Nano Letters

Silica-encapsulated gold nanostars (AuNStar-SiO2) are a widely used plasmonic nanoparticle platform for surface-enhanced resonance Raman scattering (SERRS) bio-applications. In this paper, we demonstrate that coupled nanostar subpopulations can dominate the ensemble-average SERRS response of the suspension and that near-neutral standard cell culture conditions are sufficient to hydrolyze the silica nanoshell and introduce variability in signal intensity following in vitro endocytosis. Monomeric and oligomeric AuNStar-SiO2 fractions were isolated using continuous density-gradient centrifugation and monomeric populations were found to exhibit significantly weaker SERRS compared to their oligomeric counterparts. Using monomer-enriched AuNStar-SiO2, we investigated the stability of the silica nanoshell under conditions representative of sequential acidification during endocytosis and characterized the subsequent changes to nanoparticle optical properties. In acidic environments, reflecting lysosomal pH, the silica shell was stable, whereas near-neutral and alkaline conditions in cell culture medium induced silica-shell hydrolysis, nanostar release, and interparticle aggregation, leading to transient SERS amplification. When cells were treated with AuNStar-SiO2 under near-neutral and acidic conditions, we observed the opposite trend in SERS signal strength. At pH 7.4, the SERRS signal was suppressed even though transmission electron microscopy (TEM) images of intracellular nanoparticles showed progressive extents of silica hydrolysis, while at pH 6.4 SERS signal was strong and the silica shell of intracellular nanoparticles remained intact. Together, these findings show how SERRS output can differ between control conditions and biological applications, highlighting the role that local environmental factors play in nanoparticle stability and performance. Our results highlight the previously overlooked role of silica nanoshell instability on SERRS signal output in physiological environments and describe opportunities to harness silica nanoshell hydrolysis to improve the biomedical application of silica-coated plasmonic probes.

Quantitative measurements performed directly in vivo are necessary to understand how forces shape living tissues, yet this remains challenging due to optical scattering and mechanical complexity. Here, we present a method for making absolute force measurements using nanoscopic optical tweezers with a sensitivity of 300 fN in optically turbid biological media. Our approach combines back focal plane interferometry operating within the optical memory effect regime with a global fluctuation-dissipation fitting framework that simultaneously calibrates position detection, trap stiffness, and viscoelastic response. This method overcomes aberration-induced biases by jointly fitting passive fluctuations and driven harmonic responses, enabling robust force reconstruction in thick, scattering tissues within the mechanically relevant frequency range below 300 Hz. We validate our approach using highly scattering Drosophila pupae and embryos, demonstrating reliable in vivo measurements of forces and mechanical properties. Operating at a 1 kHz acquisition bandwidth, the system captures relevant mechanical dynamics without requiring extended high-frequency detection. Using this framework, we quantify the increase in cortical tension during pupal morphogenesis, characterize tissue viscoelasticity, and reveal stage-dependent variations in nuclear membrane tension during embryogenesis, even in the presence of strong ATP-driven fluctuations. Beyond bulk measurements, our method enables the quantitative mechanical characterization of single cells within mechanically coupled tissues.

Accurate characterization of the nanoparticle (NP) protein corona is essential for predicting biological fate, safety, and therapeutic efficacy, and for enabling robust biomarker discovery. Standard isolation techniques, most commonly centrifugation and magnetic separation, are widely used, yet they rarely account for co-isolating endogenous biological NPs such as extracellular vesicles (EVs). This oversight can distort the apparent "biological identity" of the NP. Here, we quantitatively demonstrate the magnitude and impact of EVs on the perceived protein corona composition. We incubated highly monodisperse polystyrene NPs (50-1000 nm) and superparamagnetic beads in either standard human plasma or plasma depleted of EVs by immunoaffinity capture targeting 37 EV surface epitopes. Mass spectrometry revealed that EV depletion reduced the number of proteins identified on polystyrene NPs by 60-75% and on magnetic beads by 45-50%. Importantly, EV depletion also altered the apparent abundance hierarchy; it restored the expected relative abundance and rank of major plasma proteins such as albumin and shifted the top-ranked proteins from intracellular cytoskeletal component, consistent with EV carryover, to genuine soluble plasma adsorbates (e.g., apolipoproteins, complement factors). These results highlight that standard corona workflows can inadvertently co-isolate a vast array of EV-associated proteins, yielding inaccurate proteomic profiles. Discriminating genuine corona proteins and EV-associated contaminants is critical for advancing nanomedicine, ensuring predictive safety and efficacy profiles, and enhancing the precision of NP-based biomarker discovery.

The dynamic behavior of cellular membranes underpins essential biological processes, including signal transduction, intracellular trafficking, and mechanotransduction. However, simultaneously quantifying lateral molecular diffusion and vertical membrane fluctuations in live cells remains challenging. Here, we present dynamic metal-induced energy transfer spectroscopy (dynaMIET), which integrates metal-induced energy transfer with fluorescence correlation spectroscopy to resolve three-dimensional membrane dynamics with nanometer axial sensitivity and microsecond temporal resolution. dynaMIET enables concurrent measurement of lateral diffusion and vertical undulations within a single acquisition. We validate the method using simulations and model membranes and demonstrate its robustness in living cells, applying it to the plasma membrane, endoplasmic reticulum, and nuclear envelope. By capturing both molecular mobility and membrane fluctuations, dynaMIET provides a powerful, non-invasive tool for probing membrane mechanics and organization. This advance opens new avenues for studying membrane-associated phenomena in health and disease, including cancer cell mechanics, protein-membrane interactions, and organelle dynamics.

Plasma membrane (PM) lipids and proteins are organized into nanoscale regions called nanodomains, which regulate essential cellular processes by controlling local membrane organization. Despite advances in super-resolution microscopy and single particle tracking, the small size and temporal instability of nanodomains make them difficult to study in living cells. To overcome these challenges, we built fluorescent DNA origami probes that insert into the PM via lipid anchors displayed on the cell. The number and spatial distribution of anchors between the origami and the cell surface were precisely defined by the origami, enabling nanometer-scale sampling of the cell surface. Inserting these DNA origami particles into the membrane with lipid anchors allowed them to passively diffuse across the membrane, and we tracked their movement using single particle tracking to survey the PM landscape. By varying the number and spatial arrangement of lipid anchors connecting the DNA origami to the cell surface, we showed that immobilization of DNA origami particles requires simultaneous interactions with multiple nanodomains. Disruption of the actin cytoskeleton reduced immobilization, confirming its role in supporting nanodomain stability. Moreover, transient mechanical stretching of cells led to reversible increases in DNA origami mobility, indicating that mechanical force can reversibly regulate PM nanodomain organization. Altogether, we present a novel membrane-integrated DNA origami approach that provides mechanistic insights into PM nanodomain architecture and dynamics in living cells.

Chimeric antigen receptor (CAR) T cell therapies have reshaped treatment for cancers and immune-mediated diseases, yet their safety and efficacy depend on both the proliferation of engineered cells and their dynamic functional state -- features that remain challenging to monitor in real-time clinical settings. Current methods require labels, extensive processing, and provide only static snapshots of cell identity and activation. Here, we introduce a surface-enhanced Raman spectroscopy and machine learning approach that enables label-free single-cell identification of engineered CAR T cells and time-resolved, semi-continuous monitoring of their functional activation state. Using the intrinsic vibrational signatures from live cells, we detect spectral differences resulting from engineered receptor expression in donor-derived CD19- and GD2-targeted CAR T cells (nine and five donors, respectively) with 81-85% donor-level accuracy and resolve dynamic antigen-specific activation trajectories with temporal precision. These capabilities stem from biochemical signatures consistent with processes such as receptor expression, tonic signalling, and immune synapse formation, demonstrating a single method that reports both cellular identity and activation state with biochemical specificity. Our results extend CAR T cell monitoring beyond static phenotyping and establish the potential of SERS-ML analysis for rapid, point-of-care assessment of engineered immune cells.

Cathodoluminescence (CL) microscopy has the potential to achieve a key goal in biological imaging: the simultaneous visualization of proteins and cellular ultrastructure. This goal can be attained by tagging proteins of interest with spectrally distinct cathodoluminescent probes for detection in electron microscopy. To this end, lanthanide nanoparticles (LNPs) are promising probe candidates due to their stability under the electron beam and their distinct ion-dependent emission spectra suitable for multiplexed detection. However, the hydrophobic surface chemistry of LNPs limits their use in biological samples and requires surface functionalization compatible with aqueous environments and EM sample preparation protocols. Here, we use a DNA-based ligand exchange strategy that renders cathodoluminescent LNPs hydrophilic and compatible with further functionalization for specific protein labeling. We characterize the CL emission of DNA-functionalized LNPs following aqueous transfer and common EM preparation steps, including osmium tetroxide staining and drying protocols based on hexamethyldisilazane and critical point drying, and show that LNPs retain their CL emission under all tested conditions. Finally, we demonstrate multicolor CL imaging of spectrally distinct, DNA-functionalized LNPs on the surface of mammalian cells, enabling simultaneous visualization of cellular ultrastructure via secondary electrons and LNPs via multiple CL color channels.

Targeted lipid nanoparticles (tLNPs) represent the next frontier in nucleic acid therapeutics, enabling cell-specific delivery through covalent attachment of targeting ligands that drive receptor-mediated uptake. tLNPs are particularly promising for pregnancy-associated applications where precise on-target delivery is required to minimize maternal toxicity and protect fetal health. Yet, their rational design is limited by an incomplete understanding of how tLNP physicochemical properties influence biological performance. Conventional LNPs already exhibit pronounced heterogeneity in size, composition, and RNA loading, which is further amplified in tLNPs by variability in ligand attachment and surface density. Because traditional analytical methods report only ensemble-averaged properties, the nanoscale diversity of tLNPs remains unresolved. Here, we find that tLNP functional behavior is governed by previously inaccessible, structurally distinct tLNP subpopulations that are not captured by bulk measurements. We utilize asymmetric flow field-flow fractionation integrated with in-line UV spectral analysis, light scattering, and synchrotron small-angle X-ray scattering (AF4-UV-DLS-MALS-SAXS) to resolve ligand-dependent tLNP subpopulations that differ in size, shape, composition, and relative abundance. We find that protein conjugation preserves the internal lipid-RNA nanostructure of base LNPs but substantially increases particle heterogeneity, particularly for larger and multivalent targeting ligands. Despite increased heterogeneity, tLNPs functionalized with higher-avidity ligands achieve more effective targeted placental RNA delivery in mice, suggesting that binding avidity can offset the functional consequences of polydispersity. Chemometric SAXS analyses reveal that only SAXS-resolved tLNP subpopulations, not ensemble-averaged parameters, correlate with targeted placental transfection in vivo, whereas bulk-derived physicochemical metrics more strongly associate with nonspecific hepatic delivery. Together, this work harnesses a separation-coupled biophysical platform to resolve previously inaccessible tLNP subpopulations and demonstrates that subpopulation nanoscale structure, rather than bulk-averaged properties, dictates targeted RNA delivery. These insights provide a mechanistic foundation for rational engineering of next-generation precision targeted RNA LNP therapeutics.

Immune cell receptor - ligand interactions are key to cancer immunotherapy. However, receptor-ligand affinities often fail to predict T-cell mediated cancer killing, while immune-target cell binding strength measurements are limited by low precision and high non-specific binding. Here we present bilayer acoustic force spectroscopy (BAFS), a method to quantify the binding strength of receptors in immune synapses that virtually eliminates non-specific binding and increases the resolving power by up to 50-fold. By replacing target cells with a supported lipid bilayer functionalized with antigens, BAFS avoids antigen-independent interactions and target cell heterogeneity, while maintaining the spatial self-organization of receptors that typifies active immune synapses. We demonstrate the high sensitivity and control by showing how CAR T-cell synapse strength depends on CD19 antigen density, and by revealing that CD8 synergistically strengthens {beta}TCR-pMHC synapses independently of Lck recruitment to CD8. BAFS is a general method that can be used broadly in immunotherapy screening and to dissect the complex molecular interactions that underpin immune synapse activation.

The bottom-up construction of synthetic cells based on giant unilamellar vesicles (GUVs) is a central goal in synthetic biology. Achieving targeted changes in membrane and cytoplasmic composition with temporal control remains challenging however. DNA-mediated fusion with small vesicles ([~]100 nm large unilamellar vesicles; LUVs) has been proposed as a strategy to deliver lipids and cytosolic contents in a programmable manner. However, in vitro, membrane fusion is generally found to be inefficient and poorly controllable for reasons that are poorly understood. Here, we present an approach based on lipid-conjugated DNA (LiNA) to mediate programmable fusion between LUVs and micron-sized GUVs, which we quantitatively monitor with confocal microscopy at the single-GUV level. We show that lipid and content mixing both occur with high efficiency over a wide range of LiNA concentrations, demonstrating that LiNAs indeed induce robust membrane fusion. Furthermore, we show that LiNA-mediated fusion provides a powerful tool to deliver cytosolic biomolecules, enabling control over internal activities. Our findings establish a quantitative framework for studying fusion-driven processes in synthetic cells and provide a versatile platform for the programmable delivery of lipids and cytosolic cargoes - thus advancing the development of synthetic cells that can grow and adapt through fusion-based uptake of molecular building blocks.

Mechanical forces at the cell-substrate interface govern processes from migration to differentiation, yet mapping these forces at high spatial resolution remains challenging. Traction force microscopy (TFM) addresses this by quantifying substrate deformations using fiducial markers, which are conventionally fluorescent beads. Here, we introduce fluorescently labeled DNA nanostructures (FluoroCubes) as alternative fiducials grafted onto polydimethylsiloxane (PDMS) substrates. Co-anchored with RGD peptides, FluoroCubes remain stably tethered, resist internalization, and enable dense, minimally perturbing labeling. This surface-functionalized platform is compatible with TIRF microscopy and leverages tunable biotin-NeutrAvidin chemistry for precise control of fiducial density. Using a modified multi-channel optical flow algorithm, we achieve improved displacement sensitivity and force reconstruction resolution compared to conventional algorithms. FluoroCube-functionalized substrates provide a reproducible, high-resolution method for traction force mapping and offer a versatile foundation for future integration with DNA-based molecular sensors to probe interfacial forces at biointerfaces.

Epithelial ovarian cancer remains one of the most lethal malignancies among women, with late-stage diagnoses yielding 5-year survival rates below 30%. The metabolic heterogeneity of the tumor microenvironment (TME) highlights the need for methods capable of rapid, chemically specific phenotyping. Stimulated Raman scattering (SRS) microscopy when combined with deuterium labeled metabolites enables the non-invasive high contrast interrogation of cellular metabolic pathways. In this study, we used SRS microscopy to profile fatty acid and glycogen metabolism in epithelial ovarian cancer (SKOV-3) and cervical cancer (HeLa) cell models. Deuterium labeled glucose revealed striking differences in glycogen synthesis and intracellular distribution, with SKOV-3 cells exhibiting markedly greater single-cell heterogeneity than HeLa. Complementary measurements of lipid droplet (LD) synthesis and turnover under nutrient starvation further revealed cell-line-specific metabolic strategies, identifying LD and glycogen dynamics as a potential diagnostic marker of cancer metabolic phenotypes. These results demonstrate that SRS microscopy in the Raman silent region, paired with metabolic labeling, can sensitively resolve metabolic diversity across cancer cell subpopulations. Such metabolic phenotyping may inform both early diagnostic strategies and therapeutic approaches that combine cytotoxic treatment with targeted metabolic disruption.

The acoustoelectric (AE) effect, in which acoustic waves modulate the electrical properties of a conductive medium, holds significant potential for biomedical imaging. While classic models describe the phenomenon through conductivity modulation, a detailed understanding of its microscopic origins, particularly the role of ion behaviours, remains lacking. This study introduces a novel electrokinetic perspective by investigating how ultrasound modulates ion-solvent interactions, thereby bridging macroscopic AE signals with underlying ion dynamics. Through finite element simulations of a dilute NaCl solution, we demonstrate that acoustic pressure waves induce local variations in ion mobility and diffusion by altering ion hydration shells and solvent viscosity. These changes disrupt the balance among Coulombic, diffusive, and frictional forces on individual ions, leading to the local conductivity modulation. Furthermore, simulations reveal that acoustic perturbation of the electrode-electrolyte interface (EEI) significantly enhances AE signal generation, highlighting the EEIs critical role in AE-related applications. By linking acoustic modulation to fundamental ion-solvent interactions, this work not only provides a foundation for more accurate, microscopically grounded models of the AE effect but also connects AE effect modelling to the active research of solvation dynamics in physical chemistry.

Affibodies are remarkably stable three-helix bundle proteins that can be engineered to selectively bind target proteins. When combined with radioactive metals, they serve as imaging agents or cancer therapeutics, depending on the metal used. Traditionally, this involves bifunctional linkers that attach large chelators to the affibody via reactive groups. Here, we present an alternative approach that eliminates the need for such linkers by burying the metal within the core of the affibody, surrounded by its three helices. A simple engineered triple cysteine motif, with one cysteine in each helix, stably binds Bi(III), Pb(II), In(III) and Ga(III), which are commonly used in imaging and radiotherapy. Quantitative metal uptake is instantaneous at room temperature and physiological pH, and all metal-affibody complexes remain fully intact for one week at 4 {degrees}C. All retain their metal cargo when challenged with cellular concentrations of glutathione, while only the bismuth-affibody complex withstands a challenge with 100 equivalents of strong chelators, even over two weeks. We demonstrate selective uptake and retention of 213Bi, a promising isotope for targeted alpha therapy.

Super-resolution microscopy with DNA-PAINT enables molecular-scale, multiplexed, and quantitative imaging, but its throughput is limited by slow binding kinetics and elevated background at high probe concentrations. Recent speed-optimized and fluorogenic probes improve performance but impose strong constraints on sequence design, revealing a fundamental tradeoff between fast binding and efficient quenching. Here, we introduce a modular probe architecture that spatially decouples binding kinetics from fluorophore-quencher interactions by integrating speed-optimized sequence motifs with PEG spacers. Using DNA origami nanostructures, we demonstrate enhanced localization rates, signal-to-background ratios, and imaging efficiency compared to state-of-the-art probes. We validate our approach in cells, demonstrating its capability to image nuclear targets and enabling three-dimensional imaging of the endoplasmic reticulum using standard widefield illumination. Our work establishes a general framework for fast, multiplexed, and low-background super-resolution imaging.

Fluorescence microscopy is a cornerstone of biological research. However, fluorescent labeling is challenging in live cells and is constrained by photobleaching and phototoxicity. Label-free methods allow cells to be studied in their native state, but most techniques have poor contrast, lack 3D capability, rely on complex optics, and fail to provide structural information. We present broadband backscattering confocal microscopy (BBCM), which employs a broadband supercontinuum laser and collects backscattered light in confocal geometry using a photomultiplier tube. Broadband illumination averages out size-dependent oscillations that confound monochromatic backscattering. This eliminates blind spots and intensity ambiguities, allowing all scatterers to be visible, with the signal increasing approximately linearly with scatterer size. BBCM is easy to retrofit to standard confocal microscopes, requires no specialized optics, and is straightforward for nonspecialists. It enables high-contrast, label-free 3D imaging of live cells with size sensitivity to subcellular structures without employing custom optics or complex data processing.

Self-assembled DNA nanostructures show great promise as functional devices, highly configurable materials, and in nanorobotics. Magnetic control can provide a powerful actuation mechanism in a broad range of contexts, since it affords a high-level of external control, it is biocompatible, and orthogonal to chemical or electrical stimuli. Here we demonstrate magnetic molecular nanoactuators by leveraging the unique site-specificity of DNA origami to assemble highly anisotropic magnetic nanocubes on high-aspect ratio DNA origami bundles. We traced and controlled 100s of our DNA origami nanorotors at the single-rotor level and demonstrated their programmable magnetic clamping and controlled rotation under uniform and rotating magnetic fields. By varying the population and inter-particle spacing of the nanocubes, magnetic torque values in the order of 10-100 pN nm are achieved at field strengths < 10 mT. Monte Carlo simulations reveal that assembly of nanocubes on DNA origami rotors leads to collective magnetic properties, with numerically estimated torque values in good agreement with the experiments. Our magnetic nanorotors offer a foundation for biocompatible nanorobotics, as well as high-throughput magnetic force and torque tweezers.

Continuous biochemical sensing provides valuable insights into an individuals physiological state and the mechanisms underlying pathophysiological changes. However, most existing bioanalytical methods are not compatible with continuous biochemical sensing. A major technical challenge lies in achieving rapid measurement readouts while maintaining high specificity and sensitivity in complex biological fluids. Sensitive molecular detection typically requires slow analyte-binder dissociation and long incubation to reach equilibrium, whereas rapid and frequent measurements demand fast association-dissociation kinetics that are difficult to reconcile for low-abundance analytes. To address this challenge, we introduce a sensing mechanism termed photothermal recycling (PTR), which mimics the thermal cycling process in polymerase chain reaction. Using plasmonic photothermal effects, PTR rapidly recycles binders to enable frequent measurements. We demonstrate a digital PTR assay capable of multi-hour biochemical monitoring with subpicomolar(pM) sensitivity in buffer, diluted serum, and saliva. This approach leverages localized thermal energy to dynamically modulate biomolecular recognition, offering a new bioanalytical paradigm for continuous biochemical sensing across diverse application settings.

Fluorescence microscopic methods are critical for spatial profiling of multiple biological targets in cells and tissues to study cell and tissue functions, but their multiplexity was limited to 3[~]5 targets under a conventional setup using different fluorescent channels because of spectra overlap. Here, we introduce a simple, rapid, multiplexed fluorescence imaging method in cells and tissues, termed DNA based plasmonic heating activated signal exchange reaction (PHASER). PHASER uses infrared light-induced plasmonic heating of gold bipyramid nanoparticles to sequentially activate thermodynamically calibrated DNA thermal probes in situ for rapid and multiplexed fluorescent imaging of biological targets. We showed that the signal exchange per round between biological targets in PHASER can be completed within 30 seconds, and that 5 irradiation pulses of photothermal heating can activate DNA thermal probes with 5 different signal temperatures in cells and tissues. To demonstrate its practical use, we applied PHASER to profile the subcellular spatial organization of different organelles in cultured cells and resolved different protein spatial expression profiles in mouse brain tissue with dimensions of millimeters in a single fluorophore channel. PHASER is expected to have broad biotechnical applications with multiplexed fluorescence imaging for a wide variety of biological targets across diverse samples.

Understanding cellular heterogeneity in human skin is crucial for regenerative medicine and tissue engineering. In this study, we applied label-free Raman imaging to visualize molecular features corresponding to the three-dimensional architecture of the epidermis. Spatially resolved Raman spectra, combined with multivariate data analysis, enabled the identification of cell-layer-specific molecular signatures. Based on the region-specific spectra analysis, component C5 was predominantly localized to the basal layer within rete ridges and was characterized by {beta}-sheet-enriched keratin features. This spatially restricted distribution reflects the molecular microenvironment of epidermal stem cell niches, suggesting that C5 may serve as a biomarker for basal stem cell populations associated with skin undulations. These findings provide insight into the molecular basis of epidermal architecture and demonstrate the potential of Raman spectroscopy as a label-free tool for evaluating stem cell localization and differentiation status.