A bright and regenerative array tag with fluorogenic ligands for long-term live-cell nanoscopy and single-molecule tracking

Domain: neuroscience
Published: 2025-12-12
Editorial outcome: major_revision

Abstract
Live-cell fluorescence nanoscopy and single-molecule tracking offer profound insights into cellular biology, but their application is limited by fluorescent tags with low brightness, large size, and irreversible photobleaching. Here, we present ReTagX, a bright, regenerative, compact array tag system enabling rapid recovery of photobleached signal through reversible, fluorogenic labeling. Newly developed ReTag1 (~18 kDa) with highly fluorogenic MaP655-TMP achieves a signal-to-noise ratio up to 204-fold and allows fluorescence recovery within 1 minute in wash-free live-cell imaging. This tag extends STED imaging of mitochondrial TOM20 from fewer than 5 to over 300 frames, enabling long-term visualization of mitochondrial dynamics at 80 nm resolution for over 100 minutes. Multimeric ReTag1 arrays, such as ReTag8 (~156 kDa), increase brightness 5.5-fold and extend TNFR2 single-molecule tracking ~2.9-fold, preserving >80% of trajectories over 5 minutes. ReTagX provides a robust platform for high-resolution, long-term molecular visualization of cellular dynamics in living cells.

Introduction
Advances in fluorescence imaging have profoundly transformed our understanding of biological systems. Super-resolution microscopy (SRM), or nanoscopy, surpasses the diffraction limit of light (~200 nm), enabling unparalleled visualization of cellular structures and protein dynamics at the nanometer scale1,2. Over the past three decades, a variety of nanoscopy techniques—such as Structured Illumination Microscopy (SIM)3, Stimulated Emission Depletion (STED)4, Single Molecule Localization Microscopy (SMLM)5–7, and Minimal Photon Fluxes (MINFLUX)8—have been developed to enable protein imaging at the nanoscale. Of these, live-cell super-resolution imaging is particularly valuable for monitoring target proteins in real-time during cellular processes, offering robust spatial and temporal resolution while avoiding artifacts introduced by cell fixation9,10. Furthermore, single-molecule tracking (SMT) in living cells enables the direct observation of individual protein dynamics and interactions, providing critical insights into cellular processes such as signal transduction, membrane organization, and intracellular trafficking11–13. However, despite significant advancements, time-lapse recording of protein dynamics at high resolution remains a key challenge due to limited brightness, large size, and irreversible photobleaching of conventional fluorescent protein tag system.

To optimize nanoscopy and SMT performance in living cells, small-molecule organic fluorophores are preferred for their superior brightness, photostability, and continuous potential for synthetic optimization. Recent advances by research groups including Johnsson14, Lavis15, Hell16, and ourselves have propelled the development of cell-permeable and fluorogenic dyes, such as JF and MaP dyes, which exist in a nonfluorescent spirocyclic form in the unbound state but transition to a fluorescent zwitterionic form upon binding to target proteins17. Meanwhile, Self-labeling protein (SLP) tags, including SNAP-tag18, CLIP-tag19, Halo-tag20, and TMP-tag21,22, have been extensively developed to enable selective protein labeling with these organic fluorophores (Fig. 1). The covalent interaction between SLP substrates and genetically fused tags allow precise labeling of intracellular proteins. This combination has enabled live-cell labeling and nanoscopy of various proteins, such as cytoskeletal components16,23, β-site APP-cleaving enzyme 1 (BACE1)24, and lysosome-associated membrane glycoprotein 1 (LAMP1)25. However, a major limitation of this approach is irreversible photobleaching, which hampers long-term tracking and imaging of dynamic cellular processes. The issue of photobleaching and irreversible signal loss is particularly acute in super-resolution techniques like STED microscopy26, where ultrahigh laser intensities (~370 mW) are routinely used, restricting imaging to only a few frames27.

Another classical protein labeling strategy is array tag that introduces tandem repeats of fluorescent reporters into a single target protein, producing significantly enhanced brightness and tracking duration (Fig. 1). For instance, in array tag systems like SunTag28 and ArrayG29, ~24 peptides or small proteins (e.g., nanobodies) are fused to the target protein, recruiting multiple fluorescent proteins to significantly boost brightness and photobleaching resistance in single-molecule tracking. The increased brightness also allows for the use of lower laser power, further reducing photobleaching and extending the tracking time (Supplementary Table 1). However, the relatively large size (~1.4 mDa) of these fusion constructs can significantly interfere with the diffusion behavior and interaction of target proteins, which typically range from 10 to 160 kDa.

To overcome these limitations, regenerative tags that enable fluorescence self-recovery have emerged as promising tools for super-resolution and long-term single-protein tracking. Heilemann’s group pioneered a reversible labeling strategy using fluorophore-labeled oligonucleotides that bind to protein-specific antibodies, extending the number of STED images from 10 to 50 frames, corresponding to the fluorescence intensity decaying by half in fixed samples30. While effective in fixed cells, this strategy presents significant challenges in living cells. Efficient fluorescence self-recovery requires four key criteria: (1) Rapid and reversible labeling: Fluorogenic ligands must bind and release within seconds to minutes to efficiently exchange bleached ligands. (2) High fluorogenicity: The fluorescence of ligands upon binding should be significantly brighter than the background from unbound ligands, enabling a large “fresh probe pool” for efficient exchange and reducing phototoxicity inside cells. (3) High brightness: The bound ligand on protein tag must be sufficiently bright for accurate localization and tracking with high temporal resolution. (4) Small tag size: A compact tag minimizes interference with protein function, expression, and diffusion. Recent developments, such as reHaloTags31, exchangeable Halo-Tag Ligands (xHTLs)32,33, and srTAG34, have been introduced for fluorescence self-recovery, yet their large tag sizes, unsatisfied brightness, or slow binding kinetics constrain their effectiveness as regenerative tags or array-forming platforms for long-term live-cell nanoscopy and single-protein tracking (Supplementary Table 1).

To achieve for long-term live-cell superresolution imaging and tracking, we developed a bright, regenerative, and compact array tag system (ReTagX) coupled with highly fluorogenic ligands. By optimizing the spirocyclization of rhodamines and linker length, we engineered highly fluorogenic ligands that provide bright labeling with minimal background, achieving up to a 203.8-fold signal-to-noise ratio in wash-free live-cell imaging. Systematic mutations (L28Q, R44L, H45Q) generated a regenerative tag (ReTag) exhibiting a 105–106-fold increase in the dissociation rate constant (Koff), enabling reversible labeling within seconds to minutes. This remarkable fluorescence recovery from ReTag1 (~18 kDa) and a pool of fluorogenic ligands, extended STED imaging of mitochondrial TOM20 from fewer than five frames to over 300, achieving 80 nm resolution for 100 min. Moreover, multimeric ReTagX arrays (e.g., ReTag8, ~156 kDa) enhance brightness by 5.5-fold and enable >80% trajectories retention over 5 min, extending single-molecule tracking of membrane proteins such as TNFR2 by ~2.9-fold compared to HaloTag in live-cell imaging. This advancement paves the way for prolonged visualization of cellular dynamics with robust spatial and temporal resolution in living biological systems.

Results
Design of a compact, bright and regenerative array tag
Fluorescence recovery is most effectively achieved through the spontaneous replacement of bleached fluorophores on a protein tag with fresh fluorescent ligands (Fig. 1). Inspired by the labeling strategies of exchangeable Halo-Tags and ArrayG, we sought to develop a bright, regenerative, and compact array tag system (ReTagX, 18-156 kDa) featuring fluorogenic ligands spanning the green to red spectrum. This system incorporates multiple repeats of a regenerative protein tag, which has been systematically engineered to enable efficient self-recovery of photobleached signals through reversible labeling with fluorogenic ligands. Beginning with the TMP-Tag, derived from Escherichia coli dihydrofolate reductase (eDHFR) due to its relatively small size (18 kDa, around half the size of the Halo-Tag at 33 kDa)35, we initially focused on developing highly fluorogenic ligands for TMP-Tag and subsequently performed systematic mutations to optimize binding and release kinetics for developing a regenerative protein tag—ReTagX. Finally, we demonstrate the feasibility of the array tag system (ReTag1-8) and its fluorogenic ligands for long-term live-cell nanoscopy and single-protein tracking.

Development of multicolor highly fluorogenic ligands
In developing highly fluorogenic ligands for the TMP-Tag, we employed sulfonamide-modified carbo-rhodamine (MaP618m) as a fluorescent reporter due to its robust brightness, photostability, and ability to transition from a nonfluorescent spirocyclic state to a fluorescent zwitterion upon protein binding. Trimethoprim (TMP)36, a drug with picomolar affinity for TMP-Tag in the presence of NADPH37 and excellent cell permeability, was conjugated with MaP618m via alkane linkers of varying lengths (C2, C4, C6), yielding MaP618m-C2TMP, MaP618m-TMP, and MaP618m-C6TMP (Fig. 2a and Supplementary Fig. 1). A control probe, CPY-TMP, was synthesized using carbo-rhodamine with a C4 linker (Supplementary Fig. 1).

Spectroscopic characterization in absorbance and fluorescence was next examined for fluorescent ligands in the presence of TMP-Tag. MaP618m-TMP (C4 linker) demonstrated high fluorogenicity, exhibiting a 30.4-fold increase in absorbance at 616 nm and a 190.2-fold enhancement in fluorescence intensity at 636 nm (Fig. 2b, c and Supplementary Fig. 2), significantly surpassing the performance of the control probe CPY-TMP (Supplementary Fig. 3). Importantly, the interaction between the TMP-Tag and MaP618m-TMP produced greater brightness and fluorogenicity than that of Halo-Tag with ligands (MaP618m-Halo) using the same fluorogenic reporter MaP618m (Fig. 2c and Supplementary Fig. 3). Additionally, it reveals that linker length critically influenced fluorogenicity: MaP618m-C2TMP and MaP618m-C6TMP showed lower fluorescence increases of 73.0-fold and 36.7-fold, respectively (Fig. 2c), attributed to differential interactions with the TMP-Tag surface. Molecular docking studies (PDB: 7NAE) supported these findings, revealing that the C4 linker allowed optimal stabilization of the zwitterionic form on the TMP-Tag, while the shorter C2 linker restricted conformation within the binding pocket, and the longer C6 linker exposed the dye to solvent (Fig. 2d).

We then evaluated the performance of TMP ligands in live-cell imaging using U2OS cells expressing a nuclear-localized SNAP-TMP-NLS fusion protein, co-cultured with untransfected U2OS cells. Notably, U2OS cells incubated with 200 nM of the control probe CPY-TMP exhibited a high nucleus-to-cytosol signal ratio of 26.3-fold (Fig. 2e, f). However, increasing the probe concentration to 2 μM resulted in significant nonspecific staining and background signals, with a significantly reduced cytosol-to-medium signal ratio of 3.7-fold (Supplementary Fig. 4c). This inability to use high-concentration probes hinders the formation of an efficient pool for fluorescence self-recovery in live-cell imaging. In contrast, the background signal from untransfected U2OS cells incubated with MaP618m-TMP was negligible with no visible aggregation and indistinguishable from the medium, at both 200 nM, 2 μM and 10 μM (Fig. 2e and Supplementary Figs. 4, 5), a result not previously achieved in the literature. Importantly, MaP618m-TMP enables bright fluorescence labeling of nucleic TMP-NLS within 6 min (Supplementary Fig. 4f), yielding a remarkable nucleus-to-cytosol signal ratio of 110.4-fold and 134.0-fold at 200 nM and 10 μM MaP618m-TMP, respectively (Fig. 2e, f).

The performance of highly fluorogenic TMP ligands in intracellular protein labeling enables the development of fluorescent ligands across the green to red spectral range. Leveraging the comparable fluorogenicity of the TMP-Tag and HaloTag17,38, we synthesized fluorogenic TMP-Tag ligands (MaP512-TMP, MaP555-TMP, SiR-TMP, and MaP655-TMP) by replacing the Halo-Tag substrate (chloroalkane) with trimethoprim, linked via a C4 alkane chain (Supplementary Figs. 1 and 3). The fluorogenicity of newly developed fluorogenic ligands was first confirmed by the increased D50 and decreased Log K L-Z, representing the shifted equilibrium from zwitterionic states to spirocyclic form (Supplementary Figs. 6 and 7). Compared to the control probe R500-TMP, the green fluorogenic ligand MaP512-TMP demonstrated substantial improvements in both absorbance (11.8-fold) and fluorescence (31.5-fold), enabling rapid intracellular TMP-Tag labeling (<15 min) with a high signal-to-background ratio of 50.1-fold at 200 nM (Supplementary Fig. 8). However, at 1 μM, some background signal and nonspecific staining were observed, reducing the signal-to-background ratio to 8.9-fold (Supplementary Fig. 8). Notably, MaP555-TMP show moderate fluorogenicity (Fig. 2g, h) and efficiently labeled nuclear TMP-Tag, yielding an excellent nuclei-to-cytosol signal ratio (Fnuc/Fcyt = 36.5) under wash-free conditions within 5 min (Fig. 2i–k). Importantly, even at 1 μM, MaP555-TMP maintained a low background signal and bright labeling of nucleic protein with a high nuclei-to-cytosol signal ratio (Fnuc/Fcyt = 32.2) (Fig. 2i, j and Supplementary Fig. 9).

Silicon-rhodamine (SiR) is well-established for its fluorogenic properties in protein labeling14. SiR-TMP exhibited moderate fluorogenicity upon binding to TMP tags (Fig. 2l and Supplementary Fig. 3), with a 9.4-fold increase in fluorescence in vitro and a favorable nuclei-to-cytosol signal ratio (Fnuc/Fcyt = 73.9) at a low concentration of 200 nM (Fig. 2n, o). However, increasing the concentration to 2 μM led to substantial background signal and nonspecific staining, limiting its effectiveness in forming an efficient “probe pool” for fluorescence recovery (Supplementary Fig. 9). To overcome this, we incorporated 3,5-difluorobenzene-1-sulfonamide into SiR-TMP, resulting in the highly fluorogenic TMP ligand MaP655-TMP (Fig. 2l). Spectroscopic characterization confirmed its superior fluorogenicity, with substantial enhancements in both absorption (36.2-fold) and emission (179.8-fold) (Fig. 2m). In live-cell imaging, MaP655-TMP at 200 nM displayed intense nuclear fluorescence with a significant contrast ratio (Fnuc/Fcyt = 179.1) within 10 min of staining (Fig. 2n–p). Crucially, even at 10 μM, MaP655-TMP maintained negligible background (Fcyt/Fmed = 1.0) and a high contrast ratio (203.8-fold) (Fig. 2n). Such high-contrast imaging and fast labeling of live U2OS cells with highly fluorogenic MaP618m-TMP or MaP655-TMP, especially with a pool of high-concentration probes, hold great potential for fluorescence self-recovery, while maintaining good biocompatibility at high concentrations and showing no adverse effect on cell proliferation activity (Supplementary Fig. 10).

Development of regenerative tag for live-cell fluorescence self-recovery
We next explored the potential of the non-covalent TMP-Tag for reversible labeling. SDS-PAGE followed by in-gel fluorescence scanning confirmed the non-covalent nature of the interaction, as fluorescent signal from the TMP-Tag protein band removed in SDS-PAGE (Supplementary Fig. 11). Confocal imaging of live U2OS cells expressing H2B-SNAP-TMP and labeled with MaP555-TMP (with SiR-BG as control) revealed a bright nuclear signal that remained stable for over 4 h after probe washout, suggesting unexpectedly high TMP–ligand complex stability (Supplementary Fig. 12). It is probably due to the ultra-strong binding affinity and stable closed conformational states of TMP-Tag with NADPH and Trimethoprim derived probe37. This was also confirmed by Biofilm Layer Interference (BLI) analysis, which revealed that MaP555-TMP exhibited a rapid association with wild-type TMP-tag (Kon = 0.925 × 105 M−1s−1) but a relatively low dissociation rate (Koff = 4.88 × 10−7 s−1) (Fig. 3a and Supplementary Fig. 13). Such stable binding prevents the release of bleached fluorophores and subsequent fluorescence recovery through reversible labeling. Consequently, we introduced multiple mutations into the TMP-Tag to establish a reversible labeling system with fast binding kinetics, while preserving high fluorogenicity by carefully balancing exchange rate, binding specificity, and signal contrast. The high fluorogenicity effectively suppresses nonspecific background fluorescence, enabling the use of high probe concentrations to promote probe exchange and maintain strong target labeling. To maintain fluorogenicity, we initially introduced R44L and H45Q mutations, known to decrease binding with trimethoprim and NADPH but avoiding affect fluorogenicity due to few interactions with the fluorogenic ligand39. Meanwhile, more mutations, including M20V40, D27E41, L28Q41, and W30R42, which contribute to trimethoprim resistance by expediting its release from eDHFR (TMP-Tag), were further introduced (Fig. 3b). Screening results showed that the mutations (L28Q, R44L and H45Q) reduced the affinity of the TMP-Tag and its ligands (KD: 0.33-2.8 μM) while greatly maintaining their fluorogenicity (11.8-161-fold) (Fig. 3c–e). Furthermore, BLI data indicated that the dissociation rate of MaP555-TMP increased from 4.88 × 10−7 s−1 of wild-type TMP-Tag to 0.13 s−1 of ReTag1 containing L28Q, R44L, and H45Q, a remarkable enhancement of 266,000-fold (Fig. 3d, e and Supplementary Fig. 14). Additionally, other fluorogenic probes, including MaP618m-TMP and MaP655-TMP, CPY-TMP, and SiR-TMP, also demonstrated significantly increased dissociation rates, enabling reversible binding within seconds to minutes (Fig. 3e and Supplementary Figs. 15–19).

We next examined the reversible binding performance in cell imaging. U2OS cells were transfected with Histone2B-SNAP-ReTag1 and stained with SiR-BG as the control probe for quantification. Consistent with in vitro results, fluorogenic ligands exhibited the rapid binding and releasing processes, enabling the reversible labeling within seconds to minutes in cellular imaging Fig. 3f, g and Supplementary Figs. 20, 21). Notably, the ReTag1 labeled with MaP555-TMP maintained its robust signal-to-noise ratio (47.8-folds) and comparably high brightness to the wild-type TMP-Tag (Fig. 3h).

We next investigated the fluorescence self-recovery of the ReTag1 with highly fluorogenic ligands in live-cell imaging. Fluorescence recovery after photobleaching (FRAP) experiments were conducted in live U2OS cells expressing Histone2B (H2B)-SNAP-ReTag1 considering slow diffusion of H2B. U2OS cells expressing wild-type TMP-Tag served as a control group, with 50% of the fluorescence signal photobleached in each cycle of a four-time repetitive bleaching process. As anticipated, more than 80% of the fluorescence signal of the wild-type TMP-Tag labeled with fluorogenic ligands was bleached, with minimal fluorescence recovery observed, confirming few interference from H2B diffusion. In contrast, ReTag1 with MaP555-TMP, MaP618m-TMP, MaP655-TMP, SiR-TMP and CPY-TMP demonstrated rapid fluorescence recovery through reversible labeling (Fig. 3i, j and Supplementary Figs. 22–24). Notably, ReTag1 labeled with the highly fluorogenic MaP618m-TMP and MaP655-TMP achieved significant fluorescence recovery within 1–2 min, retaining 68.7% and 72.9% of the fluorescence signal under identical bleaching conditions. In contrast, low fluorogenic ligands CPY-TMP and SiR-TMP showed considerably reduced recovery of 37.6% and 55.4%, respectively (Supplementary Figs. 22, 23). Similarly, an exchangeable Halo-Tag system labeled with SiR-Hy532 was also compared under the same condition, whereas 49.9% of the original signal retained (Supplementary Fig. 25). These results highlight the critical role of high fluorogenicity for efficient fluorescence recovery. The reduced performance is likely due to the high population of the zwitterionic form in unbound CPY-TMP, SiR-TMP, and SiR-Hy5, which can absorb excitation light and generate reactive oxygen species (ROS), leading to photodamage of both the protein tag and fluorophore.

Importantly, the ultra-low background signal of highly fluorogenic ligands allowed for a substantial increase in concentration, forming a larger ligand pool to enhance the efficiency of replenishing bleached fluorophores. The bleached fluorescence of the regenerative ReTag1 could recover from 72.9% to 89.9% when the concentration of MaP655-TMP was increased from 200 nM to 2 µM (Fig. 3i, j). Similar enhanced fluorescence self-recovery in higher concentration of probe was also observed in MaP618m-TMP (Supplementary Fig. 22). We thus believe that both ligand reversibility and high fluorogenicity are crucial for achieving efficient fluorescence regeneration in live-cell imaging.

Long-term live-cell nanoscopy with ReTag1 and highly fluorogenic ligands
STED nanoscopy creates fluorescent images with up to 40 nm resolution in living cells2, but requires an intense depletion laser that causes severe photobleaching, typically limiting the number of image frames to fewer than five. This has been a persistent bottleneck in applying STED nanoscopy for tracking cellular dynamics43. We thus examined the performance of ReTag1 and highly fluorogenic ligands in live-cell STED nanoscopy.

STED imaging of live U2OS cells expressing Vimentin-ReTag1-Halo stained with MaP655-TMP enabled the capture of individual filament structures with a full-width at half maximum (FWHM) of 43±5 nm, approximately five times lower than that from confocal imaging (Fig. 4a and Supplementary Fig. 27a). Moreover, the saturated stimulated emission power of MaP655-TMP labeled ReTag1 was determined to be Psat = 2.28 ± 0.03 mW by measuring fluorescence decay under varying STED laser powers at 775 nm (Fig. S27b)44,45. Importantly, over 15 frames of STED images of Vimentin were obtained using ReTag1 and MaP655-TMP, whereas less than 5 frames could be acquired with conventional Halo-Tag under the same conditions (Supplementary Fig. 26).

We further investigated the 3D STED microscopy of translocase of the outer membrane (TOM) 20 in the outer mitochondria membrane with ReTag1 and explored its potential in time-lapse live-cell STED microscopy (Fig. 4b and Supplementary Fig. 27c, d). The fluorescence of Halo-Tag with covalently bound SiR-HTL was severely bleached under the depletion laser of ~70 mW at 775 nm, limiting imaging to fewer than five frames. In contrast, the fluorescence intensity of MaP655-TMP remained at approximately 65% after 30 frames, surpassing the 50% retention observed with the reported Halo-Tag coupled with exchangeable ligand SiR-Hy5 (Fig. 4b, c). Remarkably, consistent with the findings from the FRAP experiment, increasing the concentration of MaP655-TMP to 2 µM maintained high contrast imaging and significantly enhanced photostability, achieving approximately 80% recovery after 30 frames. This resistance to photobleaching even allowed us to track mitochondria for more than 300 frames, revealing processes of fission, fusion and elongation of mitochondria with 80 nm resolution within 100 min (Fig. 4d, e and Supplementary Fig. 28; Supplementary Movie 1). To the best of our knowledge, this represents a distinctive achievement in the field.

To assess STED induced photodamage, we used a calcium sensitive fluorescent probe and observed no significant differences in intracellular calcium levels or cell morphology between STED irradiated cells and non-irradiated controls, indicating minimal acute cytotoxicity (Supplementary Fig. 29). Interestingly, we found a clear increase in ROS accumulation during prolonged STED imaging, particularly in cells labeled with fluorescent tags, compared to non-irradiated controls (Supplementary Fig. 30), probably causing mitochondrial morphological change (Fig. 4e). To support longer term STED imaging with reduced photodamage, the inclusion of ROS scavengers may help protect live cell structures from oxidative stress26.

We also investigate the performance of ReTag1 in SIM imaging, which provides super-resolution imaging (<100 nm) with ultrahigh speed (~ms) to track cellular dynamics46,47. The wash-free imaging of live U2OS cells expressing Vimentin-ReTag1-Halo labeled with MaP555-TMP, MaP618m-TMP, and MaP655-TMP revealed linear vimentin structures and dynamic movement at 91±14 nm FWHM, significantly higher than wide-field imaging (512±19 nm) (Supplementary Fig. 31 and Supplementary Movie 2). Additionally, the ReTag1 performs well with reported fluorogenic probes, enabling wash-free multi-color confocal and SIM imaging of various organelles, such as nucleus, F-actins, mitochondria, and vimentin simultaneously (Fig. 5a and Supplementary Figs. 32, 33).

We further confirmed the ReTag1’s utility in time-lapse SIM imaging by tracking mitochondrial translocase of the outer membrane (TOM20). The reversible and fluorogenic labeling of the ReTag1 with MaP618m-TMP provided better photobleaching resistance than Halo-Tag with covalently bound MaP618m-Halo (Fig. 5b). SIM imaging of live U2OS cells expressing TOM20-Halo-ReTag1 labeled with MaP655-TMP was successfully obtained with the FWHM of 118±17 nm (Fig. 5c). Labeling with MaP618m-TMP not only revealed the specific distribution of TOM20 on the outer mitochondrial membrane with the FWHM of 123±12 nm, but also showed mitochondrial dynamics such as fission, fusion, and elongation (Fig. 5d and Supplementary Fig. 34).

Notably, the reversible and fluorogenic labeling of the ReTag1 with highly fluorogenic ligands could be employed in MINFLUX, a microscopy method capable of achieving nanoscale localization based on the fluorescence “on-off” switching of fluorophores48. MINFLUX imaging of fixed U2OS cells expressing vimentin-ReTag1 with MaP655-TMP reveals individual vimentin filaments with an 5.0 nm localization precision (Fig. 5e and Supplementary Fig. 35).

Developing ReTagX for single-protein tracking
Single-protein studies are invaluable for understanding the operating principles of cellular machines but are hindered in living cells by limited brightness and tracking time due to photobleaching, even using small organic fluorophores49,50. Tandem repeat fluorescent reporters, such as SunTag28 and ArrayG29, have greatly advanced single-molecule tracking, but their large size (~1.3 MDa) may affect diffusion behavior and protein interaction. Inspired by Vale’s and Liphardt’s work28,29, we introduced a multiple-repeat ReTag1 fusion (ReTagX) with the highly fluorogenic MaP655 to enhance imaging contrast and tracking performance while retaining small tag size. We constructed ReTag4 (~78 kDa) by fusing four ReTag1 repeats with a (GGGGS)3 linker (Fig. 6a), and its incubation with MaP655-TMP resulted in a 55.5-fold increase in absorbance and a 170-fold increase in emission intensity, about four times higher than the original ReTag1 (Fig. 6b). The increased brightness was confirmed by live-cell U2OS imaging of H2B-GFP-ReTagX-expressing cells (Fig. 6c, d), where low laser power was sufficient for high-contrast imaging and improved long-term SIM imaging (Fig. 6e, f).

We further evaluated ReTag4’s performance in wash-free single-molecule imaging by purifying the protein and fixing it on a glass surface. The average brightness of single ReTag4 is significantly higher than ReTag1, especially at 500 nM, where it showed 2.9-fold higher brightness (Fig. 6g, h). Different from fluorescent imaging of bulk H2B–ReTag1, individual fluorophores bound to ReTag1 are visible for only tens of frames, after which the signal is lost due to dissociation or photobleaching, limiting its utility for long-term tracking due to signal loss (Fig. 6i). In contrast, ReTagX supports labeling with multiple fluorophores, showing a linear increase in fluorescence intensity with repeat number (Supplementary Fig. 36). ReTag4 exhibited high-frequency reversible labeling, particularly at 500 nM MaP655-TMP, enabling efficient multi-fluorophore exchange (Fig. 6i). Even when one fluorophore was bleached or exchanged, other fluorophores on ReTag4 continued to fluoresce, enabling extended single molecule tracking of over 100 frames in a streaming acquisition (Fig. 6j).

We next evaluated the performance of ReTag8 (~156 kDa), comprising eight ReTag1 repeats, for single-molecule tracking in live cells. To target the plasma membrane, ReTag8 was fused to a CAAX motif derived from the C-terminus of H-Ras (Supplementary Fig. 37a). In single-molecule imaging, the combination of ReTag8 with fluorogenic MaP655-TMP yielded a 5.2-fold increase in brightness compared to monomeric constructs (Supplementary Figs. 37b, c, and 38), enabling continuous maximum tracking of membrane proteins from approximately 50 frames to over 200 frames in living cells (Supplementary Fig. 37b–f).

We further fused ReTag8 to a membrane protein Tumor Necrosis Factor Receptor 2 (TNFR2) and conducted single-molecule tracking in live U2OS cells, using Halo-Tag and TMP-Tag8 (8xTMP-Tag repeats) as controls (Fig. 6k). Under identical imaging conditions (60% 640 nm laser, 60.4 W/cm², 50 ms exposure), TNFR2-ReTag8 exhibited a 5.5-fold increase in brightness relative to TNFR2-HaloTag (Fig. 6i and Supplementary Fig. 39). The measured diffusion coefficient of TNFR2–ReTag8 (0.28 ± 0.019 µm²/s) closely matched that of TNFR2-HaloTag (0.30 ± 0.028 µm²/s), indicating minimal perturbation to native mobility (Figs. 6m and S40). Moreover, ReTag8’s reversible fluorogenic labeling maintained a constant number of trajectories (>80%) for over 5 min observation, whereas fewer than 20% in Halo-Tag and TMP-Tag8 labeled proteins remained visible beyond 1 min under the same conditions (Fig. 6n and Supplementary Fig. 40; Supplementary Movie 3). The increased brightness and exchangeability of ReTag8 also enabled ~2.9-fold longer tracking durations compared to Halo-Tag (Fig. 6o and Supplementary Movie 4), thereby facilitating high-precision, long-term single-molecule imaging of membrane proteins in live cells.

Methods
The detailed procedures for the synthesis and characterization of all compounds, as well as their in vitro characterization and imaging experiments, are detailed in the Supplementary notes.

General experimental information for synthesis
All chemical reagents and dry solvents were purchased from commercial suppliers (Sigma-Aldrich, Bide Pharmatech, Adamas) and were used without further purification or distillation. Reactions were monitored by thin-layer chromatography (TLC) performed on TLC-aluminum sheets (Silica gel 60 F254, Merck) or liquid chromatograph-mass spectrometry (LC-MS 1260 Infinity II-6125, Agilent). Preparative RP-HPLC was performed on a 1260 Infinity II system (Agilent) on a C18 5 µm, 21.2 × 250 mm column (Supelco, flow rate 6 mL/min), solvent A: 0.1% v/v TFA in H2O, solvent B: acetonitrile. Typical gradient was from 10% to 95% B within 60 min. 1H NMR and high-resolution mass spectrometry (HRMS) were used to characterize all isolated compounds. Nuclear magnetic resonance spectra were recorded on a Bruker 400 MHz and 600 MHz instrument. All 1H NMR experiments are reported in units, parts per million (ppm), and were measured relative to the signals for residual dimethyl sulfoxide (2.51 ppm) in the deuterated solvent, unless otherwise stated. Multiplicities are shown as follows: s (singlet), d (doublet), t (triplet), q (quartet), dd (doublet of doublets), dt (doublet of triplets), and m (multiplets). HRMS were measured using ESI-QTOF mass spectrometer.

Protein-ligand docking simulation
The binding mode of the ligand to the protein was simulated using the CDDOCK module of Discovery Studio 4.0. The protein and ligand were assigned the CHARMM force field for the docking procedure. The crystal structure of eDHFR (PDB ID: 7NAE) was obtained from the Protein Data Bank (PDB), and the protein was prepared by removing water molecules and adding polar hydrogens. The protonation states of residues were carefully considered, with His residues protonated to HID, and Asp, Arg, Glu, and Lys residues set to their deprotonated states. The binding pocket was defined as the amino acid residues within 10 Å of the ligand. The three-dimensional structure of the ligand was generated using ChemBio3D Ultra software and energetically minimized with MMFF94, applying 5000 iterations and an RMS gradient of 0.05. Random ligand conformations were generated via high-temperature molecular dynamic simulations and random rotations. Following docking, the top 10 ranked protein-ligand conformations were selected, and the binding modes of the docked poses were visualized using PyMOL 1.1. All other docking parameters were set to default values.

General spectroscopic methods
Fluorescent probes for spectroscopy were prepared as stock solutions in DMSO and diluted such that the DMSO concentration did not exceed 1% (v/v). The measurements were taken in HEPES buffer (50 mM HEPES, 50 mM NaCl, pH 7.3) at ambient temperature (25 ± 2 °C) unless otherwise noted. Absorption spectra and emission spectra were measured on BioTek Cytation 5 Cell Imaging Multimode Reader using 384-well plate (Greiner, #781090) with optical bottom.

Plasmids and transfection
The plasmid pET51b(+) eDHFR (wt)-P30-Halo, pET51b(+) Halo-GGSX4-eDHFR (mt), pET51b(+) EGFP-ReTag1 and pET51b(+) EGFP-ReTag4 were constructed for protein expression and purification in vitro experiment. Plasmid pcDNA5/FRT/TO-SNAP-GGSX4-TMP-NLS and plasmid pcDNA5/FRT/TO-H2B-SNAP-ReTag1/pcDNA5/FRT/TO-H2B-SNAP-TMP host a C-terminal SNAP-tagged and eDHFR-tagged NLS and a N-terminal SNAP-tagged and eDHFR-tagged human H2B, respectively. Plasmid pLVX-Vimentin-ReTag1-Halo carries N-terminal ReTag1-tagged and Halo-tagged human vimentin, driven by the CMV promoter. Plasmid pcDNA5/FRT/TO-TOM20-HaloTag7-ReTag1 hosts a N-terminal Halo-tagged and human TOM20. Plasmid pcDNA5/FRT/TO-EGFP-TMP-CAAX/ pcDNA5/FRT/TO-EGFP-ReTag8-CAAX host a N-terminal ReTagX-tagged CAAX.
The appropriate fragments were inserted into the plasmid according to the protocol of Gibson. The synthesis of all primers and the subsequent sequencing of plasmids were executed by Sangon Biotech (Shanghai, China) following the standard protocol. The transfections were utilized the Lipofectamine 3000 transfection reagent (#L3000008). U2OS cells were seeded on the confocal plate (#781092) at an amount of 5 × 105 cells per well. After culturing for 24 h, plasmids were transfected into the cells. After 6 h, transfected cells were washed two times with 1X PBS and were maintained at fresh phenol red DMEM media with 10% FBS at least 12 h for the imaging. In single-molecule tracking, U2OS cells were imaged ~3 h post-transfection to maintain low protein expression levels. Specific plasmid sequences and construction methods are described in Supplementary notes.

General cell culture
U2OS cells (ATCC, BFN60700128) were cultured in Dulbecco’s Modified Eagle Medium (Gibco, #11960069) supplemented with 10% (v/v) FBS (Gibco #10099141C). The cells were incubated at 37 °C and 5% CO2. Cell lines were split every 3–4 days or at confluence and underwent regular mycoplasma testing. The cells were incubated for 1–2 days before imaging. Imaging medium contains phenol red-free Dulbecco’s Modified Eagle Medium (DMEM) media with 10% FBS.

General no-wash live cell confocal microscopy
After transfection, the U2OS cells were incubated with 200-500 nM probes for 0.5-2 h at 37 °C in a 5% CO2 atmosphere and directly imaged using confocal microscopy without washing steps. A Leica SP8 confocal scanning microscope equipped with a white light laser, HC PL APO CS2 ×20.0 objective lens and spectral HyD detector was used. Microscopy conditions: R500 derivatives, λex 505 nm, detection range 515–600 nm; TMR derivatives, λex 555 nm, detection range 565–650 nm; CPY derivatives, λex 615 nm, detection range 625–700 nm; SiR derivatives, λex 640 nm, detection range 650–750 nm. U2OS cells were seeded on the confocal plate and then transfected with pcDNA5/FRT/TO-SNAP-GGSX4-TMP-NLS, pcDNA5/FRT/TO-H2B-SNAP-ReTag1, pcDNA5/FRT/TO-TOM20-HaloTag7-ReTag1, pLVX-Vimentin-ReTag1-Halo, and pcDNA5/FRT/TO-H2B-GFP-ReTagX.

Live cell STED microscopy
Live cells STED nanoscopy was performed using an Abberior STEDYCON 775/RESOLFT QUAD scanning microscope (excitation lines: 405 nm, 485 nm, 561 nm, 640 nm; STED lines: 775 nm) equipped with a UPlanSApo 100x/1.4 oil immersion objective lens (Abberior Instruments, Germany). Imaging was performed with avalanche photodiodes (APD) and spectral detection. Fluorophores, microscope and imaging parameters are described in Supplementary notes. For STED imaging, “Red Hot” LUTs were applied for data representation. STED performance was assessed by measuring the Full Width at Half Maximum (FWHM) of single intermediate filament fibers revealed by Vimentin-ReTag1-Halo and TOM20-Halo-ReTag1. STED images were acquired with a pixel dwell time of 10 µs and a pixel size of 25 nm with 5 average line scans. Image analysis was performed with ImageJ by extracting fluorescence intensity profiles perpendicular to vimentin filaments and mitochondria outer membrane. Mean diameters were calculated from at least three individual images (n≥3) by fitting a Gaussian function, yielding FWHM.

Live cell SIM microscopy
Live-cell imaging was performed using a super-resolution structured illumination microscopy (SIM) system (Multi-SIM from Beijing NanoInsights-tech Co., Ltd.). Cells were imaged in a temperature-controlled chamber (37 °C) with a CO2 concentration maintained at 5%. The SIM system utilized a laser-based light source with multiple excitation wavelengths appropriate for the fluorophores used (488 nm, 561 nm, 594 nm and 640 nm). Super-resolution imaging was achieved using structured illumination in three dimensions (xy, xz, and yz planes). A total of 9 phases of structured illumination patterns were projected onto the sample, and each image stack was acquired sequentially at 100 nm lateral resolution. For z-stack imaging, optical sectioning was performed at 200–300 nm increments to obtain high-resolution 3D reconstructions. The SIM system used a high numerical aperture (NA) objective lens, typically 100x oil immersion (NA=1.4). SIM images were reconstructed using HVL1 constrained deconvolution algorithm or DFCAN and DFGAN deep learning algorithms. The final reconstructed images were processed with a resolution of approximately 100 nm in the lateral direction and 300–400 nm in the axial direction. For analysis, images were further processed using ImageJ, and customized plugins were applied to enhance signal-to-noise ratio and correct for drift. Image analysis was performed with ImageJ by extracting fluorescence intensity profiles perpendicular to vimentin filaments and mitochondria outer membrane. Mean diameters were calculated from at least three individual images (n≥3) by fitting a Gaussian function, yielding FWHM. More detailed experimental procedures are described in Supplementary notes.

MINFLUX microscopy
For MINFLUX, U2OS Vimentin-ReTag1-expressing cells were seeded on 18 mm #1.0 coverslips (41001118, Deckglaser), grown until a confluence of 60% was reached and chemically fixed by 2.4% paraformaldehyde (157-8, Electron Microscopy Sciences) and 0.1% glutaraldehyde (16220, Electron Microscopy Sciences) for 10 min. The cells were rinsed three times with PBS, followed by an additional brief wash in PBS and immersion in a 10 mM sodium borohydride (71320, Sigma) solution in PBS for 5 min. Cells were afterwards incubated with 2.5 nM MaP655-TMP in PBS with 100 µM NADPH. MINFLUX imaging was performed on an Abberior MINFLUX (Abberior Instruments GmbH, Göttingen, Germany) built on a motorized inverted microscope IX83 (Olympus, Tokyo, Japan) and equipped with 642 nm and 560 nm MINFLUX laser lines, 642 nm, 560 nm, 488 nm and 405 nm confocal lines. Imaging was performed with APDs in the spectral windows 650–685 nm and 685-720. Images were acquired using the default 2D imaging sequence, with an L in the last iteration step of 40 nm, and photon limit of 150 photons. Pinhole was set to 0.78 A.U., and excitation power at the periscope for the first iteration was ~ 96 µW. Four iterations were applied in the post-processing step to avoid false single-molecule emission events. To exclude detections originated from transient background, we set the maximum thresholds of 0.8 for the center frequency ratio (CFR) test and 1 kHz for the fluorescence imaging.39 Localizations from the same emission trace, i.e., with same TiD, further than three standard deviations with respect to the mean trace position were considered outliers and excluded from the trace. Only the traces containing at least 3 localizations within the first 180 min of the measurement were considered to calculate the localization precision. The experimental localization precision was estimated by co-aligning the mean values of all localizations obtained from individual emission traces fitted with a Gaussian function to estimate the overall standard deviation. For MINFLUX imaging, “Hot” LUTs were applied for data representation.

Sample preparation for single-molecule imaging
Clean coverslips were incubated with a mixture of PLL-(g)-PEG-biotin (50%, Susos) and PLL-(g)-PEG at 1:100,000 ratio for at least 2 h, followed by incubation with 1 mg/mL NeutrAvidin (Thermo Fisher Scientific) for 30 min to bind the biotin. For in vitro single-molecule imaging, the surfaces were then blocked with 1% BSA for 2 h, and finally, biotinylated purified protein was added and incubated for 15 min. In live-cell single-molecule imaging, the surfaces were incubated with 1 mg/mL RGD (pepnet) for 30 min, followed by blocking with 1% BSA for 2 h (cite https://doi.org/10.7554/eLife.67379). After each incubation step, the coverslips were washed three times with 1X TBS.
Cells were plated in 35 mm dishes and, upon reaching ~70% confluence, were transfected with 1 µg of plasmid DNA per dish for 5 h. The cells were then detached with trypsin and transferred onto pre-treated coverslips at a 1:3 split ratio. After 12 h of incubation in DMEM supplemented with 10% FBS at 37 °C and 5% CO2, live-cell imaging was performed.
For covalent Halo-Tag labeling, cells were stained with 1 nM SiR-HTL for 30 min at 37 °C and 5% CO2. Excess dye was removed by three washes with PBS, and the cultures were transferred to phenol-red–free DMEM supplemented with 10% FBS before imaging. For non-covalent TMP-Tag and ReTag labeling, Cells were washed three times with PBS and then incubated in phenol-red–free DMEM supplemented with 10% FBS and 200 nM MaP655-TMP for 30 min at 37 °C and 5% CO2 prior to experiments.

Single-molecule super-resolution microscopy and analysis
For in vitro single-molecule imaging, purified proteins ReTag1 and ReTag4 immobilized on the substrate were tracked using an Olympus IX86 TIRF microscope. The imaging was conducted at 30% 640 nm laser intensity (30.4 W/cm2) and 200 ms exposure time to monitor their brightness after binding with 200 nM MaP655-TMP. The average of the 10 minimum values from each trajectory was used as the background, which was then subtracted from the signal.
Live-cell single-molecule imaging was performed on the same TIRF setup with a 640 nm excitation laser. For ReTag8-CAXX tracking and TMP-CAXX, 10% laser intensity (10.3 W/cm2) and 50 ms exposure time were used after binding with 200 nM MaP655-TMP. For TNFR2 imaging and tracking, acquisition was carried out at 60% laser intensity (60.4 W/cm2) with 50 ms exposure time. For the MSD experiment the imaging parameters the exposure time was shortened to 30 ms (still at 60% laser power) to improve temporal resolution. To quantify the fluorescence intensities of individual foci, a circular region of interest (ROI) with a diameter of 0.5 µm was selected. The average background intensity, measured in distinct areas of the cell that lacked fluorescent foci, was then subtracted from the signal. In signal processing, the following trajectories were excluded: those with fewer than 5 frames or maximal pairwise displacement < 0.6 µm, as they were considered to represent non-specific dye binding signals.

Statistics and reproducibility
Samples sizes were chosen as indicated in the figure legends. No data were excluded from the analyses. The heatmap, bar and line charts in the figures were generated using GraphPad Prism 9.4.1 software. Figures 2f, j, o and 3h were analyzed by the two-tailed Student’s t-test. Significance was defined as a not significant (n.s.), **p < 0.01 and ****p < 0.0001. Figure 6d, h, l were analyzed by One-way ANOVA test. At least two independently replicates were performed for all experiments with similar results.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript introduces ReTagX, a bright, regenerative, and compact array tag system designed for rapid self-recovery of photobleached signals through reversible and fluorogenic labeling. It demonstrates the system’s effectiveness in long-term live-cell imaging and single-molecule tracking, addressing key challenges such as photobleaching, limited brightness, and large tag size.\nHowever, the approach of reversible labeling and extending fluorescence duration through molecular modifications are not novel (e.g., Zheng, et al. Recruiting Rate Determines the Blinking Propensity of Rhodamine Fluorophores for Super-Resolution Imaging. Journal of the American Chemical Society 2023, 145 (9), 5125-5133. DOI: 10.1021/jacs.2c11395.), In addition, the image quality (resolution and signal-to-noise ratio) of super resolution is not good enough. Given this, the manuscript lacks sufficient originality for publication in Nature Communications. A more suitable venue would be a specialized chemistry journal focused on advancements in fluorogenic probes and labeling techniques. Some suggestions would help further clarify the manuscript and improve the Discussion\n1. The manuscript outlines the limitations of conventional self-labeling and array tags (e.g., HaloTag, SNAP-tag, SunTag, ArrayG), but a more precise discussion of these drawbacks would strengthen the rationale for ReTagX.\n2. While the brightness enhancement is well demonstrated, a direct quantitative comparison of fluorescence recovery efficiency between ReTagX and existing exchangeable HaloTag ligands would provide stronger support for its benefits.\n3. The improved dissociation rate of ReTag1 compared to the wild-type TMP-tag is notable. However, it is unclear whether this increase affects binding specificity, particularly in live-cell environments. A discussion on potential trade-offs between rapid exchange and specificity is needed.\n4. It would be helpful to assess whether ReTag1 exhibits off-target interactions or unintended intracellular aggregation at high probe concentrations.\n5. The study briefly mentions phototoxicity but does not include data quantifying potential cellular stress from prolonged STED imaging.\n6. A viability assay (e.g., measuring cell survival after extended STED imaging with ReTagX compared to conventional tags) would confirm whether the system induces cytotoxic effects.\n7. The manuscript primarily focuses on mitochondrial and cytoskeletal proteins. Demonstrating ReTagX’s performance with proteins of varying diffusion rates, such as nuclear transcription factors or membrane receptors, would highlight its broader applicability.\n8. The study claims that ReTagX’s small size minimizes interference with protein function. However, functional assays are needed to confirm that tagged proteins retain their biological activity. Given the significant brightness enhancement of ReTag8, its potential impact on protein diffusion and cellular function should also be examined.\n9. The fluorescence enhancement of ReTag8 (~5.2×) is promising, but its potential for Förster resonance energy transfer (FRET) or other photophysical interactions is not addressed. Could the close spatial arrangement of fluorophores cause quenching or spectral crosstalk? This could affect quantitative fluorescence measurements. Fluorescence lifetime imaging microscopy (FLIM) should be performed to assess potential FRET effects. Additionally, testing the linearity of fluorescence intensity relative to fluorophore number would confirm accurate single-molecule quantification.\n10. Some figures (e.g., Fig. 3i) lack statistical analysis or error bars, making it difficult to assess reproducibility; the Fig. 2 appears oversaturated, which is not conducive to observing image details.\n11. The distinction between different fluorogenic ligands in figures should be clearer, such as through color-coding.\n12. From the Figure S10, we can see that the cell positions in the second to fourth images are consistent. Why did the position of the cell image change after a time interval of 1 hour, and the signal became stronger at an interval of 4 hours? The light radiation experiment results at different positions are not convincing, and it is unreasonable that the signal does not weaken but becomes stronger after 4 hours\n13. The ReTag1 probe does show good anti-bleaching properties in STED super-resolution imaging, but the resolution improvement is not very obvious (Figure 4). The author should measure the saturated stimulated emission power of the ReTag1 probe and compare it with the commercial probe ATTO647.\n14. Although the authors mentioned the toxicity of the probe in the conclusion, they did not provide a cell toxicity experiment. It is recommended that the authors add a cell toxicity experiment.", "reviewer_2": "In their work “A Bright and Regenerative Array Tag with Fluorogenic Ligands for Long-Term Live-Cell Nanoscopy and Single Molecule Tracking,” Si et al. investigate the potential of their newly developed ReTag system, which is based on a mutagenized TMP-tag system in combination with fluorogenic ligands. Unlike the commonly used SNAPtag or HaloTag, this TMP system has the advantage of being noncovalent from the outset. Newly tested mutations decreased the binding duration of the labeled protein while maintaining the high association rates of the TMP-tag. Combined with its small size, this system could represent a significant advancement in transient protein labeling, especially when used alongside more established systems for multi color approaches. The authors emphasize their ambition for the ReTag system across various modern microscopy techniques, which pose demanding requirements for any fluorescent protein or dye. I am excited about every new protein labeling system, especially when it allows for efficient intracellular fluorescent labeling in combination with transient binding. However, before I can fully recommend accepting this manuscript, I have some lingering questions, comments, and recommendations for experimental additions:\n  Perhaps I'm missing something, but the values in Figure 3e don't make any sense to me. The dissociation constant (KD) should simply be koff/kon, yet this does not appear to be reflected in the table.\n  Could the authors comment on why they chose to conduct FRAP experiments in live cells (Fig. 3i), especially considering they demonstrated in Figure 3f that it is possible to stain the same construct in fixed cells? While H2B's diffusion is slow, shouldn't there be concerns about how this might impact the results?\n  In lines 338 to 344, the authors present their first STED experiments using Vimentin-ReTag1-Halo and claim that this system, labeled with MaP655-TMP, outperforms the conventional HaloTag, producing 15 consecutive super-resolution images compared to one image with poor resolution obtained using the HaloTag under the same conditions. However, Figure S23, which is referenced as evidence, mentions Tom20-Halo on the left side, not Vimentin. Furthermore, while I do not doubt that the ReTag outperforms the standard HaloTag in long-term STED experiments, I find it hard to believe that a combination of a commonly used tag for STED, paired with one of the most widely used dyes for STED, would fail to yield even a single STED image with good resolution.\n  In lines 355 to 359, the authors assert that they can track mitochondria with a resolution of 80 nm over 100 minutes or more than 300 frames. However, upon reviewing Movie 1, which is intended to showcase this “unprecedented achievement,” I found it somewhat confusing. The movie cuts off after 50 minutes without any explanation, and it is only mentioned in the supplementary material that the cell had to be refocused after several time points. Does this not imply that the exposure was also reduced during those intervals? Furthermore, the entire sample appeared to drift out of the region of interest (ROI), resulting in only a small fraction of the mitochondria being illuminated between approximately 30 and 41 minutes. However, this sample drift is not evident in the figure. Additionally, there is a significant jump in frames shown in the figure between frames 130 to 200 and frame 270, indicating a different interval than previously used. Based on the appearance of the mitochondria after 270 frames, the cell appeared stressed due to phototoxicity and likely dying, yet this observation was not mentioned by the authors.\n  Regarding the authors' tracking experiments, I have several questions:\no There is no comparison to other more commonly used intracellular tagging systems, such as the HaloTag, SNAPtag, or even the reHaloTag and xHTL labeling systems. This absence makes it difficult to assess the acquired results effectively.\no As stated, imaging was conducted with 30% laser intensity and 200 ms exposure for brightness evaluation, and with 10% laser intensity and 50 ms exposure for fluorescence lifetime (this term is normally used in another context). These values lack context. The authors should include measured power densities or power at the back focal plane. Additionally, they should provide a range of power densities for the actual tracking experiments; currently, it's only noted as \"set to optimize signal-to-noise.\"\no Fusing multiple ReTags into an array is a good idea in principle, but it contradicts the general aim of having a small tag that does not hinder diffusion. Once the authors added the second ReTag, the size of their system became larger than that of the HaloTag (36 kDa compared to 33 kDa). What is the advantage here?\no In this context, diffusion coefficients or mean square displacements were never measured in any of the experiments.\no No software is mentioned for the tracking analysis.\no CAAX or other farnesylated proteins tend to diffuse quite rapidly, especially at 37 °C. The motion blur present at the reported 20 Hz acquisition speed may hinder tracking fidelity. I recommend repeating the experiments with a faster exposure time to improve the results.\no The increase in tracking duration appears to stem mainly from the ability to reduce laser power due to the enhanced brightness of multiple bound MaP-TMP ligand. At least, this is what can be inferred from reading the manuscript. I strongly recommend comparing ReTag8 to an 8xTMP-tag to demonstrate that the improved performance is due to rebinding. Additionally, please ensure that the same laser power is used for both experiments.\no Transiently binding probes show their greatest strengths in prolonged imaging experiments lasting minutes or even hours. The authors choose to label CAAX proteins after a very short expression time point, when only a few proteins are present inside the cell. The standard practice would be to wait for higher expression levels and label only a subfraction of all CAAX proteins. Following this, one could conduct prolonged imaging to demonstrate that, with the same ligand concentrations, ReTag8 outperforms the TMP-tag or even the 8xTMP-tag, with a much milder decrease in the number of localizations over time. This approach has been successfully employed with xHTL dyes and the reHaloTag, for example. I highly recommend that the authors conduct this experiment.", "reviewer_3": "This paper describes a bright and regenerative tag system (ReTag) and highly fluorogenic ligands that provide bright labeling with minimal background. The authors show how this tag allows greatly extended STED imaging or time-lapse SIM imaging of mitochondrial TOM20, revealing mitochondrial dynamics such as fission, fusion, and elongation. STED, multi-color SIM and MINFLUX images of Vimentin-ReTag1 all show impressive resolution. In addition, the compact ReTag8 array tag fused to CAAX is used as an example of tracking a membrane protein in living cells, which extends imaging duration by ~4-fold (up to 10 seconds) compared to the conventional TMP-Tag.\nThe images generated with the ReTag are impressive, so this tag and the ligands will be useful to the field. However, the paper is very dense, especially the first half, with 22 supplementary figures described along with only 3 main figures. It is difficult to navigate the first half of the results section as the many supplementary figures are typically very poorly described, e.g. lines 251 and 253 show single sentence descriptions of Figs S9 and S10. It would help the reader to explain the supplementary data properly in the text and give more information in the legend. For me, Fig. S9 needs more explanation.\nMy major concern relates to the idea that multimerization of ReTag could be a potential alternative to e.g. the SunTag array, which would be advantageous based on the smaller size of ReTag4/8. As the SunTag is primarily used for imaging translation, ideally translation would be imaged with ReTag4/8. However, my bigger concern relates to the idea that ReTag8 can be used to quantitate fluorescence as a proxy for e.g. translation when there is reversible exchange of bleached fluorophores. As far as I understand it, quantitation of translation using SunTag relies on stable binding of scFv so that, after correcting for bleaching, the changes in fluorescence can be directly attributed to changes in ribosome density. If ReTag8 is used and there is bleaching, then exchange of bleached fluorophores, how can variations in signal due to this exchange be separated from changes in e.g. ribosome density? In addition, it is described how ReTag8 extends the imaging time to ~10 seconds in living cells, but typical SunTag imaging durations are >30 mins for translation. Therefore, it is unclear to what extent multimerising ReTag will be useful as an alternative array, other than for short-term bright imaging. If the authors only mean ReTag4/8 is better than the conventional TMP-Tag, it would be better not to mention SunTag.\nMinor comments\nThe first Results section on the design seems a bit superfluous as it is background info rather than describing results.\nAre the fusion proteins in Fig 2e and Fig S4, referred to as SNAP-TMP-NLS and H2B-SNAP-TMP-NLS the same? It appears so from the text but the nomenclature is different.\nLine 227 – ‘MaP555-TMP show nice fluorogenicity (Fig. 2g and 2h)’ – is it not quite weak?\nFig 4, S23 – indicate the time between frames in the legend (and in other figures showing time frames)\nFig 4e – point out more clearly where the evidence of fission and fusion is (e.g. highlighted regions as in Fig 5d).\nFig. S26 – is it not supposed to also show the Hoechst staining in the Vimentin labelled cells?\nLine 431 – ‘ReTag1 displayed only transiently reversible labeling, limiting its utility for long-term tracking due to signal loss (Fig. 6i)’. How is this finding reconciled with the analysis of ReTag1 in Fig. 3, which describes how it allows reversible labeling within seconds to minutes in live-cell imaging, and fluorescence recovery within 1-2 minutes in the FRAP experiments? Perhaps this relates to the timing of the 400 frames, but again I couldn’t find information about this in the legend or methods.\nFig. 6i – explain how the number of fluorophores on individual proteins was calculated.\nLine 435 – ‘Even when one fluorophore was bleached or exchanged, other fluorophores on ReTag4 continued to fluoresce, enabling extended single molecule tracking to ~200 frames in a streaming acquisition (Fig. 6j)’. It looks like 100 frames for reliable tracking for 100 frames, so better to say this or tracking up to 200 frames.\nHow does ReTag8 perform relative to ReTag4 in terms of fluorescence intensity and brightness?\nFig. 6k-o – representative images of cells used for the analysis should also be shown."}, "author_response": "Point-By-Point Response to the Comments on the Manuscript “A Bright\nand Regenerative Array Tag with Fluorogenic Ligands for Long-Term\nLive-Cell Nanoscopy and Single-Molecule Tracking”\nWe are grateful to the reviewers for their insightful comments and\nsuggestions. In the revised manuscript we have attempted to address all\nthese points and changed the manuscript accordingly. Below we provide\na point-by-point response to the comments of the reviewers.\nReviewer #1 (Remarks to the Author):\n1. The manuscript introduces ReTagX, a bright, regenerative, and compact array tag\nsystem designed for rapid self-recovery of photobleached signals through reversible\nand fluorogenic labeling. It demonstrates the system’s effectiveness in long-term live-\ncell imaging and single-molecule tracking, addressing key challenges such as\nphotobleaching, limited brightness, and large tag size.\nResponse: We thank the reviewer’s comments on our bright, regenerative and compact\narray tag.\n2. However, the approach of reversible labeling and extending fluorescence duration\nthrough molecular modifications are not novel (e.g., Zheng, et al. Recruiting Rate\nDetermines the Blinking Propensity of Rhodamine Fluorophores for Super-Resolution\nImaging. Journal of the American Chemical Society 2023, 145 (9), 5125-5133. DOI:\n10.1021/jacs.2c11395.), In addition, the image quality (resolution and signal-to-noise\nratio) of super resolution is not good enough. Given this, the manuscript lacks sufficient\noriginality for publication in Nature Communications. A more suitable venue would be\na specialized chemistry journal focused on advancements in fluorogenic probes and\nlabeling techniques. Some suggestions would help further clarify the manuscript and\nimprove the Discussion.\nResponse: We thank the reviewer for the thoughtful comments and for highlighting the\nstudy by Zheng et al. (J. Am. Chem. Soc. 2023, 145, 5125). We fully acknowledge that\nreversible labeling and fluorophore engineering have been explored previously,\nincluding by the groups of Johnsson, Chu, and others (as summarized in Supplementary\nTable S1). However, these existing strategies often suffer from key limitations, such as\nlarge tag sizes, modest fluorogenic contrast, or slow labeling kinetics, which restrict\ntheir suitability for long-term, high-resolution imaging in live cells. Notably, none of\nthe existing reversible labeling platforms have been applied in a regenerative array-tag\nformat to achieve improved brightness and signal recovery for enhanced single-\nmolecule tracking. Additionally, while the work by Zheng et al. made valuable\ncontributions to modulating rhodamine dye blinking for single-molecule localization\nmicroscopy, their approach does not address the need for reversible, regenerative\ntracking or dynamic live-cell imaging without cumulative photobleaching.\nIn contrast, except for the optimization of reversible and fluorogenic labeling, our\nstudy introduces a bright, compact, and regenerative array tag system (ReTagX)\ndesigned for rapid fluorescence recovery following photobleaching via reversible and\nfluorogenic labeling. Our lead construct, ReTag1 (~18 kDa), when paired with the\nhighly fluorogenic MaP655-TMP, achieves a signal-to-noise ratio of up to 204-fold\nand enables full fluorescence recovery within 1 minute, even under wash-free live-cell\nconditions. We further developed a compact multimeric variant, ReTag8, which\nsignificantly enhanced brightness and extended single-molecule tracking durations and\nobservation windows, while minimizing perturbation to protein behavior.\nCrucially, the unique combination of high fluorogenicity, reversibility, and small\ntag size allows ReTagX to operate seamlessly across advanced imaging modalities (e.g.\nSIM, STED, and MINFLUX) without the need for washing. Specifically, the system\nreaches super-resolution benchmarks: STED FWHM of 43 ± 5 nm, SIM FWHM of 118\n± 17 nm, and MINFLUX localization precision of ~5.0 nm. Importantly, we\ndemonstrate that ReTagX enables: (1) Prolonged STED imaging of mitochondrial\nTOM20, extending usable imaging frames from fewer than 5 to over 300, thus allowing\nreal-time visualization of mitochondrial dynamics at ~80 nm resolution for over 100\nminutes; (2) Approximately 2.9-fold longer single-molecule tracking durations of\nmembrane proteins with over 5 min observation window compared to current methods\n(< 1 min). These advances collectively address long-standing challenges in live-cell\nsuper-resolution imaging and dynamic molecular tracking.\nIn response to the reviewer’s suggestions, we have further clarified these points in\nthe revised Discussion and included additional validation experiments, including cell\nviability under MaP655-TMP labeling (Fig. S10), comparison of fluorescence self-\nrecovery among ReTag1, ReTag4, and XHaloTag (Fig. S25), effects of STED laser\nillumination on cellular Ca²⁺ and ROS levels (Fig. S29 and S30), wash-free STED\nnanoscopy of vimentin filaments (Fig. 4a), brightness comparison between ReTag4 and\nReTag8 (Fig. S37), detailed single-molecule tracking of membrane protein TNFR2 (e.g.\nbrightness, diffusion, and duration) (Fig. 6k-o, Fig. S40). We sincerely thank the\nreviewer for their insightful and constructive comments, which have significantly\nimproved the quality and clarity of our manuscript. We respectfully hope that the\nadditional data and detailed responses provided in this revised version will merit\nfavorable reconsideration for publication in Nature Communications.\n3. The manuscript outlines the limitations of conventional self-labeling and array tags\n(e.g., HaloTag, SNAP-tag, SunTag, ArrayG), but a more precise discussion of these\ndrawbacks would strengthen the rationale for ReTagX.\nResponse: We sincerely thank the reviewer for this insightful comment. In response,\nwe have provided a detailed discussion of the limitations of conventional self labeling\nand array tag systems to better clarify the rationale for developing ReTagX (line 83-\n102 in revised manuscript and Supplementary Table S1).\nConventional self labeling tags such as HaloTag and SNAP tag attach fluorophores\ncovalently to the target protein in a one to one dye to tag ratio. Although widely used,\nthis irreversible labeling approach limits long term imaging, as the tagged protein\nbecomes nonfluorescent once the fluorophore is photobleached, particularly under high\nlaser power used in super resolution microscopy. To overcome this limitation, reversible\nlabeling systems such as reHaloTags, exchangeable HaloTag ligands (xHTLs), and\nsrTAGs with emissions from green to red have been developed to allow fluorescence\nsignal recovery. However, these systems often suffer from insufficient brightness,\nunsatisfied fluorogenicity, or slow binding kinetics (Table S1). In particular, the\nrelatively large size of reHaloTags and xHTLs limits their ability to be assembled into\nmultivalent array tag, reducing their applicability in regenerative imaging strategies.\nIn addition, another powerful tag system, array tags (e.g. SunTag, ArrayG, ArrayD,\nand HA-Tag) can increase fluorescence intensity by 10 to 18 fold and support short\ntracking durations of 1 to 30 seconds (Table S1). However, these systems typically\nexceed 1 mDa in molecular weight, which significantly increases the size of the protein\ncomplex. This can alter the diffusion behavior and functional properties of the target\nprotein, especially when fusing with the native protein size ranges from 10 to 160 kDa.\nIn contrast, ReTag1 is a compact tag of approximately 18 kDa that, when paired\nwith the highly fluorogenic MaP655-TMP dye, provides a signal to noise ratio of up\nto 204-fold and enables full fluorescence recovery within 1 minute in live cell imaging\nwithout the need for washing. This system significantly prolongs imaging, as\ndemonstrated by STED imaging of mitochondrial TOM20, which was extended from\nfewer than 5 frames to over 300 frames, allowing real time visualization of\nmitochondrial dynamics at approximately 80 nanometer resolution for over 100\nminutes. Furthermore, ReTagX array tags, constructed from multiple ReTag1 units,\nsupport exchangeable multi fluorophore labeling for regenerative imaging. Notably,\nReTag8 (~156 kD), provides a 5.5-fold increase in brightness during 5 minute imaging\nand enables single molecule tracking of membrane proteins such as TNFR2 with\napproximately 2.9-fold longer durations compared to HaloTag in live cell conditions.\nWe believe this revised and detailed comparison in the revised Table S1 now more\nclearly illustrates the motivation and distinct advantages of ReTagX over existing\nlabeling platforms.\n4. While the brightness enhancement is well demonstrated, a direct quantitative\ncomparison of fluorescence recovery efficiency between ReTagX and existing\nexchangeable HaloTag ligands would provide stronger support for its benefits.\nResponse: We thank the reviewer for the insightful comment. In accordance with the\nreviewers’ suggestion, we performed fluorescence recovery after photobleaching\n(FRAP) experiments using previously reported exchangeable Halo-Tag ligands (J. Am.\nChem. Soc., 2023, 145, 3075). As shown in Figure S25, the fluorescence recovery\nefficiency of SiR-Hy5 was only 49.9% after four rounds of photobleaching. In contrast,\nReTag1 and ReTag4 exhibited a markedly higher recovery efficiency of 88.9% and\n76.5%. We hypothesize that the lower recovery observed with SiR-Hy5 and similar\nprobes such as CPY-TMP and SiR-TMP (Fig. S24) may be attributed to the abundant\nzwitterionic population in the unbound state. This state can absorb light and generate\nreactive oxygen species, which may in turn damage the protein tag or fluorophore and\nreduce recovery efficiency. These results further support the advantages of highly\nfluorogenic ReTagX in delivering robust fluorescence recovery. The new data have\nbeen added and highlighted in the revised manuscript (line 307) and Supplementary\nFigure S25.\n5. The improved dissociation rate of ReTag1 compared to the wild-type TMP-tag is\nnotable. However, it is unclear whether this increase affects binding specificity,\nparticularly in live-cell environments. A discussion on potential trade-offs between\nrapid exchange and specificity is needed.\nResponse: We sincerely thank the reviewer for this thoughtful and important comment.\nAs correctly pointed out, enhancing the dissociation rate (k ) through TMP-Tag\noff\nmutations could potentially compromise binding affinity and specificity, which is an\nessential factor for reliable fluorescent labeling in live-cell environments. We fully\nagree that a careful balance between rapid exchange and binding specificity is critical.\nTo address this, we systematically screened several TMP-Tag mutations and\nidentified an optimized variant, ReTag1 with L28Q, R44L, and H45Q mutations, that\nachieves a remarkable 266,000-fold increase in k while with suitable binding affinity\noff\nwith TMP-ligands with hundreds of nM. In addition, the fluorogenic property is key\nwhen modifying specific binding and exchange binding. The exceptional fluorogenicity\n(>100) minimizes nonspecific background and produces fluorescence only upon\nspecific binding to ReTag1, thereby preserving labeling specificity even under wash-\nfree imaging conditions. Furthermore, the high fluorogenicity permits the use of\nelevated probe concentrations to promote efficient probe exchange, without introducing\nbackground signal.\nImportantly, we validated the specificity of ReTag1 in live-cell imaging\nexperiments. Using fluorogenic MaP555-TMP, MaP618-TMP, and MaP655-TMP\nconjugates, we observed clear, high-contrast labeling of nuclear H2B, mitochondrial\nTOM20, and cytoskeletal vimentin targets in confocal, SIM, and STED imaging, all\nperformed without washing steps (Fig. 3-5 and S31-S34). These results demonstrate\nthat ReTag1 maintains excellent labeling specificity despite its enhanced exchange rate.\nAs requested, we have added a detailed discussion on the trade-offs between rapid\nexchange and specificity, along with supporting data, in the discussion of revised\nmanuscript (line 265-270).\n6. It would be helpful to assess whether ReTag1 exhibits off-target interactions or\nunintended intracellular aggregation at high probe concentrations.\nResponse: We sincerely thank the reviewer for this valuable comment. To address this,\nwe have conducted live-cell imaging at high probe concentrations (2-10 μM) using\nconfocal microscopy (Fig. 2e and Fig. S4: 2-10 μM MaP618m-TMP; Fig. 2n: 10 μM\nMaP655-TMP) and STED imaging (Fig. 4e: 2 μM MaP655-TMP) under wash-free\nconditions. Importantly, we did not observe detectable off-target interactions or\nintracellular aggregation, even at the highest probe concentrations tested. We attribute\nthis to the high fluorogenicity of ReTag and MaP probes, which effectively minimize\nnonspecific background and aggregation. As requested, we have added a more explicit\ndescription of this point in the revised manuscript (Line 181 and line 367).\n7. The study briefly mentions phototoxicity but does not include data quantifying\npotential cellular stress from prolonged STED imaging.\nResponse: We sincerely thank the reviewer for this thoughtful and important comment.\nWe fully agree that prolonged STED imaging can induce phototoxic stress in live cells,\nas illustrated in Fig. 4e of our study and previously reported in the literature (Nat.\nMethods, 2010, 7, 731; Cell Rep., 2015, 10, 778).\nIn response to the reviewer’s suggestion, we performed additional experiments to\nassess potential phototoxic effects following extended STED imaging sessions using\nReTag1 labeling. Specifically, we monitored cellular morphology and stress by\nmeasuring intracellular calcium and reactive oxygen species using a calcium sensitive\nfluorescent probe and a ROS sensitive dye respectively (Fig. S10, S29, and S30).\nOur results showed no significant changes in intracellular calcium levels or\ncellular morphology between STED irradiated cells (imaged with STED laser power at\n100 percent, approximately 520 milliwatts) and non-irradiated controls, suggesting\nminimal acute cytotoxicity (Fig. S29). However, we observed a clear increase in ROS\nlevels following prolonged STED illumination, particularly in cells labeled with\nfluorescent tags, compared to non-irradiated controls (Fig. S30). While this ROS\nincrease did not result in overt cell death or morphological disruption, it may impact\ncellular function. For example, elevated ROS could contribute to the mitochondrial\nchanges observed during long term STED imaging, as seen in Figure 4e. Notably, the\nimplementation of reactive oxygen species (ROS) scavengers, advanced imaging\nplatforms (e.g., Abberior Facility STED), and optimized cell culture conditions has\nbeen shown to mitigate phototoxicity and better preserve cellular integrity during\nprolonged imaging sessions (Nat. Methods, 2018, 15, 755). We have incorporated new\ndata and discussion and the new supporting data into the revised manuscript (line 373)\nand Figures S10, S29, and S30.\n8. A viability assay (e.g., measuring cell survival after extended STED imaging with\nReTagX compared to conventional tags) would confirm whether the system induces\ncytotoxic effects.\nResponse: We sincerely thank the reviewer for this thoughtful and important\nsuggestion. A previous study by Bewersdorf’s group has demonstrated that although\nSTED illumination does not alter Ca²⁺ levels, ER morphology, or cell shape, it can\nincrease the cell death rate from 30% to 60% after 24 hours (Nat. Methods, 2018, 15,\n755). To investigate whether ReTagX induces cytotoxic effects under extended STED\nimaging, we performed additional experiments to assess cell viability and cellular stress\nresponses (Fig. S10, S29, and S30).\nAs shown in the viability assay (Fig. S10), MaP655-TMP at concentrations up to\n10 μM exhibited no detectable cytotoxicity, confirming the low toxicity of the\nfluorogenic probe even at high concentrations. To further evaluate cellular stress under\nSTED conditions, we assessed intracellular calcium and reactive oxygen species (ROS)\nlevels, which are sensitive indicators of early cellular stress responses. Using a calcium\nsensitive fluorescent probe, we found no significant changes in cytoplasmic Ca²⁺ levels\nor cellular morphology in cells illuminated with the STED laser at 520 milliwatts for\n40 minutes (Fig. S29). These findings are consistent with previously reported results\n(Nat. Methods, 2018, 15, 755), and suggest minimal acute cytotoxicity under our\nimaging conditions.\nHowever, we observed an increase in ROS levels following prolonged STED\nimaging, particularly in cells labeled with fluorescent tags, compared to non-irradiated\ncontrols (Fig. S30). While this elevation in ROS did not result in overt morphological\nchanges or immediate cell death, it may affect cellular function over time. This\nobservation underscores the importance of considering oxidative stress in long term\nlive cell imaging. As previously reported (Nat. Methods, 2018, 15, 755), the use of\nreactive oxygen species (ROS) scavengers, advanced imaging platforms (e.g., Abberior\nFacility STED), and optimized cell culture conditions can effectively mitigate\nphototoxicity and preserve normal cell viability during prolonged imaging. We have\nincorporated these new data and corresponding discussion into the revised manuscript\n(line 373), as well as Figures S10, S29, and S30.\n9. The manuscript primarily focuses on mitochondrial and cytoskeletal proteins.\nDemonstrating ReTagX’s performance with proteins of varying diffusion rates, such as\nnuclear transcription factors or membrane receptors, would highlight its broader\napplicability.\nResponse: We sincerely thank the reviewer for this valuable suggestion. To broaden the\nscope of our study as the reviewer suggested, we fused the ReTag8 array to Tumor\nNecrosis Factor Receptor 2 (TNFR2, with the intracellular domain deleted) and\nconducted single molecule tracking in live U2OS cells. Under identical imaging\nconditions (60% 640 nm laser intensity, 60.4 W/cm², 50 ms exposure time), TNFR2\nlabeled with ReTag8 exhibited a 5.5-fold increase in brightness compared to TNFR2\nfused to Halo-Tag, a widely used self-labeling tag conjugated with small molecule\nfluorophores.\nImportantly, the diffusion coefficient of TNFR2 ReTag8 was measured at 0.28 ±\n0.019 µm²/s, which is in close agreement with the previously reported value of ~0.30\nµm²/s (Biochim. Biophys. Acta, 2010, 1798, 1081). This indicates that ReTag8 does not\nappreciably perturb the diffusion dynamics of the membrane protein. In contrast,\nalthough systems like ArrayG can enhance signal brightness and tracking duration,\nthey often interfere with native protein mobility with significantly reduced diffusion\ncoefficients (~0.14 µm²/s). Moreover, ReTag8’s reversible fluorogenic labeling\nmaintained a constant number of trajectories (>80%) for over 5 minutes of continuous\nobservation without any wash or exchange steps, whereas fewer than 20% of Halo-Tag-\nlabeled proteins remained visible under the same conditions (new added Fig. 6n and\nS40b).\nThese new results highlight that ReTag8 provides superior brightness for protein\nlabeling with minimal interference to protein mobility. The new data and discussion\nhave been incorporated into the revised manuscript (line 470) and Supplementary\nFigure 40.\n10. The study claims that ReTagX’s small size minimizes interference with protein\nfunction. However, functional assays are needed to confirm that tagged proteins retain\ntheir biological activity. Given the significant brightness enhancement of ReTag8,\nits potential impact on protein diffusion and cellular function should also be examined.\nResponse: We sincerely thank the reviewer for this important and constructive\nsuggestion. We fully agree that ensuring the functional compatibility of tagged proteins\nis essential, especially for array systems. While ReTag4 (~78 kD) and ReTag8 (~156\nkD) is indeed larger than commonly used tags such as GFP or HaloTag, it remains\nsignificantly smaller than existing array systems (e.g., SunTag, ArrayG; >1 MDa),\nwhich are known to perturb protein dynamics (Nat. Chem. Biol. 2019, 15, 401).\nTo support the biocompatibility of ReTagX, we have fused ReTag 1 and ReTag4 to\nseveral representative proteins across cellular compartments, e.g. H2B (nucleus),\nTOM20 and MitoNEET (mitochondria), and vimentin (cytoskeleton), and confirmed\ncorrect subcellular localization without observable disruption to protein function or\ncellular phenotype (Fig. 4-5 and Fig. 6a-d). To further evaluate the impact on protein\ndynamics, we tested ReTag8 in the context of TNFR2, a membrane protein with well-\ncharacterized diffusion behavior. Compared to TNFR2-HaloTag, TNFR2-ReTag8\nshowed a 5.5-fold increase in brightness (Fig. 6i). Importantly, single-molecule tracking\nrevealed a diffusion coefficient of 0.28 ± 0.019 µm²/s for TNFR2–ReTag8, closely\nmatching the reported value of 0.30 ± 0.028 µm²/s for TNFR2–HaloTag (Fig. 6m–n;\nFig. S40), indicating minimal perturbation to mobility. This contrasts with existing\nmultivalent tags such as ArrayG, which—despite increased brightness—have been\nshown to significantly impair protein diffusion (~0.14 µm²/s) due to their large size.\nTogether, these results suggest that ReTagX achieves substantial brightness\nenhancement while minimizing disruption to protein dynamics and cellular function.\nWe have revised the manuscript and Supplementary Figure S40 to incorporate this\ndiscussion (line 470).\n11. The fluorescence enhancement of ReTag8 (~5.2×) is promising, but its potential for\nFörster resonance energy transfer (FRET) or other photophysical interactions is not\naddressed. Could the close spatial arrangement of fluorophores cause quenching or\nspectral crosstalk? This could affect quantitative fluorescence measurements.\nFluorescence lifetime imaging microscopy (FLIM) should be performed to assess\npotential FRET effects. Additionally, testing the linearity of fluorescence intensity\nrelative to fluorophore number would confirm accurate single-molecule quantification.\nResponse: We sincerely thank the reviewer for this insightful comment. We fully agree\nthat labeling with multiple dyes can potentially lead to self-quenching via mechanisms\nsuch as homo-FRET, particularly when fluorophores are spaced within <10 nm (Chem\nSoc Rev, 2020, 49, 5110). We appreciate the reviewer’s suggestion to use FLIM to\nevaluate potential FRET effects; however, this measurement is currently limited by the\nlack of available instrumentation in our facility. Nevertheless, it is worth noting that\nprevious studies using systems like SunTag, ArrayG, and fluorescent light-up aptamers\n(FLAPs) have shown that introducing multiple dyes at sub-10 nm distances can\nachieve >10-fold brightness enhancement without significant quenching. This is\nprobably because self-quenching depends not only on distance but also on the dipole\norientation of fluorophores (Phys. Chem. B 1999, 103, 6839; J. Biophotonics. 2018, 11,\ne201700366.). In our study, ReTag1, ReTag2, and ReTag4 exhibited a linear increase\nin fluorescence intensity (Fig. S36). Furthermore, single-molecule imaging of purified\nReTag4 immobilized on glass revealed discrete stepwise intensity increments,\nconsistent with minimal quenching effects (Fig. 6i). Additionally, ReTag8 fusions\nexhibited a 5.5-fold increase in brightness at the single-protein level compared to the\nwidely used Halo-Tag, although we cannot fully exclude the possibility of quenching\ndue to FRET effects.\nWe agree with reviewer that precise stoichiometric quantification remains\nchallenging due to the reversible labeling of ReTagX and potential quenching. However,\nit is worth noting that we mainly focus on the increased brightness and long-term\nimaging due to the unique fluorogenic and reversible binding property. A discussion of\npotential FRET effects and their implications has been added to the revised manuscript\n(Line 544-546) and supplementary file (Fig. S36).\n12. Some figures (e.g., Fig. 3i) lack statistical analysis or error bars, making it difficult\nto assess reproducibility; the Fig. 2 appears oversaturated, which is not conducive to\nobserving image details.\nResponse: We thank the reviewer for pointing this out. In Fig. 3i, the error bars were\npreviously displayed using a light color, which may have made them difficult to\ndistinguish. We have now recolored them in the revised Fig. 3j to improve visibility and\nfacilitate assessment of data reproducibility.\nThe images in Fig. 2 were acquired under standard imaging conditions. The\napparent oversaturation resulted from post-acquisition brightness enhancement in\nImageJ, which was applied uniformly across the image to better visualize low\nbackground signals in the cytosol. Importantly, no nonspecific labeling or signal\naggregation was observed, even after increasing brightness. To address the reviewer’s\nconcern and ensure transparency, we have now included the original, unprocessed\nimages in the revised Supplementary Information (Fig. S4, S5 and S9) for direct\ncomparison and clearer assessment of image details and background.\n13. The distinction between different fluorogenic ligands in figures should be clearer,\nsuch as through color-coding.\nResponse: We thank the reviewer for this helpful suggestion to improve clarity. In the\nrevised figures, the different fluorogenic ligands have been redrawn with clearer\ndistinction. Key structural differences, such as O, C(CH₃)₂, and Si(CH₃)₂, are now color-\ncoded to make them easier to differentiate and improve overall readability.\n14. From the Figure S10, we can see that the cell positions in the second to fourth\nimages are consistent. Why did the position of the cell image change after a time\ninterval of 1 hour, and the signal became stronger at an interval of 4 hours? The light\nradiation experiment results at different positions are not convincing, and it is\nunreasonable that the signal does not weaken but becomes stronger after 4 hours\nResponse: We sincerely thank the reviewer for pointing this out. The cell imaging\nshown in Figure S10 was performed in fixed cells. Due to limited instrument\navailability and scheduling constraints, we imaged different cells at each time point\nrather than repeatedly imaging the same cell over the 4-hour period. Additionally, some\ncells appeared brighter, likely due to variability in expression levels. To minimize the\ninfluence of expression variability of different cells on fluorescence quantification, we\nhave co-expressed ReTag with SNAP-Tag and stained the cells with SiR-BG (shown in\nred) as an internal control. In the revised manuscript, we present the normalized\nfluorescence ratio, which demonstrates that the signal remains stable even after 4 hours.\nThe relatively stable fluorescence signal from TMP-Tag is likely due to a\nconformational change that forms a closed state in the presence of NADPH and TMP\nderivatives, resulting in extremely high binding affinity (~pM) and slow release kinetics\n(Biochim. Biophys. Acta, 1994, 1207, 74; Structure, 2009, 17, 386). This mechanism\nhelps maintain signal stability over extended periods, as reflected in our new\nquantification data included in the revised Supplementary file (Fig. S12). We have\nadded this explanation to the revised manuscript (Line 256).\n15. The ReTag1 probe does show good anti-bleaching properties in STED super-\nresolution imaging, but the resolution improvement is not very obvious (Figure 4). The\nauthor should measure the saturated stimulated emission power of the ReTag1 probe\nand compare it with the commercial probe ATTO647.\nResponse: We sincerely thank the reviewer for this insightful comment. In response,\nwe have repeated the STED images in Fig. 4a by applying stronger laser power (~219\nmW) to improve image quality. With this optimized approach, we achieved a full width\nat half maximum (FWHM) of 43 ± 5 nm for vimentin filaments in live cells under wash-\nfree STED conditions, representing the leading enhanced spatial resolution and imaging\nquality compared with reported work (Nat Commun, 2022, 13, 2264; J. Am. Chem. Soc.\n2023, 145, 3075). In this work, we intentionally avoided using saturated STED power\nto minimize photobleaching and enable longer-term imaging at nanoscale resolution (~\n80 nm) from 5 frames to 300 frames, which is a key advantage of our system.\nAs ATTO647, due to its hydrophilic structure, is not suitable for live-cell\napplications, we instead compared MaP655-TMP with SiR-Halo, a widely used\ncommercial STED probe compatible with live-cell imaging (J. Am. Chem. Soc. 2019,\n141, 7, 2770). As shown in the revised Fig. 4b and S26, under wash-free conditions,\nSiR-Halo exhibited poor resolution, rapid photobleaching, and high background, with\nsignal typically detectable for only 1–5 frames. In contrast, ReTag1 labeled with\nMaP655-TMP enabled improved spatial resolution (FWHM: 43–80 nm) and sustained\nhigh-contrast imaging for 15-300 frames under wash-free conditions. We have added\nnew data in the revised supplementary file (Fig. S26).\n16. Although the authors mentioned the toxicity of the probe in the conclusion, they did\nnot provide a cell toxicity experiment. It is recommended that the authors add a cell\ntoxicity experiment.\nResponse: We thank the reviewer for this valuable suggestion. In response, we\nconducted a cytotoxicity evaluation using the CCK-8 assay. As shown in Fig. S10,\nU2OS cells incubated with the probe at 2 μM and 10 μM for 12 and 24 hours exhibited\nno significant reduction in cell viability, indicating negligible cytotoxicity under the\ntested conditions. These results confirm that the fluorogenic probe is highly\nbiocompatible and suitable for live-cell applications. The new data have been added in\nthe revised supplementary file (Fig. S10).\nReviewer #2 (Remarks to the Author):\n1. In their work “A Bright and Regenerative Array Tag with Fluorogenic Ligands for\nLong-Term Live-Cell Nanoscopy and Single Molecule Tracking,” Si et al. investigate\nthe potential of their newly developed ReTag system, which is based on a mutagenized\nTMP-tag system in combination with fluorogenic ligands. Unlike the commonly used\nSNAPtag or HaloTag, this TMP system has the advantage of being noncovalent from\nthe outset. Newly tested mutations decreased the binding duration of the labeled protein\nwhile maintaining the high association rates of the TMP-tag. Combined with its small\nsize, this system could represent a significant advancement in transient protein labeling,\nespecially when used alongside more established systems for multi color approaches.\nThe authors emphasize their ambition for the ReTag system across various modern\nmicroscopy techniques, which pose demanding requirements for any fluorescent protein\nor dye. I am excited about every new protein labeling system, especially when it allows\nfor efficient intracellular fluorescent labeling in combination with transient binding.\nHowever, before I can fully recommend accepting this manuscript, I have some\nlingering questions, comments, and recommendations for experimental additions:\nResponse: We sincerely thank the reviewer for their thoughtful and encouraging\nsummary of our work. We truly appreciate the reviewer’s enthusiasm for new protein\nlabeling systems and their constructive comments. In response, we have conducted\nadditional experiments, cell viability under MaP655-TMP labeling (Fig. S10),\ncomparison of fluorescence self-recovery among ReTag1, ReTag4, and XHaloTag (Fig.\nS25), effects of STED laser illumination on cellular Ca²⁺ and ROS levels (Fig. S29 and\nS30), wash-free STED nanoscopy of vimentin filaments (Fig. 4a), brightness\ncomparison between ReTag4 and ReTag8 (Fig. S38), detailed single-molecule tracking\nof TNFR2 (e.g. brightness, diffusion, and duration) (Fig. 6k-o, Fig. S39-40). We have\nalso expanded our discussion to improve clarity on points as suggested. We believe that\nthe revised manuscript has been substantially strengthened thanks to the reviewer’s\ninsightful recommendations.\n2. Perhaps I'm missing something, but the values in Figure 3e don't make any sense to\nme. The dissociation constant (KD) should simply be koff/kon, yet this does not appear\nto be reflected in the table.\nResponse: We thank the reviewer for this insightful comment. The KD values presented\nin Figure 3e were determined through steady-state binding analysis using the Sartorius\nOctet R8 system, a widely used instrument for measuring binding affinities. In this\nmethod, equilibrium responses (Req) at various analyte concentrations are fit to a 1:1\nLangmuir binding isotherm using the following equation:\n𝑅𝑚𝑎𝑥 ×[𝐶𝑜𝑛𝑐.]\n𝑅𝑒𝑠𝑝𝑜𝑛𝑠𝑒 =\n𝐾D+[𝐶𝑜𝑛𝑐.]\nwhere Response is the equilibrium binding signal, Rmax is the maximum binding\ncapacity, [Conc.] is the analyte concentration, and KD is the equilibrium dissociation\nconstant.\nThis steady-state model allows for direct extraction of KD from equilibrium\nbinding levels across multiple concentrations, independent of the kinetic parameters\nkon and koff. As such, the KD values reported in Figure 3e are derived from equilibrium\nisotherms and not calculated from k /k . To prevent further confusion, we have now\noff on\nclarified this KD derivation method in the Methods section and added a corresponding\nnote in the figure legend.\n3. Could the authors comment on why they chose to conduct FRAP experiments in live\ncells (Fig. 3i), especially considering they demonstrated in Figure 3f that it is possible\nto stain the same construct in fixed cells? While H2B's diffusion is slow, shouldn't there\nbe concerns about how this might impact the results?\nResponse: We thank the reviewer for this thoughtful and important comment. We chose\nto conduct FRAP experiments in live cells because the primary goal of our study is to\nestablish a reversible labeling platform that functions robustly under live-cell imaging\nconditions, where labeling kinetics, probe concentration, environmental variability, and\ncellular toxicity are critical considerations.\nAs the reviewer correctly notes, histone H2B is a slow-diffusing nuclear protein,\nmaking it a suitable model for evaluating fluorescence recovery driven by probe\nexchange without significant interference from protein mobility. To further ensure that\nthe observed recovery is due to probe dynamics rather than protein diffusion, we\nincluded controls using irreversible labeling systems such as SNAP-Tag with SiR-BG\nand eDHFR-tag with TMP-based probes, which showed minimal fluorescence recovery\nunder identical imaging conditions. These results highlight that the robust recovery\nobserved with ReTag is indeed due to its dynamic and reversible labeling mechanism,\nnot from H2B mobility. We have clarified this rationale and added appropriate\ndiscussion in the revised manuscript (Line 295 and 300).\n4. In lines 338 to 344, the authors present their first STED experiments using Vimentin-\nReTag1-Halo and claim that this system, labeled with MaP655-TMP, outperforms the\nconventional HaloTag, producing 15 consecutive super-resolution images compared to\none image with poor resolution obtained using the HaloTag under the same conditions.\nHowever, Figure S23, which is referenced as evidence, mentions Tom20-Halo on the\nleft side, not Vimentin. Furthermore, while I do not doubt that the ReTag outperforms\nthe standard HaloTag in long-term STED experiments, I find it hard to believe that a\ncombination of a commonly used tag for STED, paired with one of the most widely used\ndyes for STED, would fail to yield even a single STED image with good resolution.\nResponse: We sincerely thank the reviewer for the careful observation and for pointing\nout the labeling error in Figure S23. The images originally labeled as Tom20-Halo\nactually depict vimentin structures. We have now corrected the figure labeling and\nreorganized the panel accordingly to avoid confusion. As shown in the previous\nsupplementary file (Fig. S23), SiR-Halo performs poorly under wash-free STED\nconditions. This limitation likely arises from residual unbound fluorophores at the 200\nnM concentration, which contribute to elevated background fluorescence under high\ndepletion laser power; therefore, washing steps are usually required in STED\nnanoscopy (Nat. Chem. 2013, 5, 132).\nTo further compare the performance between Halo-Tag and ReTag1, we repeated\nthe STED experiments with SiR-Halo under optimized conditions, including a thorough\nwashing step after probe incubation. As shown in revised Fig. S26, even under these\noptimized conditions, Vimentin-Halo labeled with SiR-Halo sustained only less than 5\nframes of STED imaging due to rapid photobleaching. In contrast, Vimentin-ReTag1\nlabeled with MaP655-TMP achieved 10-15 consecutive super-resolution frames,\nowing to the exchangeable and fluorogenic nature of the ReTag system, which\nminimizes background signal and enables longer imaging durations under the same\nlaser conditions. We have updated the figure, corrected the labeling, and added this\ndiscussion to the revised manuscript (Line 354) and supplementary file (Fig. S26).\n5. In lines 355 to 359, the authors assert that they can track mitochondria with a\nresolution of 80 nm over 100 minutes or more than 300 frames. However, upon\nreviewing Movie 1, which is intended to showcase this “unprecedented achievement,”\nI found it somewhat confusing. The movie cuts off after 50 minutes without any\nexplanation, and it is only mentioned in the supplementary material that the cell had to\nbe refocused after several time points. Does this not imply that the exposure was also\nreduced during those intervals? Furthermore, the entire sample appeared to drift out\nof the region of interest (ROI), resulting in only a small fraction of the mitochondria\nbeing illuminated between approximately 30 and 41 minutes. However, this sample drift\nis not evident in the figure. Additionally, there is a significant jump in frames shown in\nthe figure between frames 130 to 200 and frame 270, indicating a different interval than\npreviously used. Based on the appearance of the mitochondria after 270 frames, the\ncell appeared stressed due to phototoxicity and likely dying, yet this observation was\nnot mentioned by the authors.\nResponse: We thank the reviewer for the careful assessment and insightful comments\nregarding the long-term STED imaging experiment presented in Movie 1. As this\nexperiment involves continuous imaging of living cells for over 100 minutes, cellular\nmovement and mitochondrial repositioning are indeed unavoidable. During imaging,\nwe observed mitochondrial migration away from the field of view, likely in response to\nphotostimulation. To maintain consistent tracking of the same mitochondria, the\nimaging position was realigned at 17, 42, and 60 minutes, with each adjustment\nrequiring approximately 5 seconds. A distinctive circular mitochondrion, marked with\na white arrow, was used as a positional reference to ensure continuity.\nAs the reviewer rightly noted, continuous imaging under STED laser can induce\nphototoxicity (Nat. Methods, 2018, 15, 755). Meanwhile, some degree of photodamage\nmay also result from the suboptimal live-cell conditions, as the STEDYCON system\nused in this study is limited to imaging buffer and does not support full cell culture\nconditions (e.g. controlled temperature, CO₂, and humidity). In addition, we conducted\nadditional experiments (Fig. S30) assessing reactive oxygen species (ROS) levels under\nprolonged STED exposure. These results confirm that continuous STED illumination\nincreases ROS levels, particularly in fluorescent labeled cells. While this ROS elevation\ndid not cause overt cell death during the imaging period, it may have induced cellular\nstress, leading to changes in mitochondrial morphology.\nWe have now discussed this limitation more explicitly in the revised manuscript and\nincluded recommendations for minimizing phototoxicity in future studies, such as the\nuse of ROS scavengers (Nat. Methods, 2018, 15, 755), improved imaging platforms\n(e.g., Abberior Facility STED), and optimized cell culture conditions. We have updated\nthe main text and figure legend (Lines 373-381 and S28). In addition, we have included\ntime-lapse STED images of live U2OS cells, shown at 20-frame intervals in Fig. S28\nof revised supplementary file.\n6. Regarding the authors' tracking experiments, I have several questions: There is no\ncomparison to other more commonly used intracellular tagging systems, such as the\nHaloTag, SNAPtag, or even the reHaloTag and xHTL labeling systems. This absence\nmakes it difficult to assess the acquired results effectively.\nResponse: We thank the reviewer for this valuable suggestion. To benchmark ReTagX\nagainst established intracellular labeling systems, we directly compared it with the\nwidely used Halo-Tag under identical imaging conditions. A single ReTag8 molecule\nexhibited a 5.5-fold higher brightness than Halo-Tag-labeled proteins. After extended\ntime-lapse acquisition, a constant number of TNFR2-ReTag8 trajectories (>80%)\nremained detectable over 5 minutes, whereas fewer than 20% of Halo-Tag-labeled\ntrajectories persisted beyond 1 minute, indicating a significantly extended observation\nwindow for ReTag8. Furthermore, track-frame distribution analysis revealed that\nReTag8 enabled a 2.9-fold longer average tracking duration compared to Halo-Tag.\nImportantly, the measured diffusion coefficient of TNFR2-ReTag8 was\n0.28 ± 0.019 µm²/s, closely matching the reported value of ~0.30 µm²/s (Biochim.\nBiophys. Acta, 2010, 1798, 1081) (Fig. 6m-n; Fig. S40), suggesting that ReTag8 does\nnot significantly perturb protein mobility.\nThese results demonstrate that ReTagX offers substantially enhanced brightness,\nprolonged tracking duration, and minimal impact on protein dynamics, thereby\nproviding significant advantages over conventional Halo-Tag for single-molecule\nimaging in live cells. The new data and discussion were added in the revised manuscript\n(Line 470-483, Fig. 6m-n; Fig. S40)\n7. As stated, imaging was conducted with 30% laser intensity and 200 ms exposure for\nbrightness evaluation, and with 10% laser intensity and 50 ms exposure for\nfluorescence lifetime (this term is normally used in another context). These values lack\ncontext. The authors should include measured power densities or power at the back\nfocal plane. Additionally, they should provide a range of power densities for the actual\ntracking experiments; currently, it's only noted as \"set to optimize signal-to-noise.\"\nResponse: We thank the reviewer for this helpful suggestion to improve the clarity and\nreproducibility of our imaging conditions. As requested, we have measured and\nincluded the laser power densities used during single-molecule imaging. We fused\nTNFR2 with Halo-Tag labeled with SiR-Halo (1 nM) and ReTag8 labeled with\nMaP655-TMP (200 nM). In vitro single-molecule imaging with 640 nm laser\nillumination was performed at the following measured intensities: 30% (30.4 W/cm²),\n60% (60.4 W/cm²), and 100% (99.1 W/cm²). We used 60% laser intensity (60.4 W/cm²),\nwhich provided an optimal balance between signal-to-noise ratio and photobleach\n(newly added Fig. S39).\nRepresentative images acquired under varying illumination powers have been\nincluded in the revised Supplementary Information (Fig. S39). Additionally, the term\n“fluorescence lifetime” has been revised to avoid potential misunderstanding.\n8. Fusing multiple ReTags into an array is a good idea in principle, but it contradicts\nthe general aim of having a small tag that does not hinder diffusion. Once the authors\nadded the second ReTag, the size of their system became larger than that of the HaloTag\n(36 kDa compared to 33 kDa). What is the advantage here?\nResponse: We sincerely thank the reviewer for this insightful comment. We agree that\nthe development of array tags inherently increases the overall molecular size, which\ncan potentially influence protein function and mobility. For instance, the Array G\nsystem (~1.0 MDa) has been shown to markedly reduce the mobility of membrane\nprotein β1 from 0.25 to 0.14 µm²/s (Nat. Chem. Biol., 2019, 15, 401). However, such\nstrategies also offer distinct advantages, particularly in enhancing fluorescence\nbrightness and extending imaging duration (Cell, 2014, 159, 635; Nat. Chem. Biol.,\n2019, 15, 401).\nIn this work, ReTag8 (~156 kDa) provides a 5.5-fold increase in brightness per\nprotein compared to Halo-Tag (~36 kDa). Moreover, the exchangeable binding of\nReTag8 allows sustained fluorescence signals even as individual fluorophores bleach,\nresulting in approximately a 2.9-fold longer single-molecule tracking duration and\nsignificantly extended observation windows (>5 min) (Fig. 6) compared to monovalent\ntags like Halo-Tag (<1 min). Notably, the measured diffusion coefficient of TNFR2-\nReTag8 was 0.28 ± 0.019 µm²/s, closely matching the reported value of ~0.30 µm²/s\n(Biochim. Biophys. Acta, 2010, 1798, 1081) (Fig. 6m-n; Fig. S40), suggesting minimal\nimpact on protein mobility.\nTaken together, these results highlight that ReTagX provides a rational compromise\nbetween tag size and imaging performance. It offers a modular, tunable platform with\nenhanced brightness and photostability, while maintaining biological compatibility. We\nare actively working to develop smaller, high-performance ReTag variants to further\nminimize potential cellular perturbations. This discussion has been incorporated into\nthe revised manuscript (lines 541-546).\n9. In this context, diffusion coefficients or mean square displacements were never\nmeasured in any of the experiments.\nResponse: We sincerely thank the reviewer for this insightful comment. To directly\naddress the concern, we performed single-molecule imaging to evaluate whether\nReTagX affects protein diffusion. Specifically, we fused ReTag8 to Tumor Necrosis\nFactor Receptor 2 (TNFR2) for single-molecule tracking in live U2OS cells. As\npresented in the revised manuscript, we quantified the diffusion coefficient of TNFR2-\nReTag8 and obtained a value of 0.28 ± 0.019 µm²/s. This is in close agreement with the\npreviously reported diffusion coefficient of ~0.30 µm²/s for TNFR2 (Biochim. Biophys.\nActa, 2010, 1798, 1081) (Fig. 6m-n; Fig. S40). These results indicate that ReTag8 does\nnot significantly alter the native mobility of membrane proteins, supporting its\nsuitability for minimally perturbative, long-term imaging. We have now included these\ndata and the corresponding analysis in the revised manuscript (line 470-483, Fig. 6k-\n6o) and Supplementary Information (Fig. S40).\n10. No software is mentioned for the tracking analysis.\nResponse: We thank the reviewer for pointing this out. The software “Python” used for\ntracking analysis has been added to the revised Image Data Analysis section of the\nSupplementary Information (line 1200-1203).\n11. CAAX or other farnesylated proteins tend to diffuse quite rapidly, especially at 37 °C.\nThe motion blur present at the reported 20 Hz acquisition speed may hinder tracking\nfidelity. I recommend repeating the experiments with a faster exposure time to improve\nthe results.\nResponse: We thank the reviewer for raising this important point. To minimize motion\nblur and enhance tracking fidelity for ReTagX, we fused ReTag8 with a membrane\nreceptor Tumor Necrosis Factor Receptor 2 (TNFR2) and performed single-molecule\ntracking at an increased frame rate using a reduced exposure time of 30 ms (~33 Hz).\nAs shown in the revised manuscript, the measured diffusion coefficient of TNFR2–\nReTag8 was 0.28 ± 0.019 µm²/s, closely matching the previously reported value of\n~0.30 µm²/s (Biochim. Biophys. Acta, 2010, 1798, 1081) (Fig. 6m-n; Fig. S40). These\nresults demonstrate that ReTag8 supports high-fidelity tracking without significantly\nperturbing the native mobility of membrane proteins, underscoring its utility for\nminimally invasive, long-term imaging applications. We have now included these data\nand the corresponding analysis in the revised manuscript (line 470, Fig. 6k-6o) and\nSupplementary Information (Fig. S40).\n12. The increase in tracking duration appears to stem mainly from the ability to reduce\nlaser power due to the enhanced brightness of multiple bound MaP-TMP ligand. At\nleast, this is what can be inferred from reading the manuscript. I strongly recommend\ncomparing ReTag8 to an 8xTMP-tag to demonstrate that the improved performance is\ndue to rebinding. Additionally, please ensure that the same laser power is used for both\nexperiments.\nResponse: We thank the reviewer for this valuable suggestion. In ReTag8, the more\nthan 5-fold increased brightness is a major advantage that allows us to track single\nproteins using lower laser power. We agree with the reviewer that reduced laser power\nfor ReTag8 can contribute to prolonged tracking time.\nAs the reviewer required, we conducted a comparison between TNFR2 labeled with\nReTag8 and with an 8×TMP-Tag construct under identical imaging conditions,\nespecially with same laser power. Interestingly, we observed that the mean tracking\nduration of TNFR2-ReTag8 was approximately 2.8-fold longer than that of TNFR2-\nTMP-Tag8. Moreover, a constant number of TNFR2-ReTag8 trajectories (>80%)\nremained detectable after 5 minutes of imaging, whereas fewer than 20% of TMP-Tag8\ntrajectories persisted beyond 1 minute (new added Fig. 6n and Fig. S40). These results\nsuggest that reversible binding plays a key role in prolonging signal longevity during single-\nmolecule tracking.\nWe acknowledge that the improvement in tracking duration between ReTag8 and\nTMP-Tag8 is modest and warrants optimization in future work, particularly of the\nbinding and dissociation kinetics, to extend residence time and enable efficient\nexchange upon photobleaching. In parallel, we are working to engineer smaller ReTag\nvariants to support higher-order assemblies while minimizing tag size. This comparison\nand the associated discussion have been added to the revised manuscript (line 541, Fig.\n6n and S40).\n13. Transiently binding probes show their greatest strengths in prolonged imaging\nexperiments lasting minutes or even hours. The authors choose to label CAAX proteins\nafter a very short expression time point, when only a few proteins are present inside the\ncell. The standard practice would be to wait for higher expression levels and label only\na subfraction of all CAAX proteins. Following this, one could conduct prolonged\nimaging to demonstrate that, with the same ligand concentrations, ReTag8 outperforms\nthe TMP-tag or even the 8xTMP-tag, with a much milder decrease in the number of\nlocalizations over time. This approach has been successfully employed with xHTL dyes\nand the reHaloTag, for example. I highly recommend that the authors conduct this\nexperiment.\nResponse: We sincerely thank the reviewer for this insightful and constructive\nsuggestion. We fully agree that transiently binding probes show their greatest utility in\nprolonged imaging under more physiological expression levels, as elegantly\ndemonstrated in the work of Prof. Mike Heilemann and colleagues with exchangeable\nHaloTag systems (Angew. Chem. Int. Ed. 2025, 64, e202413117). In those studies,\nmembrane proteins were allowed to express at higher levels (~ 1d), and sub-\nstoichiometric labeling with fluorogenic dyes (e.g., SiR-S5, JF585-Hy4) enabled\nefficient single-molecule tracking with low background and minimal perturbation.\nTaking inspiration from this approach, we conducted additional experiments\nwhere cells were transfected for 6 h and then allowed to express the TNFR2-ReTag8\novernight (~12 h) before imaging. In single molecule imaging, we observed a markedly\nslower decrease in localization density over time as the reviewer assumed. Specifically,\na constant number of TNFR2-ReTag8 trajectories (>80%) remained detectable after 5\nminutes of imaging, whereas fewer than 20% of TMP-Tag8 and Halo-Tag-SiR\ntrajectories persisted beyond 1 minute under identical conditions. These results suggest\nthat the dynamic and reversible nature of the ReTag system contributes to more\nsustained signal retention. We have included these new data and discussions in the\nrevised manuscript (line 470, Fig. 6m-6o).\nReviewer #3 (Remarks to the Author):\n1. This paper describes a bright and regenerative tag system (ReTag) and highly\nfluorogenic ligands that provide bright labeling with minimal background. The authors\nshow how this tag allows greatly extended STED imaging or time-lapse SIM imaging\nof mitochondrial TOM20, revealing mitochondrial dynamics such as fission, fusion,\nand elongation. STED, multi-color SIM and MINFLUX images of Vimentin-ReTag1 all\nshow impressive resolution. In addition, the compact ReTag8 array tag fused to CAAX\nis used as an example of tracking a membrane protein in living cells, which extends\nimaging duration by ~4-fold (up to 10 seconds) compared to the conventional TMP-\nTag.\nResponse: We sincerely thank the reviewer for their positive summary and\nencouraging evaluation of our work.\n2. The images generated with the ReTag are impressive, so this tag and the ligands will\nbe useful to the field. However, the paper is very dense, especially the first half, with 22\nsupplementary figures described along with only 3 main figures. It is difficult to\nnavigate the first half of the results section as the many supplementary figures are\ntypically very poorly described, e.g. lines 251 and 253 show single sentence descriptions\nof Figs S9 and S10. It would help the reader to explain the supplementary data properly\nin the text and give more information in the legend. For me, Fig. S9 needs more explanation.\nResponse: We sincerely thank the reviewer for their positive remarks on the image\nquality and for recognizing the potential utility of the ReTag system and ligands for the\nfield. As the reviewer required, we have carefully revised the manuscript to improve\nclarity and readability. Specifically, we have expanded the descriptions of\nsupplementary figures within the main text to provide more context and explanation\n(see revised line 181 for S4 and S5; line 217 for S7; lines 252-258 for S11 and S12; line\n306 for S24 and S25). Additionally, considering the limited context provided in the\nmain manuscript, we have substantially updated the legends of the supplementary\nfigures to give more detailed information on the experimental conditions, results, and\ntheir relevance to the main findings. These improvements will make the manuscript\neasier to navigate and help readers better appreciate the supporting data.\n3. My major concern relates to the idea that multimerization of ReTag could be a\npotential alternative to e.g. the SunTag array, which would be advantageous based on\nthe smaller size of ReTag4/8. As the SunTag is primarily used for imaging translation,\nideally translation would be imaged with ReTag4/8. However, my bigger concern\nrelates to the idea that ReTag8 can be used to quantitate fluorescence as a proxy for\ne.g. translation when there is reversible exchange of bleached fluorophores. As far as I\nunderstand it, quantitation of translation using SunTag relies on stable binding of scFv\nso that, after correcting for bleaching, the changes in fluorescence can be directly\nattributed to changes in ribosome density. If ReTag8 is used and there is bleaching, then\nexchange of bleached fluorophores, how can variations in signal due to this exchange\nbe separated from changes in e.g. ribosome density?\nResponse: We sincerely thank the reviewer for this insightful and important comment.\nWe fully agree that the SunTag system, with its stable and irreversible binding of scFv,\nis ideally suited for quantifying translation by correlating fluorescence intensity with\nribosome density after correcting for photobleaching. Indeed, due to its fast ligand\nexchange, ReTag8 is not intended to replace SunTag for translation quantification.\nRather, our goal with ReTag8 is to address different limitations, primarily those related\nto tag size, photobleaching, and brightness in live-cell single-molecule imaging.\nSpecifically, the smaller size of individual ReTag units enables construction of\nmultivalent arrays with reduced steric hindrance compared to large systems like SunTag\nor ArrayG (>1 MDa), which have been reported to perturb protein mobility.\nTo explore the performance of ReTag8 in this context, we fused it to the\ntransmembrane protein TNFR2 and performed single-molecule tracking in live U2OS\ncells. Compared with TNFR2-Halo-Tag, TNFR2-ReTag8 showed a 5.5-fold increase in\nbrightness and enabled longer-term tracking (Fig. 6i). Importantly, the measured\ndiffusion coefficient of TNFR2-ReTag8 was 0.28 ± 0.019 µm²/s, closely matching the\nknown diffusion coefficient of ~0.30 µm²/s for this receptor (Biochim. Biophys. Acta,\n2010, 1798, 1081) (Fig. 6m-n; Fig. S40), suggesting that ReTag8 does not significantly\ninterfere with protein dynamics. In contrast, large tag arrays like ArrayG have been\nshown to reduce mobility considerably (e.g., to 0.14 µm²/s) (Nat. Chem. Biol., 2019,\n15, 401), which may limit their applicability for studying dynamic membrane or\ncytoplasmic proteins.\nIn summary, while ReTag8 is not ideal for quantitative translation imaging due to\nits exchangeable binding, it offers key advantages for minimally perturbative, extended-duration single-molecule imaging. We have clarified this distinction and\nadded relevant discussion in the revised manuscript (line 476 and line 544, Fig. 6m-60,\nand S40).\n4. In addition, it is described how ReTag8 extends the imaging time to ~10 seconds in\nliving cells, but typical SunTag imaging durations are >30 mins for translation.\nTherefore, it is unclear to what extent multimerising ReTag will be useful as an\nalternative array, other than for short-term bright imaging. If the authors only mean\nReTag4/8 is better than the conventional TMP-Tag, it would be better not to mention\nSunTag.\nResponse: We sincerely thank the reviewer for this thoughtful and important comment.\nSunTag system is well established for long-term translation imaging, with reported\ndurations exceeding 30 minutes. Such experiments are typically performed with\nrelatively long acquisition intervals and exposure times (e.g., 20-500 ms), resulting in\nlower imaging frequencies (~1 Hz) (Nat. Protoco., 2020, 15, 1371)\nIn contrast, our study focused on fast single-molecule imaging under higher\ntemporal resolution (33 Hz), where fluorophore exchange and photobleaching present\ndifferent challenges. Under these conditions, we observed that ReTag8 supports single-\nmolecule tracking durations of 1-10 seconds, which is in a comparable range to other\nsystems used for high-speed acquisition—for example, ~1.9 s at 5 Hz for kinesin-1 with\nSunTag, ~34 s at 20 Hz for integrin β1 with ArrayG, and ~1.6 s at 33 Hz for H2B with\nHA-tag (Table S1). These comparisons suggest that multimeric ReTag constructs can\nperform competitively under fast imaging regimes, particularly for tracking highly\ndynamic targets. Moreover, we found that ReTag8 does not noticeably perturb the\nmobility of the fused protein. Specifically, TNFR2-ReTag8 exhibited a diffusion\ncoefficient of 0.28 ± 0.019 µm²/s, closely matching the literature value of ~0.30 µm²/s\nfor this receptor (Biochim. Biophys. Acta, 2010, 1798, 1081) (Fig. 6m-n; Fig. S40). This\nsuggests that ReTag8 is minimally invasive in terms of molecular dynamics. In contrast,\nlarger arrays such as ArrayG, while offering long observation times, have been reported\nto reduce mobility significantly (e.g., to ~0.14 µm²/s), potentially limiting their\napplication in certain contexts.\nThus, we agree with the reviewer that direct comparisons to SunTag in the context\nof translation imaging may lead to confusion, as ReTag8 currently lacks the stable\nbinding necessary for quantitative translation assays. We have therefore revised the\nmanuscript to clarify the intended application scope of ReTag8 and have adjusted the\ndiscussion to focus on its advantages in single-molecule tracking, rather than long-term\ntranslation monitoring. We have incorporated these clarifications into the revised\nmanuscript (line 550).\nMinor comments\n5. The first Results section on the design seems a bit superfluous as it is background\ninfo rather than describing results.\nResponse: We appreciate the reviewer’s comment. We agree that some portions of the\nfirst Results section may read more like background. To address this, we have revised\nthe section to focus more explicitly on the experimental design process and the key\nfindings that guided the development. Background information has been moved to the\nIntroduction to maintain a clear distinction between context and results.\n6. Are the fusion proteins in Fig 2e and Fig S4, referred to as SNAP-TMP-NLS and\nH2B-SNAP-TMP-NLS the same? It appears so from the text but the nomenclature is\ndifferent.\nResponse: We thank the reviewer for pointing this out. The correct construct used in\nboth Figure 2e and Figure S8 is SNAP-TMP-NLS. The designation H2B-SNAP-TMP-\nNLS was a labeling error in the text and has now been corrected to maintain consistency\nthroughout the manuscript and SI (line 77 and 139).\n7. Line 227 – ‘MaP555-TMP show nice fluorogenicity (Fig. 2g and 2h)’ – is it not quite\nweak?\nResponse: We thank the reviewer for the careful reading and thoughtful comment.\nMaP555-TMP exhibits a 13.3-fold increase in fluorescence intensity and achieves\napproximately a 36.5-fold signal-to-noise ratio in live-cell imaging. Even at an elevated\nprobe concentration of 1 µM, only minimal nonspecific signal was detected. While its\nfluorogenicity is somewhat lower compared to MaP618m-TMP (190-fold) and\nMaP655-TMP (179-fold), MaP555-TMP still demonstrates reasonably robust\nfluorogenic performance. To clarify this point and facilitate comparison with\nMaP618m-TMP and MaP655-TMP, we have revised the manuscript to describe the\nfluorogenicity as \"moderate\" rather than \"nice\". (line 226).\n8. Fig 4, S23 – indicate the time between frames in the legend (and in other figures\nshowing time frames)\nResponse: We thank the reviewer for the careful reading and helpful suggestion. We\nhave now updated the figure legends to clearly indicate the time interval between\nframes, using appropriate units (min, h, or s). Additionally, image frame numbers are\nnow marked with a \"#\" symbol for clarity.\n9. Fig 4e – point out more clearly where the evidence of fission and fusion is (e.g.\nhighlighted regions as in Fig 5d).\nResponse: We thank the reviewer for the helpful suggestion. In the revised version of\nFig. 4e, we have highlighted the regions where fission and fusion events occur in Fig.\n5d and S28.\n10. Fig. S26 – is it not supposed to also show the Hoechst staining in the Vimentin\nlabelled cells?\nResponse: We thank the reviewer for the helpful suggestion. In the revised version of\nFig. S26, we have repeated the confocal imaging to include Hoechst staining in the\nvimentin-labeled cells. The updated data have been incorporated into the revised\nSupplementary Information (Fig. S32).\n11. Line 431 – ‘ReTag1 displayed only transiently reversible labeling, limiting its utility\nfor long-term tracking due to signal loss (Fig. 6i)’. How is this finding reconciled with\nthe analysis of ReTag1 in Fig. 3, which describes how it allows reversible labeling\nwithin seconds to minutes in live-cell imaging, and fluorescence recovery within 1-2\nminutes in the FRAP experiments? Perhaps this relates to the timing of the 400 frames,\nbut again I couldn’t find information about this in the legend or methods.\nResponse: We thank the reviewer for this insightful comment. The distinction the\nreviewer highlights arises from the differing contexts and timescales of the experiments.\nIn the FRAP experiments, the recovery of bulk H2B-ReTag1 fluorescence occurs within\n1-2 minutes, representing ensemble-level dynamics in a high-density context. This\nrecovery reflects the exchange of many labeled proteins and may also be influenced by\nfactors such as protein localization, local environment, and probe concentration. By\ncontrast, in single-molecule tracking (Fig. 6i), individual fluorophores bound to ReTag1\nare typically visible for only tens of frames at 20 Hz (approximately 1-2 seconds), after\nwhich the signal is lost due to dissociation or photobleaching. This transient labeling\nshown in Fig. 6i limits the continuous observation time of single molecules. To\novercome this limitation, we implemented multimeric constructs (e.g., ReTag4 or\nReTag8), which enable multi-fluorophore labeling and dynamic exchange. These arrays\nhelp maintain signal despite fluorophore loss, thus extending the duration of single-\nmolecule tracking. We have clarified this distinction between ensemble fluorescence\nrecovery (FRAP) and single-molecule dwell time in the revised discussion and figure\nlegends (Lines 455).\n12. Fig. 6i – explain how the number of fluorophores on individual proteins was\ncalculated.\nResponse: We sincerely thank the reviewer for this insightful comment. To determine\nthe number of fluorophores bound to individual proteins in Fig. 6i, we performed\nsingle-molecule brightness analysis using purified ReTag1 and ReTag4 constructs\nimmobilized on glass. Imaging was conducted under identical conditions (30% 640 nm\nlaser power, 30.4 W/cm²; 200 ms exposure). After background subtraction, the\nfluorescence intensity of individual spots exhibited discrete levels corresponding to\ninteger multiples of a baseline intensity.\nWe first calibrated the signal using ReTag1, which binds exactly one MaP655-TMP\nmolecule. The fluorescence plateau intensity for ReTag1 was ~40 arbitrary units (a.u.),\nwhich we define as the brightness of a single fluorophore. Since rhodamine dyes scale\nlinearly with copy number below saturation, we estimated fluorophore stoichiometry\nby dividing the pre-bleach plateau intensity of each trace by 40 a.u. and rounding to the\nnearest integer.\nFor ReTag4 labeled with 100 nM probe, the plateau intensities were typically ~80–\n100 a.u., indicating ~2 bound fluorophores per protein. At 500 nM probe, the intensities\nreached ~150–160 a.u., consistent with full occupancy of all four TMP-binding sites.\nThese discrete intensity levels and stepwise bleaching events further validated our copy\nnumber estimation. This explanation has now been included in the revised Image data\nanalysis section (Lines 1200) for clarity.\n13. Line 435 – ‘Even when one fluorophore was bleached or exchanged, other\nfluorophores on ReTag4 continued to fluoresce, enabling extended single molecule\ntracking to ~200 frames in a streaming acquisition (Fig. 6j)’. It looks like 100 frames\nfor reliable tracking for 100 frames, so better to say this or tracking up to 200 frames.\nResponse: We thank the reviewer for pointing this out. We have revised the manuscript\nto state \"tracking over 100 frames\" as suggested.\n14. How does ReTag8 perform relative to ReTag4 in terms of fluorescence intensity and\nbrightness?\nResponse: We sincerely thank the reviewer for this insightful and constructive\nsuggestion. Under 60% 640 nm laser intensity (99.1 W/cm2) with 50 ms exposure time,\nwe measured the single-molecule brightness of ReTag8-CAAX and ReTag4-CAAX.\nThe results show that ReTag8-CAAX is, on average, 1.5 times brighter than ReTag4-\nCAAX. The data has been added in Fig. S38 of revised supplementary file.\n15. Fig. 6k-o – representative images of cells used for the analysis should also be shown.\nResponse: We thank the reviewer for the helpful suggestion. In the revised manuscript,\nwe have added representative images of the cells used for the analysis in Fig. S37.\nPoint-By-Point Response to the Comments on the Manuscript “A Bright\nand Regenerative Array Tag with Fluorogenic Ligands for Long-Term\nLive-Cell Nanoscopy and Single-Molecule Tracking”\nWe are grateful to the reviewers for their insightful comments and\nsuggestions. In the revised manuscript we have attempted to address all\nthese points and changed the manuscript accordingly. Below we provide\na point-by-point response to the comments of the reviewers.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\n1. The authors mentioned that the maximum FWHM of STED super-resolution\nimaging is 43± 5 nm, and the laser energy used is 219 mW. However, the\nauthors also mentioned that the maximum laser energy of STED can reach 520\nmW. According to STED imaging theory, the higher the STED light energy, the\nhigher the resolution. Why is the maximum resolution of this probe only 43 nm,\nand not higher?\nResponse: We thank the reviewer for this insightful question. In principle,\nSTED resolution improves with increasing depletion laser power. However, in\nbiological imaging the achievable resolution is constrained by several factors,\nincluding imperfect overlap of excitation and depletion beams, residual intensity\nat the central zero of the STED doughnut, detector signal-to-noise limitations,\nand biological factors such as motion blur and sample heterogeneity. These\ncollectively prevent unlimited resolution gains in live-cell contexts [e.g., FWHM\nof vimentin filaments reported as 39–52 nm (Angew. Chem. Int. Ed. 2016, 55,\n3290), 93–102 nm (J. Am. Chem. Soc. 2023, 145, 3075), and 37 ± 4 nm (Nat.\nCommun. 2022, 13, 2264)].\nRegarding laser power, the maximum 520 mW value refers to the Leica SP8\nSTED system, whereas the STEDYCON system used in our study provides a\nmaximum depletion power of 370 mW. Under our conditions, we observed a\nFWHM of ~43 ± 5 nm at 219 mW. Upon increasing the depletion power to 350\nmW, the resolution improved modestly to ~40 ± 7 nm in living cells (new data\nin Supplementary Fig. S27c). This value is close to the practical resolution limit\nof the STEDYCON system (~30 nm in fixed samples under ideal conditions).\nThese new results and clarifications have been added to the revised\nSupplementary Information.\n2. If 219 mW laser energy is directly applied to the sample, it will directly cause\ncarbonization of the sample, not to mention the living cell experiment. The\nauthor should provide the measurement position of the 219 mW laser energy\nand also provide the laser energy directly acting on the sample.\nResponse: We sincerely thank the reviewer for pointing out this important issue.\nAs requested, we calculated the actual laser power at the sample surface using\nthe STEDYCON system specifications:\n𝑷 = 𝑷 ×𝜂 ×𝜂 ×𝜂\n{𝒔𝒂𝒎𝒑𝒍𝒆} {𝒍𝒂𝒔𝒆𝒓} {𝑠𝑦𝑠𝑡𝑒𝑚} {𝑝𝑎𝑡ℎ} {𝑜𝑏𝑒𝑗𝑒𝑐𝑡𝑖𝑣𝑒}\nWhere P = 1.98 W (laser output power), η =59.137% (measured\nlaser system\nmodulation ratio of the laser system), η =25% (measured transmission\npath\nefficiency from laser exit to the objective entrance), η =75% (transmission\nobjective\nefficiency of the objective lens). Substituting these values gives: P =1.98\nsample\nW×0.59137×0.25×0.75 ≈ 219 mW.\nWe fully agree with the reviewer that continuous irradiation with such high\npowers could indeed damage biological samples. However, it is important to\nnote that STEDYCON employs pulsed STED excitation rather than\ncontinuous-wave depletion (https://abberior.rocks/superresolution-confocal-\nsystems/stedycon/). Pulsed STED delivers depletion photons only within the\neffective time window following excitation, thereby enhancing resolution while\nminimizing photodamage. This pulsed approach also ensures tight temporal\nsynchronization between excitation and depletion pulses, maximizing depletion\nefficiency while minimizing unnecessary light exposure (Proc. Natl. Acad. Sci.\nU.S.A. 2000, 97, 8206–8210).\nIndeed, STEDYCON has been widely applied in live-cell imaging without\nobservable sample carbonization (Angew Chem Int Ed Engl. 2022, 36,\ne202207905; Angew. Chem. Int. Ed. 2016, 55, 3290–3294). Consistent with\nthese reports, in our experiments we did not observe any carbonization or\nirreversible photodamage in live-cell samples.\n3. The STED and SIM imaging of mitochondria using FS27 and FS34 in the\narticle show that the probes target the outer mitochondrial membrane rather\nthan the inner membrane, and SIM imaging is better than STED. However,\nSTED imaging has a higher resolution than SIM. Why does this problem occur?\nResponse: We thank the reviewer for this insightful comment. While STED\nmicroscopy inherently provides superior spatial resolution (as reflected by\nnarrower full width at half maximum, FWHM, values compared to SIM),\nin our initial submission the SIM images appeared visually clearer than the STED\nimages. This apparent discrepancy stems from differences in image processing:\nSIM data undergo computational reconstruction, including contrast\nenhancement and noise reduction (using the HVL1 constrained deconvolution\nalgorithm or DFCAN/DFGAN deep learning algorithms, as described in the\nMethods), whereas the STED images were presented as raw data without post-\nprocessing.\nTo clarify this point, we have now (i) supplemented Fig. S27 with quantitative\nFWHM analyses confirming the superior resolution of STED over SIM, and (ii)\napplied deconvolution to the STED data to improve visual clarity. These\nupdates allow for a more balanced comparison between SIM and STED and\nare explicitly noted in the revised figure legends and Methods section.\n4. The authors mention that this probe has great advantages among STED\nsuper-resolution imaging probes. However, according to the literature that has\nbeen reported (Nature Communication 2020.11:3699, Proc. Natl Acad. Sci.\nUSA 116, 15817–15822 (2019). Biomedical Optics Express. 2024, 15(3):1595-\n1604), the highest resolution of STED super-resolution imaging probes can\nreach 35 nm and the longest imaging time can reach 60 minutes. Compared\nwith these probes, the probe mentioned by the authors does not have much\nadvantage in the field of STED super-resolution imaging. It can only be said\nthat this probe can be used for STED imaging, but it does not have a leading\nadvantage.\nResponse: We thank the reviewer for this constructive comment. We fully\nagree that several excellent mitochondria-targeting dyes have been developed\nfor long-term STED imaging. For example, TPERh demonstrates high\nphotostability and low saturation power for cristae visualization (Biomed. Opt.\nExpress 2024, 15, 1595), while MitoESq-635 achieves ~35 nm resolution\nand >50 min imaging of cristae dynamics in live cells (Nat. Commun. 2020, 11,\n3699). These probes, similar to our reversible-binding approach, enable signal\nrecovery and prolonged imaging performance. However, their utility is largely\nconfined to mitochondria, particularly cristae structures, and cannot be\ngeneralized to other protein targets in living cells.\nBy contrast, the goal of our work is not to introduce another mitochondria-\nspecific dye but to establish a generalizable protein tagging platform for\nregenerative labeling and nanoscopy of diverse targets. Our ReTagX system\nextends STED imaging of protein TOM20 from <5 frames to >300 frames while\nmaintaining ~80 nm resolution for >100 min. Crucially, ReTag1 is genetically\nencodable and can be fused to diverse proteins (e.g., vimentin, H2B, TNFR),\nenabling regenerative STED imaging beyond mitochondrial proteins.\nFurthermore, multimeric ReTag variants (e.g., ReTag8, ~156 kDa) enhance\nbrightness by 5.5-fold and extend single-molecule tracking of TNFR2 by ~2.9-\nfold, retaining >80% stable trajectories for 5 min—substantially outperforming\nconventional self-labeling tags such as HaloTag.\nThus, while TPERh and MitoESq-635 represent important advances in dye\nchemistry for mitochondria-focused imaging, our contribution lies in engineering\na versatile, regenerative protein tagging system. ReTagX complements these\ndye-based tools by broadening the application scope of live-cell nanoscopy to\ndiverse proteins and cellular contexts, rather than being limited to mitochondrial\nimaging.\n5. In STED super-resolution imaging, the saturated stimulated emission power\n(Is) of the probe is an important parameter that determines the performance of\nthe probe for STED imaging. Could the authors provide the Is value of the\nMaP655-TMP probe?\nResponse: We thank the reviewer for this important suggestion. As requested,\nwe have now measured and reported the saturated stimulated emission power\n(Is) of the MaP655-TMP probe in the revised manuscript. Briefly, the Is value\nwas determined by recording the fluorescence intensity of MaP655-TMP–\nlabeled samples as a function of the depletion laser power. A Gaussian beam\nprofile was applied to the sample, and the fluorescence–power dependence\nwas fitted with a single-exponential decay function to extract the saturation\npower.\nThe measured Is of MaP655-TMP was 2.28 ± 0.03 mW at 775 nm STED\nexcitation, which is consistent with previously reported values for comparable\nSTED probes (2.44 ± 0.03 mW) (Biomed. Opt. Express 2024, 15, 1595–1604).\nThis result highlights the favorable depletion efficiency and photophysical\nstability of MaP655-TMP under our experimental conditions. The experimental\ndetails have been added to the Supplementary Notes section, and the\ncorresponding data are now presented in the revised Fig. S27.\nReviewer #2 (Remarks to the Author):\n1. Please consider including a movie (or the second 50-minute movie)\nshowing the complete 100 minutes of STED imaging of TOM20–HALO–\nReTag1, corresponding to Figure 4e.\nResponse: We thank the reviewer for this helpful suggestion. As requested,\nwe have now provided a supplementary movie showing the complete 100\nminutes of STED imaging of TOM20–HALO–ReTag1 corresponding to Figure\n4e. This has been included in the revised manuscript as Movie 1.\n2. In Supplementary Figure 40c–e, both ReTag8 and TMP–Tag8 appear to\nreduce the diffusion coefficient of TNFR2 to slightly below 0.30 µm²/s. While I\nagree that this difference is small, it appears consistent and reproducible. It is\nnot clear whether the authors have tested whether this decrease is statistically\nsignificant; if not, such an analysis would strengthen the interpretation.\nResponse: We thank the reviewer for this valuable suggestion. To address this\npoint, we performed a Dunn’s multiple comparisons test on the diffusion\ncoefficients of TNFR2–ReTag8, TNFR2–TMP–Tag8, and TNFR2–Halo. The\nanalysis revealed no significant difference between ReTag8 and TMP–Tag8,\nwhile both exhibited small but statistically significant reductions compared with\nthe Halo–Tag group. Importantly, the impact on TNFR2 mobility was minimal,\nas ReTag8 and TMP–Tag8 retained ~92% and ~93% of the diffusion rate of the\nHalo–Tag control, respectively (Supplementary Fig. 40). The corresponding\nstatistical analysis has been added to the revised Supplementary Figure S40."}, {"round": 2, "reviewer_comments": {"reviewer_1": "I would like to thank the authors for answering most of my questions, but some issues still need to be addressed.\n1. The authors mentioned that the maximum FWHM of STED super-resolution imaging is 43± 5 nm, and the laser energy used is 219 mW. However, the authors also mentioned that the maximum laser energy of STED can reach 520 mW. According to STED imaging theory, the higher the STED light energy, the higher the resolution. Why is the maximum resolution of this probe only 43 nm, and not higher?\n2. If 219 mW laser energy is directly applied to the sample, it will directly cause carbonization of the sample, not to mention the living cell experiment. The author should provide the measurement position of the 219 mW laser energy and also provide the laser energy directly acting on the sample.\n3. The STED and SIM imaging of mitochondria using FS27 and FS34 in the article show that the probes target the outer mitochondrial membrane rather than the inner membrane, and SIM imaging is better than STED. However, STED imaging has a higher resolution than SIM. Why does this problem occur?\n4. The authors mention that this probe has great advantages among STED super-resolution imaging probes. However, according to the literature that has been reported (Nature Communication 2020.11:3699, Proc. Natl Acad. Sci. USA 116, 15817–15822 (2019). Biomedical Optics Express. 2024, 15(3):1595-1604), the highest resolution of STED super-resolution imaging probes can reach 35 nm and the longest imaging time can reach 60 minutes. Compared with these probes, the probe mentioned by the authors does not have much advantage in the field of STED super-resolution imaging. It can only be said that this probe can be used for STED imaging, but it does not have a leading advantage.\n5. In STED super-resolution imaging, the saturated stimulated emission power (Is) of the probe is an important parameter that determines the performance of the probe for STED imaging. Could the authors provide the Is value of the MaP655-TMP probe?", "reviewer_2": "In their revised manuscript “A Bright and Regenerative Array Tag with Fluorogenic Ligands for Long-Term Live-Cell Nanoscopy and Single Molecule Tracking,” Si et al. address many of the open questions raised in my initial review. This Enables a more rigorous comparison with established tagging systems. I definitely recommend the manuscript for publication.\nI would, however, encourage the authors to address the following two minor points:\n1. Please consider including a movie (or the second 50-minute movie) showing the complete 100 minutes of STED imaging of TOM20–HALO–ReTag1, corresponding to Figure 4e.\n2. In Supplementary Figure 40c–e, both ReTag8 and TMP–Tag8 appear to reduce the diffusion coefficient of TNFR2 to slightly below 0.30 µm²/s. While I agree that this difference is small, it appears consistent and reproducible. It is not clear whether the authors have tested whether this decrease is statistically significant; if not, such an analysis would strengthen the interpretation.", "reviewer_3": "The authors have fully addressed my comments. I have no further concerns."}, "author_response": "Point-By-Point Response to the Comments on the Manuscript “A Bright\nand Regenerative Array Tag with Fluorogenic Ligands for Long-Term\nLive-Cell Nanoscopy and Single-Molecule Tracking”\nWe are grateful to the reviewers for their insightful comments and\nsuggestions. In the revised manuscript we have attempted to address all\nthese points and changed the manuscript accordingly. Below we provide\na point-by-point response to the comments of the reviewers.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\n1. The authors mentioned that the maximum FWHM of STED super-resolution\nimaging is 43± 5 nm, and the laser energy used is 219 mW. However, the\nauthors also mentioned that the maximum laser energy of STED can reach 520\nmW. According to STED imaging theory, the higher the STED light energy, the\nhigher the resolution. Why is the maximum resolution of this probe only 43 nm,\nand not higher?\nResponse: We thank the reviewer for this insightful question. In principle,\nSTED resolution improves with increasing depletion laser power. However, in\nbiological imaging the achievable resolution is constrained by several factors,\nincluding imperfect overlap of excitation and depletion beams, residual intensity\nat the central zero of the STED doughnut, detector signal-to-noise limitations,\nand biological factors such as motion blur and sample heterogeneity. These\ncollectively prevent unlimited resolution gains in live-cell contexts [e.g., FWHM\nof vimentin filaments reported as 39–52 nm (Angew. Chem. Int. Ed. 2016, 55,\n3290), 93–102 nm (J. Am. Chem. Soc. 2023, 145, 3075), and 37 ± 4 nm (Nat.\nCommun. 2022, 13, 2264)].\nRegarding laser power, the maximum 520 mW value refers to the Leica SP8\nSTED system, whereas the STEDYCON system used in our study provides a\nmaximum depletion power of 370 mW. Under our conditions, we observed a\nFWHM of ~43 ± 5 nm at 219 mW. Upon increasing the depletion power to 350\nmW, the resolution improved modestly to ~40 ± 7 nm in living cells (new data\nin Supplementary Fig. S27c). This value is close to the practical resolution limit\nof the STEDYCON system (~30 nm in fixed samples under ideal conditions).\nThese new results and clarifications have been added to the revised\nSupplementary Information.\n2. If 219 mW laser energy is directly applied to the sample, it will directly cause\ncarbonization of the sample, not to mention the living cell experiment. The\nauthor should provide the measurement position of the 219 mW laser energy\nand also provide the laser energy directly acting on the sample.\nResponse: We sincerely thank the reviewer for pointing out this important issue.\nAs requested, we calculated the actual laser power at the sample surface using\nthe STEDYCON system specifications:\n𝑷 = 𝑷 ×𝜂 ×𝜂 ×𝜂\n{𝒔𝒂𝒎𝒑𝒍𝒆} {𝒍𝒂𝒔𝒆𝒓} {𝑠𝑦𝑠𝑡𝑒𝑚} {𝑝𝑎𝑡ℎ} {𝑜𝑏𝑒𝑗𝑒𝑐𝑡𝑖𝑣𝑒}\nWhere P = 1.98 W (laser output power), η =59.137% (measured\nlaser system\nmodulation ratio of the laser system), η =25% (measured transmission\npath\nefficiency from laser exit to the objective entrance), η =75% (transmission\nobjective\nefficiency of the objective lens). Substituting these values gives: P =1.98\nsample\nW×0.59137×0.25×0.75 ≈ 219 mW.\nWe fully agree with the reviewer that continuous irradiation with such high\npowers could indeed damage biological samples. However, it is important to\nnote that STEDYCON employs pulsed STED excitation rather than\ncontinuous-wave depletion (https://abberior.rocks/superresolution-confocal-\nsystems/stedycon/). Pulsed STED delivers depletion photons only within the\neffective time window following excitation, thereby enhancing resolution while\nminimizing photodamage. This pulsed approach also ensures tight temporal\nsynchronization between excitation and depletion pulses, maximizing depletion\nefficiency while minimizing unnecessary light exposure (Proc. Natl. Acad. Sci.\nU.S.A. 2000, 97, 8206–8210).\nIndeed, STEDYCON has been widely applied in live-cell imaging without\nobservable sample carbonization (Angew Chem Int Ed Engl. 2022, 36,\ne202207905; Angew. Chem. Int. Ed. 2016, 55, 3290–3294). Consistent with\nthese reports, in our experiments we did not observe any carbonization or\nirreversible photodamage in live-cell samples.\n3. The STED and SIM imaging of mitochondria using FS27 and FS34 in the\narticle show that the probes target the outer mitochondrial membrane rather\nthan the inner membrane, and SIM imaging is better than STED. However,\nSTED imaging has a higher resolution than SIM. Why does this problem occur?\nResponse: We thank the reviewer for this insightful comment. While STED\nmicroscopy inherently provides superior spatial resolution (as reflected by\nnarrower full width at half maximum, FWHM, values compared to SIM),\nin our initial submission the SIM images appeared visually clearer than the STED\nimages. This apparent discrepancy stems from differences in image processing:\nSIM data undergo computational reconstruction, including contrast\nenhancement and noise reduction (using the HVL1 constrained deconvolution\nalgorithm or DFCAN/DFGAN deep learning algorithms, as described in the\nMethods), whereas the STED images were presented as raw data without post-\nprocessing.\nTo clarify this point, we have now (i) supplemented Fig. S27 with quantitative\nFWHM analyses confirming the superior resolution of STED over SIM, and (ii)\napplied deconvolution to the STED data to improve visual clarity. These\nupdates allow for a more balanced comparison between SIM and STED and\nare explicitly noted in the revised figure legends and Methods section.\n4. The authors mention that this probe has great advantages among STED\nsuper-resolution imaging probes. However, according to the literature that has\nbeen reported (Nature Communication 2020.11:3699, Proc. Natl Acad. Sci.\nUSA 116, 15817–15822 (2019). Biomedical Optics Express. 2024, 15(3):1595-\n1604), the highest resolution of STED super-resolution imaging probes can\nreach 35 nm and the longest imaging time can reach 60 minutes. Compared\nwith these probes, the probe mentioned by the authors does not have much\nadvantage in the field of STED super-resolution imaging. It can only be said\nthat this probe can be used for STED imaging, but it does not have a leading\nadvantage.\nResponse: We thank the reviewer for this constructive comment. We fully\nagree that several excellent mitochondria-targeting dyes have been developed\nfor long-term STED imaging. For example, TPERh demonstrates high\nphotostability and low saturation power for cristae visualization (Biomed. Opt.\nExpress 2024, 15, 1595), while MitoESq-635 achieves ~35 nm resolution\nand >50 min imaging of cristae dynamics in live cells (Nat. Commun. 2020, 11,\n3699). These probes, similar to our reversible-binding approach, enable signal\nrecovery and prolonged imaging performance. However, their utility is largely\nconfined to mitochondria, particularly cristae structures, and cannot be\ngeneralized to other protein targets in living cells.\nBy contrast, the goal of our work is not to introduce another mitochondria-\nspecific dye but to establish a generalizable protein tagging platform for\nregenerative labeling and nanoscopy of diverse targets. Our ReTagX system\nextends STED imaging of protein TOM20 from <5 frames to >300 frames while\nmaintaining ~80 nm resolution for >100 min. Crucially, ReTag1 is genetically\nencodable and can be fused to diverse proteins (e.g., vimentin, H2B, TNFR),\nenabling regenerative STED imaging beyond mitochondrial proteins.\nFurthermore, multimeric ReTag variants (e.g., ReTag8, ~156 kDa) enhance\nbrightness by 5.5-fold and extend single-molecule tracking of TNFR2 by ~2.9-\nfold, retaining >80% stable trajectories for 5 min—substantially outperforming\nconventional self-labeling tags such as HaloTag.\nThus, while TPERh and MitoESq-635 represent important advances in dye\nchemistry for mitochondria-focused imaging, our contribution lies in engineering\na versatile, regenerative protein tagging system. ReTagX complements these\ndye-based tools by broadening the application scope of live-cell nanoscopy to\ndiverse proteins and cellular contexts, rather than being limited to mitochondrial\nimaging.\n5. In STED super-resolution imaging, the saturated stimulated emission power\n(Is) of the probe is an important parameter that determines the performance of\nthe probe for STED imaging. Could the authors provide the Is value of the\nMaP655-TMP probe?\nResponse: We thank the reviewer for this important suggestion. As requested,\nwe have now measured and reported the saturated stimulated emission power\n(Is) of the MaP655-TMP probe in the revised manuscript. Briefly, the Is value\nwas determined by recording the fluorescence intensity of MaP655-TMP–\nlabeled samples as a function of the depletion laser power. A Gaussian beam\nprofile was applied to the sample, and the fluorescence–power dependence\nwas fitted with a single-exponential decay function to extract the saturation\npower.\nThe measured Is of MaP655-TMP was 2.28 ± 0.03 mW at 775 nm STED\nexcitation, which is consistent with previously reported values for comparable\nSTED probes (2.44 ± 0.03 mW) (Biomed. Opt. Express 2024, 15, 1595–1604).\nThis result highlights the favorable depletion efficiency and photophysical\nstability of MaP655-TMP under our experimental conditions. The experimental\ndetails have been added to the Supplementary Notes section, and the\ncorresponding data are now presented in the revised Fig. S27.\nReviewer #2 (Remarks to the Author):\n1. Please consider including a movie (or the second 50-minute movie)\nshowing the complete 100 minutes of STED imaging of TOM20–HALO–\nReTag1, corresponding to Figure 4e.\nResponse: We thank the reviewer for this helpful suggestion. As requested,\nwe have now provided a supplementary movie showing the complete 100\nminutes of STED imaging of TOM20–HALO–ReTag1 corresponding to Figure\n4e. This has been included in the revised manuscript as Movie 1.\n2. In Supplementary Figure 40c–e, both ReTag8 and TMP–Tag8 appear to\nreduce the diffusion coefficient of TNFR2 to slightly below 0.30 µm²/s. While I\nagree that this difference is small, it appears consistent and reproducible. It is\nnot clear whether the authors have tested whether this decrease is statistically\nsignificant; if not, such an analysis would strengthen the interpretation.\nResponse: We thank the reviewer for this valuable suggestion. To address this\npoint, we performed a Dunn’s multiple comparisons test on the diffusion\ncoefficients of TNFR2–ReTag8, TNFR2–TMP–Tag8, and TNFR2–Halo. The\nanalysis revealed no significant difference between ReTag8 and TMP–Tag8,\nwhile both exhibited small but statistically significant reductions compared with\nthe Halo–Tag group. Importantly, the impact on TNFR2 mobility was minimal,\nas ReTag8 and TMP–Tag8 retained ~92% and ~93% of the diffusion rate of the\nHalo–Tag control, respectively (Supplementary Fig. 40). The corresponding\nstatistical analysis has been added to the revised Supplementary Figure S40."}, {"round": 3, "reviewer_comments": {"reviewer_1": "I thank the author for responding to all my issues and recommend that the manuscript be accepted for publication in current state."}, "author_response": ""}]

Decision letter