Statistical analysis
Each experiment was performed at least three independent times. No statistical method was used to predetermine sample size. No data were excluded from the analyses. The experiments were not randomized. The Investigators were not blinded to allocation during experiments and outcome assessment. Statistical results were analyzed using GraphPad Prism 8.0 and shown as the mean ± s.d. Comparisons between two groups were conducted using unpaired two-tailed Student’s t-test (Figs. 4m–o, 5k), and among three or more groups using one-way ANOVA with Bonferroni test correction (Figs. 4h, 6b) as indicated. If a data set did not pass the normality tests, the significances were calculated with Mann-Whitney (Figs. 2e, 3g, 3i, 5b, d, i and 6b) for two-group comparisons or Kruskal-Wallis corrected with Dunn’s test (Figs. 2b–c, 3c–e, 4b, 4f, and 5f) for multiple groups. For all analyses, *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001 were considered significant. P>0.05 was considered statistical non-significance.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This is an elegant study, the authors used aptamers to replace traditional protein/peptide ligands conjugated to DNA force probes. This allowed them to target proteins beyond classical mechanoreceptors with existing aptamers. Additionally, the construction of the aptamer DNA force probes became significantly simplified, requiring no conjugation to proteins. This advancement broadens the receptors that can be targeted by force probes and giving researchers the ability to control the interactions using established DNA reactions.\nSpecifically, the study showed several examples applications. First, they targeted nucleolin to establish the concept with both unzipping and shearing DNA geometries. They showed that both Blebbistatin and Cytochalasin D reduced the nucleolin-AS1411 tension signal. The authors elucidated the tension mechanisms using different drugs (CPZ, MbCD, and EIPA) and showed specific signal reduction due to EPIA but not the other 2, linking the source of tension to macropinocytosis. Additionally, the demonstration of ring-like structures using PTK7 force probes further showed PTK7 tension signal is directly related to macropinocytosis. The authors showed that different cells with and without certain receptor expression can be detected by the presence and absence of aptamer tension signals. Lastly, they demonstrated using various reaction schemes (e.g., DNA strand displacement, RNaseH) to reversibly activate and deactivate the aptamer force probes.\nOverall, the paper did an excellent job demonstrating the utility and advantages of aptamer in DNA tension probes. Both the text and figure are clearly presented, I believe this is an important step in the evolution of DNA-based force probes.\nI have a few comments that I hope the authors could address to strengthen the manuscript. Two major comments first:\n1. An important aspect of the manuscript is to enable studying tension from non-canonical mechanoreceptors. While I believe this is of significant interest, the authors should elaborate on how investigations using non-native interactions (i.e. with aptamers) yield biological relevance for native interactions. For instance, if a receptor’s native ligand binds weakly and only allow low force transmission, a high affinity aptamer would report high force transmission, particularly if the receptor is tightly associated with… say, the focal adhesion complex. The tension signal here would report the capacity of tension generation, rather than the receptor’s functional tension with its native ligands. I think this distinction could be discussed in more detail, and claims related to the utilities using aptamers could be tightened accordingly. That said, I also I very much appreciate the accuracy of statements such as “the AS1411‐nucleolin couple transmits substantially lower force than RGD‐integrin engagement.”, which highlights the context of the comparison is with the specific interaction pairs.\n2. The data on cell-type specific mechanoresponses is clean. But it is also kind of obvious that cells with and without receptors would and wouldn’t produce tension signals. By testing simple presence or absence, this section seems more like a control for aptamer specificity. The wording of claim for being able to discern “cell-type specific mechanoresponse” would suggest that given the same existing mechanoreceptor on different cell types, one can differentiate tension signals (e.g. magnitude, spatial patterns, etc) as a result of their different cellular physiology. The “mechano-response” should produce additional details that examining receptor expression level alone (e.g. with immunofluorescence with FC) cannot reveal. I would strongly encourage an additional experiment with 2 cell types that both express say, PTK7, but show that they produce different tension signal – that would be very nice! In the absence of such experiment, the claim would be more limited, and the text should be tightened accordingly.\nMinor comments:\n3. Could the authors elaborate why in the co-culture settings (Fig 5i,k), the differences between the mean RRX signals aren’t as tight as in separate monoculture (Fig 5b,d)? Additionally, could you also elaborate on the variability in RRX intensity? Esp for cells that do not express the receptor, some show significant tension signal (Fig 5k) in comparison to the MUC1(+) cells. The authors should also refer to their flow cytometry data in Supp. Fig 9 in this discussion (maybe I missed it).\n4. The manuscript presents mean and integrated RRX intensity in various places, how should we interpret the difference between them (e.g. Fig 3c,d)? And why is integrated intensity included in some places but not others (Fig 5b,d,f)? If it’s not adding useful information, I would suggest removing integrated intensity from Fig 5b,d,f, to keep consistency throughout.\n5. Nucleases in the live-cell media (either from serum or cell-release) have been problematic for long-term studies using DNA-based probes. Have the authors observed any problems with DNA degradation and non-specific fluorescence signals in this system? I think a short discussion on protecting native aptamers from such degradation could be useful to elevate its impact. There have been several strategies using PNA (https://www.sciencedirect.com/science/article/pii/S095656631931036X), your L-DNA work (https://onlinelibrary.wiley.com/doi/10.1002/anie.202413983), PS mod (https://pubs.acs.org/doi/10.1021/acsami.3c04826), and probably more relevant for protecting unmodified nucleic acid suitable for aptamers with a decoy DNA (https://onlinelibrary.wiley.com/doi/10.1002/anie.202506590).\n6. Maybe I missed it, could the authors indicate the time frame in the RHaseH study (Fig 6e,f)? Could you also comment on the reaction rate using RHaseH or strand-displacement (using a reasonable amount of reagent)? Which one is faster?\n7. There are some existing work on using strand displacement to control junctional tension such as (https://pubs.acs.org/doi/10.1021/acsabm.4c00142) I think a short discussion of existing literature on force mediation/modulation using DNA reactions could strengthen the manuscript and help highlight differences and your advantages – modulating function without destroying the probe.", "reviewer_2": "In this manuscript, Xu et al. present a modular all-DNA aptamer-based mechanosensing platform. The authors integrate receptor-specific recognition via aptamers with the tunable force sensitivity of DNA mechanoprobes. A key strength of the study is the demonstration of dual selectivity, discriminating cellular forces based on both cell type and discrete force thresholds. The investigation of non-canonical mechanoreceptors such as nucleolin and PTK7 provides valuable new insights into how cells mechanically interact with their microenvironment. Moreover, the integration of the platform with DNA reaction networks to achieve programmable temporal control represents a significant methodological advance for synthetic mechanobiology. Overall, this work is well executed and presents a notable contribution to the field. However, several mechanistic claims and aspects of validation should be further substantiated to strengthen the rigor and overall impact of the work. Therefore, I recommend minor revision prior to acceptance for publication in Nature Communications.\nMajor concerns and suggestions\n1.The claim that nucleolin-mediated forces are “relayed via focal adhesions” is central to the mechanistic interpretation. The current evidence—paxillin colocalization and sensitivity to myosin II inhibition—supports spatial and functional correlation but not direct physical linkage. An alternative explanation, that nucleolin and integrins are independently coupled to the same actomyosin network, cannot be excluded. It is recommended that the authors either moderate this claim or propose additional experiments (e.g., FRET, co-immunoprecipitation) to establish a direct link.\n2.In Figure 4, the mechanistic investigation relies exclusively on small-molecule inhibitors, which are prone to off-target effects. The distinction between myosin-driven (nucleolin) and actin-polymerization-driven (PTK7) forces would be more convincing if validated with orthogonal genetic perturbations, such as shRNA knockdown of N-WASP.\n3.A critical control is missing to validate the claim that the aptamer-based force sensing is functionally independent of the RGD-mediated adhesion used throughout the study. To robustly prove this independence, the assay should also be performed on a surface functionalized with the aptamer MPs alone (i.e., without co-immobilized RGD). This would directly test whether baseline integrin adhesion is a prerequisite for the aptamer probes to function, making the conclusion in Figure 2c more convincing.\n4.The actual surface density and stoichiometry of the immobilized RGD and MP molecules were not quantified. These parameters are known to significantly influence cell adhesion and force generation. Providing this characterization would greatly enhance the rigor and reproducibility of the platform.\n5.The SDR module is currently demonstrated with only a single block–activate cycle, which is not sufficient to establish reversibility. It is recommended that the authors present at least two or more consecutive suppression–recovery cycles. In addition, the RNase H-based “timer” shows tunable activation rates but not a clearly defined lag phase. Including earlier and more frequent time points would help demonstrate a controllable delay before signal onset, and a simple kinetic model correlating decoy concentration with delay would further substantiate the claim of programmability.\n6.The statement that aptamer MPs exhibit “enhanced force selectivity” compared to RGD MPs may be misleading. A more accurate interpretation is that AS1411–nucleolin interactions occur within a lower physiological force regime (consistently below 54 pN), while RGD–integrin forces readily exceed this threshold. This distinction reflects biological force domains rather than an intrinsic property of the probe.\n7.Regarding the citation of Yang et al. (Ref. 20), the manuscript currently describes this study as “force being sensed using a DNA MP”, which may not fully reflect its contribution. Specifically, Yang et al. provided the proof-of-concept that a DNA aptamer, taking Cl-M6RP-targeting aptamer as the example, can function as a force-reception module to sense weak endocytic forces with fluorescent signal readout and then translate these force cues into intracellular signals via DNA dynamic reactions. Thus, a more balanced description should acknowledge this advance in the revision, and it is recommended to add a brief discussion clarifying how the present work extends this concept.\nMinors\n1.Use more precise terminology such as “receptor-low” or “low-expressing” instead of “receptor-negative,” as the supplementary flow cytometry data indicate non-zero expression.\n2.Clarify in the text or figure legend for Figure 3 why both “mean” and “integrated” intensity are presented, and how each metric provides distinct insights.\n3.In Figure 6, the restored force signal distribution appears inconsistent across conditions (e.g., recovery mainly at the cell edge vs. both edge and center). The authors should clarify whether these differences reflect mechanistic variations or are attributable to experimental variability.\n4.For PTK7-mediated forces, it remains unclear whether the signal reflects protrusive pushing or contractile pulling. The schematic suggests pulling, whereas the text emphasizes polymerization-driven pushing. This ambiguity should be explicitly acknowledged and discussed.\n5.In the discussion, briefly acknowledge the limitations of DRN-controlled systems for potential in vivo applications, including delivery challenges, nuclease stability, and cytotoxicity.\n6.While the manuscript identifies novel force pathways, the discussion could be enriched by addressing their biological significance. Why do cells exert forces through these non-canonical receptors? Furthermore, since detailed analysis was limited to nucleolin and PTK7, it would be valuable to comment on the potential generalizability to other possible aptamer–receptor pairs with biological functions."}, "author_response": "Thank you for your very positive evaluation of our work.\nComment 1-1. An important aspect of the manuscript is to enable studying tension from non-canonical mechanoreceptors. While I believe this is of significant interest, the authors should elaborate on how investigations using non-native interactions (i.e. with aptamers) yield biological relevance for native interactions. For instance, if a receptor’s native ligand binds weakly and only allow low force transmission, a high affinity aptamer would report high force transmission, particularly if the receptor is tightly associated with… say, the focal adhesion complex. The tension signal here would report the capacity of tension generation, rather than the receptor’s functional tension with its native ligands. I think this distinction could be discussed in more detail, and claims related to the utilities using aptamers could be tightened accordingly. That said, I also I very much appreciate the accuracy of statements such as “the AS1411-nucleolin couple transmits substantially lower force than RGD-integrin engagement.”, which highlights the context of the comparison is with the specific interaction pairs.\nResponse: We thank the reviewer for this insightful comment. While aptamer-based platforms represent a promising and accessible tool for the pre-screening of non-canonical mechanoreceptors, we fully agree that extrapolating mechanoevents detected using aptamer-receptor pairs to native ligand-receptor interactions must be done with caution. Several key factors require careful consideration:\n1. Immobilization state of the ligand is critical, as it determines the magnitude of the reaction force that can be supported. In our system, aptamer probes are immobilized via biotin-streptavidin interaction, which provides sufficient reaction force to resist cellular pulling. Mobile or weakly immobilized ligands may lead to inefficient force transmission.\n2. Binding affinity and bond mechanics. As the reviewer noted, differences in binding affinity may affect force transmission. Overly stable aptamer-receptor interactions may lead to an overestimation of the force transmitted under native conditions. In addition, while aptamer-receptor interactions typically exhibit slip bond behavior, native ligand-receptor pairs, especially classical mechanoreceptors such as RGD-integrin, pMHC-TCR, etc., may involve complex catch bonds. This difference in bond lifetime under load may significantly change the mechanical behaviors.\n3. Multivalency and cooperative effects. Synthetic aptamers often lack the multi-domain architecture of native ligands (e.g., RGD and LDV motifs present in fibronectin).1 These differences may lead to variations in inside-out force transmission, potentially influencing unknown outside-in mechanotransduction pathways and thus affecting result interpretation. These limitations are inherent to studies using simplified synthetic ligands.\nRegarding the effect of binding affinity as the reviewer mentioned, the AS1411 aptamer exhibits a Kd of 69.1 nM for nucleolin, whereas its native ligand endostatin possesses a Kd of 23.2 nM. Since the endostatin's affinity is approximately 3-fold higher, it is highly possible that our platform underestimates the native capacity for tension generation. Despite this, we still observe the mechanosignals, suggesting such affinity is sufficient for partial force transmission. The ability of nucleolin to transmit force is indicated by existing literature: (i) Nucleolin mediates the internalization of endostatin in endothelial cells by forming a receptor complex with integrin α5β1,2,3 a process we consider a mechanoevent, which cell survival and proliferation. (ii) Nucleolin is known to associate with cytoskeletal components, including integrins,2,4,5 focal adhesion proteins,6 and MYH9,7 which provides a structural basis for its role in force transmission. Notably, endostatin is derived from the C-terminal non-collagenous (NC1) domain of type XVIII collagen in the extracellular matrix and is generated by MMP-mediated cleavage.8 Whether nucleolin can transmit force through this precursor protein in the native ECM context, and whether endostatin-associated biological functions are linked to the non-canonical mechanical properties of nucleolin, remain important questions for future investigation. Addressing these questions may reveal how cryptic motifs within the ECM regulate cellular homeostasis through non-canonical mechanotransduction pathways.\nConversely, as the reviewer noted, if an aptamer possesses a higher affinity than the native ligand, the resulting over-stabilized mechanical linkage might overestimate natural tension capacity. In such cases, beyond serving as a detection probe, the aptamer MPs may be better viewed as functional actuators. This relative independence and orthogonality make it well suited for integration with DNA nanotechnology and artificial extracellular matrix materials for tissue engineering purposes to construct artificial signaling pathways for controlled modulation of cellular behaviors.\nWe have revised the related discussion in the text:\nThe identification of noncanonical forces highlight the importance of revisiting the biological functions of both these aptamers and receptors from a mechanobiological perspective. Determining whether these receptors are directly coupled to classical adhesion structures or are independently linked to the same actomyosin network via intermediary components will be essential for uncovering more general principles. Whether similar interactions occur under native conditions remains to be elucidated. A prime candidate is the precursor of the nucleolin ligand endostatin, located in the NC1 domain of type XVIII collagen,60 which may serve as a native anchor for nucleolin despite potential differences in binding affinity. Addressing these questions could reveal how cryptic motifs within the ECM regulate cellular homeostasis through non-canonical mechanotransduction pathways. Meanwhile, caution is required when extrapolating from aptamer-receptor pairs to native ligand-receptor interaction, particularly regarding the immobilization state of the ligand, binding affinity and bond mechanics, and the multivalency and cooperative effects inherent to native ligands. For example, the AS1411 aptamer binds nucleolin with a Kd of 69.1 nM,28 which is lower than that of its native ligand endostatin (23.2 nM),61 suggesting that our measurements may slightly underestimate the native capacity for tension generation.\nComment 1-2. The data on cell-type specific mechanoresponses is clean. But it is also kind of obvious that cells with and without receptors would and wouldn’t produce tension signals. By testing simple presence or absence, this section seems more like a control for aptamer specificity. The wording of claim for being able to discern “cell-type specific mechanoresponse” would suggest that given the same existing mechanoreceptor on different cell types, one can differentiate tension signals (e.g. magnitude, spatial patterns, etc) as a result of their different cellular physiology. The “mechano-response” should produce additional details that examining receptor expression level alone (e.g. with immunofluorescence with FC) cannot reveal. I would strongly encourage an additional experiment with 2 cell types that both express say, PTK7, but show that they produce different tension signal – that would be very nice! In the absence of such experiment, the claim would be more limited, and the text should be tightened accordingly.\nResponse: Thanks for the suggestion. We indeed identified two different cell lines that both express the same receptor but exhibit different force patterns. Compared with PTK7 high-expressing HepG2 and PTK7 low-expressing A549 cells, flow cytometry data demonstrate that HeLa cells display an intermediate expression level of PTK7 (Supplementary Fig. 11e,f). Interestingly, mechanosensing using Sgc8 MPs reveals not only an expression-level-dependent signal magnitude but also different force patterns. Unlike the characteristic ring-like mechanosignals observed in HepG2 cells, HeLa cells exhibit a stripe-like force pattern (Fig. 5e,f). These results indicate that even when the same receptor is expressed, different cell types may transmit forces through that receptor via distinct pathways, and thus leads to different mechanoresponses in both signal magnitude and force patterns. We added this new data to Fig. 5e,f and Supplementary Fig. 11e,f and discussed it in the text.\nFig. 5 e Representative brightfield and fluorescence images of HepG2 (PTK7-high-expressing), HeLa (PTK7-intermediate-expressing) and A549 (PTK7-low-expressing) cells on Sgc8 MP surfaces. f Corresponding quantification of mean and integrated fluorescence intensity per cell of HepG2, HeLa and A549 cells on Sgc8 MP surfaces. n = 36, 31, 27 cells from 3 replicates. Statistics: Kruskal-Wallis test with Dunn’s multiple comparisons.\nSupplementary Figure 11. e Representative flow cytometry histogram of HepG2, HeLa, and A549 cells stained with anti-PTK7 primary antibody followed by Alexa Fluor™ 488-conjugated secondary antibody. Mock controls represent unstained cells, indicating cellular autofluorescence. f Quantification of PTK7 expression levels on HepG2, HeLa, and A549 cells based on median fluorescence intensity. n = 3 independent experiments. Statistics: One-way ANOVA with Bonferroni post-hoc tests.\nA discussion for this result has been added to the main text:\nInterestingly, beyond expression level-dependent signal magnitude, different cell types expressing the same receptor can exhibit distinct, cell-type-dependent force patterns. HeLa cells express PTK7 at an intermediate level (Supplementary Fig. 11e,f). Mechanosensing using Sgc8 MPs reveals a stripe-like force pattern in contrast to the characteristic ring-like mechanosignals observed in HepG2 cells (Fig. 5e, f). These results indicate that different cell types may transmit forces through the same receptor via distinct pathways, a phenomenon that cannot be inferred from receptor expression levels alone.\nMinor comments:\nComment 1-3. Could the authors elaborate why in the co-culture settings (Fig 5i,k), the differences between the mean RRX signals aren’t as tight as in separate monoculture (Fig 5b,d)? Additionally, could you also elaborate on the variability in RRX intensity? Esp for cells that do not express the receptor, some show significant tension signal (Fig 5k) in comparison to the MUC1(+) cells. The authors should also refer to their flow cytometry data in Supp. Fig 9 in this discussion (maybe I missed it).\nResponse: Thank you for the question. We are sorry that our original terminology, “receptor-negative” and “receptor-positive”, are not precise enough. As noted by Reviewer#2 (question 2-8), and shown in Supplementary Fig. 11 (original Supp. Fig 9), A549 and HepG2 cells do express basal levels of PTK7 and MUC1 receptors, respectively, meaning they are not truly “negative” cell lines. The mechanosignal distribution is strongly influenced by the overlap in receptor expression level between these populations. Specifically, the higher mechanosignals observed in “low-expressing” groups may come from those few cells that happen to have a high expression of receptors. Consequently, the breadth of the signal distribution depends on whether these overlapped cells are captured during sampling. This makes the results sensitive to sample size, and batch-to-batch variations in expression. While the transition from monoculture to co-culture conditions may have some effect, we do not wish to speculate excessively on this behavior beyond what is directly supported by the data. To reflect this, we have updated our terminology to “low-expressing” and “high-expressing” throughout the manuscript. We also discussed the original Supp. Fig 9 (now Supplementary Fig. 11) in the text.\nComment 1-4. The manuscript presents mean and integrated RRX intensity in various places, how should we interpret the difference between them (e.g. Fig 3c,d)? And why is integrated intensity included in some places but not others (Fig 5b,d,f)? If it’s not adding useful information, I would suggest removing integrated intensity from Fig 5b,d,f, to keep consistency throughout.\nResponse: Thank you for the question. Our analysis follows the well-established methodology for RGD-based MPs developed by the Salaita group.9,10 We distinguish between mean intensity, which represents the average mechanosignal per unit area within the cell footprint, and integrated intensity, which reflects the cumulative mechanosignal across the entire footprint and is thus inherently dependent on cell area.11\nUnder idealized conditions where cell spreading remains invariant across different probe surfaces, the trends for mean and integrated intensities should converge: higher tension tolerance Ttol probes offer greater resistance to rupture, theoretically yielding lower values for both metrics. Previous literatures confirm that mean intensity captures these Ttol-dependent shifts.11 Cells on 12 pN RGD-MP surfaces typically exhibit robust, uniform rupture, whereas cells on 54 pN RGD-MP surfaces yield a streak pattern, resulting in a lower mean intensity.\nHowever, the magnitude of integrated intensity is often inconsistent across studies because it is cross-regulated by both the intrinsic Ttol of the probe and the cell spreading area.10,11 The later is highly sensitive to surface chemistry or probe geometries themselves. In the absence of exogenous adhesion enhancers, cells on 12 pN RGD-MP surfaces often fail to achieve adhesion maturation as the reaction force required typically exceeds 43 pN. This results in restricted cell area, in which the integrated intensity may paradoxically appear lower than that of cells on 54 pN surfaces. Conversely, when spreading is promoted by adhesion enhancers (e.g., fibronectin, poly-L-lysine) or mixed-Ttol surfaces, cells on 12 pN surfaces exhibit greater sensitivity to these cues to increase cell area, while 54 pN surfaces inherently provide sufficient mechanical resistance to support adhesion. As a result, when cells are equally well-spread on both surfaces, the 12 pN surfaces yield a higher integrated fluorescence, reflecting the lower threshold for probe rupture.\nGiven these complexities, we believe it is essential to present integrated intensity whenever variations in cell area cannot be neglected. This is particularly relevant when comparing different probes that may alter spreading behavior (Fig. 3c, d) or when examining different cell lines with distinct basal morphologies (Fig. 5b, d, f), such as the smaller MDA-MB-231 cells versus the larger HeLa cells. Furthermore, the integrated intensity provides the most accurate representation of the total stimulatory input per individual cell, which is the primary driver to design downstream reactions.\nA discussion for this result has been added to the main text:\nThe integrated intensity reflects the cumulative mechanosignal across the entire cell footprint and is thus not only regulated by intrinsic Ttol of the probe but inherently dependent on cell area.19\nIn addition, this discussion provided now in the answer letter (that is available to readers in Nat. Commun.) provides additional clarification.\nWe also updated the figure captions:\nFig.3 c-e Quantification of (c) mean fluorescence intensity per cell, (d) integrated fluorescence intensity per cell, and (e) cell spreading area for HeLa cells on each MP type and geometry. n = 49, 47, 45, 36 (left to right) cells from 3 replicates. Statistics: Kruskal-Wallis test with Dunn’s multiple comparisons. Note that both mean and integrated intensity are shown because mean intensity mainly captures the Ttol-dependent shifts while integrated intensity is cross-regulated by both the intrinsic Ttol of the probe and the cell spreading area.\nFig.5 b Corresponding quantification of mean and integrated fluorescence intensity per cell of HeLa and HepG2 cells on MUC1 S2.2 MP surfaces. n = 31, 27 cells from 3 replicates. Statistics: Two-tailed, Mann-Whitney test. Note that both mean and integrated intensity are shown here because different cell lines may have distinct basal morphologies with different cell area.\nComment 1-5. Nucleases in the live-cell media (either from serum or cell-release) have been problematic for long-term studies using DNA-based probes. Have the authors observed any problems with DNA degradation and non-specific fluorescence signals in this system? I think a short discussion on protecting native aptamers from such degradation could be useful to elevate its impact. There have been several strategies using PNA (https://www.sciencedirect.com/science/article/pii/S095656631931036X), your L-DNA work (https://onlinelibrary.wiley.com/doi/10.1002/anie.202413983), PS mod (https://pubs.acs.org/doi/10.1021/acsami.3c04826), and probably more relevant for protecting unmodified nucleic acid suitable for aptamers with a decoy DNA (https://onlinelibrary.wiley.com/doi/10.1002/anie.202506590).\nResponse: Thank you for the question and providing these useful papers for reference. In our study, we adopted a protocol like those reported in the literature,11,12 using low-serum media (1% FBS/DMEM) to mitigate the effects of exogenous nucleases. This approach with appropriate background subtraction provides reliable results during short-term observations. Regarding endogenous nucleases secreted by cells, Fig. 2b demonstrates that the signal from the “null probe” accounts for both nuclease-mediated degradation and non-specific interactions (such as dye-membrane interactions). Compared to the robust mechanosignals observed for AS1411 MPs, these nonspecific signal levels remain within an acceptable range.\nBut we fully agree that additional nucleic acid protection strategies are essential for long-term practical applications. In our RNA-RNase H kinetic assays extending up to 7 h, we introduced actin as a competitive nuclease inhibitor to preserve the structural integrity of DNA MPs.13 While actin offers effective protection, its high cost limits broader application. Moving forward, we believe that integrating a combination of the recommended strategies will significantly enhance the stability of aptamer-based platforms. Regarding the duplex-based force probes, incorporating peptide nucleic acid or phosphorothioate modifications into the double-stranded regions or directly using L-DNA could indeed provide resistance against nuclease-mediated degradation. But the situation differs for single-stranded aptamers, which are inherently more susceptible to degradation. Any chemical modification intended to increase stability must be carefully balanced against potential losses in targeting affinity. In this context, we believe that employing decoy DNA is a better solution, as it provides an optimal balance between protective efficacy and cost-effectiveness. Please also note that the sequence-to-force relationships (well established for pristine D-DNA and L-DNA) would also need to be re-established by single molecule force measurements when applying modifications such as PDNA or locked nucleic acids.\nWe cited these references and discussed in the main text:\nHowever, long-term application in these environments still faces stability challenges. While modifications like peptide nucleic acids,63 phosphorothioate linkages,64 or our recent L-DNA strategy protect the MP structure,65 single-stranded aptamers remain inherently degradation-prone. Balancing their structural integrity with binding affinity is critical. In this context, using nuclease inhibitors such as actin or decoy DNA may be more effective.66,67\nComment 1-6. Maybe I missed it, could the authors indicate the time frame in the RHaseH study (Fig 6e,f)? Could you also comment on the reaction rate using RHaseH or strand-displacement (using a reasonable amount of reagent)? Which one is faster?\nResponse: Thank you for the question. (1) Time frame: We immobilized the RNA blocker-locked AS1411 probes on the surface, seeded the HeLa cells and added RNase H at varying concentration (0-500 U/mL) simultaneously. Cells were imaged after 4 h of incubation. We updated these experimental details to the figure caption.\n(2) Speed of reactions (new experiments): We monitored the reaction kinetic of strand displacement reaction (SDR) and RNA-RNase H modules to assess reaction rates. A FRET pair (Atto488/BHQ1) was incorporated into both the AS1411 strand and the DNA/RNA blocker so that we can quantify the ratio of AS1411 aptamer reconfiguration. We selected RNase H concentrations of 10 U/mL and 100 U/mL for comparison with SDR using 200 nM DNA activator, as no obvious signal recovery was observed at lower RNase H concentration, whereas signal saturation occurred at higher RNase H concentration (Fig.6e). The reaction rate of SDR with 200 nM DNA activator (2.14×10-2 min-1) lies between those observed for RNase H at 10 U/mL (0.71 ×10-2 min-1) and 100 U/mL (5.26 ×10-2 min-1). We added this new data to Supplementary Fig. 12d, Supplementary Fig. 13a, and Supplementary Table 3, and discussed it in the text.\nCombined representation of Supplementary Figure 12d and 13a. Plot of normalized surface atto488 intensity versus time to compare the reconfiguration rate of AS1411 aptamer in SDR and RNA-RNAse H modules. For the SDR module, surface-immobilized AS1411 MPs are initially blocked with DNA blocker and subsequently reconfigured by adding 200 nM DNA activator. For the RNA-RNAse H module, surface-immobilized AS1411 MPs are blocked with RNA blocker and degraded by RNase H at different concentrations (10 or 100 U/mL). At each reaction time point, the surface Atto488 intensity is normalized to that of unblocked AS1411 MPs labeled with atto488, which serves as the positive control. Reaction medium: 1 % FBS, 1 % P/S, 100 µg/mL actin protein, DMEM. n = 15 from 3 replicates.\nSupplementary Table 3. Reaction rate constant of SDR and RNA-RNase H modules. To understand the reconfiguration efficiency and tunability of different DNRs, we quantified the surface reconfiguration kinetics. The data were analyzed assuming pseudo-first-order kinetics and fitted using one phase exponential decay function. For the SDR module, reactivation with 200 nM DNA activator yields a reconfiguration rate of 2.14 × 10⁻² min⁻¹. The RNA-RNase H module enables degradation rate tuning by adjusting the initial RNase H concentration, resulting in accelerated (100 U/mL, 5.26 × 10⁻² min⁻¹) or decelerated (10 U/mL, 0.71 × 10⁻² min⁻¹) reconfiguration kinetics. Moreover, at a fixed RNase H concentration (10 U/mL), the RNA-RNase H module allows the introduction of non-enzymatic RNA/DNA decoys, providing an additional orthogonal layer of regulation.\nk (×10-2 min-1) R2\nSDR 200 nM DNA activator 2.14 0.9926\nRNase H 100U/mL 5.26 0.9925\nRNase H 10U/mL 0.71 0.9842\nRNase H 10U/mL+1 nM rdm RNA/DNA decoy 0.73 0.9782\nRNase H 10U/mL+10 nM rdm RNA/DNA decoy 0.42 0.9603\nRNase H 10U/mL+100 nM rdm RNA/DNA decoy 0.11 0.9831\nA discussion for this result has been added to the main text:\nSubsequent medium exchange with the equimolar DNA activator restores ~ 50 % of the signal within 30 min with a reaction rate of 2.14×10-2 min-1, confirming aptamer reactivation (see Methods, Supplementary Fig. 12a-d, Supplementary Table 3).\nMonitoring the aptamer reconfiguration kinetics with simple fluorescence assays reveals RNase H concentration-dependent reaction rates of 0.71 ×10-2 min-1 at 10 U/mL and 5.26 ×10-2 min-1 at 100 U/mL (see Methods, Supplementary Fig. 13a, Supplementary Table 3).\nComment 1-7. There are some existing work on using strand displacement to control junctional tension such as (https://pubs.acs.org/doi/10.1021/acsabm.4c00142) I think a short discussion of existing literature on force mediation/modulation using DNA reactions could strengthen the manuscript and help highlight differences and your advantages – modulating function without destroying the probe.\nResponse: Thanks for the suggestion of more references. A discussion for has been added to the main text:\nMoving forward, pioneering work using protein-ligand functionalized MPs has shown that strand displacement can control intracellular mechanics in 3D spheroids,62 although at the cost of ligand release. Introducing aptamer-based MPs further enables non-invasive, tunable modulation without compromising probe integrity.\nReviewer #2 (Remarks to the Author)\nIn this manuscript, Xu et al. present a modular all-DNA aptamer-based mechanosensing platform. The authors integrate receptor-specific recognition via aptamers with the tunable force sensitivity of DNA mechanoprobes. A key strength of the study is the demonstration of dual selectivity, discriminating cellular forces based on both cell type and discrete force thresholds. The investigation of non-canonical mechanoreceptors such as nucleolin and PTK7 provides valuable new insights into how cells mechanically interact with their microenvironment. Moreover, the integration of the platform with DNA reaction networks to achieve programmable temporal control represents a significant methodological advance for synthetic mechanobiology. Overall, this work is well executed and presents a notable contribution to the field. However, several mechanistic claims and aspects of validation should be further substantiated to strengthen the rigor and overall impact of the work. Therefore, I recommend minor revision prior to acceptance for publication in Nature Communications.\nThank you for your very positive evaluation of our work.\nMajor concerns and suggestions\nComment 2-1. The claim that nucleolin-mediated forces are “relayed via focal adhesions” is central to the mechanistic interpretation. The current evidence—paxillin colocalization and sensitivity to myosin II inhibition—supports spatial and functional correlation but not direct physical linkage. An alternative explanation, that nucleolin and integrins are independently coupled to the same actomyosin network, cannot be excluded. It is recommended that the authors either moderate this claim or propose additional experiments (e.g., FRET, co-immunoprecipitation) to establish a direct link.\nResponse: We thank the reviewer for this insightful suggestion. We agree that establishing a direct physical link between nucleolin and focal adhesions is essential for a complete molecular picture. However, fully characterizing these protein-protein interactions requires a substantial body of work, which we consider to rather warrant dedicated follow up studies.\nPrevious studies have reported interactions between nucleolin and multiple focal adhesion-related or cytoskeletal components, including integrins αvβ3,4 α5β1, 2and β1,5 the focal adhesion protein talin,6 and MYH9,7 as identified by Co-IP or Co-IP/MS. These findings indicate that nucleolin and focal adhesion-associated proteins can coexist within the same molecular complexes; however, they also do not exclude the presence of intermediary linker proteins that may independently connect nucleolin to the complexes or actomyosin network.\nAs suggested by the reviewer, definitive validation of direct interactions would require approaches such as in situ intermolecular FRET or in vitro pull-down assays using purified proteins. However, both methods present significant challenges: (i) FRET require extensive protein engineering to generate fluorescent fusion constructs, which is technically demanding, and may sterically hinder the native interaction or alter the localization of nucleolin. (ii) In vitro pull-down using purified proteins cannot fully recapitulate the post-translational modifications or the specific membrane microenvironment that may be critical for nucleolin’s binding affinity in live cells. This is critical as nucleolin exists in both cytoplasmic and membrane-bound forms, which may exhibit distinct binding affinities for the same partner protein.\nGiven the complexity of high-throughput screening and the rigorous requirements of in situ validation, we cannot provide a definitive conclusion within the current revision timeframe. We have therefore softened our claim and acknowledged this limitation in the Discussion Section:\nDetermining whether these receptors are directly coupled to classical adhesion structures or are independently linked to the same actomyosin network via intermediary components will be essential for uncovering more general principles.\nComment 2-2. In Figure 4, the mechanistic investigation relies exclusively on small-molecule inhibitors, which are prone to off-target effects. The distinction between myosin-driven (nucleolin) and actin-polymerization-driven (PTK7) forces would be more convincing if validated with orthogonal genetic perturbations, such as shRNA knockdown of N-WASP.\nResponse: Thanks for this insightful suggestion. We used a commercially available N-WASP (human) siRNA and successfully knocked down intracellular N-WASP in both HeLa cells (nucleolin high-expressing) and HepG2 cells (PTK7 high-expressing). After detachment, these cells were seeded onto AS1411 MPs or Sgc8 MPs surface, respectively, to probe the corresponding mechanosignals.\nFor the AS1411-nucleolin pair, compared with cells transfected with scrambled siRNA, N-WASP knockdown HeLa cells still exhibit a stripe-like force pattern, although the signal amplitude is reduced by 27.9% (Fig. 4n, Supplementary Fig. 10a). This result indicates that force transmission through the AS1411-nucleolin pair is weakly dependent on N-WASP. In contrast, for the Sgc8-PTK7 pair, N-WASP knockdown in HepG2 cells leads to an almost complete loss of the ring-like pattern, with a 57.8% reduction in signal intensity relative to scrambled siRNA controls, suggesting a strong dependence of force transmission on N-WASP (Fig. 4o, Supplementary Fig. 10b).\nN-WASP is a key activator of Arp2/3-mediated branched actin polymerization, which has been reported to contribute distinctly to myosin-driven versus actin-polymerization-driven behaviors.14 Our observations are consistent with these prior reports. Moreover, the different effects of N-WASP knockdown in both cell lines closely mirror the results observed upon EIPA treatment. This is consistent with the fact that EIPA inhibits Rac1/Cdc42 activity via suppression of the Na⁺/H⁺ exchanger, and Cdc42 is a key upstream regulator of N-WASP. Together, these results further demonstrate that the AS1411-nucleolin and Sgc8-PTK7 pairs exhibit distinct dependencies on N-WASP and downstream Arp2/3-mediated branched actin polymerization, which we attribute to fundamentally different force transmission pathways. We added this new data to Fig. 4n-o and Supplementary Fig. 10.\nSupplementary Figure 1 a Representative immunostaining images and quantification of N-WASP expression level of HeLa cells after siRNA knockdown. HeLa cells transfected with either scrambled or N-WASP siRNA were stained with anti-N-WASP primary antibody followed by Alexa Fluor™ 647-conjugated secondary antibody. n = 1372 and 1304 cells from 3 replicates for scrambled and siRNA groups. Statistics: unpaired two-tailed Student’s t-test. b Representative immunostaining images and quantification of N-WASP expression level of HepG2 cells after siRNA knockdown. HepG2 cells transfected with either scrambled or N-WASP siRNA were stained with anti-N-WASP primary antibody followed by Alexa Fluor™ 647-conjugated secondary antibody. n = 1222 and 1198 cells from 3 replicates for scrambled and siRNA groups. Statistics: unpaired two-tailed Student’s t-test. Scale bars = 20 µm.\nFig. 4 n-o Effect of N-WASP on nucleolin- and PTK7-mediated force. n Representative brightfield and fluorescence images of HeLa cells after N-WASP siRNA knockdown on AS1411 MP surfaces. Mean fluorescence intensity per cell was quantified. n = 85, 83 cells from 3 replicates for scrambled and siRNA groups. Statistics: unpaired two-tailed Student’s t-test. o Representative brightfield and fluorescence images of HepG2 cells after N-WASP siRNA knockdown on Sgc8 MP surfaces. Mean fluorescence intensity per cell was quantified. n = 45 cells from 3 replicates. Statistics: unpaired two-tailed Student’s t-test. Scale bar = 20 µm.\nA discussion for this result has been added to the main text:\nTo further distinguish between two force transmission mechanisms, we knocked down the orthogonal regulator neuronal Wiskot–Aldrich syndrome protein (N-WASP). As a critical cytoskeletal regulator, N-WASP interacts with Arp2/3 complex to drive branched actin polymerization,52 thereby regulating essential cellular processes such as membrane ruffling, endocytosis, and cytoskeletal remodeling. Inhibition of the Arp2/3 complex has been shown to inhibit Arp2/3-mediated actin polymerization in phagocytic adhesion rings of macrophages but have little effect on integrin-mediated tensions in FAs.53 The later depends on myosin II-loaded stress fibers which lack Arp2/3 complex. For the AS1411-nucleolin pair, compared with cells transfected with scrambled siRNA, N-WASP knockdown HeLa cells still exhibit a stripe-like force pattern, although the signal amplitude is reduced by 27.9% (Fig. 4n, Supplementary Fig. 10a). This result indicates that force transmission through the AS1411-nucleolin pair is weakly dependent on N-WASP. In contrast, for the Sgc8-PTK7 pair, N-WASP knockdown in HepG2 cells leads to an almost complete loss of the ring-like pattern, with a 57.8% reduction in signal intensity relative to scrambled controls, suggesting a strong dependence on N-WASP-mediated actin dynamics (Fig. 4o, Supplementary Fig. 10b). Moreover, the different effects of N-WASP knockdown in both cell lines closely mirror the results observed upon EIPA treatment (Fig. 4b,h). This is consistent with the fact that EIPA inhibits Rac1/Cdc42 activity, and Cdc42 is a key upstream activator of N-WASP.54 Together, these results demonstrate that the AS1411-nucleolin and Sgc8-PTK7 pairs exhibit distinct dependencies on N-WASP and downstream Arp2/3-mediated branched actin polymerization, reflecting fundamentally different force transmission pathways.\nComment 2-3. A critical control is missing to validate the claim that the aptamer-based force sensing is functionally independent of the RGD-mediated adhesion used throughout the study. To robustly prove this independence, the assay should also be performed on a surface functionalized with the aptamer MPs alone (i.e., without co-immobilized RGD). This would directly test whether baseline integrin adhesion is a prerequisite for the aptamer probes to function, making the conclusion in Figure 2c more convincing.\nResponse: Thank you. We prepared surfaces functionalized with AS1411 MPs or Sgc8 MPs in the absence of co-immobilized RGD to directly compare unzipping- and shearing-mode of two probes under minimal adhesion conditions. As shown in Fig. 3f, HeLa cells are largely unable to spread on surfaces presenting AS1411 MPs alone, regardless of whether unzipping or shearing probes are used. Unlike the RGD-integrin, the AS1411-nucleolin interaction by itself is insufficient to promote cell adhesion. Compared with conditions in which RGD is co-immobilized, the AS1411 unzipping-mediated mechanosignals are significantly reduced (Fig. 3g), implying that baseline cell adhesion is a prerequisite for AS1411-based force sensing. We do observe a small fraction of cells that adhered to the surface via nonspecific interactions; notably, these cells are able to activate AS1411 unzipping MPs, further supporting the requirement for baseline adhesion.\nInterestingly, Sgc8 MP-functionalized surfaces exhibit a different behavior (Fig.3h). While Sgc8 MPs alone also fail to promote HepG2 cell spreading, the mechanoactivation remains robust (Fig.3i). Even in a non-spread, quasi-suspended state, HepG2 cells effectively activate Sgc8 unzipping MPs, yielding ring-like mechanosignals with intensities comparable to those observed on RGD co-immobilized surfaces. This indicates that for Sgc8-PTK7, baseline cell adhesion is not a prerequisite for mechanoactivation.\nTogether, the differential dependence of these two aptamer MPs on baseline cell adhesion further supports the existence of distinct force transmission mechanisms. For the AS1411-nucleolin pair, force generation driven by myosin contractility requires mature cell adhesion and myosin-loaded stress fibers. In contrast, for the Sgc8-PTK7 pair, as further elucidated in the mechanistic analysis, force generation arises primarily from membrane ruffling/actin polymerization, with much less dependence on cell adhesion. We have therefore softened the claim of functional independence and added this new data to Fig. 3f-i.\nFig. 3 f Representative brightfield and fluorescence images of HeLa cells cultured on surfaces functionalized with AS1411 MPs alone featuring different force geometries. No co-immobilized RGD-biotin is present to support baseline cell adhesion. Most HeLa cells remain non-adherent and fail to activate the unzipping AS1411 MPs. Only few cells adhering to the surface via nonspecific interactions can activate MPs. g Quantification of mean fluorescence intensity per cell for HeLa cells on each geometry. n = 68, 67 cells from 3 replicates. Statistics: Two-tailed, Mann-Whitney test. h Representative brightfield and fluorescence images of HepG2 cells cultured on surfaces functionalized with Sgc8 MPs alone featuring different force geometries. No co-immobilized RGD-biotin is present to support baseline cell adhesion. Non-spread HepG2 cells activate Sgc8 MPs. i Quantification of mean fluorescence intensity per cell for HepG2 cells on each geometry. n = 71, 68 cells from 3 replicates. Statistics: Two-tailed, Mann-Whitney test. Scale bars = 20 µm\nA discussion for this result has been added to the main text:\nSince the above results suggest that aptamers cannot promote cell adhesion, it raises the question to what extent baseline adhesion is required for aptamer MPs to function. To address this, we prepared surfaces functionalized solely with aptamer MPs without co-immobilized RGD to investigate mechanosensing under minimal adhesion conditions. HeLa cells are largely unable to spread on surfaces presenting AS1411 MPs alone, regardless of whether unzipping or shearing MPs are used (Fig.f). Compared with RGD co-immobilized conditions, AS1411 unzipping-mediated mechanosignals are significantly reduced to below 24.4 % (Fig.g), implying that baseline cell adhesion is a prerequisite for AS1411-based force sensing.\nConsidering that we aim to develop receptor-orthogonal aptamer MPs, we further investigated the Sgc8-aptamer as a potential MP for the protein tyrosine kinase 7 (PTK7), using HepG2 cells that overexpress this receptor. Interestingly, Sgc8 MPs-functionalized surfaces exhibit a somewhat different behavior (Fig.h). Even though Sgc8 MPs alone also fail to promote HepG2 cell spreading, robust mechanoactivation occurs (Fig.i). Even in a non-spread, quasi-suspended state (see brightfield images), HepG2 cells effectively activate Sgc8 unzipping MPs, yielding ring-like mechanosignals. This indicates that for Sgc8-PTK7, baseline cell adhesion is not a prerequisite for mechanoactivation. The differential adhesion dependence of these two aptamer MPs suggests that they transmit forces through fundamentally distinct mechanisms, which we will further discuss below.\nComment 2-4. The actual surface density and stoichiometry of the immobilized RGD and MP molecules were not quantified. These parameters are known to significantly influence cell adhesion and force generation. Providing this characterization would greatly enhance the rigor and reproducibility of the platform.\nResponse: We performed a reported surface density quantification assay that converts raw fluorescence intensity on our surfaces to the density of fluorescent molecules.12,15 Briefly, we used supported lipid bilayers (SLBs) prepared with known stoichiometry of fluorescent lipid as fluorescence standards because their dye density is well defined. We determined the relative molecular brightness (F factor) between the fluorophores by comparing the fluorescence intensities of labeled oligonucleotides and small unilamellar vesicles (SUVs) in solution. Using this F factor together with a fluorescence calibration curve obtained by imaging SLBs of known dye density, we converted measured fluorescence on our samples to surface concentrations of the oligonucleotide probes.\nWe generated two independent fluorescence calibration curves (using Texas Red™ DHPE and Oregon Green™ 488 DHPE as standards) and calculated the corresponding F factors (Supplementary Fig. 1); these are then applied to independently quantify the surface densities of immobilized RGD and MP molecules. However, direct quantification of the pristine c[RGDfK(Biotin)] molecule is still unfeasible due to its lack of a fluorophore. While using a fluorophore-conjugated RGD-biotin variant is an option, the substantial increase in size and steric hindrance could distort the actual RGD density. Thus, we chose FITC biotin (Mw 831.01 g/mol) as a proxy to estimate the RGD density, given its similar molecular weight to c[RGDfK(Biotin)] (Mw 829.98 g/mol). It should be noted that this approach ignores potential effects arising from differences in charge and hydrophilicity/hydrophobicity. The surface densities of aptamer MPs and RGD molecules are listed in Supplementary Table 2 now. The different MPs exhibit comparable surface densities in the range of 3500-4500 molecules/µm², and the overall stoichiometric ratio of MPs to RGD is approximately 3:1.\nSupplementary Figure 1. Quantification of surface density of aptamer MPs and RGD molecules on the surface. a-c Standard curves for quantifying density of aptamer MPs. a SLB calibration: Intensity of Texas Red DHPE (Texas Red™ 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt) to the number of molecules on the surface. n = 15 from 3 replicates. b-c F-factor plot estimation using concentrations of TR-DHPE-derived SUVs (b) and RRX-labelled oligonucleotide in solution (c) to compare the fluorescence intensity with density. The ratio of the calibration curve slopes was used to determine the “F factor” for the labelled oligonucleotide and the SUV samples. n = 15 from 3 replicates. d-f Standard curves for quantifying density of RGD molecules. Direct quantification of the pristine c[RGDfK(Biotin)] molecule is unfeasible due to its lack of a fluorophore. Using a fluorophore-conjugated RGD-biotin variant may distort the actual RGD density due to the increase in size and steric hindrance. Thus, we chose FITC-biotin (Fluorescein-5(6)-biotinamidohexanoylamidopentylthiourea, Mw 831.01 g/mol) as a proxy to estimate the RGD density, given its similar molecular weight to c[RGDfK(Biotin)] (Mw 829.98 g/mol). It should be noted that this approach ignores potential effects arising from differences in charge and hydrophilicity/hydrophobicity. d SLB calibration: Intensity of Oregon Green 488 DHPE (Oregon Green™ 488 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine) to the number of molecules on the surface. n = 12 from 3 replicates. e-f F-factor plot estimation using concentrations of OG-DHPE-derived SUVs (e) and FITC-biotin in solution (f) to compare the fluorescence intensity with density. The ratio of the calibration curve slopes was used to determine the “F factor” for the labelled oligonucleotide and the SUV samples. n = 12 from 3 replicates.\nSupplementary Table 2. Surface density of aptamer MPs and RGD molecules. n = 18 from 3 replicates.\nMP density (molecule/µm2) RGD-bio density (molecule/µm2)a\nAS1411 unzipping 4329±394 1297±155\nSgc8 unzipping 4480±239 1320±159\nMUC1 S2.2 unzipping 4040±216 1322±101\nSYL3c unzipping 4072±166 1356±134\nAS1411 shearing 4141±269 1254±82\nSgc8 shearing 3918±609 1321±61\nRGD unzipping 3495±201 1374±92\nRGD shearing 3581±241 1318±155\na) mimicked by FITC-biotin molecules with similar molecular weight.\nA discussion for this result has been added to the main text:\nSupported lipid bilayer-based surface density calibration assays reveal that the densities of different aptamer MPs are in the range of 3500-4500 molecules/µm², and the overall stoichiometric ratio of MPs to RGD is approximately 3:1 (Supplementary Fig. 1, Supplementary Table 2).\nComment 2-5. The SDR module is currently demonstrated with only a single block–activate cycle, which is not sufficient to establish reversibility. It is recommended that the authors present at least two or more consecutive suppression-recovery cycles. In addition, the RNase H-based “timer” shows tunable activation rates but not a clearly defined lag phase. Including earlier and more frequent time points would help demonstrate a controllable delay before signal onset, and a simple kinetic model correlating decoy concentration with delay would further substantiate the claim of programmability.\nResponse: Thank you for these suggestions.\n(i) SDR module.\nTo further demonstrate the dynamic reversibility, we conducted a secondary suppression-recovery cycle following the initial cycle. Specifically, after adding the first-round activator for 1 h, the medium was replaced with the second-round blocker to suppress further mechanoactivation. After an additional 0.5 h, a second-round activator was introduced (Fig. 6a). Consistent with the first cycle, the system was incubated for 3 hours prior to imaging. In the second suppression-recovery cycle, addition of the blocker again effectively suppresses the mechanosignals, whereas the subsequent addition of the activator restores the force transmission, mirroring the behavior observed in the initial cycle. The signal level in the second-round blocker condition is higher than that in the first-round blocker group, which we attribute to the cumulative mechanoactivation accumulated over the additional activation period. We added this new data to Fig. 6b and Supplementary Fig. 12e, and discussed it in the text.\nFig. 1 a Schematic of switchable mechanosensing using SDRs for two suppression-recovery cycles. In the 1st cycle, HeLa cells were seeded on surfaces grafted with AS1411 MP for 30 min, followed by addition of 200 nM DNA blocker strands to inhibit aptamer-receptor interaction. After another 30 min, 200 nM DNA activator strands were introduced to displace the blockers via toehold-mediated SDR, restoring mechanosensing. Mechanosiganls were quantified after 3 h. In the 2nd cycle, another round of blocker strands was introduced 1 h after adding activator strands in the 1st cycle. After an additional 30 min, a second-round activator was introduced. Consistent with the 1st cycle, the system was incubated for 3 h prior to imaging. b Quantification of mean fluorescence intensity per cell of HeLa cells on AS1411 MP surfaces after sequential blocker and activator treatment for two suppression-recovery cycles. 1st cycle, n = 54 cells from 3 replicates. 2nd cycle, n = 48 cells from 3 replicates. Statistics: 1st cycle, One-way ANOVA with Bonferroni post-hoc tests. 2nd cycle, Two-tailed, Mann-Whitney test.\nSupplementary Figure 12 e Representative brightfield and fluorescence images of HeLa cells on AS1411 MP surfaces after sequential blocker and activator treatment. Scale bars = 20 µm.\nA discussion for this result has been added to the main text:\nThe second blocker-activator cycle closely mirrors the suppression-recovery behavior observed in the initial cycle (Supplementary Fig. 12e). The overall higher signal level can be attributed to cumulative mechanoactivation accumulated during the additional activation period.\n(ii) RNA-RNase H module.\nWe monitored the reaction kinetic of RNA-RHase H module to assess reaction rates. A FRET pair (Atto488/BHQ1) was incorporated into the AS1411 strand and RNA blocker so that we can quantify the ratio of AS1411 aptamer reconfiguration. We assumed pseudo first-order kinetics and fitted the data using one phase exponential decay function. The reconfiguration of aptamer MPs does not reach saturation by 2 h (the onset of measurable mechanoactivation) due to a relatively low RNase H concentration used (10 U/mL) before. Therefore, no clearly defined lag phase is observed (Supplementary Fig. 13b and Supplementary Table 3). Nevertheless, the introduction of decoy strand exerts a clear, dose-dependent inhibitory effect on the reaction rate of RNA blocker degradation: Consistent with this trend, we monitored mechanoactivation on the cell level at 0.5 h and 1 h after cell seeding and find that the decoy-induced reduction in reaction rate and the extent of MP reconfiguration correlates with the onset of force signals (Fig. 6h, Supplementary Fig. 13c). Compared with 1 nM decoy, 10 nM decoy slightly delays the rate of signal increase, whereas 100 nM decoy markedly prolonged the time required to reach a signal amplitude comparable to that achieved with 1 nM decoy at 2 h, extending it to approximately 7 h.\nSupplementary Figure 13. b Plot of normalized surface atto488 intensity versus time to compare the reconfiguration rate of AS1411 aptamer in RNA-RNase H module in the presence of decoy strands. Surface-immobilized AS1411 MPs are blocked with RNA blocker and degraded by RNase H at 10 or 100 U/mL and decoy strands at 1-100 nM. Reaction medium: 1 % FBS, 1 % P/S, 100 µg/mL actin protein, DMEM. n = 15 from 3 replicates. At each reaction time point, the surface Atto488 intensity is normalized to that of unblocked AS1411 MPs labeled with atto488, which serves as the positive control. c Representative brightfield and fluorescence images of HeLa cells on RNA-blocked AS1411 MP surfaces after adding 10 U/mL RNase H and varying RNA/DNA duplex concentrations. Scale bars = 20 µm.\nSupplementary Table 3. Reaction rate constant of SDR and RNA-RNase H modules. To understand the reconfiguration efficiency and tunability of different DNRs, we quantified the surface reconfiguration kinetics. The data were analyzed assuming pseudo-first-order kinetics and fitted using one phase exponential decay function. For the SDR module, reactivation with 200 nM DNA activator yields a reconfiguration rate of 2.14 × 10⁻² min⁻¹. The RNA-RNase H module enables degradation rate tuning by adjusting the initial RNase H concentration, resulting in accelerated (100 U/mL, 5.26 × 10⁻² min⁻¹) or decelerated (10 U/mL, 0.71 × 10⁻² min⁻¹) reconfiguration kinetics. Moreover, at a fixed RNase H concentration (10 U/mL), the RNA-RNase H module allows the introduction of non-enzymatic RNA/DNA decoys, providing an additional orthogonal layer of regulation.\nk (×10-2 min-1) R2\nSDR 200 nM DNA activator 2.14 0.9926\nRNase H 100U/mL 5.26 0.9925\nRNase H 10U/mL 0.71 0.9842\nRNase H 10U/mL+1 nM rdm RNA/DNA decoy 0.73 0.9782\nRNase H 10U/mL+10 nM rdm RNA/DNA decoy 0.42 0.9603\nRNase H 10U/mL+100 nM rdm RNA/DNA decoy 0.11 0.9831\nFig. 2 h Corresponding quantification of mean fluorescence intensity per cell. Reaction medium: 1 % FBS, 1 % P/S, 100 µg/mL actin protein, DMEM. n = 27 cells from 3 replicates. Scale bars = 20 µm.\nA discussion for this result has been added to the main text:\nThe introduction of decoy strand exerts a clear, dose-dependent inhibitory effect on the reaction rate of RNA blocker degradation (Supplementary Fig. 13b, Supplementary Table 3). Consistent with this trend, 10 nM decoy is sufficient to slightly delay the mechanoresponse, whereas 100 nM nearly abolishes the response within 4 h and a significant response is observed only at 7 h (Fig. 6g,h, Supplementary Fig. 13c).\nComment 2-6. The statement that aptamer MPs exhibit “enhanced force selectivity” compared to RGD MPs may be misleading. A more accurate interpretation is that AS1411–nucleolin interactions occur within a lower physiological force regime (consistently below 54 pN), while RGD–integrin forces readily exceed this threshold. This distinction reflects biological force domains rather than an intrinsic property of the probe.\nResponse: Thanks for bringing this up. We have revised the subsection title and re-written the corresponding text to make it clearer.\nComment 2-7. Regarding the citation of Yang et al. (Ref. 20), the manuscript currently describes this study as “force being sensed using a DNA MP”, which may not fully reflect its contribution. Specifically, Yang et al. provided the proof-of-concept that a DNA aptamer, taking Cl-M6RP-targeting aptamer as the example, can function as a force-reception module to sense weak endocytic forces with fluorescent signal readout and then translate these force cues into intracellular signals via DNA dynamic reactions. Thus, a more balanced description should acknowledge this advance in the revision, and it is recommended to add a brief discussion clarifying how the present work extends this concept.\nResponse: We thank the reviewer for pointing this out and have properly credited this work in the Introduction Section:\nIndeed, a recent proof-of-concept using the CI-M6PR-targeting aptamer demonstrated that an aptamer can function as a force-reception module to sense weak endocytic forces (~4 pN)24 and then translate this force cue into intracellular signals via DNA-mediated receptor dimerization.20\nWe included a comparison in the Discussion Section:\nThese newly uncovered mechanisms also expand the potential of aptamers as versatile force-reception modules for bidirectional mechano-biochemical signal transduction. Beyond relying on weak DNA hairpins to sense clathrin-mediated endocytic forces (~4 pN),20 cells can exert forces >12 pN on aptamer MPs. Such forces allow the irreversible exposure of cryptic ssDNA domains that can trigger downstream reactions such as transcription, hairpin chain reactions,21 or rolling circle amplification.59 This expanded force range substantially broadens the design space for intelligent, force-responsive material interfaces capable of cellular adaptation.\nMinors\nComment 2-8. Use more precise terminology such as “receptor-low” or “low-expressing” instead of “receptor-negative,” as the supplementary flow cytometry data indicate non-zero expression.\nResponse: Thank you. We have replaced the terminology “receptor-negative” and “receptor-positive” with “low-expressing” and “high-expressing,” respectively.\nComment 2-9. Clarify in the text or figure legend for Figure 3 why both “mean” and “integrated” intensity are presented, and how each metric provides distinct insights.\nResponse: Thank you. Please see Response 1-4 above.\nA discussion for this result has been added to the main text:\nThe integrated intensity reflects the cumulative mechanosignal across the entire cell footprint and is thus not only regulated by intrinsic Ttol of the probe but inherently dependent on cell area.19\nWe also updated the figure captions:\nFig.3 c-e Quantification of (c) mean fluorescence intensity per cell, (d) integrated fluorescence intensity per cell, and (e) cell spreading area for HeLa cells on each MP type and geometry. n = 49, 47, 45, 36 (left to right) cells from 3 replicates. Statistics: Kruskal-Wallis test with Dunn’s multiple comparisons. Note that both mean and integrated intensity are shown because mean intensity mainly captures the Ttol-dependent shifts while integrated intensity is cross-regulated by both the intrinsic Ttol of the probe and the cell spreading area.\nFig.5 b Corresponding quantification of mean and integrated fluorescence intensity per cell of HeLa and HepG2 cells on MUC1 S2.2 MP surfaces. n = 31, 27 cells from 3 replicates. Statistics: Two-tailed, Mann-Whitney test. Note that both mean and integrated intensity are shown here because different cell lines may have distinct basal morphologies with different cell area.\nComment 2-10. In Figure 6, the restored force signal distribution appears inconsistent across conditions (e.g., recovery mainly at the cell edge vs. both edge and center). The authors should clarify whether these differences reflect mechanistic variations or are attributable to experimental variability.\nResponse: Thank you for the question. We have carefully checked the spatial distribution of the force signals in HeLa cells. The characteristic force pattern mediated by the AS1411-nucleolin interaction is defined by strong stripe-like signals at the cell edge, like focal adhesions, whereas signals in the central region are consistently weaker. We examined all images showing restored force signals. Under comparable restoration conditions, the overall force patterns are consistent. For example, the force distributions observed in Fig. 6d (RNase H, 10 U/mL, 4 h) and Fig. 6g (RNase H, 10 U/mL+ 1 nM decoy, 4 h) are largely similar. We do acknowledge that the intensity of the central signal can vary between individual cells. For instance, in Fig. 6g, the 1 nM decoy group (7h) exhibits central signals below the average, while the 10 nM decoy group (7h) shows relatively higher central intensity. Despite these fluctuations, the fundamental spatial logic where peripheral stripe-like signals dominate over central signals remains conserved across most of the groups. We therefore attribute these differences to individual cellular heterogeneity and experimental variability rather than a shift in the mechanisms.\nComment 2-11. For PTK7-mediated forces, it remains unclear whether the signal reflects protrusive pushing or contractile pulling. The schematic suggests pulling, whereas the text emphasizes polymerization-driven pushing. This ambiguity should be explicitly acknowledged and discussed.\nResponse: Thank you for pointing this out, and we apologize for this mistake in the schematic. Based on our previous data and the new analyses presented in Responses 2-2 and 2-3, PTK7-mediated forces arise predominantly from actin polymerization-driven pushing. We have corrected the schematic accordingly now.\nAll related discussions have been revised to consistently emphasize this actin polymerization-driven pushing.\nComment 2-12. In the discussion, briefly acknowledge the limitations of DRN-controlled systems for potential in vivo applications, including delivery challenges, nuclease stability, and cytotoxicity.\nResponse: Thank you. We have added the related discussion in the text:\nHowever, long-term application in these environments still faces stability challenges. While modifications like peptide nucleic acids,63 phosphorothioate linkages,64 or our recent L-DNA strategy protect the MP structure,65 single-stranded aptamers remain inherently degradation-prone. Balancing their structural integrity with binding affinity is critical. In this context, using nuclease inhibitors such as actin or decoy DNA may be more effective.66,67 Additionally, for potential in vivo applications, deep-tissue diffusion, off-target effects arising from protein corona formation, and potential immunotoxicity remain significant challenges.\nComment 2-13. While the manuscript identifies novel force pathways, the discussion could be enriched by addressing their biological significance. Why do cells exert forces through these non-canonical receptors? Furthermore, since detailed analysis was limited to nucleolin and PTK7, it would be valuable to comment on the potential generalizability to other possible aptamer-receptor pairs with biological functions.\nResponse: Thank you. The transient coupling between these non-canonical receptors and the cytoskeleton provides the fundamental structural basis for force transmission, while their biological functions are still unknown at this stage and should not become subject to excessive speculation in this manuscript. Endocytic force is a classical form of actively applied cellular force acting on membrane receptors and is associated with well-defined biological functions. During endocytosis, receptors are coupled to endocytic machinery, such as clathrin and the cytoskeleton, generating forces of ~4.7 pN. Through this canonical process, cells actively internalize and recycle membrane proteins to maintain signaling homeostasis.\nIn our study, however, the unzipping probes selectively filter out forces below 12 pN, indicating that the forces we observe are unlikely to arise from receptor internalization or recycling. Instead, the relatively high forces associated with nucleolin or PTK7 likely originate from their coupling to more mechanically active, high-force cytoskeletal structures. Whether this force transmission reflects an active, regulated cellular process or a passive mechanical linkage remains unclear, and to our knowledge, there is currently no direct evidence demonstrating a definitive function associated with such forces. This highlights the importance of revisiting the biological functions of both these aptamers and receptors from a mechanobiological perspective. For example, whether AS1411-mediated processes such as angiogenesis are modulated by mechanical forces remains an open question. Although we are highly interested in these directions, they extend beyond the primary scope of the present work. Notably, in our experiments, unlike mobile native ligands, aptamer MPs are stably immobilized on the surface, thereby providing sufficient reaction force to support force transmission. It is also of significant interest to explore whether cryptic motifs within the native extracellular matrix can interact with nucleolin with similar force transmission, and regulate cellular homeostasis through non-canonical mechanotransduction pathways. A prime candidate for such an interaction is the precursor of the nucleolin ligand endostatin, located in the NC1 domain of Type XVIII collagen,8 which may serve as a native mechanical anchor despite potential differences in binding affinity.\nFinally, as suggested by the reviewer, the generalizability of the aptamer-based platform provides opportunities to probe more established aptamer–receptor pairs with known biological functions, such as PD-L1-targeting aptamers involved in immune checkpoint regulation,16,17 or CD28 costimulatory aptamers in T cells,18 to determine whether their biological activities are also regulated by mechanical force. Exploring these questions represents an important and promising direction for future research.\nWe have revised the related discussion in the text:\nThe identification of noncanonical forces highlight the importance of revisiting the biological functions of both these aptamers and receptors from a mechanobiological perspective. Determining whether these receptors are directly coupled to classical adhesion structures or are independently linked to the same actomyosin network via intermediary components will be essential for uncovering more general principles. Whether similar interactions occur under native conditions remains to be elucidated. A prime candidate is the precursor of the nucleolin ligand endostatin, located in the NC1 domain of type XVIII collagen,60 which may serve as a native anchor for nucleolin despite potential differences in binding affinity. Addressing these questions could reveal how cryptic motifs within the ECM regulate cellular homeostasis through non-canonical mechanotransduction pathways. Meanwhile, caution is required when extrapolating from aptamer-receptor pairs to native ligand-receptor interaction, particularly regarding the immobilization state of the ligand, binding affinity and bond mechanics, and the multivalency and cooperative effects inherent to native ligands. For example, the AS1411 aptamer binds nucleolin with a Kd of 69.1 nM,28 which is lower than that of its native ligand endostatin (23.2 nM),61 suggesting that our measurements may slightly underestimate the native capacity for tension generation. Finally, the generalizability of the aptamer-based platform provides opportunities to probe other established aptamer-receptor pairs with well-defined biological functions, such as PD-L1-targeting aptamers involved in immune checkpoint regulation,62,63 or CD28 costimulatory aptamers in T cells,64 to determine whether their biological activities are similarly regulated by mechanical force."}, {"round": 2, "reviewer_comments": {"reviewer_1": "I appreciate the careful revision from the authors. Excellent work!\nI have reviewed all the responses to my previous questions and the revised manuscript. I believe the manuscript is suitable for publication in Nat. Comms. without further revision.", "reviewer_2": "The authors have adequately addressed the concerns raised in my previous review by providing substantial additional experimental data and analyses. The revised manuscript presents strengthened mechanistic support for the main conclusions.Overall, the data support the claims, the methodology is sound, and the current version meets the standards for publication in Nature Communications. I therefore support acceptance of the manuscript in its present form."}, "author_response": ""}]

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