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 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.
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.
This protocol follows the guidelines of and has been approved by the human research ethics committee of Georgia Institute of Technology.
1. Human RBCs Isolation, Biotinylation and Osmolarity Adjustment
Note: Step 1.1 should be performed by a trained medical professional such as a nurse, with an Institutional Review Board approved protocol.
2. Glass Bead Silanization
3. Bead Functionalization
4. Cell Preparation
Note: To purify the cells, follow standard cell purification protocols corresponding to the type of cells in use, for example T-cells27 or certain cell lines21,29.
5. Preparation for Micropipettes and a Cell Chamber
6. BFP experiment
Figure 1: fBFP assembly. (A) An overview picture of the fBFP hardware system. (B) A schematic drawing of the fBFP hardware system. (C) The dual-cam system “DC2” (orange) onto which the high-speed camera (blue) and a fluorescence camera (white) were mounted. (D) The microscope stage that adapts an experiment chamber and three micropipette manipulation systems. (E and F) Micrographs of BFP setting in an experimental chamber. (E) Micropipettes assembly showing the probe pipette (left), target pipette (upper right) and helper pipette (lower right). (F) Probe bead placement. A probe bead was manipulated by a helper pipette and attached to a RBC apex to form a force probe. Please click here to view a larger version of this figure.
Figure 2: BFP scheme and its test cycle. (A) Video-micrograph depicting a force probe (left) and a target T-cell (right) aspirated by their respective pipettes.The stationary force probe consists of a swollen RBC and an attached ligand-bearing bead. The receptor-bearing T-cell (target) is mounted to a piezotranslator aligned opposite to the probe. The ROI is indicated in green. The edge tracker is indicated in a blue line. The insert depicts the ligand (pMHC, bead side) and receptor (TCR, T-cell side) pair on the two opposing surfaces in the area marked in orange. (B) The intensity profile of the bead edge in (A). The ROI region in the x-direction is plotted as x-axis (in pixel number) and the light intensity (in gray scale value) averaged by binning 30 pixels along the y-direction. (C) The deflection of the RBC and the position of the bead and the target (T-cell) in a test cycle of force clamp assay. The vertical and horizontal dashed lines indicate the zero-force position of the RBC apex and the time course, respectively. The line edge tracker of the RBC deformation is shown in blue in each panel. The same yet less steps are adopted in adhesion frequency assay (which lacks the steps of “clamp” and “dissociate”) and thermal fluctuation assay (which lacks the step of “dissociate”).
7. Fluorescence BFP (fBFP) experiment
8. Data Analysis
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 (Table 1) is added (Figure 1A–C).
The fBFP system consists of a hardware system (optical, mechanical and electrical components) and a software system (Table 1 and Figure 1A, B). An inverted microscope (Figure 1A, middle) with optical components enabling bright-field and epi-fluorescence microscopy makes up the main body of the fBFP system, onto which the experiment stage and pipette manipulators are mounted. A fluorescence light source controller (Figure 1A, upper left corner) is used to deliver alternating excitation lights. A dual-cam system “DC2” splits the light into two and transmits them to a high-speed camera (blue) and a fluorescence camera (white) (Figure 1A, lower left corner). The former collects the probe image for position determination and the latter collects fluorescence images emitted from the target cell at two wave lengths. The experiment is monitored and controlled by two computers of which Host PC 1 (Figure 1A, upper right corner, cropped in the picture due to space limitation) is connected to both the high-speed camera and the fluorescence camera and is responsible for experiment execution and data acquisition, and Host PC 2 is connected to the monitoring camera and responsible for presenting a full view of the BFP experiment (Figure 1A, B).
During an experiment, the experimentalist uses three micropipettes to grab and manipulate cells and microspheres respectively (Figure 1E, F). A typical image of a BFP experiment consists of a force probe, which is assembled by attaching a probe bead onto the apex of an aspirated RBC, and a target cell/bead (Figure 2A). A region of interest (ROI), which contains the edge area of the bead on the RBC apex, is tracked by a fast-speed camera (2,000 fps) to monitor the deformation of the RBC in real-time. The tracked displacement of the bead then can be used to calculate the external force exerted on the force probe (see the note in Protocol section 6.6), given the spring constant of the BFP (Figure 2A, B).
Adhesion frequency assay, thermal fluctuation assay, and force clamp assay are the three commonly used experimental modes on a BFP system. By adopting any of these three modes, a BFP experiment is composed of repeated testing cycles that are performed sequentially.
In adhesion frequency assay, the target approaches and contacts the probe bead at a given contact time (for example, 1 sec) and retracts directly back to the original position and start the next cycle. An adhesion event is reflected by a tensile force signal during the retraction phase. This approach-contact-retraction cycle is repeated 50 times for at least 3 target-probe pairs to calculate a mean SEM of adhesion frequency, Pa.
Thermal fluctuation assay is used for lifetime measurements under zero-force. In each cycle, after touching the probe bead, the target is retracted to the zero-force position and clamped in contact with the bead without either compression or tension for 20 sec, and then returns to the original position to start the next cycle. Bond association/dissociation under zero-force is manifested as a sudden drop/increase in the thermal fluctuations of the bead16,30.
Force clamp assay is used for lifetime measurements under forces. A desired clamping force (for example, 20 pN) is set prior to the experiment. The target is driven repeatedly to contact the probe bead for a given contact time (for example, 1 sec) to allow the formation of a receptor/ligand bond (Figure 2C, Panels 2 and 3) and then retract (Figure 2C, Panel 4). If no bond is formed, reflected by no tensile force signal on the RBC, or if the bond ruptures before reaching the preset clamping force, the target will return to the original position and start the next cycle. In the binding events that survive the retraction phase, the target is held at the clamping force with an corresponding elongation of the RBC by d until the bond dissociates (Figure 2C, Panels 5 and 6), and then returns to the original position to start the next cycle (Figure 2C, Panel 7). Lifetime is defined as the time interval from the instant when the force reached the desired level to the instant of bond dissociation20.
For all these three BFP experimental modes, the experimentalist should be able to see the approach-impinge-contact-retract-(clamp-)(dissociate-)return testing cycle (Figure 2C), which will be repeated for multiple times16,20,21.
Data collected from each experimental mode of BFP could be analyzed in various ways to derive desired results. The average lifetime curve is one representative result from the force clamp assay (Figure 4). It reflects the reciprocal off-rates of the receptor-ligand dissociation under forces. Thermal fluctuation assay allows for the characterization of 2D on-rate and off-rate of a receptor-ligand pair under zero-force16; while adhesion frequency assay renders the 2D on-rate, off-rate and affinity under zero-force13.
The add-in of the fluorescence imaging function allows for monitoring the intracellular Ca2+ level of a single cell, which is used as the readout of cell signaling in this system. To use this function during an fBFP experiment, one needs to pre-load the cells with a Ca2+ indicator. In the case of Fura2-AM being the fluorescent dye, the fluorescence images of the same cell under 340 nm and 380 nm excitation channels were recorded alternately in the experiment and inspected pair-by-pair in the analysis. An intensity threshold was assigned to each channel’s fluorescent image to remove background noise and recognize the cell contour (Figure 5). Comparison of each pair of the fluorescence images allows the calculation of the intracellular Ca2+ level. Clear fluorescent images of the cell under both 340 nm and 380 nm excitation channels are necessary for accurate measurement of the Ca2+ level (Figure 5A, B), while poor images with non-negligible background noise will result in biased results and should be avoided (Figure 5C).
By introducing controllable mechanical stimulations onto the cell and record cell Ca2+ level simultaneously, the fBFP provides a powerful single-molecule tool to study mechano-transduction on a living cell. It is worth noting that, the platform described here is quite versatile; in principle, one can design numerous ways of applying force to the cell to suit particular purposes and concurrently monitoring various signaling events of interest. The force-dependent kinetic details are then analyzed in the context of the chosen signaling readout to extract characteristics of force-regulated receptor-ligand kinetics, the induced signaling, and their correlation. In the example of TCR signaling, an agonist-specific TCR-pMHC catch bond was first discovered and then its relevance to T-cell Ca2+ flux was investigated27. The strategy was to take the peak value of Ca2+ flux as the signaling readout to seek its best predictor among various kinetic parameters, including the number of adhesions, the force amplitude of the binding, the average lifetime, the longest lifetime and the cumulative lifetime of the sequentially formed bindings. Shown in Figure 6 is an example of simultaneously recorded individual bond lifetimes (where a clamping force of 10 pN was applied) of a T-cell and their accumulation together with the corresponding Ca2+ signal curve. In this case, four lifetime events occurred prior to the time when Ca2+ level began to elevate at 55 sec, accumulating a sum of 10-second bond lifetime. This level of lifetime accumulation triggered a Ca2+ signal with a peak of normalized Fura2 ratio of 1.8 that occurred at 65 sec. A systematic mathematical analysis of such data collected from many individual cells revealed that the best correlate of Ca2+ signaling strength is lifetimes cumulated in the 1st minute of repeated TCR-pMHC interactions (refer to Reference 27 for technical details).
Figure 3: Exemplary time-force raw data curves of a no-adhesion event (A), a rupture force event (B) and a lifetime event (C). Various phases of the cycle and the corresponding curve segments are respectively marked in each panel. The force (y-axis) is derived from tracking the position change of the probe bead, as shown in Figure 2C. (A) No adhesion: the compressive (negative) force in the contact phase returns to zero upon retraction. (B) A rupture force event: a tensile (positive) force pulls via the receptor-ligand bond to elongate the RBC, which ruptures during the retraction phase. (C) A lifetime event: the bond persists until the clamping force is reached and dissociates thereafter. The duration of the lifetime event is indicated. The rupture force event (B) and the beginning of the lifetime event (C) are highlighted by an asterisk.
Figure 4: “Average lifetime vs. force” curve of OT1 T-cell interacting with its agonist OVA (green) and antagonist G4 (blue). The pooled data are grouped into different force bins, and the mean ± SEM of bond lifetimes is plotted versus force.
Figure 5: Representative Ca2+ images excited at two wavelengths. (A, B) Correct image recognition of a T-cell (indicated in red) in 340 nm (A) and 380 nm (B) channels based on point-to-point screening using a properly assigned intensity threshold. (C) Inability to recognize the fluorescence image of a T-cell (indicated in red) in the 340 nm excitation channel, due to poor Fura2 loading.
Figure 6: Superimposition of the “intracellular Ca2+ level (relative Fura2 ratio) vs. time” and the “cumulative lifetime vs. time” curves of an OT1 T-cell, generated by repeatedly touching the T-cell with an OVA-coated bead over 300 seconds. (A) A force curve showing a sequence of non-adhesion, rupture force, and lifetime events generated by repeated contacts over time. (B) Epi-fluorescence pseudo-color images of intracellular Ca2+ signals in the T-cell at different time points. The normalized Ca2+ level is indicated by the pseudo-color scale on the right. (C) Superimposition of the Ca2+ signal curve (red) and the cumulative lifetime curve (yellow) on the same time course. The Ca2+ curve was plotted based on the Ca2+ imaging. A Ca2+ flux is signified by a sharp elevation in the normalized Fura2 ratio. The time when Ca2+ reaches the peak is indicated by a dashed line. The onset time of each lifetime event is marked on the cumulative lifetime curve (solid triangle).
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 µm, which yields a best fit of Equation 117,30. Second, to ensure measurements in force clamp assay and thermal fluctuation assay are mostly for single bonds, an adhesion frequency of under 20% has to be maintained10,13,30. Here, non-specific adhesions should be carefully controlled (usually < 5% by sufficient blocking with BSA). In addition, data should be collected only from adhesion events with a single force-drop – hallmark of a single-molecule behavior (analogous to a single-step disappearance of fluorescence in single-molecule FRET). Third, loading concentration of the fluorescence dye should be optimized on a case-by-case basis to achieve the best imaging quality. Fourth, the toxicity of dye loading to T-cell or other cells of interest needs to be carefully examined before each experiment. For example, the binding affinity of receptor-ligand interactions upon dye loading need to be measured and compared to that without dye loading. If the binding affinity is change dramatically due to dye loading, a different dye or a different loading concentration has to be considered. Fifth, terminally biotin-tagged proteins are preferred to allow convenient, specific, and strong coupling that preserves the protein’s native orientation.
A major strength of the fBFP is that it performs a single-bond assay on a single cell. Despite the low throughput, single-bond analysis often uncovers important features which are inaccessible by conventional ensemble methods. For example, by examining the lifetime distribution at each force bin, one can correlate different binding characteristics with protein conformational states, providing insight on how force regulates protein conformational changes20,32. The BFP can also be used to measure molecular stiffness from the force and piezo-translator displacement data of the retraction phase, which can be used to investigate protein conformational dynamics20,21.
Many methods have been developed to study receptor-mediated cell adhesion and signaling. Receptor-ligand binding kinetics is generally measured with recombinant proteins in complete isolation from the cellular environment. Such practice is potentially problematic. For instance, it has recently been shown that in situ kinetics measured on live cells drastically differs from that measured using the corresponding recombinant proteins14, revealing novel insights of the receptor’s cellular functions. Not only is the fBFP capable of quantifying receptor-ligand kinetics in situ, but more importantly, it can also simultaneously record the binding-induced cell signaling. As demonstrated for the TCR27, the rich information of binding characteristics and cell signaling obtained using fBFP provides an unprecedented opportunity for analyzing their relations and understanding the molecular mechanisms of mechano-transduction. It is likely that fBFP will find more applications in other important receptor-ligand systems.
The authors have nothing to disclose.
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.
Table 1: Reagents/Equipment | |||
Name of Material/ Equipment | Company | Catalog Number | Comments/Description |
Sodium Phosphate Monobasic Monohydrate (NaH2PO4•H2O) | Sigma-Aldrich | S9638 | Phosphate buffer preparation |
Anhy. Sodium Phosphate Dibasic (Na2HPO4) | Sigma-Aldrich | S7907 | Phosphate buffer preparation |
Sodium Carbonate (Na2CO3) | Sigma-Aldrich | S2127 | Carbonate/bicarbonate buffer preparation |
Sodium Bicarbonate (NaHCO3) | Sigma-Aldrich | S5761 | Carbonate/bicarbonate buffer preparation |
Sodium chloride (NaCl) | Sigma-Aldrich | S7653 | N2-5% buffer preparation |
Potassium chloride (KCl) | Sigma-Aldrich | P9541 | N2-5% buffer preparation |
Potassium phosphate monobasic (KH2PO4) | Sigma-Aldrich | P5655 | N2-5% buffer preparation |
Sucrose | Sigma-Aldrich | S0389 | N2-5% buffer preparation |
MAL-PEG3500-NHS | JenKem | A5002-1 | Bead functionalization |
Biotin-PEG3500-NHS | JenKem | A5026-1 | RBC biotinylation |
Nystatin | Sigma-Aldrich | N6261 | RBC osmolarity adjustment |
Ammonium Hydroxide (NH4OH) | Sigma-Aldrich | A-6899 | Glass bead silanization |
Methanol | BDH | 67-56-1 | Glass bead silanization |
30% Hydrogen Peroxide (H2O2) | J. T. Barker | Jan-86 | Glass bead silanization |
Acetic Acid (Glacial) | Sigma-Aldrich | ARK2183 | Glass bead silanization |
3-MERCAPTOPROPYLTRIMETHOXYSILANE(MPTMS) | Uct Specialties, llc | 4420-74-0 | Glass bead functionalization |
Borosilicate Glass beads | Distrilab Particle Technology | 9002 | Glass bead functionalization |
Streptavidin−Maleimide | Sigma-Aldrich | S9415 | Glass bead functionalization |
BSA | Sigma-Aldrich | A0336 | Ligand functionalizing |
Fura2-AM | Life Technologies | F-1201 | Intracellular calcium fluorescence dye loading |
Dimethyl sulfoxide (DMSO) | Sigma-Aldrich | D2650 | Intracellular calcium fluorescence dye loading |
Quantibrite PE Beads | BD Biosciences | 340495 | Density quantification |
Flow Cytometer | BD Biosciences | BD LSR II | Density quantification |
Capillary Tube 0.7-1.0mm x 30" | Kimble Chase | 46485-1 | Micropipette making |
Flaming/Brown Micropipette Puller | sutter instrument | P-97 | Micropipette making |
Pipette microforce | Narishige | MF-900 | Micropipette making |
Mineral Oil | Fisher Scientific | BP2629-1 | Chamber assembly |
Microscope Cover Glass | Fisher Scientific | 12-544-G | Chamber assembly |
Micro-injector | World Precision Instruments | MF34G-5 | Chamber assembly |
1ml Syringe | BD | 309602 | Chamber assembly |
Micropipette holder | Narishige | HI-7 | Chamber assembly |
Home-designed mechanical parts and adaptors fabrications using CNC machining. | Biophysics Instrument | All parts are customized according to the CAD designs. | BFP system |
Microscope (TiE inverted) | Nikon | MEA53100 | BFP system |
Objective CFI Plan Fluor 40x (NA 0.75, WD 0.72mm, Spg) | Nikon | MRH00401 | BFP system |
Camera, GE680, 640×480, GigE, 1/3" CCD, mono | Graftek Imaging | 02-2020C | BFP system |
Prosilica GC1290 – ICX445, 1/3", C-Mount, 1280×960, Mono., CCD, 12 Bit ADC | Graftek Imaging | 02-2185A | BFP system |
Manual submicron probehead with high resolution remote control | Karl Suss | PH400 | BFP system |
Anti-vibration table (5’ x 3’) | TMC | 77049089 | BFP system |
3D manual translational stage | Newport | 462-XYZ-M | |
SolidWorks 3D CAD software | SOLIDWORKS Corp. | Version 2012 SP5 | BFP system |
LabVIEW software | National Instruments | Version 2009 | BFP system, BFP program |
3D piezo translational stage | Physik Instrumente | M-105.3P | BFP system |
Linear piezo accuator | Physik Instrumente | P-753.1CD | BFP system |
Micromanager software | Version 1.4 | fBFP system, fluorescence imaging program | |
Dual Cam (DC-2) | Photometrics | 77054724 | fBFP system |
Dual Cam emission filter (T565LPXR) | Photometrics | 77054725 | fBFP system |
Fluorescence Camera | Hamamatsu | ORCA-R2 C10600-10B | fBFP system |
Plastic paraffin film (Parafilm) | Bemis Company, Inc | PM996 | bottle sealing |
Table 2: Buffer solutions | |||
Carbonate/bicarbonate buffer (pH 8.5) | |||
Sodium Carbonate (Na2CO3) | 8.4g/L | ||
Sodium Bicarbonate (NaHCO3) | 10.6g/L | ||
Phosphate buffer (pH 6.5-6.8) | |||
NaPhosphate monobasic NaH2PO4•H2O | 27.6g/L | ||
Anhy. NaPhosphate dibasic Na2HPO4 | 28.4g/L | ||
N2-5% buffer (pH 7.2) | |||
Potassium chloride (KCl) | 20.77g/L | ||
Sodium chloride (NaCl) | 2.38g/L | ||
Potassium phosphate monobasic (KH2PO4) | 0.13g/L | ||
Anhy. Sodium Phosphate Dibasic (Na2HPO4) | 0.71g/L | ||
Sucrose | 9.70g/L |