Biophysical Reports

For decades, the photon counting histogram (PCH) was used as the sole method to quantify fluorophore numbers in a diffraction-limited focal volume. This technique combines spatial excitation profiles, and the distribution of photon counts to register the photon emission statistics of individual fluorophores. However, this approach has not yet been transferred to widefield fluorescent imaging due to the lack of fast and single photon sensitive camera sensors which can capture the photon emission statistics of a single fluorophore. Here, we explore avenues towards quantitative analysis of the active fluorophore number by leveraging recent advancements in single photon avalanche diode (SPAD) array technology. Binary exposures of a SPAD array can be synchronized with picosecond laser pulses to measure the PCH in a widefield setting. Then, by modeling the statistical relationship between the active fluorophore number and the PCH in a region of interest following a laser pulse, we can perform Bayesian inference of this number. The model is demonstrated experimentally by counting quantum dots and various numbers of fluorescent dye molecules bound to DNA origamis. We find that this method has several important applications in widefield microscopy, including enhanced localization microscopy and constrained fitting of multiple unresolvable fluorescent emitters.

Fluorescent voltage-sensitive dyes (VSDs) enable non-invasive, high-throughput optical measurement of membrane potential in living cells, but the analytical reliability of such measurements depends critically on whether the dye and associated imaging conditions perturb the system under study. Here, we systematically characterise the photophysical performance and cell-perturbing effects of FluoVolt, a widely adopted VSD, across cancer cell lines (GIN31 glioblastoma and SK-MEL-30 melanoma) and primary human macrophages. Photobleaching kinetics were strongly cell-type-dependent, with SK-MEL-30 cells exhibiting complete fluorescence loss within 400 seconds under standard widefield conditions. FluoVolt staining combined with laser excitation caused an approximately 2.5-fold increase in cell detachment relative to unstained controls, and dual-wavelength excitation (488 + 405 nm) reduced GIN31 cell viability by approximately 17.5%. Critically, morphological changes, a transition from elongated to amoeboid-like phenotypes, were detected under staining conditions alone, prior to any laser exposure, indicating baseline dye-induced perturbation independent of phototoxicity. Halving dye concentration and loading time significantly attenuated these effects while preserving measurable fluorescence signal. These findings identify FluoVolt staining and excitation as previously uncharacterised sources of systematic measurement artefact and provide practical, actionable guidance for protocol design, control selection, and data interpretation in optical membrane potential imaging.

Fluorescence lifetime imaging microscopy (FLIM) of endogenous NAD(P)H enables the label-free assessment of cellular metabolic state. Although metabolic imaging is increasingly combined with fluorescent protein (FP) reporters to enhance biological specificity, the potential cross-talk between the intrinsic and extrinsic labels remain ill-defined. Here, we systematically evaluate cross-talk from FPs in metabolic FLIM using phasor analysis of two-photon fluorescence microscopy. The results clearly show that many widely used fluorescent proteins are excited under the conditions used for NADH imaging; they emit blue-shifted, short-lifetime fluorescence that can interfere with imaging NADH metabolic signatures. This overlap persists across excitation wavelengths and FP classes, posing a significant challenge for multiplexed metabolic imaging. This cautionary tale argues against unvalidated multiplexing strategies in metabolic FLIM studies. Our study aims to identify acceptable imaging partners, offer a pipeline for assaying potential cross-talk, and provide practical guidance for experimental design. SignificanceFluorescence lifetime imaging of NADH autofluorescence is a powerful, label-free approach to map cellular metabolism in living tissues. A growing number of studies combine NADH imaging with fluorescent protein (FP) reporters to simultaneously identify specific cell types or subcellular compartments. This study reveals that many FPs, spanning the visible spectrum, are unexpectedly excited under NADH conditions. Commonly used green, yellow, and red variants produce short-lifetime, blue-shifted fluorescence that directly overlaps with metabolic NADH signals. This cross-excitation can be falsely interpreted as a shift in cellular metabolic state, posing a significant risk for multiplexed metabolic imaging studies. Our studies establish a pipeline to assess and manage this risk. We identify StayGoldE138D and mNeonGreen as the most compatible FPs for co-imaging with NADH, and provide a practical framework to guide experimental design and control strategies for multiplexed metabolic FLIM.

Temperature is a fundamental parameter governing all molecular processes, including those that define life. Fluorescence microscopy is a powerful tool to observe molecular processes in living systems in real time. Precise control and measurement of temperature during fluorescence microscopy is therefore essential. We present here a robust temperature measurement based on the excited-state lifetime of the widely available and relatively inexpensive fluorescent dye pentamethine cyanine (Cy5). The excited-state lifetime of Cy5 shows a monotonic decline in the measurement range of 0 {degrees}C - 80 {degrees}C. The measured dependency is linear until 39 {degrees}C and monoexponential above. The dependance of excited-state lifetime upon temperature is used to measure temperature up to a precision of 0.5 {degrees}C or less, a temporal resolution down to <1 millisecond and to resolve temperature gradients with spatial resolutions that are only diffraction-limited. The far-red excitation and emission of Cy5 leaves bandwidth to simultaneously measure at least 3 additional spectral channels in standard fluorescent microscopes simultaneously. We demonstrate determination of temperature during 4-color live-cell fluorescence microscopy for a temperature-controlled experiment. We also show its applicability in measuring temperature gradients and laser-induced sample heating such as during STED nanoscopy.

Quantum effects in biology are unavoidable at the molecular scale; the unresolved question is whether they can remain functionally relevant across the timescale gap between femtosecond molecular dynamics and microsecond-to-millisecond biological function. Here we formalize this mismatch as an equilibrium-to-functionality gap and use tubulin as a stringent open-system test case. We combine secular Lindblad, Redfield, and hierarchical equations of motion (HEOM) treatments to quantify decoherence, non-perturbative relaxation, and the physical amplification required for functional relevance. Equilibrium dephasing yields a conservative [Formula] fs at 310 K, with a generic protein-bath baseline of {approx} 13 fs. A completed 30 ps HEOM trajectory for the full 1JFF tryptophan network shows distributed non-Markovian relaxation, with terminal purity Pur = 0.210 and stretched-exponential exponent {beta}KWW {approx} 0.44, confirming that Redfield is useful as a short-time perturbative comparator but not quantitatively interchangeable with HEOM in this intermediate-coupling regime. We introduce a coherence-utility criterion [U] = [K]{tau}coh/{tau}func, separating required amplification from empirically bounded gain. A thermodynamic uncertainty relation closure shows that neural-scale cascade amplification would require Pmin [~] 10-7 W, about five orders of magnitude above the local microtubule GTP budget. Frohlich pumping is found to be linewidth-gated rather than generically micron-scale; ordered-water cavity QED and geometric subradiance remain experimentally testable but severely constrained candidates. The result is not a model of consciousness, but a reproducible physical benchmark framework for evaluating biological quantum-coherence claims under explicit open-system, energetic, and experimental constraints. Six falsifiable experimental programmes are prioritized, and the full computational framework is released with a validation ledger, cryptographic audit trail, and living supplementary material. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=107 SRC="FIGDIR/small/724047v1_ufig1.gif" ALT="Figure 1"> View larger version (20K): org.highwire.dtl.DTLVardef@19e4f42org.highwire.dtl.DTLVardef@65a719org.highwire.dtl.DTLVardef@1bd63beorg.highwire.dtl.DTLVardef@df77d8_HPS_FORMAT_FIGEXP M_FIG O_FLOATNOGraphical abstract.C_FLOATNO Equilibrium tubulin coherence lies in the femtosecond regime, while functional neural timescales lie in the millisecond regime. Frohlich pumping, QED-cavity protection, and geometric subradiance remain experimentally discriminable non-equilibrium candidates requiring independently bounded amplification. C_FIG FundingThis research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors. Versioned computational scope of this releaseThis manuscript reports the theoretical framework, calibrated equilibrium baseline, Redfield/HEOM validation ledger, stratified Bayesian evidence synthesis, classical comparators, and falsifiable experimental design. The release-specific reproduction audit, including the current validation-check total and the SHA-256 fingerprints of the binary production artefacts (.npz, .pkl), is documented in LIVING_SI.md and outputs_data/raw_json/structur al/validation_report.json. A completed 30 ps HEOM production trajectory has been validated on constrained hardware; the master dataset contains the full 8-site population trajectory. A summary of those results is provided in [&#167;]2.2.5. All claims made below are restricted to the numerical and theoretical evidence reported in this manuscript and its associated repository artefacts. The public repository ships the calibrated phenomenological baseline for accessibility; the HEOM production artefacts serve as the non-perturbative validation benchmark. All source figure outputs associated with this release are maintained in the public repository under outputs_data/figures_final/.

Fluorescence imaging of dense cellular regions of interest (ROIs) in cells using fluorescence microscopy provides detailed images with pixels that report ensemble- and time-averaged biomolecular data, due to the diffraction limit when super-resolution modalities are not used and acquisition times are slower than typical biomolecular mobilities. The fluorescently-tagged biomolecules that are undergoing imaging can be more heterogeneous and dynamic, all within the dimensions of a single acquired image pixel. The ability to acquire data one biomolecule at a time within a given ROI can help recover some of the underlying biomolecular subpopulations that are otherwise averaged out. In this work, we present a relatively simple approach to achieving single-biomolecule photon bursts, BLeaching In-cell Single-molecule burstS (BLISS). We reveal millisecond photon bursts arising from clusters of mCherry-tagged heterochromatin protein 1 (mCherry-HP1) within heterochromatin biomolecular condensates in undifferentiated mouse embryonic stem cells (ESCs). Fluorescence lifetimes of these bursts are substantially lower than the averaged-out values observed per pixel in fluorescence lifetime imaging microscopy (FLIM), attributed to higher density in mCherry-HP1 clusters. These higher density clusters are observed primarily in undifferentiated ESCs. Two days after retinoic acid (RA) induction of differentiation, these bursts are rarely observed. In summary, using BLISS, we revealed a rare subpopulation of dense mCherry-HP1 clusters characterized by rapid, millisecond dynamics. These clusters are part of heterochromatin biomolecular condensates in ESCs at the pluripotent state, which would be otherwise averaged out in diffraction-limited fluorescence microscopy.

Protein oligomerization equilibria are central to many biological processes and are often highly sensitive to environmental conditions such as ionic strength, pH, and ligand binding. Quantitative characterization of these equilibria remains experimentally challenging because stable protein complexes frequently dissociate only at concentrations that are difficult to access with conventional biophysical methods. Fluorescence correlation spectroscopy (FCS) is uniquely suited to this problem, as it provides direct access to diffusion coefficients of fluorescently labeled proteins at nanomolar concentrations. However, the quantitative interpretation of FCS data from oligomeric systems requires a rigorous mathematical framework and careful experimental practice that have not previously been described in sufficient detail to guide implementation. Here, we provide a comprehensive description of the experimental workflow and analytical framework for determining dissociation equilibrium constants by FCS, covering instrument calibration, sample preparation, data quality control, after-pulse correction, and nonlinear least-squares fitting. We discuss common sources of error and provide practical guidance on critical experimental considerations including surface passivation, buffer preparation, equilibration time, and the role of labeling efficiency. Using the homotrimeric sliding clamp PCNA as a model system, we demonstrate the complete workflow under a range of KCl concentrations and show that moderate ionic strength stabilizes the PCNA trimer while very high salt partially destabilizes the complex. The approach is general and applicable to any reversible protein self-association reaction accessible by fluorescence detection at low protein concentrations.

Optical microscopy is fundamental to modern life-science research, yet interpreting its results requires precise modelling of point spread functions (PSFs) within complex environments. This manuscript introduces a versatile and efficient approach to wave-optical PSF calculations that extends existing frameworks by incorporating detection PSF modelling through the principle of reciprocity. Accompanying this work is a free MATLAB software package centred on a single, minimalistic core function, PlaneWaveExc.m, which utilizes a plane-wave superposition based on the Richards-Wolf model. Despite its simplicity, the framework accounts for "real-life" complexities such as systemic aberrations, arbitrary amplitude and phase modulations, and stratified media with complex-valued refractive indices. We demonstrate the softwares broad applicability through diverse case studies, including single-molecule imaging, STED microscopy, the segmented aperture of the James Webb Space Telescope, and coherent wide-field iSCAT microscopy. Each example is supported by dedicated scripts to facilitate adaptation for specific research needs.

Biomolecular condensates are important regulators of cellular compartmentalization and biochemical processes. Understanding their material properties is critical to elucidate how they control molecular organization and dynamics within cells. However, quantitatively probing these properties remains challenging due to the wide range of length scales, concentrations, and timescales over which condensates operate, as well as the limited force ranges accessible to current nanoscale mechanical mapping methods. We explored the use of a non-contact 3D imaging tool Scanning Ion Conductance Microscopy (SICM) for stiffness measurements of liquid-liquid phase-separated biomolecular condensates. We focus on the Dhh1 protein, which is a regulator of cytoplasmic processing bodies (PBs) membrane-less cytoplasmic condensates that control the storage and degradation of untranslated mRNA. In our study, we investigate the properties of mCherry2- or His-mCherry2- tagged full-length Dhh1 and N- or C-terminus tail-deletion constructs, as well as the catalytically inactive mutant DQAD, under different pH and incubation times. We mapped both spatial and temporal changes in the material properties of the condensates, highlighting the capabilities of the instrument. We found that the removal of either of the two tails led to an increase in condensate stiffness upon shifting the pH from a stress-associated cellular environment (pH 6.5) to physiological conditions (pH 7.5). Additionally, the choice of protein tags led to vastly different results depending on the pH where mCherry2-Dhh1 exhibited a stiffening going from pH 6.0 to 6.5 while the double-tagged His-mCherry2 did not. Our measurements are verified and corroborated by established techniques such as optical tweezer-based fusion assays and fluorescence recovery after photobleaching (FRAP). Furthermore, we were able to track the same biomolecular condensate sample for up to 20 days getting insights on the ageing and evolution of the condensates. Overall, our study demonstrates the applicability of SICM for direct measurement of the material properties of biomolecular condensate.

Photoactivated intermolecular electron transfer (ET) in cryptochromes proceeds along chains of aromatic residues and creates a spatially separated pair of radical electrons. Ultrafast time-dependent spectroscopy can provide experimental insight into this process and theoretical estimates of charge transfer rates are commonly obtained via Marcus theory. Here, we present a new perspective on the ET in European robin cryptochrome 4a (ErCry4a) that synthesizes insights from real-time ET calculations, ultrafast spectroscopic measurements and analytical derivations. The simulations exemplify that molecular vibrations play an essential role in enabling the ET dynamics, which was further rationalized through analytical derivations. Ultrafast pump-probe spectroscopy provided experimental access to the first 1.5 ns of the ET cascade, where multiple radical pair recombination rates arise due to the dynamic equilibrium along the ET chain. We show that the motions of the protein environment and the ET dynamics are inseparably coupled, violating the timescale separation required for Marcus theory. The presented results highlight that non-equilibrium coupling between electronic and nuclear motion dominates ET kinetics in ErCry4a during the first nanosecond after photo-excitation. The findings exemplify the limits of Marcus theory and refine the interpretation of ultrafast spectroscopic signatures in cryptochromes.

Many biological macromolecules are activated upon ligand binding at multiple specific binding domains. However, how these domains interact and the transient intermediate conformations that connect binding events to protein activity are typically unknown. Ensemble-averaged measures over stochastic binding events are challenged to resolve the underlying asynchronous dynamics. Single-molecule resolution of these dynamics offers an attractive approach to investigate the ligand-activation process. Optical methods using fluorescently labeled ligands enable observation of individual binding events that report on the energetics of early ligand-bound conformational changes. However, diffraction-limited microscopy limits these methods to low ligand concentrations, often below what is required for physiologically relevant activation. Here, we overcome this limitation using nanophotonic zero-mode waveguides to observe the sequential binding of a fluorescent cyclic nucleotide to each of four subunits in TAX-4 cyclic nucleotide-gated ion channels in cell-derived vesicles. Our observations suggest that binding at one domain positively promotes binding at other domains, and that binding induces an isomerization of the binding domain in individual subunits which we attribute to a sequence of pre-activated intermediate states. This approach provides a broadly applicable tool to dissect the energetic landscape of ligand-binding at macromolecules in native cell membranes.

Forster resonance energy transfer (FRET) is a physicochemical phenomenon involving non-radiative energy transfer between donor and acceptor fluorophores. While FRET efficiency primarily depends on the proximity between fluorophores, additional factors also substantially influence the efficiency in living cells. However, how non-distance factors modulate live-cell FRET efficiency remains poorly understood. Here, we report the significant role of N- and C-terminal topology in determining live-cell FRET efficiency, independent of fluorophore proximity, donor variants, and subcellular compartment. Using acceptor photobleaching and sensitized emission measurements in living cells, we found that FRET efficiencies of mCherry-EGFP or mCherry-EYFP (acceptor-donor) were significantly higher than those of EGFP-mCherry or EYFP-mCherry (donor-acceptor), respectively. These efficiencies were comparable between the nucleus and cytoplasm. An orientation index analysis showed that the acceptor-donor configuration is more favorable than the donor-acceptor configuration regardless of donor variants and subcellular localization. FRET efficiencies were also higher with EYFP than with EGFP as the donor. AlphaFold2-based structural modeling suggested similar proximity with structurally heterogeneous and loosely constrained geometry of donor and acceptor fluorophores. Collectively, these results demonstrate that topological arrangement, rather than simple distance considerations, plays a significant role in FRET efficiency in living cells, providing molecular implications for the design of intramolecular FRET-based biosensors.

Following ER Ca2+ depletion, Ca2+ release-activated Ca2+ (CRAC) channels are activated by STIM1 at ER-plasma membrane junctions. The restricted localization and low conductance of the CRAC channel (<40 fS) precludes single-channel recordings, limiting studies of CRAC channel gating. Here we describe an optical approach to characterize the gating of HaloTag-fused Orai1 channels labeled with JF646-BAPTA, a Ca2+-sensitive fluorescent dye. While Ca2+ influx through single channels generates fluorescence fluctuations, identifying true gating events is complicated by stochastic transitions of JF646-BAPTA to a non-fluorescent state. To overcome this, we combine TIRF microscopy with whole-cell voltage clamp to control the driving force for Ca2+ entry. We show the open channel intensity at -100 mV reflects Ca2+ saturation of the dyes on each channel, while the closed-channel intensity is defined by the fluorescence at +30 mV, where influx is absent. True gating events can be identified from transitions between the open- and closed-channel levels, distinguishing them from transitions to a non-fluorescent state. We describe the gating behavior of CRAC channels activated by STIM1 after store depletion. Dwell time distributions indicate at least two open and closed states with durations of 0.1 to several seconds, with most channels having an open probability of [&ge;]0.7. We also detect silent channels that colocalize with STIM1 but show no activity over tens of seconds, a population that would be undetectable by whole-cell electrophysiology alone. This method offers an approach to explore CRAC channel gating mechanisms and may be applicable to other Ca2+- permeable channels not amenable to patch-clamp techniques.

The spatial resolution of optical imaging systems is fundamentally restricted by the diffraction limit. However, in widefield live-cell microscopy, the achievable resolution is further constrained by the specimen motion, which indicates the existence of a fundamental spatio-temporal resolution trade-off between signal accumulation during the full frame integration and the resulting motion blur. To improve the fidelity with which moving objects can be imaged, a quantitative understanding of this spatio-temporal trade-off is necessary. Here, we present a systematic analysis of motion-induced resolution dynamics measured with spectral signal-to-noise ratio (SSNR). We developed a simulation framework which models the image formation of objects undergoing arbitrary motion, to evaluate the degradation of the spatial resolution under translational and rotational dynamics. Our results demonstrate that for translating objects, the spatial resolution is anisotropically reduced as a function of the orientation of the object relative to the motion vector, leading to the spectral signal-to-noise ratio degrading by up to 50% and the resolution by up to 40% for a 90{degrees} change in the motion direction. Furthermore, we show that for rotational motion, conventional radially averaged metrics such as the Fourier Ring Correlation are not able to quantify the effects of angular blur. On the other hand, the SSNR is able to accurately quantify this degradation. These findings underscore the necessity of an object-oriented imaging approach, in which acquisition parameters such as exposure time are tuned to specific biological spatio-temporal characteristics to optimize the trade-off between motion blur and spatial fidelity.

Fluorescent dyes are critical to visualizing nucleic acids in many applications. SYTOX Orange and SYTOX Green are cyanine dyes, used in dead cell staining and increasingly in single-molecule assays to probe DNA supercoiling and processing. However, their structures and effects on DNA mechanics are not or only partially known. We determine the structure of SYTOX Orange to be (E)-2-((2-(4 ((diethyl(methyl)ammonio)methyl)phenyl)-6-methoxy-1-methylquinolin-4(1H)-ylidene)methyl)-4-methyloxazolo[4,5-b]pyridin-4-ium, identical to SYBR Gold except for an aza-benzoxazol core that is fundamentally different from other dyes in the SYTOX and SYBR families. We report SYTOX Green to be (Z)-2-(bis(3-(trimethylammonio)propyl)amino)-4-((3-methylbenzo[d]thiazol-2(3H)-ylidene)methyl)-1-phenylquinolin-1-ium, similar to PicoGreen. Using magnetic tweezers, we characterize the effect of SYTOX Orange and SYTOX Green on DNA mechanics. They lengthen and unwind DNA consistent with intercalation and the DNA unwinding angles per dye are 21.1(1) degree and 20.5(1) degree for SYTOX Orange and Green, respectively. Both dyes leave the DNA bending persistence length and plectoneme size almost unaltered (<10% change up to 1 uM), which is advantageous in assays probing DNA supercoiling. Their photophysical properties reveal close agreement between single-molecule manipulation and optical absorbance and fluorescence spectroscopy. Our comprehensive set of complementary measurements relates mechanical and optical properties to the molecular structures and provides recommendations for their use in applications.

Mechanotransduction via the actin cytoskeleton is linked to fundamental cellular processes such as morphogenesis, cell division, and motility, requiring the control of tensile forces mediated by the motor protein non-muscle myosin 2 (NM2). Formins such as mDia1 have been shown to elongate actin structures that are under mechanical tension; conversely, mDia1s elongation rates are modulated by the applied force. Despite their relevance at the membrane/cortex interface, reported values for tension in formin-elongated actin filaments stem from theoretical estimates and simulations, but have not been amenable experimentally so far. Thus, we developed a Forster resonance energy transfer (FRET)-based, tension-sensitive probe (mDia1TS) and quantified the measured tension in live U2OS cells using fluorescence lifetime imaging microscopy (FLIM). Through whole-cell ROI analysis we show a short and long lifetime component, reporting an intensity-weighted, averaged lifetime corresponding to [~]3.5 pN. Upon mitogen stimulation of cells using EGF, we show that the tension homeostasis changed significantly, with a measurable increase in tension in the cells periphery and relaxation in its center. Furthermore, the reported average tension relaxed by 2 pN after adding the NM2 inhibitor para-nitroblebbistatin. We utilized siRNA knockdowns of individual NM2 paralogs (NM2-A, NM2-B, or NM2-C) to measure their individual contribution, revealing NM2-A as the main paralog to produce tensile force in this system. Taken together, we demonstrate that mDia1TS is able to directly determine that active mDia1 in cells is under tension, and that subcellular quantification with pN precision is possible. SignificanceDespite the fundamental importance of formins in regulating actin-based processes, reported values for tension in formin-mediated actin structures stem from simulations and theoretical estimates. In this study we developed a FRET-based, tension-sensitive reporter probe for formin mDia1, which we termed mDia1TS. Given the expanding clinical spectrum of DIAPH1/mDia1 mutations, our tool mDia1TS provides a quantitative tool for elucidation of changes in cytoskeletal assemblies.

GPCR oligomerization has been reported for decades, yet its extent and functional relevance in living cells remain unresolved because existing approaches, often done in bulk, are poorly account for local receptor density, a major determinant of intermolecular interactions. Here, we establish a generic quantitative imaging framework that links spatially resolved FRET measurements describing protein oligomerization to local membrane protein in living cells. Using automated high-throughput analysis of fluorescence images, the method generates large density-resolved datasets that enable direct quantification of receptor oligomerization parameters, including apparent affinity, oligomerization state, and monomer/dimer populations at the submicrometer scale. Applied to class A GPCRs in HEK293 cells, the approach reveals receptor-specific density-dependent equilibria between monomers and dimers over physiologically relevant expression ranges, with no evidence for stable higher-order oligomers under basal conditions. The receptors studied exhibit distinct apparent affinities for dimerization, ranging from predominantly monomeric to dynamic monomer-dimer equilibria, indicating that local membrane density strongly influences receptor organization and that it is receptor dependent. The agreement between our measurements and low-density single-molecule studies further suggests that previously reported higher-order oligomers may partly reflect density-driven receptor proximity effects. By bridging single-molecule and ensemble measurements within a unified quantitative framework, this work reconciles conflicting observations in the GPCR oligomerization literature and provides a broadly applicable strategy for investigating membrane protein organization in living cells. SignificanceGPCR oligomerization in living cells is strongly influenced by the local protein density, yet most approaches do not quantitatively account for this parameter. Here, we introduce a quantitative high-throughput imaging framework that directly relates membrane protein local density to local oligomerization state in living cells. Applied to distinct GPCRs over physiologically relevant density ranges, the method reveals distinct density-dependent monomer-dimer equilibrium and apparent affinities for self-association. These results help reconcile longstanding discrepancies, where distinct oligomerization states have been measured depending on experimental conditions. More broadly, this work establishes local membrane protein density as a key determinant of membrane protein organization, and provides a quantitative framework applicable to membrane protein complexes in their native cellular context.

Imaging the structural organization inside cells in their native state is essential for understanding how the arrangement and interactions among molecular components give rise to biological function. Fluorescence microscopy is one of the pivotal techniques that provides molecular specificity for imaging in real space, however, the technique is limited to labeled components. X-rays, on the contrary, are sensitive to electron density contrast and therefore to label-free samples, and probe structure in reciprocal space. In particular, scanning small-angle X-ray scattering (SAXS) combines information from real and reciprocal space and enables access to intact cells, owing to the high penetration power of the X-rays. Combining both imaging modalities in a synergistic manner promises powerful tools for cellular imaging, but remains challenging, because of the differing requirements the complementary methods introduce. Here we present a correlative imaging platform that integrates a modular, compact and beamline-compatible fluorescence microscope with scanning SAXS, to enable fast sequential imaging of the identical cellular regions. We developed a dedicated microfluidics flow chamber enabling measurements under hydrated, near-native conditions. We demonstrate the utility of our methodology by investigating two different relevant cellular components, i.e., thick keratin bundles in epithelial cells that contribute to cell mechanics, and force-generating actomyosin in cardiomyocytes. Employing adapted data analysis methods, we find a good agreement between the fluorescence-derived and the SAXS-derived orientation maps. This result demonstrates that the label-free approach with SAXS captures cytoskeletal organization through-out the cell, and can be directly linked to specific molecular information provided by the complementary fluorescence imaging, in a physiologically relevant cellular environment. Our work establishes a general strategy for multimodal imaging of cellular architecture and opens ways to investigate living cells under the influence of drugs and chemical manipulation experiments.

Proteins in solution adsorb to the corona of nanoparticles such as single-walled carbon nanotubes (SWCNTs), but these interactions are difficult to predict and analyze due to ambiguities in the structure of the latter. In this work, we employ ss(GT)15-DNA wrapped SWCNTs, a commonly used fluorescent sensor construct, to examine protein adsorption by quantifying binding dissociation constants and characterizing the corresponding photophysical effects. A library of 20 proteins are used to evaluate adsorption-induced changes in photoluminescence (PL) intensity ({Delta}I/I0) and emission wavelength upon solution phase binding. We find that 15 proteins produce monotonic dose-response behavior well described using a single-site Langmuir model. Alternatively, five proteins exhibited more complex, non-monotonic behavior consistent with a two-step binding model representing protein-protein interactions coupled to adsorption. The study reveals that metalloproteins, which comprised 12 of the 20 proteins in the library, induced greater PL quenching compared with metal-free proteins for this system, with maximum binding-associated quenching ({Delta}I/I0) of 94% for metalloproteins versus 20% for metal-free proteins. For metalloproteins, we introduce a proximity-based quenching framework in which protein size provides a coarse proxy for cofactor-SWCNT separation, offering a mechanistic interpretation of the observed quenching variation across proteins. Together, these results establish the use of metal coordination sites, such as those in metalloproteins, to assist the transduction of certain nanoparticle fluorescent sensors, helping with sensor probe design and interpretation in biological environments.

Stimulated emission depletion (STED) microscopy is a super-resolution fluorescence imaging technique that achieves high spatial and temporal resolution by exploiting stimulated emission to induce fluorescence depletion (FD) and is expected to have substantial utility for imaging applications using fluorescent proteins. However, the compatibility of fluorescent proteins with STED microscopy systems has been understood primarily through empirical observations, and there is no established methodology for the rational selection of fluorescent proteins for STED microscopy. In this study, we systematically evaluated the compatibility of commonly used fluorescent proteins with STED microscopy systems by measuring FD properties using transient absorption spectroscopy and fluorescence dip spectroscopy, both of which are classified as two-color spectroscopy (TCS). Fluorescent proteins identified as compatible with the STED microscopy system based on the TCS measurements were employed for three-dimensional STED imaging of cellular samples expressing each protein. In all samples, three-dimensional spatial resolution was improved relative to confocal laser microscopy, with particularly marked improvements in z-axis resolution. These findings demonstrate that measurements of FD properties via TCS provide a robust approach for evaluating the compatibility of fluorescent proteins with the STED microscopy system and for selecting suitable fluorescent proteins for STED imaging.