Kinetics of small molecule interactions with membrane proteins in single cells measured with mechanical amplification

Berberine inhibits intracellular Ca2+ signals in mouse pancreatic acinar cells through M3 muscarinic receptors: Novel target, mechanism, and implication

Berberine, a natural isoquinoline alkaloid, exhibits a variety of pharmacological effects, but the pharmacological targets and mechanisms remain elusive. Here, we report a novel finding that berberine inhibits acetylcholine (ACh)-induced intracellular Ca2+ oscillations, mediated through an inhibition of the muscarinic receptor subtype 3 (M3) receptor. Patch-clamp recordings and confocal Ca2+ imaging were applied to acute dissociated pancreatic acinar cells prepared from CD1 mice to examine the effects of berberine on ACh-induced Ca2+ oscillations. Whole-cell patch-clamp recordings showed that berberine (from 0.1 to 10 µM) reduced ACh-induced Ca2+ oscillations in a concentration-dependent manner, and this inhibition also depended on ACh concentrations. The inhibitory effect of berberine neither occurred in intracellular targets nor extracellular cholecystokinin (CCK) receptors, chloride (Cl-) channels, and store-operated Ca2+ channels. Together, the results demonstrate that berberine directly inhibits the muscarinic M3 receptors, further confirmed by evidence of the interaction between berberine and M3 receptors in pancreatic acinar cells.

Functional Plasmonic Microscope: Characterizing the Metabolic Activity of Single Cells via Sub-nm Membrane Fluctuations

Metabolic abnormalities are at the center of many diseases, and the capability to film and quantify the metabolic activities of a single cell is important for understanding the heterogeneities in these abnormalities. In this paper, a functional plasmonic microscope (FPM) is used to image and measure metabolic activities without fluorescent labels at a single-cell level. The FPM can accurately image and quantify the subnanometer membrane fluctuations with a spatial resolution of 0.5 μm in real time. These active cell membrane fluctuations are caused by metabolic activities across the cell membrane. A three-dimensional (3D) morphology of the bottom cell membrane was imaged and reconstructed with FPM to illustrate the capability of the microscope for cell membrane characterization. Then, the subnanometer cell membrane fluctuations of single cells were imaged and quantified with the FPM using HeLa cells. Cell metabolic heterogeneity is analyzed based on membrane fluctuations of each individual cell that is exposed to similar environmental conditions. In addition, we demonstrated that the FPM could be used to evaluate the therapeutic responses of metabolic inhibitors (glycolysis pathway inhibitor STF 31) on a single-cell level. The result showed that the metabolic activities significantly decrease over time, but the nature of this response varies, depicting cell heterogeneity. A low-concentration dose showed a reduced fluctuation frequency with consistent fluctuation amplitudes, while the high-concentration dose showcased a decreasing trend in both cases. These results have demonstrated the capabilities of the functional plasmonic microscope to measure and quantify metabolic activities for drug discovery.

A fluorescence-based binding assay for proteins using the cell surface as a sensing platform

Protein-protein interaction (PPI) analysis is very important for elucidating the functions of proteins because many proteins execute their functions in living cells by interacting with one another. In PPI analysis, methods using the sensor chips are widely employed to obtain quantitative data. However, these methods require that the target proteins be immobilized on the sensor chips, and the immobilization processes can affect the binding of the target proteins to their binding partners. In the present work, we propose a PPI analysis system in which the surface of the living cells is utilized as a sensing platform. In our approach, the target protein is displayed on the cell surface by expressing it as a fusion protein with a membrane protein, and the PPI analysis is then conducted by applying its binding partner labeled with a fluorescent dye to the cell surface. We have constructed a model of this binding analysis system using the interaction between biotin protein ligase (BPL) and biotin carboxyl carrier protein (BCCP), where BCCP was displayed on the cell surface and BPL labeled with fluorescein was applied to the cell surface. Here, a red fluorescent protein, mApple, was attached to the C-terminus of the fusion protein of BCCP with a membrane protein. We evaluated the binding level of the labeled BPL by using the intensity ratios of fluorescence from fluorescein to that from mApple. We found that the binding level of the labeled BPL was stably evaluated at least across 60 min observation period and estimated the binding dissociation constant between BPL and BCCP by equilibrium analysis to be 0.33 ± 0.05 μM.

Electric field modulation of ERK dynamics shows dependency on waveform and timing

Different exogenous electric fields (EF) can guide cell migration, disrupt proliferation, and program cell development. Studies have shown that many of these processes were initiated at the cell membrane, but the mechanism has been unclear, especially for conventionally non-excitable cells. In this study, we focus on the electrostatic aspects of EF coupling with the cell membrane by eliminating Faradaic processes using dielectric-coated microelectrodes. Our data unveil a distinctive biphasic response of the ERK signaling pathway of epithelial cells (MCF10A) to alternate current (AC) EF. The ERK signal exhibits both inhibition and activation phases, with the former triggered by a lower threshold of AC EF, featuring a swifter peaking time and briefer refractory periods than the later-occurring activation phase, induced at a higher threshold. Interestingly, the biphasic ERK responses are sensitive to the waveform and timing of EF stimulation pulses, depicting the characteristics of electrostatic and dissipative interactions. Blocker tests and correlated changes of active Ras on the cell membrane with ERK signals indicated that both EGFR and Ras were involved in the rich ERK dynamics induced by EF. We propose that the frequency-dependent dielectric relaxation process could be an important mechanism to couple EF energy to the cell membrane region and modulate membrane protein-initiated signaling pathways, which can be further explored to precisely control cell behavior and fate with high temporal and spatial resolution.

Ligand Binding-Induced Cellular Membrane Deformation is Correlated with the Changes in Membrane Stiffness

Study interaction between ligands and protein receptors is a key step for biomarker research and drug discovery. In situ measurement of cell surface membrane protein binding on whole cells eliminates the cost and pitfalls associated with membrane protein purification. Ligand binding to membrane protein was recently found to induce nanometer-scale cell membrane deformations, which can be monitored with real-time optical imaging to quantify ligand/protein binding kinetics. However, the insight into this phenomenon has still not been fully understood. We hypothesize that ligand binding can change membrane stiffness, which induces membrane deformation. To investigate this, cell height and membrane stiffness changes upon ligand binding are measured using atomic force microscopy (AFM). Wheat germ agglutinin (WGA) is used as a model ligand that binds to the cell surface glycoprotein. The changes in cell membrane stiffness and cell height upon ligand bindings are determined for three different cell lines (human A431, HeLa, and rat RBL-2H3) on two different substrates. AFM results show that cells become stiffer with increased height after WGA modification for all cases studied. The increase in cell membrane stiffness is further confirmed by plasmonic scattering microscopy, which shows an increased cell spring constant upon WGA binding. Therefore, this study provides direct experimental evidence that the membrane stiffness changes are directly correlated with ligand binding-induced cell membrane deformation.

A LIGHTFUL nanomedicine overcomes EGFR-mediated drug resistance for enhanced tyrosine-kinase-inhibitor-based hepatocellular carcinoma therapy

Targeting the activated epidermal growth factor receptor (EGFR) via clustered regularly interspaced short palindromic repeat (CRISPR) technology is appealing to overcome the drug resistance of hepatocellular carcinoma (HCC) towards tyrosine kinase inhibitor (TKI) therapy. However, combining these two distinct drugs using traditional liposomes results in a suboptimal synergistic anti-HCC effect due to the limited CRISPR/Cas9 delivery efficiency caused by lysosomal entrapment after endocytosis. Herein, we developed a liver-targeting gene-hybridizing-TKI fusogenic liposome (LIGHTFUL) that can achieve high CRISPR/Cas9 expression to reverse the EGFR-mediated drug resistance for enhanced TKI-based HCC therapy efficiently. Coated with a galactose-modified membrane-fusogenic lipid layer, LIGHTFUL reached the targeting liver site to fuse with HCC tumor cells, directly and efficiently transporting interior CDK5- and PLK1-targeting CRISPR/Cas9 plasmids (pXG333-CPs) into the HCC cell cytoplasm and then the cell nucleus for efficient expression. Such membrane-fusion-mediated pXG333-CP delivery resulted in effective downregulation of both CDK5 and PLK1, sufficiently inactivating EGFR to improve the anti-HCC effects of the co-delivered TKI, lenvatinib. This membrane-fusion-participant codelivery strategy optimized the synergetic effect of CRISPR/Cas9 and TKI combinational therapy as indicated by the 0.35 combination index in vitro and the dramatic reduction of subcutaneous and orthotopic TKI-insensitive HCC tumor growth in mice. Therefore, the established LIGHTFUL provides a unique co-delivery platform to combine gene editing and TKI therapies for enhanced synergetic therapy.

Label-free analysis of membrane protein binding kinetics and cell adhesions using evanescent scattering microscopy

Measuring ligand interactions with membrane proteins in single live cells is critical for understanding many cellular processes and screening drugs. However, developing such a capability has been a difficult challenge. Here, we employ evanescent scattering microscopy (ESM) to show that ligand binding to membrane proteins can change the cell adhesion properties, which are intrinsic cell properties and independent of random cell micromotions and ligand mass, thus allowing the kinetics analyses of both proteins and small molecules binding to membrane proteins in both single fixed and live cells. In addition, utilizing the high spatiotemporal resolution of ESM, the positions of cell adhesion sites can be tracked in real-time to analyze the cell deformations and migrations, thus providing a potential approach for understanding the cell activity during the ligand binding process in detail. The presented method may pave the road for developing a versatile and easy-to-use label-free detection strategy for in situ analysis of molecular interaction dynamics in living biosystems with single-cell resolution.

Label-free imaging and biomarker analysis of exosomes with plasmonic scattering microscopy

Exosome analysis is a promising tool for clinical and biological research applications. However, detection and biomarker quantification of exosomes is technically challenging because they are small and highly heterogeneous. Here, we report an optical approach for imaging exosomes and quantifying their protein markers without labels using plasmonic scattering microscopy (PSM). PSM can provide improved spatial resolution and distortion-free image compared to conventional surface plasmon resonance (SPR) microscopy, with the signal-to-noise ratio similar to objective coupled surface plasmon resonance (SPR) microscopy, and millimeter-scale field of view as a prism-coupled SPR system, thus allowing exosome size distribution analysis with high throughput. In addition, PSM retains the high specificity and surface sensitivity of the SPR sensors and thus allows selection of exosomes from extracellular vesicles with antibody-modified sensor surfaces and in situ analyzing binding kinetics between antibody and the surface protein biomarkers on the captured exosomes. Finally, the PSM can be easily constructed on a popular prism-coupled SPR system with commercially available components. Thus, it may provide an economical and powerful tool for clinical exosome analysis and exploration of fundamental issues such as exosome biomarker binding properties.

In Situ Analysis of Membrane-Protein Binding Kinetics and Cell-Surface Adhesion Using Plasmonic Scattering Microscopy

Surface plasmon resonance microscopy (SPRM) is an excellent platform for in situ studying cell-substrate interactions. However, SPRM suffers from poor spatial resolution and small field of view. Herein, we demonstrate plasmonic scattering microscopy (PSM) by adding a dry objective on a popular prism-coupled surface plasmon resonance (SPR) system. PSM not only retains SPRM's high sensitivity and real-time analysis capability, but also provides ≈7 times higher spatial resolution and ≈70 times larger field of view than the typical SPRM, thus providing more details about membrane protein response to ligand binding on over 100 cells simultaneously. In addition, PSM allows quantifying the target movements in the axial direction with a high spatial resolution, thus allowing mapping adhesion spring constants for quantitatively describing the mechanical properties of the cell-substrate contacts. This work may offer a powerful and cost-effective strategy for upgrading current SPR products.

Label-Free Quantification of Molecular Interaction in Live Red Blood Cells by Tracking Nanometer Scale Membrane Fluctuations

Molecular interactions in live cells play an important role in both cellular functions and drug discovery. Current methods for measuring binding kinetics involve extracting the membrane protein and labeling, while the in situ quantification of molecular interaction with surface plasmon resonance (SPR) imaging mainly worked with fixed cells due to the micro-motion related noises of live cells. Here, an optical imaging method is presented to measure the molecular interaction with live red blood cells by tracking the nanometer membrane fluctuations. The membrane fluctuation dynamics are measured by tracking the membrane displacement during glycoprotein interaction. The data are analyzed with a thermodynamic model to determine the elastic properties of the cell observing reduced membrane fluctuations under fixatives, indicating cell fixations affect membrane mechanical properties. The binding kinetics of glycoprotein to several lectins are obtained by tracking the membrane fluctuation amplitude changes on single live cells. The binding kinetics and strength of different lectins are quite different, indicating the glycoproteins expression heterogeneity in single cells. It is anticipated that the method will contribute to the understanding of mechanisms of cell interaction and communication, and have potential applications in the mechanical assessment of cancer or other diseases at the single-cell level, and screening of membrane protein targeting drugs.

Emerging Optical Microscopy Techniques for Electrochemistry

An optical microscope is probably the most intuitive, simple, and commonly used instrument to observe objects and discuss behaviors through images. Although the idea of imaging electrochemical processes operando by optical microscopy was initiated 40 years ago, it was not until significant progress was made in the last two decades in advanced optical microscopy or plasmonics that it could become a mainstream electroanalytical strategy. This review illustrates the potential of different optical microscopies to visualize and quantify local electrochemical processes with unprecedented temporal and spatial resolution (below the diffraction limit), up to the single object level with subnanoparticle or single-molecule sensitivity. Developed through optically and electrochemically active model systems, optical microscopy is now shifting to materials and configurations focused on real-world electrochemical applications.

Critical angle reflection imaging for quantification of molecular interactions on glass surface

Quantification of molecular interactions on a surface is typically achieved via label-free techniques such as surface plasmon resonance (SPR). The sensitivity of SPR originates from the characteristic that the SPR angle is sensitive to the surface refractive index change. Analogously, in another interfacial optical phenomenon, total internal reflection, the critical angle is also refractive index dependent. Therefore, surface refractive index change can also be quantified by measuring the reflectivity near the critical angle. Based on this concept, we develop a method called critical angle reflection (CAR) imaging to quantify molecular interactions on glass surface. CAR imaging can be performed on SPR imaging setups. Through a side-by-side comparison, we show that CAR is capable of most molecular interaction measurements that SPR performs, including proteins, nucleic acids and cell-based detections. In addition, we show that CAR can detect small molecule bindings and intracellular signals beyond SPR sensing range. CAR exhibits several distinct characteristics, including tunable sensitivity and dynamic range, deeper vertical sensing range, fluorescence compatibility, broader wavelength and polarization of light selection, and glass surface chemistry. We anticipate CAR can expand SPR′s capability in small molecule detection, whole cell-based detection, simultaneous fluorescence imaging, and broader conjugation chemistry.

Here, the authors present a method for quantifying molecular interactions on a glass surface, based on measuring surface refractive index changes via the reflectivity near the critical angle. They demonstrate tunable sensitivity and dynamic range, deep vertical sensing range, also for intracellular signals.

Topological Encoded Vector Beams for Monitoring Amyloid-Lipid Interactions in Microcavity

Lasers are the pillars of modern photonics and sensing. Recent advances in microlasers have demonstrated its extraordinary lasing characteristics suitable for biosensing. However, most lasers utilized lasing spectrum as a detection signal, which can hardly detect or characterize nanoscale structural changes in microcavity. Here the concept of amplified structured light-molecule interactions is introduced to monitor tiny bio-structural changes in a microcavity. Biomimetic liquid crystal droplets with self-assembled lipid monolayers are sandwiched in a Fabry-Pérot cavity, where subtle protein-lipid membrane interactions trigger the topological transformation of output vector beams. By exploiting Amyloid β (Aβ)-lipid membrane interactions as a proof-of-concept, it is demonstrated that vector laser beams can be viewed as a topology of complex laser modes and polarization states. The concept of topological-encoded laser barcodes is therefore developed to reveal dynamic changes of laser modes and Aβ-lipid interactions with different Aβ assembly structures. The findings demonstrate that the topology of vector beams represents significant features of intracavity nano-structural dynamics resulted from structured light-molecule interactions.

Label-Free Probing of Molecule Binding Kinetics Using Single-Particle Interferometric Imaging

Probing molecular interactions is critical for screening drugs, detecting pollutants, and understanding biological processes at the molecular level, but these interactions are difficult to detect, especially for small molecules. A label-free optical imaging technology that can detect molecule binding kinetics is presented, in which free-moving particles are driven into oscillations with an alternating electrical field and the interferometric scattering patterns of the particles are imaged via an optical imaging method. By tracking the charge-sensitive variations in the oscillation amplitude with sub-nanometer precision, the small molecules and metal ions binding to the surface as well as protein-protein binding kinetics were measured. The capability of the label-free measurement of molecular interactions can provide a promising platform for screening small-molecule drugs, probing conformational changes in proteins, and detecting environmental pollutants.

Optical Imaging of Electrical and Mechanical Couplings between Cells

Intercellular communication plays a pivotal role in multicellular organisms. Studying the electrical and mechanical coupling among multiple cells has been a difficult task due to the lack of suitable techniques. In this study, we developed a label-free imaging method for monitoring the electrical-induced communications between connected cells. The method was based on monitoring subtle mechanical motions of the cells under electrical modulation of the membrane potential. We observed that connected cells responded to electrical modulation of neighboring cells with mechanical deformation of the membrane. We further investigated the mechanism of the coupling and confirmed that this mechanical response was induced by electrical signal communicated through the gap junction. Blocking the gap junction can temporally cease the mechanical signal, and this inhibition can be rescued after removing the inhibitor. This study sheds light on the mechanism of electrical coupling between neurons and provides a new method for studying intercellular communications.

Cell surface thermal proteome profiling tracks perturbations and drug targets on the plasma membrane

Numerous drugs and endogenous ligands bind to cell surface receptors leading to modulation of downstream signaling cascades and frequently to adaptation of the plasma membrane proteome. In-depth analysis of dynamic processes at the cell surface is challenging due to biochemical properties and low abundances of plasma membrane proteins. Here we introduce cell surface thermal proteome profiling for the comprehensive characterization of ligand-induced changes in protein abundances and thermal stabilities at the plasma membrane. We demonstrate drug binding to extracellular receptors and transporters, discover stimulation-dependent remodeling of T cell receptor complexes and describe a competition-based approach to measure target engagement of G-protein-coupled receptor antagonists. Remodeling of the plasma membrane proteome in response to treatment with the TGFB receptor inhibitor SB431542 leads to partial internalization of the monocarboxylate transporters MCT1/3 explaining the antimetastatic effects of the drug.

Moving Electrons Purposefully through Single Molecules and Nanostructures: A Tribute to the Science of Professor Nongjian Tao (1963-2020)

Electrochemistry intersected nanoscience 25 years ago when it became possible to control the flow of electrons through single molecules and nanostructures. Many surprises and a wealth of understanding were generated by these experiments. Professor Nongjian Tao was among the pioneering scientists who created the methods and technologies for advancing this new frontier. Achieving a deeper understanding of charge transport in molecules and low-dimensional materials was the first priority of his experiments, but he also succeeded in discovering applications in chemical sensing and biosensing for these novel nanoscopic systems. In parallel with this work, the investigation of a range of phenomena using novel optical microscopic methods was a passion of his and his students. This article is a review and an appreciation of some of his many contributions with a view to the future.

Simultaneous Quantification of Protein Binding Kinetics in Whole Cells with Surface Plasmon Resonance Imaging and Edge Deformation Tracking

Most drugs work by binding to receptors on the cell surface. Quantification of binding kinetics between drug and membrane protein is an essential step in drug discovery. Current methods for measuring binding kinetics involve extracting the membrane protein and labeling, and both have issues. Surface plasmon resonance (SPR) imaging has been demonstrated for quantification of protein binding to cells with single-cell resolution, but it only senses the bottom of the cell and the signal diminishes with the molecule size. We have discovered that ligand binding to the cell surface is accompanied by a small cell membrane deformation, which can be used to measure the binding kinetics by tracking the cell edge deformation. Here, we report the first integration of SPR imaging and cell edge tracking methods in a single device, and we use lectin interaction as a model system to demonstrate the capability of the device. The integration enables the simultaneous collection of complementary information provided by both methods. Edge tracking provides the advantage of small molecule binding detection capability, while the SPR signal scales with the ligand mass and can quantify membrane protein density. The kinetic constants from the two methods were cross-validated and found to be in agreement at the single-cell level. The variation of observed rate constant between the two methods is about 0.009 s^-1, which is about the same level as the cell-to-cell variations. This result confirms that both methods can be used to measure whole-cell binding kinetics, and the integration improves the reliability and capability of the measurement.

Detecting a single nanoparticle by imaging the localized enhancement and interference of surface plasmon polaritons

Label-free single-nanoparticle detection is crucial for the fast detection of nanoparticles and viruses in environmental monitoring and biological sciences. In this Letter, benefiting from the leakage radiation that transforms the near-field surface plasmon polariton (SPP) distribution along the interface to the far field, we demonstrated the plasmonic imaging of single polystyrene nanoparticles with a particle size down to 39 nm. The imaging is composed of the localized enhancement and interference of SPPs. The localized enhancement is the result of the accumulation of charges around the nanoparticle, and it is connected to the size and refractive index of nanoparticles. The interference is induced by the coupling between the incident SPPs and the scattered SPPs, verified by extracting the interference fringe periodicity to be half of the SPP wavelength. Our study provides an in-depth physical understanding of plasmonic imaging of single nanoparticles, which paves the way for a fast identification of nanomaterials.

Quantifying Ligand-Protein Binding Kinetics with Self-Assembled Nano-oscillators

Measuring ligand-protein interactions is critical for unveiling molecular-scale biological processes in living systems and for screening drugs. Various detection technologies have been developed, but quantifying the binding kinetics of small molecules to the proteins remains challenging because the sensitivities of the mainstream technologies decrease with the size of the ligand. Here, we report a method to measure and quantify the binding kinetics of both large and small molecules with self-assembled nano-oscillators, each consisting of a nanoparticle tethered to a surface via long polymer molecules. By applying an oscillating electric field normal to the surface, the nanoparticle oscillates, and the oscillation amplitude is proportional to the number of charges on the nano-oscillator. Upon the binding of ligands onto the nano-oscillator, the oscillation amplitude will change. Using a plasmonic imaging approach, the oscillation amplitude is measured with subnanometer precision, allowing us to accurately quantify the binding kinetics of ligands, including small molecules, to their protein receptors. This work demonstrates the capability of nano-oscillators as an useful tool for measuring the binding kinetics of both large and small molecules.

An integrated platform for the capture of circulating tumor cells and in situ SERS profiling of membrane proteins through rational spatial organization of multi-functional cyclic RGD nanopatterns

An integrated platform was established for capture of cancer cells and SERS detection of HER2 activity via multifunctional RGD nanopatterns.

Chemical Sensing in Real Time with Plants Using a Webcam

It has been established that plants can smell and respond to chemicals in order to adapt to and survive in a changing chemical environment. Here we show that a plant responds to chemicals in air, and the response can be detected rapidly to allow tracking of air pollution in real time. We demonstrate this capability by detecting subtle color and shape changes in the leaves of mosses upon exposure to sulfur dioxide in air with a simple webcam and an imaging-processing algorithm. The leaves of mosses consist of a monolayer of cells, providing a large surface-to-volume ratio for highly sensitive chemical sensing. The plant sensor responds linearly to sulfur dioxide within a wide concentration range (0-180 ppm), and it can tolerate humidity variation (15-85% relative humidity) and chemical interference and regenerate itself. We envision that plants can help alert chemical exposure danger as a part of our living environment using low-cost CMOS imagers, and their chemical-sensing capabilities may be further improved with genetic engineering.

Label-free imaging of epidermal growth factor receptor-induced response in single living cells

Epidermal growth factor receptor (EGFR), which belongs to the second-largest protein family for cell signal transduction, plays crucial roles in homeostasis, cellular organized patterns and most human cancers. In EGFR-activated signaling networks, the detection of the spatial and temporal dynamics of cascades that encode the many cell fates is still a challenge. Here, we report real-time imaging of epidermal growth factor (EGF)-induced EGFR activation and its signaling cascade in single A431 cells using surface plasmon resonance (SPR) microscopy. A two-phase SPR response pattern was observed within 30 min after EGF treatment, including a positive SPR response that was related to the EGFR-activated mass redistribution in the first 600 s, and a subsequent negative SPR signal caused by the morphological change of the cells. Furthermore, the inhibitor analysis verified that AG1478 inhibited the response from the whole the cell, whereas cytochalasin B strongly inhibited the response from the cell edge region.

Measuring Ligand Binding Kinetics to Membrane Proteins Using Virion Nano-oscillators

Membrane proteins play vital roles in cellular signaling processes and serve as the most popular drug targets. A key task in studying cellular functions and developing drugs is to measure the binding kinetics of ligands with the membrane proteins. However, this has been a long-standing challenge because one must perform the measurement in a membrane environment to maintain the conformations and functions of the membrane proteins. Here, we report a new method to measure ligand binding kinetics to membrane proteins using self-assembled virion oscillators. Virions of human herpesvirus were used to display human G-protein-coupled receptors (GPCRs) on their viral envelopes. Each virion was then attached to a gold-coated glass surface via a flexible polymer to form an oscillator and driven into oscillation with an alternating electric field. By tracking changes in the oscillation amplitude in real-time with subnanometer precision, the binding kinetics between ligands and GPCRs was measured. We anticipate that this new label-free detection technology can be readily applied to measure small or large ligand binding to any type of membrane proteins and thus contribute to the understanding of cellular functions and screening of drugs.

Imaging Action Potential in Single Mammalian Neurons by Tracking the Accompanying Sub-Nanometer Mechanical Motion

Action potentials in neurons have been studied traditionally by intracellular electrophysiological recordings and more recently by the fluorescence detection methods. Here we describe a label-free optical imaging method that can measure mechanical motion in single cells with a sub-nanometer detection limit. Using the method, we have observed sub-nanometer mechanical motion accompanying the action potential in single mammalian neurons by averaging the repeated action potential spikes. The shape and width of the transient displacement are similar to those of the electrically recorded action potential, but the amplitude varies from neuron to neuron, and from one region of a neuron to another, ranging from 0.2-0.4 nm. The work indicates that action potentials may be studied noninvasively in single mammalian neurons by label-free imaging of the accompanying sub-nanometer mechanical motion.

Gradient-Based Colorimetric Sensors for Continuous Gas Monitoring

Colorimetry detects a color change resulted from a chemical reaction or molecular binding. Despite its widespread use in sensing, continuous monitoring of analytes with colorimetry is difficult, especially when the color-producing reaction or binding is irreversible. Here, we report on a gradient-based colorimetric sensor (GCS) to overcome this limitation. Lateral transport of analytes across a colorimetric sensor surface creates a color gradient that shifts along the transport direction over time, and GCS tracks the gradient shift and converts it into analyte concentration in real time. Using a low cost complementary metal-oxide semiconductor imager and imaging processing algorithm, we show submicrometer gradient shift tracking precision and continuous monitoring of ppb-level ozone.

Tracking fast cellular membrane dynamics with sub-nm accuracy in the normal direction

Cellular membranes are important biomaterials with highly dynamic structures. Membrane dynamics plays an important role in numerous cellular processes, but precise tracking it is challenging due to the lack of tools with a highly sensitive and fast detection capability. Here we demonstrate a broad bandwidth optical imaging technique to measure cellular membrane displacements in the normal direction at sub-nm level detection limits and 20 μs temporal resolution (1 Hz-50 kHz). This capability allows us to study the intrinsic cellular membrane dynamics over a broad temporal and spatial spectrum. We measured the nanometer-scale stochastic fluctuations of the plasma membrane of HEK-293 cells, and found them to be highly dependent on the cytoskeletal structure of the cells. By analyzing the fluctuations, we further determine the mechanical properties of the cellular membranes. We anticipate that the method will contribute to the understanding of the basic cellular processes, and applications, such as mechanical phenotyping of cells at the single-cell level.

Label-Free Quantification of Small-Molecule Binding to Membrane Proteins on Single Cells by Tracking Nanometer-Scale Cellular Membrane Deformation

Measuring molecular binding to membrane proteins is critical for understanding cellular functions, validating biomarkers, and screening drugs. Despite the importance, developing such a capability has been a difficult challenge, especially for small-molecule binding to membrane proteins in their native cellular environment. Here we show that the binding of both large and small molecules to membrane proteins can be quantified on single cells by trapping single cells with a microfluidic device and detecting binding-induced cellular membrane deformation on the nanometer scale with label-free optical imaging. We develop a thermodynamic model to describe the binding-induced membrane deformation, validate the model by examining the dependence of membrane deformation on cell stiffness, membrane protein expression level, and binding affinity, and study four major types of membrane proteins, including glycoproteins, ion channels, G-protein coupled receptors, and tyrosine kinase receptors. The single-cell detection capability reveals the importance of local membrane environment on molecular binding and variability in the binding kinetics of different cell lines and heterogeneity of different cells within the same cell line.