Research related to this paper and the development of the fBFP technology in the Zhu lab were supported by NIH grants AI044902, AI077343, AI038282, HL093723, HL091020, GM096187, and TW008753. We thank Evan Evans for inventing this empowering experimental tool, and members of the Evans lab, Andrew Leung, Koji Kinoshita, Wesley Wong, and Ken Halvorsen, for helping us to build the BFP. We also thank other Zhu lab members, Fang Kong, Chenghao Ge and Kaitao Li, for their helps in the instrumentation development.

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</ol>

The authors have nothing to disclose.

A successful fBFP experiment entails a few critical considerations. First, for the force calculation to be reliable, the micropipette, the RBC, and the probe bead should be aligned as close to coaxial as possible. The projection of the RBC inside the pipette should be about one probe pipette diameter so that the friction between the RBC and the pipette is negligible. For a typical human RBC, the optimal pipette diameter is 2.0-2.4 &#181;m, which yields a best fit of Equation 117,30. Second, to ensure measureme...

Cell-to-cell and cell-to-extracellular matrix (ECM) adhesion is mediated by binding between cell surface receptors, ECM proteins, and/or lipids1. Binding allows cells to form functional structures1, as well as recognize, communicate, and react to the environment1-3. Unlike soluble proteins (e.g., cytokines and growth factors) that bind from a three-dimensional (3D) fluid phase onto the cell surface receptors, cell adhesion receptors form bonds with their ligands across a narrow junctional gap to bridge two opposing surfaces that constrain molecular diffusion in a two dimensional (2D) interface4-7. In contrast to 3D kinetics that are commonly measured by traditional binding assays (e.g., surface plasmon resonance or SPR), 2D kinetics have to be quantified with specialized techniques such as atomic force microscopy (AFM)8-10, flow chamber11,12, micropipette13,14, optical tweezers15 and biomembrane force probe (BFP)16-21.
More than merely providing physical linkage for cellular cohesion, adhesion molecules are a major component of the signaling machinery for the cell to communicate with its surroundings. There has been increasing interest in understanding how ligand engagement of adhesion molecules initiates intracellular signaling and how the initial signal is transduced inside the cell. Intuitively, properties of receptor-ligand binding can impact the signals it induces. However, it is difficult to dissect mechanistic relationships between the extracellular interaction and intracellular signaling events using traditional ensemble of biochemical assays because of their many limitations, e.g., a poor temporal resolution and the complete lack of spatial resolution. Existing methods that allow both biophysical (2D receptor-ligand binding kinetics) and biochemical (signaling) observations on live cells include substrates of tunable rigidity22, elastomer pillar arrays23 and flow chamber/microfluidic devices incorporated with fluorescence capability24-26. However, readouts of signaling and receptor-ligand binding have to be obtained separately (most often by different methods), making it difficult to dissect temporal and spatial relations of bond characteristics with signaling events.
Conventional BFP is an ultrasensitive force spectroscopy with high spatiotemporal resolution17. It uses a flexible red blood cell (RBC) as a force sensor, enabling measurement of single-molecule 2D kinetics, mechanical properties and conformational changes14,16,19-21,27-29. A fluorescent imaging based BFP (fBFP) correlates the receptor-ligand binding kinetics with the binding-triggered cell signaling at single-molecule scale. With this setup, in situ cell signaling activities in the context of surface mechanical stimulation was observed in T-cells27. The fBFP is versatile and can be used for studies of cell adhesion and signaling mediated by other molecules in other cells.

<table><tbody><tr><td>Sodium Phosphate Monobasic Monohydrate (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; &amp;bull; H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>S9638</td><td>Phosphate buffer preparation</td></tr><tr><td>Anhy. Sodium Phosphate Dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S7907</td><td>Phosphate buffer preparation</td></tr><tr><td>Sodium Carbonate (Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S2127</td><td>Carbonate/bicarbonate buffer preparation</td></tr><tr><td>Sodium Bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S5761</td><td>Carbonate/bicarbonate buffer preparation</td></tr><tr><td>Sodium chloride (NaCl)</td><td>Sigma-Aldrich</td><td>S7653</td><td>N2-5% buffer preparation</td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma-Aldrich</td><td>P9541</td><td>N2-5% buffer preparation</td></tr><tr><td>Potassium phosphate monobasic (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>P5655</td><td>N2-5% buffer preparation</td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S0389</td><td>N2-5% buffer preparation</td></tr><tr><td>MAL-PEG3500-NHS</td><td>JenKem</td><td>A5002-1</td><td>Bead functionalization</td></tr><tr><td>Biotin-PEG3500-NHS</td><td>JenKem</td><td>A5026-1</td><td>RBC biotinylation</td></tr><tr><td>Nystatin</td><td>Sigma-Aldrich</td><td>N6261</td><td>RBC osmolarity adjustment</td></tr><tr><td>Ammonium Hydroxide (NH&lt;sub&gt;4&lt;/sub&gt;OH)</td><td>Sigma-Aldrich</td><td>A-6899</td><td>Glass bead silanization</td></tr><tr><td>Methanol</td><td>BDH</td><td>67-56-1</td><td>Glass bead silanization</td></tr><tr><td>30% Hydrogen Peroxide (H&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;)</td><td>J. T. Barker</td><td>Jan-86</td><td>Glass bead silanization</td></tr><tr><td>Acetic Acid (Glacial)</td><td>Sigma-Aldrich</td><td>ARK2183</td><td>Glass bead silanization</td></tr><tr><td>3-Mercaptopropyltrimethoxysilane (MPTMS)</td><td>Uct Specialties, llc</td><td>4420-74-0</td><td>Glass bead functionalization</td></tr><tr><td>Borosilicate Glass beads</td><td>Distrilab Particle Technology</td><td>9002</td><td>Glass bead functionalization</td></tr><tr><td>Streptavidin&amp;minus;Maleimide</td><td>Sigma-Aldrich</td><td>S9415</td><td>Glass bead functionalization</td></tr><tr><td>BSA</td><td>Sigma-Aldrich</td><td>A0336</td><td>Ligand functionalizing</td></tr><tr><td>Fura2-AM</td><td>Life Technologies</td><td>F-1201</td><td>Intracellular calcium fluorescence dye loading</td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D2650</td><td>Intracellular calcium fluorescence dye loading</td></tr><tr><td>Quantibrite PE Beads</td><td>BD Biosciences</td><td>340495</td><td>Density quantification</td></tr><tr><td>Flow Cytometer</td><td>BD Biosciences</td><td>BD LSR II</td><td>Density quantification</td></tr><tr><td>Capillary Tube 0.7-1.0 mm x 30 inches</td><td>Kimble Chase</td><td>46485-1</td><td>Micropipette making</td></tr><tr><td>Flaming/Brown Micropipette Puller</td><td>sutter instrument</td><td>P-97</td><td>Micropipette making</td></tr><tr><td>Pipette microforce</td><td>Narishige</td><td>MF-900</td><td>Micropipette making</td></tr><tr><td>Mineral Oil</td><td>Fisher Scientific</td><td>BP2629-1</td><td>Chamber assembly</td></tr><tr><td>Microscope Cover Glass</td><td>Fisher Scientific</td><td>12-544-G</td><td>Chamber assembly</td></tr><tr><td>Micro-injector</td><td>World Precision Instruments</td><td>MF34G-5</td><td>Chamber assembly</td></tr><tr><td>1 ml syringe</td><td>BD&amp;nbsp;</td><td>309602</td><td>chamber assembly</td></tr><tr><td>Micropipette holder</td><td>Narishige</td><td>HI-7</td><td>Chamber assembly</td></tr><tr><td>Home-designed mechanical parts and adaptors fabrications using CNC machining.&amp;nbsp;</td><td>Biophysics Instrument</td><td>All parts are customized according to the CAD designs.</td><td>BFP system</td></tr><tr><td>Microscope (TiE inverted)</td><td>Nikon</td><td>MEA53100</td><td>BFP system</td></tr><tr><td>Objective CFI Plan Fluor 40x (NA 0.75, WD 0.72 mm, Spg)</td><td>Nikon</td><td>MRH00401</td><td>BFP system</td></tr><tr><td>Camera, GE680, 640 x 480, GigE, 1/3&quot; CCD, mono</td><td>Graftek Imaging</td><td>02-2020C</td><td>BFP system</td></tr><tr><td>Prosilica GC1290 - ICX445, 1/3&quot;, C-Mount, 1280 x 960, Mono., CCD, 12 Bit ADC</td><td>Graftek Imaging</td><td>02-2185A</td><td>BFP system</td></tr><tr><td>Manual submicron probehead with high resolution remote control</td><td>Karl Suss</td><td>PH400</td><td>BFP system</td></tr><tr><td>Anti-vibration table (5&amp;rsquo; x 3&amp;rsquo;)</td><td>TMC</td><td>77049089</td><td>BFP system</td></tr><tr><td>3D manual translational stage</td><td>Newport</td><td>462-XYZ-M</td><td></td></tr><tr><td>SolidWorks 3D CAD software</td><td>SOLIDWORKS Corp.</td><td>Version 2012 SP5</td><td>BFP system</td></tr><tr><td>LabVIEW software</td><td>National Instruments</td><td>Version 2009</td><td>BFP system, BFP program</td></tr><tr><td>3D piezo translational stage</td><td>Physik Instrumente</td><td>M-105.3P</td><td>BFP system</td></tr><tr><td>Linear piezo accuator</td><td>Physik Instrumente</td><td>P-753.1CD</td><td>BFP system</td></tr><tr><td>Micromanager software</td><td></td><td>Version 1.4</td><td>fBFP system, fluorescence imaging program</td></tr><tr><td>Dual Cam (DC-2)</td><td>Photometrics</td><td>77054724</td><td>fBFP system</td></tr><tr><td>Dual Cam emission filter (T565LPXR)</td><td>Photometrics</td><td>77054725</td><td>fBFP system</td></tr><tr><td>Fluorescence Camera</td><td>Hamamatsu</td><td>ORCA-R2 C10600-10B</td><td>fBFP system</td></tr><tr><td>Plastic paraffin film (Parafilm)</td><td>Bemis Company, Inc</td><td>PM996</td><td>bottle sealing</td></tr><tr><td>Carbonate/bicarbonate buffer (pH 8.5)</td><td></td><td></td><td>8.4 g/L sodium carbonate (Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;), 10.6 g/L sodium bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td></tr><tr><td>Phosphate buffer (pH 6.5-6.8)</td><td></td><td></td><td>27.6 g/L NaPhosphate monobasic (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; &amp;bull; H&lt;sub&gt;2&lt;/sub&gt;O), 28.4 g/L Anhy. NaPhosphate dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td></tr><tr><td>N2-5% buffer (pH 7.2)</td><td></td><td></td><td>20.77 g/L potassium chloride (KCl), 2.38 g/L sodium chloride (NaCl), 0.13 g/L potassium phosphate monobasic (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;), 0.71 g/L anhy. sodium phosphate dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;), 9.70 g/L sucrose</td></tr></tbody></table>

fluorescence biomembrane force probe: concurrent quantitation of receptor-ligand kinetics and binding-induced intracellular signaling on a single cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

The BFP technique was pioneered by the Evans laboratory in 1995 17. This picoforce tool has been extensively used to measure interactions of proteins immobilized on surfaces, so as to analyze two-dimensional kinetics of adhesion molecules interacting with their ligands16,19,20,30, to measure molecular elasticity21,29, and to determine protein conformational changes21. For an fBFP, an additional set of epi-fluorescence-related devices with the corresponding software system (<str...

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Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

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Research

JoVE Journal

Bioengineering

12.4K Views.  Georgia Institute of Technology. We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Video: Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Biochemistry

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Measuring Nucleotide Binding to Intact, Functional Membrane Proteins in Real Time

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. This method combines expression of proteins tagged with the fluorescent non-canonical amino acid ANAP, and FRET between ANAP and fluorescent (trinitrophenyl) nucleotide derivatives. We present examples of nucleotide binding to ANAP-tagged KATP ion channels measured in unroofed plasma membranes and excised, inside-out membrane patches under voltage clamp. The latter allows for simultaneous measurements of ligand binding and channel current, a direct readout of protein function. Data treatment and analysis are discussed extensively, along with potential pitfalls and artefacts. This method provides rich mechanistic insights into the ligand-dependent gating of KATP channels and can readily be adapted to the study of other nucleotide-regulated proteins or any receptor for which a suitable fluorescent ligand can be identified.

This protocol presents a method for measuring adenine nucleotide binding to receptors in real time in a cellular environment. Binding is measured as F&#246;rster resonance energy transfer (FRET) between trinitrophenyl nucleotide derivatives and protein labeled with a non-canonical, fluorescent amino acid.

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Biology

Dual-Color Fluorescence Cross-Correlation Spectroscopy to Study Protein-Protein Interaction and Protein Dynamics in Live Cells

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques. In dual-color FCCS, where the fluctuations in fluorescence intensity represent the dynamic &#34;fingerprint&#34; of the respective fluorescent biomolecule, we can probe co-diffusion or binding of the receptors. FRET, with its high sensitivity to molecular distances, serves as a well-known &#34;nanoruler&#34; to monitor intramolecular changes. Taken together, conformational changes and key parameters such as local receptor concentrations and mobility constants become accessible in cellular settings.
Quantitative fluorescence approaches are challenging in cells due to high noise levels and the vulnerability of the sample. Here we show how to perform this experiment, including the calibration steps using dual-color labeled &#946;2-adrenergic receptor (&#946;2AR) labeled with eGFP and SNAP-tag-TAMRA. A step-by-step data analysis procedure is provided using open-source software and templates that are easy to customize.
Our guideline enables researchers to unravel molecular interactions of biomolecules in live cells in situ with high reliability despite the limited signal-to-noise levels in live cell experiments. The operational window of FRET and particularly FCCS at low concentrations allows quantitative analysis at near-physiological conditions.

We present an experimental protocol and data analysis workflow to perform live cell dual-color fluorescence cross correlation spectroscopy (FCCS) combined with F&#246;rster Resonance Energy transfer (FRET) to study membrane receptor dynamics in live cells using modern fluorescence labeling techniques.

We present a protocol and workflow to perform live cell dual-color fluorescence cross-correlation spectroscopy (FCCS) combined with F&#246;rster Resonance ...

A Protocol for Using F&#246;rster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. Nesprin-2G is a key component of the LINC complex that connects the actin cytoskeleton to membrane proteins (SUN domain proteins) in the perinuclear space. These membrane proteins connect to lamins inside the nucleus. Recently, a F&#246;rster Resonance Energy Transfer (FRET)-force probe was cloned into mini-Nesprin-2G (Nesprin-TS (tension sensor)) and used to measure tension across Nesprin-2G in live NIH3T3 fibroblasts. This paper describes the process of using Nesprin-TS to measure LINC complex forces in NIH3T3 fibroblasts. To extract FRET information from Nesprin-TS, an outline of how to spectrally unmix raw spectral images into acceptor and donor fluorescent channels is also presented. Using open-source software (ImageJ), images are pre-processed and transformed into ratiometric images. Finally, FRET data of Nesprin-TS is presented, along with strategies for how to compare data across different experimental groups.

A number of FRET-based force biosensors have recently been developed, enabling the protein-specific resolution of intracellular force. In this protocol, we demonstrate how one of these sensors, designed for the linker of the nucleoskeleton-cytoskeleton (LINC) complex protein Nesprin-2G can be used to measure actomyosin forces on the nuclear LINC complex.

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. ...

Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An important first step in any membrane-dependent reaction is binding of protein on the phospholipid membrane. An approach to studying protein interaction with lipid membranes has been developed using fluorescently labeled proteins and flow cytometry. This method allows the study of protein-membrane interactions using live cells and natural or artificial phospholipid vesicles. The advantage of this method is the simplicity and availability of reagents and equipment. In this method, proteins are labeled using fluorescent dyes. However, both self-made and commercially available, fluorescently labeled proteins can be used. After conjugation with a fluorescent dye, the proteins are incubated with a source of the phospholipid membrane (microvesicles or cells), and the samples are analyzed by flow cytometry. The obtained data can be used to calculate the kinetic constants and equilibrium Kd. In addition, it is possible to estimate the approximate number of protein binding sites on the phospholipid membrane using special calibration beads.

Here, we describe a set of methods for characterizing the interaction of proteins with membranes of cells or microvesicles.

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure single-molecular binding kinetics, assess mechanical properties of ligand-receptor interactions, visualize protein dynamic conformational changes and more excitingly elucidate receptor mediated cell mechanosensing mechanisms. More recently, BFP has been used to measure the spring constant of molecular bonds. This protocol describes the step-by-step procedure to perform molecular spring constant DFS analysis. Specifically, two BFP operation modes are discussed, namely the Bead-Cell and Bead-Bead modes. This protocol focuses on deriving spring constants of the molecular bond and cell from DFS raw data.

A biomembrane force probe (BFP) is an in situ dynamic force spectroscopy (DFS) technique. BFP can be used to measure the spring constant of molecular interactions on living cells. This protocol presents spring constant analysis for molecular bonds detected by BFP.

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

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