Abstract Spontaneous curvature characterizes the propensity of a membrane to bend in a specific direction. It is therefore crucial in the multitude of cellular processes that involve membrane shape remodelling. Yet, experimentally quantifying the spontaneous curvature remains a significant challenge in complex biological membranes, as their heterogeneity causes ambiguities in spontaneous curvature’s physical interpretation. Here, we introduce a general experiment-simulation framework to measure an effective spontaneous curvature using dual-direction tether pulling from cell-attached giant plasma membrane vesicles (GPMVs) and mesoscale simulations. For homogeneous membranes, the force difference between inward and outward pulls yields a tension-independent readout of spontaneous curvature. We show that this continuum observable can be generalized to the mean of the spontaneous curvature in a heterogeneous membrane, independent of the underlying microscopic spontaneous curvature distribution. Applied to HEK-derived GPMVs, a baseline negative spontaneous curvature of the plasma membrane is revealed. Sucrose treatment and extracellular addition of Annexin A5 systematically shift the effective spontaneous curvature, while mucin reporter overexpression does not measurably alter it under the conditions tested. We also measure the curvature imprint of individual fluorescently tagged proteins through a sorting index. Benchmarked with Annexin A5, our scheme recovers curvature imprints very similar to previous atomistic molecular dynamics simulations. Taken together this makes spontaneous curvature accessible as a directly measurable material property of native membranes and membrane-proteins, enabling quantitative studies of membrane remodelling across diverse cellular processes.

Subcellular protein complexes and organelles exhibit diverse dynamic behaviors that reflect the mechanical constraints and organization of the intracellular environment. Although some structures follow classical Brownian motion, many display anomalous dynamics. The transitions between these regimes are increasingly recognized as critical for subcellular organization, yet how they influence pattern formation remains unclear. Here, we investigate the spatial arrangement of cilia on the apical surface of multiciliated cells (MCCs) in developing Xenopus laevis embryos, where coordinated ciliary beating depends on the precise organization of hundreds of centriole-derived basal bodies (BBs). Using quantitative confocal, high-resolution and high-speed TIRF imaging together with theoretical modeling, we show that BB trajectories undergo time-resolved transitions between diffusive and anomalous motion, with distinct regimes that correlate with apical surface expansion. During the early stages, actin remodeling facilitates the dispersal of BBs by providing a permissive, low-confinement environment. As development progresses, the actin network becomes increasingly cross-linked that constrains BB movement and promotes uniform spacing across the apical domain. Disruption of α-actinin-1, a major actin cross-linking protein, impairs the integrity of the apical actin meshwork, weakens BB confinement, and disrupts regular spatial patterning, ultimately compromising the arrangement of BBs required for proper cilia alignment. Together, we show that progressive apical actin cross-linking coordinates BB positioning and regulates their dynamic state, guiding the shift from diffusive to confined motion. This transition in dynamics enables the emergence of a uniform BB pattern, which in turn ensures the aligned deployment of motile cilia necessary for effective directional fluid flow.

A thorough understanding of starch structure from crystal distribution level is crucial for production, processing, and application of starch. In this study, we utilized a custom-built polarization-based quantitative phase microscopy based on digital holography as well as a multimodal polarization microscope to investigate the crystalline structure of starch granules at the level of the Maltese cross. Through quantitative analysis, we observed that crystalline regions are distributed on both sides of the granules, with non-crystalline material extending radially in between, with amylose playing a significant role in disrupting this crystal alignment. This offers insights into why starch granules exhibit low overall crystallinity, often below 20%, as observed in maize and potato starches. Additionally, in situ enzymatic hydrolysis and digestion analysis of starch granules revealed the rapid breakdown of granular structures at specific points in the process. The crystalline heterogeneity, quantified as variation in birefringence intensity (with standard deviations of 25-50% for maize starch and 13-25% for potato starch), which was observed across the granule population of the same starch type, resulted in asynchronous granule disruption, emphasizing the structural complexity within starch granules. These findings contribute to a deeper understanding of current models of starch structure, particularly in terms of how crystalline and amorphous regions interact during gelatinization and enzymatic processes. This study offers deeper insights into starch structure-function relationships and underscores the value of digital holographic quantitative phase microscopy for starch analysis.

Abstract 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. Significance GPCR 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.

The cell membrane is crucial for cell survival, and ensuring its integrity is essential as the cell experiences injuries throughout its entire life cycle. To prevent damage to the membrane, cells have developed efficient plasma membrane repair mechanisms. These repair mechanisms can be studied by combining confocal microscopy and nanoscale thermoplasmonics to identify and investigate the role of key proteins, such as annexins, involved in surface repair in living cells and membrane model systems. The puncturing method employs a laser to induce highly localized heating upon nanoparticle irradiation. The use of near-infrared light minimizes phototoxicity in the biological sample, while the majority of the absorption takes place in the near-infrared resonant plasmonic nanoparticle. This thermoplasmonic method has been exploited for potential photothermal and biophysical research to enhance the understanding of intracellular mechanisms and cellular responses through vesicle and cell fusion studies. The approach has shown to be complementary to existing methods for membrane disruption, such as mechanically, chemically, or optically induced injuries, and provides a high level of control by inflicting extremely localized injuries. The extent of the injury is limited to the vicinity of the spherical nanoparticle, and no detrimental damage occurs along the beam path as opposed to pulsed lasers using different wavelengths. Despite certain limitations, such as the formation of nanobubbles, the thermoplasmonic method offers a unique tool for investigating cellular responses in plasma membrane repair in an almost native environment without compromising cell viability. When integrated with confocal microscopy, the puncturing method can provide a mechanistic understanding of membrane dynamics in model membrane systems as well as quantitative information on protein responses to membrane damage, including protein recruitment and their biophysical function. Overall, the application of this method to reduced model systems can enhance our understanding of the intricate plasma membrane repair machinery in living cells.

Optical trapping of plasmonic nanoparticles for controlled nanoscopic damage of cellular plasma membranes can be used to gain deeper insight into the role of plasma membrane repair proteins. Here we present a synthetic platform of giant unilamellar vesicles (GUVs) in the vicinity of trapped nanoplasmonic particles as a proposed model assay to characterize the permeability of a damaged GUV membrane, i.e. size of an inflicted hole. Water soluble fluorescent molecules with different sizes are used to characterize the extent of the membrane lesion since their differential permeability will provide information about the size of the rupture. We find that trapped gold nanoparticles can create substantial holes, observed via the discriminating influx of various sized molecules across the membrane. The technique, yet unrefined, provides groundwork for future investigations of annexin repair proteins, using nanoscopic heating of plasmonic particles to create quantifiable membrane damage.

Filopodia, widely distributed on cell surfaces, are distinguished by their dynamic extensions, playing pivotal roles in a myriad of biological processes. Their functions span from mechanosensing and guidance to cell-cell communication during cellular organization in the early embryo. Filopodia have significant roles in pathogenic processes, such as cancer invasion and viral dissemination. Molecular mapping of the filopodome has revealed generic components essential for filopodia functions. In parallel, recent insights into biophysical mechanisms governing filopodia dynamics have provided the foundation for broader investigations of filopodia's biological functions. We highlight recent discoveries of engagement of filopodia in various stages of development and pathogenesis and present an overview of intricate molecular and physical features of these cellular structures across a spectrum of cellular activities.

During the last century, fluorescence microscopy has played a pivotal role in a range of scientific discoveries. The success of fluorescence microscopy has prevailed despite several shortcomings like measurement time, photobleaching, temporal resolution, and specific sample preparation. To bypass these obstacles, label-free interferometric methods have been developed. Interferometry exploits the full wavefront information of laser light after interaction with biological material to yield interference patterns that contain information about structure and activity. Here, we review recent studies in interferometric imaging of plant cells and tissues, using techniques such as biospeckle imaging, optical coherence tomography, and digital holography. These methods enable quantification of cell morphology and dynamic intracellular measurements over extended periods of time. Recent investigations have showcased the potential of interferometric techniques for precise identification of seed viability and germination, plant diseases, plant growth and cell texture, intracellular activity and cytoplasmic transport. We envision that further developments of these label-free approaches, will allow for high-resolution, dynamic imaging of plants and their organelles, ranging in scales from sub-cellular to tissue and from milliseconds to hours.

Biomembranes are fundamental to our understanding of the cell, the basic building block of all life. An intriguing aspect of membranes is their ability to assume a variety of shapes, which is crucial for cell function. Here, we review various membrane shaping mechanisms with special focus on the current understanding of how local curvature and local rigidity induced by membrane proteins leads to emerging forces and consequently large-scale membrane deformations. We also argue that describing the interaction of rigid proteins with membranes purely in terms of local membrane curvature is incomplete and that changes in the membrane rigidity moduli must also be considered.

Many cellular processes involve the lateral organization of integral and peripheral membrane proteins into nanoscale domains. Despite the biological significance, the mechanisms that facilitate membrane protein clustering into nanoscale lipid domains remain enigmatic. In cells, the analysis of membrane protein phase affinity is complicated by the size and temporal nature of ordered and disordered lipid domains. To overcome these limitations, we developed a method for delivering membrane proteins from transfected cells into phase-separated model membranes that combines optical trapping with thermoplasmonic-mediated membrane fusion and confocal imaging. Using this approach, we observed clear phase partitioning into the liquid disordered phase following the transfer of GFP-tagged influenza hemagglutinin and neuraminidase from transfected cell membranes to giant unilamellar vesicles. The generic platform presented here allows investigation of the phase affinity of any plasma membrane protein which can be labeled or tagged with a fluorescent marker.

Biological systems are regulated by molecular interactions which are tuned by the concentrations of each of the molecules involved. Cells exploit this feature by regulating protein expression, to adapt their responses to overstimulation. Correlating events in single cells to the concentrations of proteins involved can therefore provide important mechanistic insight into cell behavior. Unfortunately, quantification of molecular densities by fluorescence imaging becomes non-trivial due to the diffraction limited resolution of the imaged volume. We show here an alternative approach to overcome this limitation in optical quantification of protein concentrations which is based on calibrating protein volume and surface densities in a model membrane system. We exploit the ability of fluorescently labeled annexin V to bind membranes in presence of calcium. By encapsulating known concentrations of annexin V, we can directly infer the membrane density of annexin V after addition of Ca2+ and correlate the density with the measured fluorescence signal. Our method, named Calmet, enables quantitative determination of the concentration of cytosolic and membrane associated proteins. The applicability of Calmet is demonstrated by quantification of a transmembrane protein receptor (beta 1 adrenergic receptor) labeled by SNAP tagged fluorophores and expressed in HEK293 cells. Calmet is a generic method suitable for the determination of a broad range of concentrations and densities and can be used on regular fluorescence images captured by confocal laser scanning microscopy.

Continuous reshaping of the plasma membrane into pleomorphic shapes is critical for a plethora of cellular functions. How the cell carries out this enigmatic control of membrane remodeling has remained an active research field for decades and several molecular and biophysical mechanisms have shown to be involved in overcoming the energy barrier associated with membrane bending. The reported mechanisms behind membrane bending have been largely concerned with structural protein features, however, in the last decade, reports on the ability of densely packed proteins to bend membranes by protein–protein crowding, have challenged prevailing mechanistic views. Crowding has now been shown to generate spontaneous vesicle formation and tubular morphologies on cell- and model membranes, demonstrating crowding as a relevant player involved in the bending of membranes. Still, current research is largely based on unnatural overexpression of proteins in non-native domains, and together with efforts in modeling, this has led to questioning the in vivo impact of crowding. In this review, we examine this previously overlooked mechanism by summarizing recent advances in the understanding of protein–protein crowding and its prevalence in cellular membrane-shaping processes.

The transcription factor NF-κB plays a vital role in the control of the immune system, and following stimulation with TNF-α its nuclear concentration shows oscillatory behaviour. How environmental factors, in particular temperature, can control the oscillations and thereby affect gene stimulation is still remains to be resolved question. In this work, we reveal that the period of the oscillations decreases with increasing temperature. We investigate this using a mathematical model, and by applying results from statistical physics, we introduce temperature dependency to all rates, resulting in a remarkable correspondence between model and experiments. Our model predicts how temperature affects downstream protein production and find a crossover, where high affinity genes upregulates at high temperatures. Finally, we show how or that oscillatory temperatures can entrain NF-κB oscillations and lead to chaotic dynamics presenting a simple path to chaotic conditions in cellular biology.

Many cellular processes involve the lateral organization of integral and peripheral membrane proteins into nanoscale domains. Despite the biological significance the mechanisms that facilitate membrane protein clustering into nanoscale lipid domains remain enigmatic. In cells the analysis of membrane protein phase affinity is complicated by the size and temporal nature of ordered and disordered lipid domains. To overcome these limitations we developed a method for delivering membrane proteins from transfected cells into phase-separated model membranes that combines optical trapping with thermoplasmonic-mediated membrane fusion and confocal imaging. Using this approach we observed clear phase partitioning following the transfer of GFP-tagged influenza hemagglutinin and neuraminidase from transfected cell membranes to giant unilamellar vesicles. The generic platform presented here allows investigation of the phase affinity of any plasma membrane protein which can be labeled or tagged with a fluorescent marker.

Maintaining the integrity of the cell plasma membrane (PM) is critical for the survival of cells. While an efficient PM repair machinery can aid survival of healthy cells by preventing influx of extracellular calcium, it can also constitute an obstacle in drug delivery and photothermal therapy. We show how nanoscopic holes can be created in a controlled fashion to the cell's plasma membrane, thus allowing identification of molecular components which have a pivotal role in PM repair. Cells are punctured by laser induced local heating of gold nanostructures at the cell surface which causes nano-ruptures in cellular PMs. Recruitment of annexin V near the hole is found to locally reshape the ruptured plasma membrane. Experiments using model membranes, containing recombinant annexin V, provide further biophysical insight into the ability of annexin V to reshape edges surrounding a membrane hole. The thermoplasmonic method provides a general strategy to monitor the response to nanoscopic injuries to the cell surface which offer new insight into how cells respond to photothermal treatment.

Filopodia are actin-rich structures, present on the surface of eukaryotic cells. These structures play a pivotal role by allowing cells to explore their environment, generate mechanical forces or perform chemical signaling. Their complex dynamics includes buckling, pulling, length and shape changes. We show that filopodia additionally explore their 3D extracellular space by combining growth and shrinking with axial twisting and buckling. Importantly, the actin core inside filopodia performs a twisting or spinning motion which is observed for a range of cell types spanning from earliest development to highly differentiated tissue cells. Non-equilibrium physical modeling of actin and myosin confirm that twist is an emergent phenomenon of active filaments confined in a narrow channel which is supported by measured traction forces and helical buckles that can be ascribed to accumulation of sufficient twist. These results lead us to conclude that activity induced twisting of the actin shaft is a general mechanism underlying fundamental functions of filopodia.