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. 2025 Jul 22;4(2):217–226. doi: 10.1021/cbmi.5c00037

Single-Molecule Localization Expansion Nanoscopy via Spontaneously Blinking Fluorophores

Lu Yang 1, Tianli Zhai 1, Song Chen 1, Daoming Guan 1, Qian Liu 1, Yunxiang Zhang 1,*
PMCID: PMC12933481  PMID: 41756770

Abstract

Expansion microscopy (ExM) combined with stochastic optical reconstruction microscopy (STORM) offers a powerful method for the super-resolution imaging of cellular ultrastructures beyond the diffraction limit. However, the integration of these two techniques is hindered by technical challenges, including hydrogel shrinkage induced by high-ionic-strength SMLM imaging buffers and severe fluorophore bleaching during expansion. Here, we employ HMSiR, a spontaneously blinking silicon rhodamine dye, to address these limitations. HMSiR exhibits intrinsic stochastic blinking at physiological pH, eliminating the need for specialized imaging buffers and significantly improving the fluorescence retention during gelation. By leveraging HMSiR, we establish a streamlined Ex-STORM workflow that achieves a stable linear expansion factor of 4.2 ± 0.3 and resolves the cellular ultrastructure with enhanced clarity. This approach resolves detailed features of mitochondrial ultrastructure, including TOM20 protein clusters with approximately 35 nm diameters on the outer mitochondrial membrane, and reveals microtubule organization with widths around 55 nm, offering nanoscale insights into protein distributions. This study not only overcomes critical barriers in Ex-STORM but also opens avenues for high-resolution imaging in cell biology and beyond, offering possibilities for exploring the nanoworld within cells.

Keywords: super-resolution imaging, expansion microscopy, stochastic optical reconstruction microscopy, spontaneously blinking dye, HMSiR

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Introduction

Expansion microscopy (ExM) is a transformative imaging technique developed in recent years that enables super-resolution imaging beyond the diffraction limit by physically expanding biological samples embedded in a swellable polyelectrolyte hydrogel. Since its introduction by Ed Boyden’s team in 2015, ExM has revolutionized the field of nanoscale imaging by isotropically separating biomolecules, allowing conventional microscope to resolve structures previously indistinguishable. − However, ExM alone is limited by its reliance on physical expansion, which does not address the molecular density constraints inherent in high-resolution imaging.

Single-molecule localization microscopy (SMLM), particularly stochastic optical reconstruction microscopy (STORM), , is also a powerful super-resolution imaging method that complements ExM by achieving ultrahigh spatial resolution through precise localization of individual fluorescent molecules. , STORM circumvents signal overlap by stochastically switching fluorophores on and off, ensuring that only a sparse subset of molecules is illuminated during each imaging cycle. ,,, However, STORM encounters limitations in scenarios with high-density fluorophore distributions, , where molecular proximity restricts localization accuracy due to overlapping signals, such as intracellular ultrastructures.

The integration of ExM with STORM (Ex-STORM) leverages the complementary strengths of both techniques. − ExM physically expands molecular distances, thereby reducing the density constraints of STORM and significantly reducing the signal overlap between fluorophores. This synergistic approach not only enhances the resolution of STORM but also extends its applicability to complex biological structures.

Despite its potential, Ex-STORM faces several critical technical challenges that require resolution. A primary limitation stems from the inherent properties of AF647, a cyanine5-based fluorophore widely employed in SMLM imaging. , Unlike spontaneously blinking fluorophores, AF647 requires a specialized SMLM imaging buffer containing sulfhydryl groups to induce stochastic fluorescence blinking essential to achieve single-molecule localization (Figure ). These sulfhydryl groups randomly bind to a fraction of the fluorophores, transitioning them into a dark state. This specialized SMLM imaging buffer, with its essential ionic composition, creates a fundamental incompatibility with expansion microscopy workflows. The high ionic strength of the SMLM buffer induces hydrogel shrinkage through charge-shielding effects on the polyelectrolyte network, thereby significantly compromising the hydrogel’s expansion factor. This ionic interference between imaging buffer components and the hydrogel matrix , represents a critical technical barrier that must be addressed for effective Ex-STORM implementation (Figure ).

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Comparison of the Ex-STORM methodologies. Conventional Ex-STORM (a) utilizes Alexa Fluor 647, a cyanine5 (Cy5) derivative, requiring specialized SMLM buffers containing thiol compounds for stochastic fluorescence blinking via transient adduct formation with free sulfhydryl groups. Our developed Ex-STORM (b) employs HMSiR, a dye exhibiting spontaneous fluorescence blinking at physiological pH through equilibrium between fluorescent open-ring and nonfluorescent closed-ring states, enabling expansion in pure water. Key performance advantages of the HMSiR-based system include complete hydrogel expansion, improved fluorescence retention rate, enhanced photostability, and a simplified workflow.

A second critical challenge stems from severe fluorophore bleaching during the expansion process. The free-radical polymerization chemistry, essential for hydrogel formation, generates reactive oxygen species that induce irreversible fluorophore degradation. This phenomenon disproportionately affects cyanine-based dyes, including Alexa Fluor 647the gold-standard fluorophore for SMLM. Experimental evidence demonstrates catastrophic dye loss during specimen expansion, with AF647 exhibiting only approximately 10% survival rates postexpansion. , This degradation mechanism likely involves radical-mediated cleavage of the dye’s polymethine backbone, fundamentally compromising its photophysical properties.

Recent advances in spontaneous blinking fluorescent dyes offer a previously unexplored promising solution to some of these limitations. , These dyes undergo stochastic fluorescence transitions via an intrinsic thermal equilibrium mechanism, eliminating the need for high-power lasers or additional chemical additives. In this study, we employ HMSiRa spontaneously blinking silicon rhodamine derivative exhibiting unique intramolecular spirocyclization dynamicsto synergize expansion microscopy (ExM) and STORM imaging through a streamlined workflow. HMSiR exhibits spontaneous blinking at physiological pH due to ground-state equilibrium between its fluorescent open form and nonfluorescent closed form, eliminating the need for additives or preactivation lasers required by conventional fluorophores (Figure ). The inherent spontaneous photoswitching capability of HMSiR addresses the two fundamental limitations of conventional Ex-STORM methodologies.

This approach not only resolves the hydrogel shrinkage caused by SMLM imaging buffers in Ex-STORM, thereby maintaining expanded hydrogel dimensions for achieving higher expansion factors, but also significantly improves fluorescence retention during gelation. Consequently, this optimized method enables superior resolution imaging of intracellular ultrastructures, particularly mitochondria and microtubules, surpassing conventional implementations. This enhanced performance is attributed to the synergistic combination of improved physical expansion stability and optimized fluorescence signal preservation.

Results and Discussion

To evaluate the compatibility of HMSiR with Ex-STORM, we labeled intracellular structures with HMSiR-conjugated antibodies and performed protein-retention expansion microscopy (pro-ExM) gelation , (Figure a). Hydrogels were expanded in pure water without requiring SMLM imaging buffer, achieving a linear expansion factor (EF) of 4.2 ± 0.3 (Figure b). Specimens were immobilized via agarose embedding and reconstructed through single-molecule localization to generate Ex-STORM super-resolution images (Figure S1). Notably, immersion of fully expanded hydrogels in conventional SMLM imaging buffer induced significant contraction, reducing the linear EF to 2.0 ± 0.2 (Figure c). Compared to expansion in pure water, this shrinkage significantly shortened the distance between the target biomolecules, which were anchored within the gel network and separated during expansion, thereby compromising the effectiveness of super-resolution imaging. In contrast, this HMSiR-based method enabled direct STORM imaging under additive-free aqueous conditions, ensuring complete hydrogel expansion and preserving enhanced spatial resolution.

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Hydrogel expansion under varying conditions. (a) Unexpanded hydrogel. (b) Hydrogel expanded in pure water. (c) Hydrogel expanded in SMLM buffer. Red dashed lines indicate gel boundaries, denoting the gel boundaries. (d) Expansion factor (EF) of hydrogels in pure water and SMLM buffer. (e) Logarithmic functional fit of hydrogel expansion factor to different buffer ionic concentration. R 2 = 0.94.

Systematic evaluation of the hydrogel swelling behavior across standardized buffer systems revealed a concentration-dependent modulation of linear EFs (Figure S2). Quantitative analysis demonstrated a strong logarithmic correlation between ionic strength (x) and swelling coefficients (y), described by y = a·log­(x + b) + c (R 2 = 0.94, Figure e). This confirmed the predictive validity of this physicochemical model for hydrogel dimensional stability, demonstrating that the ionic concentration significantly influences gel expansion. An increase of solution concentration results in a logarithmically related cliff-like decrease in expansion factor. Especially in the low concentration interval, a small concentration perturbation can trigger a dramatic contraction response of the expanded gel.

The gelation process in expansion microscopy (ExM) necessitates the incorporation of ammonium persulfate (APS) and tetramethylethylenediamine (TEMED) as radical initiators within the monomer solution. TEMED catalyzes APS decomposition, generating sulfate radicals that subsequently initiate free-radical chain polymerization, forming a cross-linked hydrogel network. Notably, the highly reactive radical speciesboth from APS/TEMED initiation and propagating polymer chainscan lead to structural damage in probe molecules or induce a nonfluorescent state, resulting in significant attenuation of the fluorescence signal.

To systematically evaluate the chemical stability of HMSiR compared with Alexa Fluor 647 (AF647) under ExM polymerization conditions, we employed commercial Cy5 as a surrogate for AF647. This substitution is considered appropriate, because Cy5 and AF647 share a similar cyanine-based fluorophore scaffold. Moreover, previous studies have reported that Cy5 and AF647 exhibit comparable single-molecule brightness and survival times. We monitored the absorption spectral dynamics of the fluorophores during polymerization (Figure a,b). Adhering to Beer–Lambert principles, temporal fluorophore retention was quantified via the absorbance at λmax. After 24 min, HMSiR maintained 64.6% ± 2.1% retention, whereas Cy5 exhibited only 24.1% ± 3.4% survival (2.7-fold enhancement, p < 0.001), demonstrating superior radical resistance of HMSiR (Figure c).

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Photophysical properties of HMSiR and Cy5 under Ex-STORM conditions. (a) Absorption spectra of 2 μM Cy5 during gelation. (b) Absorption spectra of 2 μM HMSiR during gelation. (c) Fluorophore retention over time, calculated from absorbance at maximum absorption wavelength. (d) Single-molecule photostability in gel (photons per second: HMSiR, 5477 ± 7; Cy5, 5414 ± 16). (e) Single-molecule fluorescence time trace of HMSiR in gel. (f) Comparison of HMSiR blinking properties in PBS, APS&TEMED solution, and gel.

Beyond chemical stability, the photophysical properties of fluorescent dyes within the gel matrix can be influenced by free radicals and other environmental factors. To evaluate the performance of HMSiR in expansion microscopy, we assessed its single-molecule photophysical properties under various buffer conditions. Single-molecule experiments were performed by imaging biotinylated HMSiR or Cy5 molecules immobilized on PEG-biotin-modified coverslips via streptavidin (Figure S5). Imaging chambers were constructed on top of the coverslip, and chambers are filled with desired buffer or gel matrix. Although the photostability of HMSiR was inevitably compromised due to irreversible photochemical damage induced by free radicals generated during the gelation process involving APS and TEMED, it nevertheless exhibited markedly superior stability compared to Cy5. Single-molecule analyses revealed that while the majority of Cy5 molecules underwent complete photobleaching within 20 s of illumination, approximately 80% of HMSiR molecules remained photostable under identical conditions, with about 20% still active even after 180 s of continuous exposure (Figure d). This superior photostability allows for prolonged imaging times in expansion microscopy, enabling a more comprehensive and accurate data collection.

Silicon rhodamine-based dyes, in general, exhibit exceptional chemical and optical stability in gels, making them ideal candidates for advanced imaging techniques. However, for successful SMLM, factors such as a high single-molecule brightness and a lower duty cycle are crucial. To ensure HMSiR was the better silicon rhodamine derivative for our Ex-STORM application, we compared HMSiR with another silicon rhodamine variant, COOH-SiR. In PBS, the duty cycle of COOH-SiR was significantly higher than that of HMSiR (Figure S3). Notably, within the gel matrix, the duty cycle of COOH-SiR increased 4-fold (Figure S7), rendering it unsuitable for SMLM. In contrast, HMSiR demonstrated better blinking properties and photostability even within the gel matrix (Figure e and Figure S6). Quantitative characterization of HMSiR photoswitching parameters revealed a duty cycle range of 0.001–0.01, consistent with literature reports. − Importantly, within the pH range of pure water (6.6–7.2), HMSiR maintained a low duty cycle below 0.01 (Figure S4). Through systematic evaluation in distinct chemical environments, including PBS, radical-initiated polymerization systems (APS/TEMED), and gel matrices, we observed environment-dependent modulation of single-molecule parameters. While HMSiR exhibited a modestly elevated duty cycle in gel matrices (0.0104 ± 0.0022) compared to PBS buffer conditions (0.0044 ± 0.0009), its single-molecule brightness increased by approximately 10% (Figure f). These findings indicate that HMSiR’s superior photophysical properties within the gel matrix make it a highly promising candidate for use in both expansion microscopy (ExM) and SMLM imaging applications. Consequently, subsequent Ex-STORM experiments were conducted using a HMSiR.

Mitochondria, double-membrane organelles essential for eukaryotic cells, typically form highly dynamic tubular networks that interact with diverse cellular structures. − Visualizing submitochondrial features presents a significant challenge due to the proximity of mitochondrial tubule diameters to the diffraction limit of conventional optical microscopy. Furthermore, the densely populated outer mitochondrial membrane, containing numerous transporters and channels, complicates detailed imaging. Among these, the translocase of the outer membrane (TOM) complex serves as the primary entry point for nuclearly encoded proteins into the mitochondria. Previous studies have reported the clustering of the TOM complexes at the mitochondrial membrane. Using stimulated emission depletion (STED) nanoscopy, researchers observed that antibody-modified TOM clusters had apparent diameters of approximately 65–70 nm in several human cell lines. However, subsequent correction for antibody-induced enlarging demonstrated that the native TOM20 cluster structures possess an actual physical diameter of 30–40 nm. This dimensional correction accounts for the cumulative spatial enlargement due to primary and secondary antibody binding, with each antibody layer contributing approximately 17.5 nm radially to the detected structure. Consequently, when combined with expansion microscopy, this ambiguous linkage error can be significantly reduced.

We immunostained the TOM20 protein using the TRITC dye, which was successfully localized to mitochondria under conventional microscopy. However, diffraction-limited resolution precludes clear distinction of membrane structures (Figure a). Following expansion, resolution significantly improved, enabling differentiation of outer mitochondrial membrane (MOM) architectures, which is outlined by the yellow dotted line in Figure b, although signal intensity was diminished due to suboptimal dye retention during expansion (Figure b). Colocalization imaging of TRITC-labeled TOM20 and MitoTracker Deep Red, a mitochondrial marker, yielded a colocalization coefficient of 0.81 before expansion, which decreased to 0.27 postexpansion, revealing distinct spatial patterns between the two markers. This separation, enabled by physical expansion and increased resolution, supports the conclusion that anti-TOM20 immunolabeling is specifically associated with the mitochondrial outer membrane (Figure S9).

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Ex-STORM imaging of TOM20 mitochondrial clusters. (a) Pre-expansion image of TOM20 immunolabeled with TRITC. (b) Postexpansion image of TOM20 immunolabeled with TRITC. (c) STORM image of unexpanded TOM20 immunolabeled with HMSiR. (d) Ex-STROM image of the TOM20 immunolabeled with HMSiR. (e–i) Enlarged views of regions in (d). (j) Scatter plot with box plots showing the full width at half-maximum (fwhm) of 68 TOM20 mitochondrial clusters in (c). (k) Scatter plot with box plots showing the fwhm of 115 TOM20 clusters in (d). White dashed lines in (a–c) indicate cell outlines; in (f), mitochondrial tubule outlines. Yellow dashed lines in (b) indicate individual mitochondria.

Immunostaining of TOM20 with HMSiR dye, followed by conventional STORM imaging, demonstrated the enhanced resolution of individual mitochondria. However, this approach failed to distinguish membrane structures along the minor axis or resolve TOM20 protein clusters (Figure c,e and Figure S10). Quantitative analysis of intensity profiles along the mitochondrial minor axis revealed an average full width at half-maximum (fwhm) of 152.8 ± 31.3 nm (mean ± SD, n = 68), consistent with diffraction-limited mitochondrial dimensions (Figure j).

In contrast, the combination of expansion microscopy (ExM) with STORM (Ex-STORM) led to a substantial enhancement in the imaging resolution. The reconstructed Ex-STORM images clearly delineated the architecture of the mitochondrial outer membrane (MOM) (Figure d,f) and further resolved nanoscale TOM20 protein clusters distributed along the periphery of tubular mitochondria (Figure g–i). Quantitative analysis of all signal clusters in the Ex-STORM images (Figure d) revealed an average full width at half-maximum (fwhm) of 34.8 ± 9.7 nm (mean ± SD, n = 115), reflecting the localized fluorescence spread of individual clusters. As previously reported by Jans et al., STED microscopy combined with density-based clustering analysis has been employed to report TOM20 cluster diameters encompassing broader spatial extents and potential molecular overlaps. In contrast to these clustering-based approaches, our Ex-STORM analysis utilized a peak-based strategy by directly measuring the fwhm of the localized fluorescence distributions corresponding to individual TOM20-positive spots in reconstructed super-resolution images. This approach captures the local fluorescence distribution of each TOM20-positive spot and provides a more direct and accurate estimation of the core physical size of protein aggregates rather than the maximal spatial range of clustered regions. Considering that our immunostaining was performed before expansion, the observed structures inherently include a cumulative antibody linkage error of approximately 17.5 nm. To account for this inherent displacement, we applied a simplified quadrature summation rule, yielding an fwhm-based TOM20 cluster size of approximately 30.1 nm after correction. While quantitatively distinct, this corrected value falls within a similar nanoscale range as the antibody-corrected cluster sizes (35–40 nm) reported in STED-based studies. This concordance in identifying nanoscale protein organization, despite methodological differences, supports the robustness and biological relevance of our analytical method. Overall, these findings highlight the effectiveness of our Ex-STORM workflow in resolving native nanoscale protein architectures. By integrating physical specimen expansion, efficient fluorophore retention, and high-precision single-molecule localization, our method enables reliable quantification of protein complex sizeseven in the presence of moderate antibody-induced displacement. This is achieved by measuring the core spatial dimension through fwhm analysis, thereby avoiding the need for complex density-based clustering algorithms and their associated posthoc corrections for cluster definition or overlap.

Having demonstrated the efficacy of HMSiR-based Ex-STORM in resolving the mitochondrial ultrastructure, we next evaluated its performance in imaging microtubules, another critical cellular component. Microtubule proteins were immunostained with TRITC, and the full width at half-maximum (fwhm) of microtubule widths was measured to assess the resolution enhancement provided by Ex-STORM (Figure e). Under conventional microscopy, the average fwhm of microtubule widths was 418.45 ± 71.80 nm (Figure a). Following expansion, the average fwhm decreased to 119.57 ± 14.19 nm, representing an approximately 3.5-fold resolution improvement (Figure b). Immunostaining with HMSiR, followed by STORM, further enhanced the resolution, yielding an average fwhm of 92.22 ± 10.75 nm (Figure c). While HMSiR exhibited commendable chemical and photophysical stability in expansion microscopy, some fluorescence and protein loss occurred postpolymerization and proteinase K digestion. Concurrently, the linear distance between fluorescent signals is expanded by over 4-fold, leading to a volume expansion exceeding 64 times and a consequently lower signal density. While this expansion mitigates the issue of overlapping signals in STORM, which can compromise single-molecule localization accuracy, the overall density of fluorescent molecules becomes insufficient, resulting in suboptimal continuity in the final super-resolution reconstruction. To address the occasional discontinuities in microtubule images arising from the stringent localization accuracy requirements of STORM imaging postexpansion, Artificial neural network (ANN) , processing was employed to generate Ex-STORM reconstructed images. As a result, the average full width at half-maximum (fwhm) of microtubules decreased to 57.37 ± 3.92 nm, closely matching the reported outer diameter of immunolabeled microtubules (∼60 nm) (Figure d and Figure S11), thereby confirming the structural fidelity of the reconstruction. To quantitatively compare the spatial resolution of STORM and Ex-STORM, we conducted Fourier ring correlation (FRC) analysis on the reconstructed super-resolution images. In unexpanded samples labeled with HMSiR, STORM achieved the best FRC resolution of 67.4 nm (Figures S8 and S13). In contrast, Ex-STORM, leveraging the spontaneous blinking properties of HMSiR in the physically expanded hydrogel matrix, reached a significantly improved resolution of 21.4 nm (Figures S8 and S12). These results demonstrate that Ex-STORM offers a threefold enhancement in effective resolution over conventional STORM, highlighting its superior capability for visualizing nanoscale cytoskeletal structures. Furthermore, this study validates the synergistic advantage of combining physical sample expansion with HMSiR-based single-molecule localization microscopy.

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Ex-STORM imaging of tubulin. (a) Pre-expansion image of tubulins immunolabeled with TRITC. (b) Expansion image of tubulins immunolabeled with TRITC. (c) STORM image of unexpanded tubulins immunolabeled with HMSiR. (d) Ex-STORM image of tubulins immunolabeled with HMSiR. (e) Scatter plots with box lines obtained from a series of randomly selected microtubules with statistical half-peak full-width fwhm in (a–d).

Conclusions

This study demonstrates a refined approach to super-resolution microscopy by synergistically combining physical sample expansion (ExM) with the molecular localization precision of STORM. This dual-resolution strategy overcomes key limitations of conventional techniques and enables high-resolution visualization of intricate cellular ultrastructures. We successfully resolved the mitochondrial outer membrane, revealing nanoscale TOM20 protein clusters (∼35 nm in size) and captured microtubule organization at 21.4 nm resolution. Quantitative analysis further showed heterogeneous protein cluster sizes ranging from 25 to 50 nm, offering statistically robust insights into the subcellular organization.

A core innovation of this study lies in the strategic integration and comprehensive photophysical characterization of a spontaneously blinking dye within the Ex-STORM framework. Through a comparative analysis of single-molecule properties between commonly used silicon rhodamine derivatives, HMSiR and COOH-SiR, we selected HMSiR for its superior blinking behavior both in PBS and within the hydrogel environment. Unlike conventional STORM dyes such as Cy5 or AF647, which require high-ionic-strength switching buffers incompatible with gel swelling, HMSiR enables single-molecule localization imaging under additive-free aqueous conditions. This compatibility preserves the hydrogel’s maximal linear expansion factor (4.2×) and eliminates shrinkage-induced artifacts. Importantly, our study complements and expands upon prior reports by systematically evaluating HMSiR’s blinking behavior, photostability, and retention across diverse chemical environments, thereby offering a practical reference for future applications in both ExM and SMLM.

We acknowledge that HMSiR, like most spontaneously blinking dyes, exhibits a higher duty cycle in water compared to that of AF647 in optimized switching buffers. This trade-off may reduce localization density and partially limit the effective resolution of Ex-STORM. However, we prioritized the preservation of the full expansion factor to maintain ExM’s physical decrowding advantage and avoid signal overlap, especially in densely packed structures. Our results suggest that even under such conditions HMSiR-based Ex-STORM achieves resolution on par with conventional STORM while offering additional spatial separation of biomolecular features.

Moving forward, future studies can build upon this foundation by further optimizing spontaneously blinking dyes with a higher photon yield and improved kinetics in aqueous conditions. Alternative strategies, such as nanobody-based labeling to minimize linkage error or gel formulations compatible with blinking buffers, may help bridge the current trade-off between expansion and imaging quality. Moreover, extending Ex-STORM to multicolor labeling and applying it to clinical or in vivo samples represents a promising direction for leveraging this technique’s full potential. Overall, our work presents a robust and accessible refinement of Ex-STORM, facilitating the high-fidelity visualization and quantitative analysis of complex intracellular architectures.

Experimental Section

Cell Culture

U2OS cells were cultured in a medium consisting of McCoy’s 5A­(PM150710) + 10% fetal bovine serum (FBS, 164210-50) + 1% penicillin/streptomycin (P/S, PB180120). The cells were maintained at 37 °C/5% CO2 and grown on 18 mm diameter coverslips or glass-bottomed dishes.

Antibody Conjugation

The carboxyl group of HMSiR was activated to an N-hydroxysuccinimide (NHS) ester, as confirmed by NMR spectroscopy (Bruker Avance III HD) and mass spectrometry (Bruker microTof-QII). The activated HMSiR-NHS was reacted with the secondary antibody (rabbit antimouse IgG, Proteintech, cat. no. B900120) at a 30:1 molar ratio in 0.15 M sodium bicarbonate buffer (pH ∼ 8.3) under dark conditions for 3 h. The labeled antibody was purified using dialysis against PBS using membranes with a 50,000 kDa cutoff to remove unreacted dye. Antibody dilutions for immunostaining are detailed in the immunostaining section.

Retention of Fluorescence in the Gelation Process

A 2 μM solution of HMSiR or Cy5 was prepared using an ExM monomer solution (8.625% (w/w) sodium acrylate, 2.5% (w/w) acrylamide, 0.15% (w/w) N,N′-methylenebis­(acrylamide), and 2 M NaCl in PBS). Baseline absorption spectra of both fluorophores were recorded from 400 to 800 nm at time zero. Subsequently, 10% ammonium persulfate (APS) and 10% N,N,N′,N′-tetramethylethylenediamine (TEMED) were rapidly added and thoroughly mixed into the solution. Absorption spectra were then measured at 3, 6, 9, 12, 15, 18, 21, and 24 min, respectively. The retention rate of each dye was determined based on absorbance values at their respective maximum absorption wavelengths, which are directly proportional to dye concentrations.

Single-Molecule Experiment

Coverslip Preparation and Dye Attachment

Coverslips were sequentially cleaned using piranha solution (a mixture of sulfuric acid and hydrogen peroxide), 1 M sodium hydroxide, and ultrapure water. After drying under nitrogen, coverslips were immersed in a 2% aminosilane solution in acetone for 15 min. A mixture of mPEG-SVA and Biotin-PEG-SVA (Laysan Bio) in 0.1 M sodium bicarbonate buffer, containing 0.55 M potassium sulfate to reach cloudy point, was applied to the coverslip surface for 2 h, followed by thorough rinsing with ultrapure water. Biotinylated dyes were attached to the PEGylated surface via streptavidin binding through sequential incubation within a freshly assembled flow channel.

Imaging Experiments in PBS

Single-molecule imaging was performed using flow channels formed by a coverslip and a microscope slide sealed with double-sided tape. The imaging medium was PBS (pH = 7.4).

Imaging Experiments in APS and TEMED

For experiments involving APS and TEMED, four chambers were created using a laser-machined quad-well slide, sealed with a coverslip, and incubated with biotinylated dyes. The imaging medium consisted of 0.01% APS and 0.01% TEMED.

Imaging Experiments in Hydrogel

Imaging in hydrogel was conducted with a similar setup using a laser-machined quad-well slide, sealed with a coverslip incubated with biotinylated dyes. The imaging medium in this case was the hydrogel formed inside the well.

Single-Molecule Fluorescence Data Acquisition

Single-molecule fluorescence signals were collected for 300 s under 638 or 532 nm excitation at a power density of 0.27 kW/cm2. The acquired single-molecule data were subsequently analyzed using custom-written code in either MATLAB (MathWorks) or Python.

Immunostaining of Unexpanded Cells

Cells were fixed with a prewarmed fixation solution (3% paraformaldehyde [PFA], 0.1% glutaraldehyde [GA]) for 15 min. The fixation solution was discarded, and the cells were washed three times with PBS for 5 min each. Cells were then permeabilized with a solution of 0.2% Triton X-100 in PBS for 10 min. After removal of the permeabilization solution, cells were treated with a blocking solution (Beyotime, P0260) for 15 min. The blocking solution was removed, and cells were incubated with the primary antibodies at room temperature for 1.5 h or kept at 4 °C overnight. The following primary antibodies were used: mouse monoclonal anti-α-tubulin (Proteintech, cat. no. 66031-1-Ig, 1:100 dilution), mouse monoclonal anti-β-tubulin (Proteintech, cat. no. 66240-1-Ig, 1:400 dilution), and mouse monoclonal anti-TOM20 (Proteintech, cat. no. 66777-1-Ig, 1:400 dilution).

After washing three times with PBS (10 min each, gentle shaking), cells were incubated with the secondary antibodies for 1 h at room temperature. The following secondary antibodies were used: goat antimouse IgG (H+L), TRITC conjugate (Proteintech, cat. no. SA00007-1, 1:100 dilution), and rabbit antimouse IgG, HMSiR conjugate (1:50 dilution).

After another three PBS washes (10 min each), the samples were ready for fluorescence imaging.

Immunostaining of Expanded Cells

After removal of the blocking solution, cells were incubated with the primary antibodies at room temperature for 1.5 h or kept at 4 °C overnight. The following primary antibodies were used: mouse monoclonal anti-α-tubulin (Proteintech, cat. no. 66031-1-Ig, 1:50 dilution), mouse monoclonal anti-β-tubulin (Proteintech, cat. no. 66240-1-Ig, 1:200 dilution), and mouse monoclonal anti-TOM20 (Proteintech, cat. no. 66777-1-Ig, 1:100 dilution).

After washing three times with PBS (10 min each, gentle shaking), cells were incubated with the secondary antibodies for 1 h at room temperature. The following secondary antibodies were used: goat antimouse IgG (H+L), TRITC conjugate (Proteintech, cat. no. SA00007-1, 1:50 dilution), and rabbit antimouse IgG, HMSiR conjugate (1:25 dilution).

After another three PBS washes (10 min each), the samples were ready for fluorescence imaging.

Cell Expansion

Cells were incubated with Acryloyl-X, SE (0.1 mg/mL in PBS) at 4 °C overnight and washed three times with PBS. Gelation was performed in the gelation chamber. A standard glass slide (base slide) was tightly wrapped with Parafilm. Two square coverslips (22 × 22 mm) were positioned as spacers at opposing ends of the Parafilm-coated slide. A cell culture coverslip was inverted (cells facing downward) and carefully placed on the spacers, forming a gelation chamber. The edges of the coverslip will rest on the spacers with the cells suspended over the gap between the spacers. ExM monomer solution (8.625% (w/w) sodium acrylate, 2.5% (w/w) acrylamide, 0.15% (w/w) N,N′-methylenebis­(acrylamide), and 2 M NaCl in PBS) supplemented with 0.1% APS and 0.1% TEMED was added to the gelation chamber. The sample was incubated at 37 °C for 1.5 h to allow gel polymerization. After gelation, the gels were soaked in a digestion buffer (0.5% Triton X-100, 0.8 M guanidine hydrochloride, and 2% 50× TAE) containing 8 U/mL proteinase K and placed in a humidified box at 37 °C overnight. The digestion buffer was aspirated and then exchanged with ultrapure water for 30 min, and this was repeated until the hydrogels were fully expanded. The expanded hydrogels were transferred to poly-l-lysine (0.1 mg/mL)-coated coverslips with the cells downward and fixed with 3% agarose gel to minimize drift of the hydrogel from the bottom of the dish.

Microscope Setup and STORM Imaging

SMLM imaging was conducted on a custom-built total internal reflection fluorescence (TIRF) microscope on a Nikon ECLIPSE TE2000-U platform. A continuous 638/532 nm laser (Cobolt 06-01, HÜBNER Photonics) was focused on the back focal plane of a 100× TIRF objective (NA 1.49, Nikon). The fluorescence emission from the sample was filtered through a band-pass filter (ZET532/640m-TRF, Chroma) and detected using an iXon 897 electron-multiplying charge-coupled device (EMCCD, Andor).

All STORM imaging was conducted using HMSiR. The effective pixel size was 106 nm, the excitation power density was adjusted between 0.1 and 7 kW/cm2, and the exposure time was set at 50 ms. For STORM imaging, a total of 3000 to 6000 frames were acquired to ensure sufficient data for high-resolution reconstruction.

STORM images were reconstructed using ThunderStorm, and the resolution was quantitatively assessed by measuring the full width at half-maximum (fwhm) in ImageJ. Additionally, Fourier ring correlation (FRC) analysis was performed using a previously established MATLAB code to further evaluate the image resolution.

The reconstructed images of microtubules were padded to a size of 512 × 512 pixels to match the network’s input dimensions. The preprocessed images were then processed through an artificial neural network (ANN) trained for reconstructing dense microtubule images from sparse localization data sets. ,

Expansion Factor in Different Buffer Systems

The gel expansion was assessed in different buffer systems, including thiol-containing SMLM buffer (0.695 M, pH 8.0), NaHCO3–Na2CO3 buffer, Tris–HCl (1 M, pH 8.0), PBS (0.01 M, pH 7.4), and soda water (0.005 M, pH 9.0). The SMLM buffer was based on that reported by Xiaowei Zhuang’s group and consisted of 10% glucose, 140 mM β-mercaptoethanol, 0.5 mg/mL glucoseoxidase, and 40 μg/mL catalase. The NaHCO3–Na2CO3 buffer (0.01 M, pH 10.1) contained 5 mM NaHCO3 and 5 mM Na2CO3. The NaHCO3–Na2CO3 buffer (0.001 M, pH 10.0) contained 0.5 mM NaHCO3 and 0.5 mM Na2CO3.

Pre-expansion gels were cut into trapezoidal shapes, and triplicate measurements of each edge length were performed to determine the mean baseline dimensions. Subsequently, the gel specimens were immersed in different buffer systems and gently swirled at 30 rpm (10 min per cycle). Following three consecutive cycles of buffer exchange (aspiration and replenishment), the edge lengths of the expanded gels were similarly quantified via triplicate measurements to establish average values. Linear expansion coefficients (ratio of postexpansion to pre-expansion dimensions) were calculated to evaluate hydrogel dimensional scalability under each buffer system.

Supplementary Material

im5c00037_si_001.pdf (1,003.7KB, pdf)

Acknowledgments

The authors are grateful for the financial support from the National Natural Science Foundation of China (Grant Nos. 22174025 and 22474025) and National Key Research and Development Program of China (Grant No. 2024YFF1206300).

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/cbmi.5c00037.

  • Expansion factors in water and various buffer systems; additional single-molecule fluorescence data; synthesis and characterization of HMSiR-NHS and HMSiR-Biotin; experimental setup for single-molecule imaging in gel; and images of pre- and postexpansion mitochondria and microtubules using STORM and Ex-STORM (PDF)

Y. Zhang and Q. Liu conceived and supervised the project. L. Yang designed and performed Ex-STORM experiments. T. Zhai performed expansion protocol optimization. S. Chen and L. Yang performed single-molecule experiments and imaging analysis. D. Guan assisted super-resolution imaging experiments. L. Yang, Q. Liu, and Y. Zhang wrote the manuscript. All authors have approved the final version of the manuscript.

The authors declare no competing financial interest.

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