Abstract
All cells maintain ionic gradients across their plasma membranes, producing transmembrane potentials (Vmem). Mounting evidence suggests a relationship between resting Vmem and the physiology of non-excitable cells with implications in diverse areas, including cancer, cellular differentiation, and body patterning. A lack of non-invasive methods to record absolute Vmem limits our understanding of this fundamental signal. To address this need, we developed a fluorescence lifetime-based approach (VF-FLIM) to visualize and optically quantify Vmem with single-cell resolution. Using VF-FLIM, we report Vmem distributions over thousands of cells, a 100-fold improvement relative to electrophysiological approaches. In human carcinoma cells, we visualize the voltage response to epidermal growth factor stimulation, stably recording a 10-15 mV hyperpolarization over minutes. Using pharmacological inhibitors, we identify the source of the hyperpolarization as the Ca2+-activated K+ channel Kca3.1. The ability to optically quantify absolute Vmem with cellular resolution will allow a re-examination of its roles as a cellular signal.
Introduction
Membrane potential (Vmem) is an essential facet of cellular physiology. In electrically excitable cells, such as neurons and cardiomyocytes, voltage-gated ion channels enable rapid changes in membrane potential. These fast membrane potential changes, on the order of milliseconds to seconds, trigger release of neurotransmitters in neurons or contraction in myocytes. The resting membrane potential of these cells, which changes over longer timescales, affects their excitability. In non-electrically excitable cells, slower changes in Vmem—on the order of seconds to hours—are linked to a variety of fundamental cellular processes1, including mitosis 2, cell cycle progression 3, and differentiation 4 At the tissue and organismal level, mounting lines of evidence point to the importance of electrochemical gradients in development, body patterning, and regeneration 5.
Despite the importance of membrane potential to diverse processes over a range of time scales, the existing methods for recording Vmem are inadequate for characterizing distributions of Vmem states in a sample or studying gradual shifts in resting membrane potential (Figure 1- supplement 1). Patch clamp electrophysiology remains the gold standard for recording cellular electrical parameters, but it is low throughput, highly invasive, and difficult to implement over extended time periods. Where reduced invasiveness or higher throughput analyses of Vmem are required, optical methods for detecting events involving Vmem changes (e.g. whether an action potential occurred) are often employed 6–8. However, optical approaches generally use fluorescence intensity values as a readout, which cannot report either the absolute values of Vmem or the absolute amount by which Vmem changed 9. Variations in dye loading, illumination intensity, fluorophore bleaching, and/or cellular morphology dramatically complicate fluorescence intensity measurements, making calibration and determination of actual membrane potential difficult or impossible. This limitation restricts optical analysis to detection of acute Vmem changes, which can be analyzed without comparisons of Vmem between cells or over long timescales. Two-component systems, with independent wavelengths for ratio-based calibration, have seen limited success 10, and they confer significant capacitive load on the cell 11. Further, their performance hinges on carefully tuned loading procedures of multiple lipophilic indicators 12, which can be challenging to reproduce across different samples and days.
To quantify a parameter such as voltage or concentration from a single-color fluorescence signal, fluorescence lifetime (τfl) imaging (FLIM) can be employed instead of conventional fluorescence microscopy. By measuring the fluorescence lifetime, an intrinsic property of the sensor, FLIM avoids many of the artifacts that confound extrinsic fluorescence intensity measurements. As a result, FLIM can be calibrated to reproducibly and quantitatively report biological properties if the analyte or property in question affects the lifetime of the probe’s fluorescent excited state. FLIM has been successfully employed to record a number of biochemical and biophysical parameters, including intracellular Ca2+ concentration 13, viscosity 14, GTPase activity 15, kinase activity 16, and redox state (NADH/NAD+ ratio)17, among others 18. Attempts to record absolute voltage with FLIM, however, have been limited in success 19–21. Previous work focused on genetically-encoded voltage indicators (GEVIs), which have complex relationships between τfl and voltage 20 and low sensitivity to voltage in lifetime 21. Because of their poor voltage resolution, the fluorescence lifetimes of these GEVIs cannot be used to detect most biologically relevant voltage changes, which are on the order of tens of millivolts.
Fluorescent voltage indicators that use photoinduced electron transfer (PeT) as a voltage-sensing mechanism are promising candidates for a FLIM-based approach to optical Vmem quantification. Because PeT affects the nonradiative decay rate of the fluorophore excited state, it has been successfully translated from intensity to τfl imaging with a number of small molecule probes for Ca2+ 22. We previously established that VoltageFluor (VF)-type dyes transduce changes in cellular membrane potential to changes in fluorescence intensity and that the voltage response of VF dyes is consistent with a photoinduced electron transfer (PeT)-based response mechanism 23,24 Changes in the transmembrane potential alter the rate of PeT 25,26 from an electron-rich aniline donor to a fluorescent reporter, thereby modulating the fluorescence intensity of VF dyes 23 (Fig. 1A,B). VoltageFluors also display low toxicity and rapid, linear responses to voltage.
Here, we develop fluorescence lifetime imaging of VoltageFluor dyes (VF-FLIM) as a quantitative, all-optical approach for recording absolute membrane potential with single cell resolution. Using patch-clamp electrophysiology as a standard, we demonstrate that the fluorescence lifetime of the VoltageFluor dye VF2.1.Cl reports absolute membrane potential with >20-fold improved accuracy over previous optical approaches. To highlight the 100-fold increase in throughput over manual patch-clamp electrophysiology, we record resting membrane potentials of thousands of cells. To our knowledge, this work represents the first broad view of the distribution of resting membrane potentials present in situ. To showcase the spatiotemporal and voltage resolution of VF-FLIM, we quantify the gradual, small voltage changes that arise from growth factor stimulation of human carcinoma cells. Through pharmacological perturbations, we conclude that the voltage changes following epidermal growth factor (EGF) stimulation arise from activation of the calcium-activated potassium channel Koa3.1. Our results show that fluorescence lifetime of VF dyes is a generalizable and effective approach for studying resting membrane potential in a range of biological contexts.
Results
VoltageFluor Fluorescence Lifetime Varies Linearly with Membrane Potential
To characterize how the photoinduced electron transfer process affects fluorescence lifetime, we compared the τfl of the voltage-sensitive dye VF2.1.Cl with its voltage-insensitive counterpart VF2.0.Cl (Fig. 1B). We recorded the τfl of bath-applied VF dyes in HEK293T cells using time-correlated single-photon counting (TCSPC) FLIM (Fig. 1C-E). VF2.1.Cl is localized to the plasma membrane and exhibits a biexponential τfl decay with decay constants of approximately 0.9 and 2.6 ns (Scheme S2). For all subsequent analysis of VF2.1.Cl lifetime, we refer to the weighted average τfl, which is approximately 1.6 ns in HEK293T cell membranes at rest. VF2.0.Cl (Fig. 1B), which lacks the aniline substitution and is therefore voltage-insensitive 24, shows a τfl of 3.5 ns in cell membranes, which is similar to the lifetime of an unsubstituted fluorescein 27 (Fig. 1-supplement 2). We also examined VoltageFluor lifetimes at a variety of dye loading concentrations to test for concentration-dependent changes in dye lifetime, which have been reported for fluorescein derivatives 28. Shortened VF lifetimes were observed at high dye concentrations (Fig. 1-supplement 3); all subsequent VF-FLIM studies were conducted at dye concentrations low enough to avoid this concentration-dependent change in lifetime.
To assess the voltage dependence of VoltageFluor τfl, we controlled the plasma membrane potential of HEK293T cells with whole-cell voltage-clamp electrophysiology while simultaneously measuring the τfl of VF2.1.Cl (Fig. 1C). Single-cell recordings show a linear τfl response to applied voltage steps, and individual measurements deviate minimally from the linear fit (Fig. 1F-H). VF2.1.Cl τfl is reproducible across different cells at the same resting membrane potential, allowing determination of Vmem from τfl images taken without concurrent electrophysiology (Fig. 1I). Voltage-insensitive VF2.0.Cl shows no τfl change in response to voltage (Fig. 1J, Fig. 1-supplement 4), consistent with a τfl change in VF2.1.Cl arising from a voltage-dependent PeT process. In HEK293T cells, VF2.1.Cl exhibits a sensitivity of 3.50 ± 0.08 ps/mV and a 0 mV lifetime of 1.77 ± 0.02 ns, corresponding to a fractional change in τfl (Δτ/τ) of 22.4 ± 0.4% per 100 mV. These values are in good agreement with the 27% ΔF/F intensity change per 100 mV originally observed for VF2.1.Cl 23,24 To estimate the voltage resolution of VF-FLIM, we analyzed the variability in successive measurements on the same cell (intra-cell resolution) and on different cells (inter-cell resolution, see Methods). We estimate that the resolution for tracking and quantifying voltage changes in a single HEK293T cell is 4 mV (intra-cell resolution), whereas the resolution for single-trial determination of a particular HEK293T cell’s absolute Vmem is 20 mV (inter-cell resolution).
We compared the performance of VF-FLIM to that of CAESR, the best previously reported GEVI for optically recording absolute membrane potential using FLIM 21. Using simultaneous FLIM and voltage-clamp electrophysiology, we determined the relationship between τfl and Vmem for the genetically encoded voltage indicator CAESR under 1 photon excitation (Fig. 1- supplement 5). We recorded a sensitivity of −1.2 ± 0.1 ps/mV and a 0 mV lifetime of 2.0 ± 0.2 ns, which corresponds to a −6.1 ± 0.8% Δτ/τ per 100 mV (mean ± SEM of 9 measurements), in agreement with the reported sensitivity of −0.9 ps/mV and 0 mV lifetime of 2.7 ns with 2 photon excitation 21. Relative to VF2.1.Cl, CAESR displays 3-fold lower sensitivity (−1.2 ps/mV vs 3.5 ps/mV in HEK293T cells) and 7-fold higher voltage-independent variability in lifetime (0.46 ns vs 0.07 ns, standard deviation of the 0 mV lifetime measurement). For CAESR in HEK293T cells, we calculate a voltage resolution of 37 ± 7 mV for quantifying voltage changes on an individual cell (intra-cell, compared to 4 mV for VF2.1.Cl, see Methods) and resolution of 390 mV for determination of a particular cell’s absolute Vmem (inter-cell, compared to 20 mV for VF2.1.Cl). Because cellular resting membrane potentials and voltage changes (e.g. action potentials) are on the order of tens of millivolts, VF-FLIM has sufficient resolution for biologically relevant Vmem recordings, whereas CAESR does not.
Evaluation of VF-FLIM across Cell Lines and Culture Conditions
The voltage-dependent τfl response of VF2.1.Cl is generalizable across different cell types. We calibrated VF-FLIM in four additional commonly used cell lines: A431, CHO, MDA-MB-231, and MCF-7 (Fig. 2, Fig. 2-supplement 1, Fig. 2- supplement 2). All cells displayed a linear relationship between VF τfl and Vmem, with average sensitivities of 3.1 to 3.7 ps/mV and average 0 mV lifetimes ranging from 1.74 to 1.87 ns. In all cases, we observed better voltage resolution for quantification of Vmem changes on a given cell versus comparisons of absolute Vmem between cells. For all cell lines tested, the changes in voltage for a given cell could be quantified with resolutions at or better than 5 mV (intra-cell resolution, Methods). For absolute Vmem determination of a single cell, we observed voltage resolutions ranging from 11 to 24 mV (intercell resolution, Fig. 2-supplement 3). The inter-cell resolution of VF-FLIM appears to be cell-type dependent; MCF-7 cells displayed greater variability than other cell lines tested (Fig. 2B, Fig. 2-supplement 3).
To verify that VF-FLIM was robust in groups of cells in addition to the isolated, single cells generally used for patch clamp electrophysiology, we determined lifetime-voltage relationships for small groups of A431 cells (Fig. 2-supplement 4A-E). We found that calibrations made in small groups of cells are nearly identical to those obtained on individual cells, indicating that VF-FLIM only needs to be calibrated once for a given type of cell. For pairs or groups of three cells we recorded a sensitivity of 3.3 ± 0.2 ps/mV and a 0 mV lifetime of 1.78 ± 0.02 ns (mean ± SEM of 5 pairs and 2 groups of 3; values are for the entire group, not just the cell in contact with the electrode), which is similar to the sensitivity of 3.55 ± 0.08 ps/mV and 0 mV lifetime of 1.74 ± 0.02 ns we observe in single A431 cells. The slight reduction in sensitivity seen in cell groups is likely attributable to space clamp error, which prevents complete voltage clamp of the cell group 29,30. Indeed, when we analyzed only the most responsive cell in the group (in contact with the electrode), we obtained a slope of 3.7 ± 0.1 ps/mV and 0 mV lifetime of 1.79 ± 0.02 ns, in good agreement with the single cell data. The space clamp error can be clearly visualized (Figure 2 – supplement 4E), where one cell in the group of 3 responded much less to the voltage command.
To test whether VF-FLIM is also extensible to cells maintained with different culture conditions, we recorded lifetime-Vmem relationship in serum-starved A431 cells (Figure 2 – supplement 4F-K), obtaining an average sensitivity of 3.6 ± 0.1 ps/mV and a 0 mV lifetime of 1.76 ± 0.01 ns (n=2 single cells, 2 pairs, 3 groups of 3 cells), in excellent agreement with the values obtained for non-serum starved cells. We also tested for concentration-dependent changes in VF lifetime in all five cell lines and in serum starvation conditions. Similar to VF2.1.Cl lifetime in HEK293T cells (Fig. 1-supplement 3), we observed shortening of VF2.1.Cl lifetimes between 200 and 500 nM dye in all cases (Figure 2-supplement 5). All subsequent experiments were carried out at VF2.1.Cl concentrations well below the regime where VF concentration-dependent lifetime changes were observed.
Optical Determination of Resting Membrane Potential Distributions
The throughput of VF-FLIM enables cataloging of resting membrane potentials of thousands of cells in only a few hours of the experimenter’s time. We optically recorded resting membrane potential distributions for A431, CHO, HEK293T, MCF-7, and MDA-MB-231 cells using VF-FLIM (Fig. 3, Fig. 3 – supplement 1, Fig. 3 – supplement 2). We report resting membrane potentials by cell group (Methods, Scheme S2) because adjacent cells in these cultures are electrically coupled to some degree via gap junctions 31. Each group of cells represents an independent sample for Vmem. In addition, the fluorescent signal originating from membranes of adjacent cells cannot be separated with a conventional optical microscope, so assignment of a region of membrane connecting multiple cells would be arbitrary. VF-FLIM images (Fig. 3, Fig. 3 – supplement 1, Fig. 3 - supplement 2) contain spatially resolved voltage information, but caution should be employed in interpreting pixel to pixel differences in lifetime. Because VF-FLIM was calibrated here using the average plasma membrane τfl for each cell, optical Vmem should be interpreted per cell or cell group.
Mean resting membrane potentials recorded by VF-FLIM range from −53 to −29 mV, depending on the cell line. These average Vmem values fall within the range reported in the literature for all of the cell lines we measured (Fig. 3 - supplement 3). We also recorded resting membrane potentials in a high K+ buffer (120 mM K+, “high K+ HBSS”), where we observed a depolarization of 15 to 41 mV, bringing the mean Vmem up to −26 mV to +4 mV, again depending on the cell line. Our optical determination of Vmem is in good agreement with theory: the Goldman-Hodgkin-Katz equation 32 predicts Vmem of −91 to −27 mV in 6 mM extracellular K+ and −25 to +2 mV in 120 mM extracellular K+, depending on ion permeability and intracellular ion concentration (see Methods).
Membrane potential dynamics in epidermal growth factor signaling
We thought VF-FLIM was a promising method for elucidating the roles of membrane potential in non-excitable cell signaling. Specifically, we wondered whether VF-FLIM might be well-suited to dissect conflicting reports surrounding changes in membrane potential during EGF/EGF receptor (EGFR)-mediated signaling. Receptor tyrosine kinase (RTK)-mediated signaling is a canonical signaling paradigm for eukaryotic cells, transducing extracellular signals into changes in cellular state. Although the involvement of second messengers like Ca2+, cyclic nucleotides, and lipids are well characterized, membrane potential dynamics and their associated roles in non-excitable cell signaling remain less well-defined. In particular, the activation of EGFR via EGF has variously been reported to be depolarizing 33, hyperpolarizing 34, or electrically silent 35,36
We find that treatment of A431 cells with EGF results in a 15 mV hyperpolarization within 60-90 seconds in approximately 80% of cells (Fig. 4A-C, Fig. 4-supplement 1, Fig. 4 – supplement 2), followed by a slow return to baseline within 15 minutes (Fig. 4D-F, Fig. 4-supplement 3). The voltage response to EGF is dose-dependent, with an EC50 of 90 ng/mL (14 nM) (Fig. 4-supplement 4). Vehicle-treated cells show very little τfl change (Fig. 4A-F). Identical experiments with voltage-insensitive VF2.0.Cl (Fig. 4G-H, Fig. 4 – supplement 1, Fig. 4 – supplement 3, Fig. 4 – supplement 5) reveal little change in τfl upon EGF treatment, indicating the drop in τfl arises from membrane hyperpolarization. We observe the greatest hyperpolarization 1 to 3 minutes after treatment with EGF, which is abolished by inhibition of EGFR and ErbB2 tyrosine kinase activity with the covalent inhibitor canertinib (Fig. 4I-J, Fig. 4-supplement 6). Blockade of the EGFR kinase domain with gefitinib, a non-covalent inhibitor of EGFR, also results in a substantial decrease in the EGF-evoked hyperpolarization (Fig. 4I-J, Fig. 4-supplement 6). Together, these results indicate that A431 cells exhibit an EGF-induced hyperpolarization, which depends on the kinase activity of EGFR and persists on the timescale of minutes.
Outward K+ currents could mediate EGF-induced hyperpolarization. Consistent with this hypothesis, dissipation of the K+ driving force by raising extracellular [K+] completely abolishes the typical hyperpolarizing response to EGF and instead results in a small depolarizing potential of approximately 3 mV (Fig. 5A, Fig. 5 – supplement 1B). Blockade of voltage-gated K+ channels (Kv) with 4-aminopyridine (4-AP) prior to EGF treatment enhances the hyperpolarizing response to EGF (Fig. 5A, 5B, Fig. 5-supplement 1C). In contrast, blockade of Ca2+-activated K+ channels (Koa) with charybdotoxin (CTX) results in a depolarizing potential of approximately 4 mV after exposure to EGF, similar to that observed with high extracellular [K+] (Fig. 5A, 5B, Fig. 5- supplement 1D). TRAM-34, a specific inhibitor of the intermediate-conductance Ca2+ activated potassium channel Kea3.1 37, also abolishes EGF-induced hyperpolarization (Fig. 5A, Fig. 5- supplement 1E). CTX treatment has little effect on the resting membrane potential, while TRAM-34 or 4-AP depolarizes cells by approximately 5-10 mV (Fig. 5-supplement 2).
To explore the effects of other components of the EGFR pathway on EGF-induced hyperpolarization, we perturbed intra- and extracellular Ca2+ concentrations during EGF stimulation. Reduction of extracellular Ca2+ concentration did not substantially alter the EGF response (Fig. 5A, Fig. 5-supplement 1F). However, sequestration of intracellular Ca2+ with BAPTA-AM disrupts the hyperpolarization response. BAPTA-AM treated cells show a small, 4 mV depolarization in response to EGF treatment, similar to CTX-treated cells (Fig. 5A, Fig. 5-supplement 1G). Perturbation of Ca2+ levels had little effect on the resting membrane potential (Fig. 5-supplement 2). Introduction of wortmannin (1 μM) to block downstream kinase activity has no effect on the membrane potential response to EGF, while orthovanadate addition (Na3VO4, 100 μM) to block phosphatase activity results in a small increase in the hyperpolarizing response (Fig. 5A, Fig. 5-supplement 1H-I). These results support a model for EGF-EGFR mediated hyperpolarization in which RTK activity of EGFR causes release of internal Ca2+ stores to in turn open KCa channels and hyperpolarize the cell (Fig. 5C).
Discussion
We report the design and implementation of a new method for optically quantifying absolute membrane potential in living cells. VF-FLIM is operationally simple, requires just a single-point calibration, and is applicable across a number of cell types. VF-FLIM exhibits a 20fold improvement in voltage resolution over previous FLIM-based approaches 20,21, achieving sufficient resolution to make biologically relevant voltage measurements. The photoinduced electron transfer mechanism of VoltageFluors 23 renders superior sensitivity and consistency of the lifetime measurement; furthermore, because VoltageFluors are applied exogenously, the vast majority of the fluorescence signal is voltage-sensitive and at the membrane. Unlike small-molecule FRET-oxonol approaches to optically estimate membrane potential values 10, VF-FLIM presents a direct relationship between τfl and Vmem with a single optical reporter and avoids complex and potentially toxic multi-dye loading protocols.
Because VF-FLIM is an optical approach, it improves upon both the throughput and spatial resolution of patch clamp electrophysiology and thereby enables new lines of inquiry in biological systems. Although individual VF-FLIM measurements have more voltage-equivalent noise than modern electrophysiology, the ability to perform thousands of recordings over the course of a few hours enables a more complete documentation of the distributions of cellular Vmem present in a cell population. In addition to throughput, another key difference between VF-FLIM and patch-clamp electrophysiology is spatial resolution. While VF-FLIM records the Vmem of an optically defined region of interest (in this case a cell or cell group), electrophysiology records Vmem at an individual cell or part of a cell where the electrode makes contact, which may or may not reflect the Vmem of the entire cell or group. In principle, VF-FLIM could record subcellular differences in Vmem that would be difficult to dissect with electrophysiology. Looking ahead, such subcellular recordings in cells with complex morphology and processes are an exciting area for future development of VF-FLIM, in conjunction with cellular and sub-cellular strategies for targeting VF dyes 38,39.
We optically documented resting membrane potential distributions in cultured cells, as VF-FLIM is well suited to address questions about Vmem states present in these samples. The presence and significance of distinct Vmem states in cell populations is mostly uncharacterized due to the throughput limitations of patch-clamp electrophysiology, but some reports suggest that distinct Vmem states arise during the various phases of the cell cycle 40,41. Vmem histograms presented in this work appear more or less unimodal, showing no clear sign of cell cycle-related Vmem states (Fig. 3A,D; Fig. 3-supplement 1A,D,G). We considered the possibility that VF-FLIM does not detect cell-cycle-related Vmem states because we report average Vmem across cell groups in cases where cells are in contact (Scheme S2). This explanation is unlikely for two reasons. First, Vmem distributions for CHO cells appear unimodal, even though CHO cultures were mostly comprised of isolated cells under the conditions tested (Fig. 3D-F). Second, theoretical work suggests that dramatically different Vmem states in adjacent cells are unlikely, as electrical coupling often leads to equilibration of Vmem across the cell group 42,43. Although we cannot rule out the possibility of poorly separated Vmem populations (i.e. with a mean difference in voltage below our resolution limit), VF-FLIM both prompts and enables a re-examination of the notion that bi- or multimodal Vmem distributions exist in cultured cells. Furthermore, VF-FLIM represents an exciting opportunity to experimentally visualize theorized Vmem patterns in culture and in more complex tissues. Studies towards this end are ongoing in our laboratory.
In the present study, we use VF-FLIM to provide the first cell-resolved, direct visualization of voltage changes induced by growth factor signaling. For long term Vmem recordings during growth-related processes, an optical approach is more attractive than an electrode-based one. Electrophysiology becomes increasingly challenging as time scale lengthens, especially if cells migrate, and washout of the cytosol with pipette solution can change the very signals under study 44,45. Previous attempts to electrophysiologically record Vmem in EGF-stimulated A431 cells were unsuccessful due to these technical challenges.34 Because whole cell voltage-clamp electrophysiology was intractable, the Vmem response in EGF-stimulated A431 cells was addressed indirectly through model cell lines expressing EGFR exogenously 34, bulk measurements on trypsinized cells in suspension 46, or cell-attached single channel recordings 47–49. By stably recording Vmem during EGF stimulation, VF-FLIM enables direct study of Vmem signaling in otherwise inaccessible pathways.
In conjunction with physiological manipulations and pharmacological perturbations, we explore the molecular mechanisms underlying EGF-induced hyperpolarization. We find that signaling along the EGF-EGFR axis results in a robust hyperpolarizing current carried by K+ ions, passed by the Ca2+-activated K+ channel Kea3.1, and mediated by intracellular Ca2+ (Fig. 5C). We achieve a complete loss of the hyperpolarizing response to EGF by altering the K+ driving force (“High K+” Fig. 5A, Fig. 5-supplement 1B), blocking calcium-activated K+ currents directly (“CTX” and “TRAM-34”, Fig. 5A, Fig. 5-supplement 1D,E), or intercepting cytosolic Ca2+ (“BAPTA-AM”, Fig. 5A, Fig. 5-supplement 1G). These results, combined with transcriptomic evidence that Kea3.1 is the major Kea channel in A431 cells 50, indicate that Kea3.1 mediates the observed hyperpolarization. Interestingly, under some conditions where K+-mediated hyperpolarization is blocked (“CTX,” “high K+”, “BAPTA-AM”), VF-FLIM reveals a small, secondary depolarizing current not visible during normal EGF stimulation. This current likely arises from initial Ca2+ entry into the cell, as previously observed during EGF signaling 51,52. Although we have obtained direct and conclusive evidence of EGF-induced hyperpolarization in A431 cells, the interactions between this voltage change and downstream targets of EGFR remain incompletely characterized. Enhancing EGF signaling by blockade of cellular tyrosine phosphatases with orthovanadate 53 correspondingly increases EGF-mediated hyperpolarization (“Na3VO4” Fig. 5A, Fig. 5-supplement 1H), but inhibition of downstream kinase activity appears to have little effect on hyperpolarization (“wortmannin” Fig. 5A, Fig. 5-supplement 1I).
In the context of RTK signaling, Vmem may serve to modulate the driving force for external Ca2+ entry 3,54 and thereby act as a regulator of this canonical signaling ion. Alternatively, Vmem may play a more subtle biophysical role, such as potentiating lipid reorganization in the plasma membrane 55. Small changes in Vmem likely affect signaling pathways in ways that are currently completely unknown, but high throughput discovery of Vmem targets remains challenging. Combination of electrophysiology with single cell transcriptomics has begun to uncover relationships between Vmem and other cellular pathways in excitable cells 56; such approaches could be coupled to higher throughput VF-FLIM methods to explore pathways that interact with Vmem in non-excitable contexts.
VF-FLIM represents a novel and general approach for interrogating the roles of membrane potential in fundamental cellular physiology. Future improvements to the voltage resolution could be made by use of more sensitive indicators, which may exhibit larger changes in fluorescence lifetime 24 VF-FLIM can be further expanded to include the entire color palette of PeT-based voltage indicators 57,58, allied with targeting methods to probe absolute membrane potential in heterogeneous cellular populations 38,39, and coupled to high-speed imaging techniques for optical quantification of fast voltage events 59.
Author Contributions
JLD performed experiments, analyzed data, and wrote the paper. AMMG performed experiments and analyzed data. EWM analyzed data and wrote the paper.
Competing Interest Statement
EWM is listed as an inventor on a patent describing voltage-sensitive fluorophores. This patent is owned by the Regents of the University of California.
Materials and Methods
Materials
VoltageFluor dyes VF2.1.Cl and VF2.0.Cl were synthesized in house according to previously described syntheses 24. Dyes were stored either as solids at room temperature or as 1000x DMSO stocks at −20°C. Dye stock concentrations were normalized to the absorption of the dichlorofluorescein dye head via UV-Vis spectroscopy.
All salts and buffers were purchased from either Sigma-Aldrich (St. Louis, MO) or Fisher Scientific (Waltham, MA). TRAM-34, 4-aminopyridine, and charybdotoxin were purchased from Sigma-Aldrich. Gefitinib, wortmannin, sodium orthovanadate, and BAPTA-AM were purchased from Fisher Scientific. Canertinib was a gift from the Kuriyan laboratory at UC Berkeley. Gefitinib, wortmannin, canertinib, and TRAM-34 were made up as 1000x-10000x stock solutions in DMSO and stored at −20°C. Charybdotoxin was made up as a 1000x solution in water and stored at −80°C. 4-aminopyridine was made up as a 20x stock in imaging buffer (HBSS) and stored at 4°C. Recombinantly expressed epidermal growth factor was purchased from PeproTech (Rocky Hill, NJ) and aliquoted as a 1 mg/mL solution in water at −80°C.
Solid sodium orthovanadate was dissolved in water and activated before use 60. Briefly, orthovanadate solutions were repeatedly boiled and adjusted to pH 10 until the solution was clear and colorless. 200 mM activated orthovanadate stocks were aliquoted and stored at −20°C.
Unless otherwise noted, all imaging experiments were performed in Hank’s Balanced Salt Solution (HBSS; Gibco/Thermo Fisher Scientific). HBSS composition in mM: 137.9 NaCl, 5.3 KCl, 5.6 D-glucose, 4.2 NaHCO3, 1.3 CaCl2, 0.49 MgCl2, 0.44 KH2PO4, 0.41 MgSO4, 0.34 Na2HPO4. High K+ HBSS was made in-house to 285 mOsmol and pH 7.3, containing (in mM): 120 KCl, 23.3 NaCl, 5.6 D-glucose, 4.2 NaHCO3, 1.3 CaCl2, 0.49 MgCl2, 0.44 KH2PO4, 0.41 MgSO4, 0.34 Na2HPO4. Nominally Ca2+/Mg2+ free HBSS (Gibco) contained, in mM: 137.9 NaCl, 5.3 KCl, 5.6 D-glucose, 4.2 NaHCO3, 0.44 KH2PO4, 0.34 Na2HPO4.
Methods
Cell Culture
All cell lines were obtained from the UC Berkeley Cell Culture Facility and discarded after twenty passages. Cells were maintained in Dulbecco’s Modified Eagle Medium (DMEM) with 4.5 g/L D-glucose supplemented with 10% FBS (Seradigm (VWR); Radnor, PA) and 2 mM GlutaMAX (Gibco) in a 5% CO2 incubator at 37°C. Media for MCF-7 cells was supplemented with 1 mM sodium pyruvate (Life Technologies/Thermo Fisher Scientific) and 1x non-essential amino acids (Thermo Fisher Scientific). Media for CHO.K1 (referred to as CHO throughout the text) cells was supplemented with 1x non-essential amino acids. HEK293T and MDA-MB-231 were dissociated with 0.05% Trypsin-EDTA with phenol red (Thermo Fisher Scientific) at 37°C, whereas A431, CHO, and MCF-7 cells were dissociated with 0.25% Trypsin-EDTA with phenol red at 37°C. To avoid potential toxicity of residual trypsin, all cells except for HEK293T were spun down at 250xg or 500xg for 5 minutes and re-suspended in fresh complete media during passaging.
For use in imaging experiments, cells were plated onto 25 mm diameter poly-D-lysine coated #1.5 glass coverslips (Electron Microscopy Sciences) in 6 well tissue culture plates (Corning; Corning, NY). To maximize cell attachment, coverslips were treated before use with 1 2 M HCl for 2-5 hours and washed overnight three times with 100% ethanol and three times with deionized water. Coverslips were sterilized by heating to 150°C for 2-3 hours. Before use, coverslips were incubated with poly-D-lysine (Sigma-Aldrich, made as a 0.1 mg/mL solution in phosphate-buffered saline with 10 mM Na3BO4) for 1-10 hours at 37°C and then washed twice with water and twice with Dulbecco’s phosphate buffered saline (dPBS, Gibco).
A431, CHO, HEK293T, and MCF-7 were seeded onto glass coverslips 16-24 hours before microscopy experiments. MDA-MB-231 cells were seeded 48 hours before use because it facilitated formation of gigaseals during whole-cell voltage clamp electrophysiology. Cell densities used for optical resting membrane potential recordings (in 103 cells per cm2) were: A431 42; CHO 42; HEK293T 42; MCF-7 63; MDA-MB-231 42. To ensure the presence of single cells for whole-cell voltage clamp electrophysiology, fast-growing cells were plated more sparsely (approximately 20% confluence) for electrophysiology experiments. Cell densities used for electrophysiology (in 103 cells per cm2) were: A431 36-52; CHO 21; HEK293T 21; MCF-7 63; MDA-MB-231 42. To reduce their rapid growth rate, HEK293T cells were seeded onto glass coverslips in reduced glucose (1 g/L) DMEM with 10% FBS, 2 mM GlutaMAX, and 1 mM sodium pyruvate for electrophysiology experiments.
Cellular Loading of VoltageFluor Dyes
Cells were loaded with 1x VoltageFluor in HBSS for 20 minutes in a 37°C incubator with 5% CO2. For most experiments, 100 nM VoltageFluor was used. Serum-starved A431 cells were loaded with 50 nM VoltageFluor. After VF loading, cells were washed once with HBSS and then placed in fresh HBSS for imaging. All imaging experiments were conducted at room temperature under ambient atmosphere. Cells were used immediately after loading the VF dye, and no cells were kept for longer than an hour at room temperature.
Whole-Cell Patch-Clamp Electrophysiology
Pipettes were pulled from borosilicate glass with filament (Sutter Instruments, Novato, CA) with resistances ranging from 4 to 7 MΩ with a P97 pipette puller (Sutter Instruments). Internal solution composition, in mM (pH 7.25, 285 mOsmol/L): 125 potassium gluconate, 10 KCl, 5 NaCl, 1 EGTA, 10 HEPES, 2 ATP sodium salt, 0.3 GTP sodium salt. EGTA (tetraacid form) was prepared as a stock solution in either 1 M KOH or 10 M NaOH before addition to the internal solution. Pipettes were positioned with an MP-225 micromanipulator (Sutter Instruments). A liquid junction potential of − 14 mV was determined by the Liquid Junction Potential Calculator in the pClamp software package 61 (Molecular Devices, San Jose, CA), and all voltage step protocols were corrected for this offset.
Electrophysiology recordings were made with an Axopatch 200B amplifier and digitized with a Digidata 1440A (Molecular Devices). Signals were filtered with a 5 kHz low-pass Bessel filter. Correction for pipette capacitance was performed in the cell attached configuration. Voltagelifetime calibrations were performed in V-clamp mode, with the cell held at the potential of interest for 15 or 30 seconds while lifetime was recorded. Potentials were applied in random order, and membrane test was conducted between each step to verify the quality of the patch. For single cell patching, recordings were only included if they maintained a 30:1 ratio of membrane resistance (Rm) to access resistance (Ra) and an Ra value below 30 MΩ throughout the recording. Due to the reduced health of HEK293T cells transfected with CAESR, recordings were used as long as they maintained a 10:1 Rm:Ra ratio, although most recordings were better than 30:1 Rm:Ra. Only recordings stable for at least 4 voltage steps (roughly 2 minutes) were included in the dataset.
For electrophysiology involving small groups of cells (Fig. 2-supplement 4), complete voltage clamp across the entire cell group was not possible. Recordings were used as long as Ra remained below 30 MΩ for at least three voltage steps. Most recordings also retained Rm:Ra ratios greater than 20:1.
Epidermal Growth Factor Treatment
A431 cells were serum starved prior to epidermal growth factor studies. Two days before the experiment, cells were trypsizined and suspended in complete media with 10% FBS. Cells were then spun down for 5 minutes at 500xg and re-suspended in reduced serum DMEM (2% FBS, 2 mM GlutaMAX, 4.5 g/L glucose). Cells were seeded onto 25 mm coverslips in 6 well plates at a density of 84 × 103 cells per cm2. 4-5.5 hours before the experiment, the media was exchanged for serum-free DMEM (0% FBS, 2 mM GlutaMAX, 4.5 g/L glucose).
After 4-5.5 hours in serum-free media, cells were loaded with 50 nM VF dye as described above. In pharmacology experiments, the drug or vehicle was also added to the VF dye loading solution. All subsequent wash and imaging solutions also contained the drug or vehicle. For changes to buffer ionic composition, VoltageFluor dyes were loaded in unmodified HBSS to avoid toxicity from prolonged incubation with high K+ or without Ca2+. Immediately prior to use, cells were washed in the modified HBSS (120 mM K+ or 0 mM Ca2+) and recordings were made in the modified HBSS.
For analysis of short-term responses to EGF (3 minute time series), VF lifetime was recorded in 6 sequential 30 second exposures. Immediately after the conclusion of the first frame (30-35 seconds into the recording), EGF or vehicle (imaging buffer only) was added to the indicated final concentration from a 2x solution in HBSS imaging buffer. For analysis of longterm responses to EGF (15 minute time series), EGF addition occurred in the same way, but a gap of 150 seconds (without laser illumination) was allotted between each 30 second lifetime recording. Times given throughout the text correspond to the start of an exposure. Voltage changes at 2.5 minutes were calculated from the difference between an initial image (taken before imaging buffer vehicle or EGF addition) and a final image (a 30 second exposure starting 2.5 minutes into the time series).
Transfection and Imaging of CAESR in HEK293T
The CAESR plasmid was obtained as an agar stab (FCK-Quasar2-Citrine, Addgene #59172), cultured overnight in LB with 100 μg/mL ampicillin, and isolated via a spin miniprep kit (Qiagen). HEK293T cells were plated at a density of 42,000 cells per cm2 directly onto a 6 well tissue culture plate and incubated at 37°C in a humidified incubator for 24 hours prior to transfection. Transfections were performed with Lipofectamine 3000 according to the manufacturer’s protocol (Thermo Fisher Scientific). Cells were allowed to grow an additional 24 hours after transfection before they were plated onto glass coverslips for microscopy experiments (as described above for electrophysiology of untransfected HEK293T cells).
Determination of EC50 for EGF in A431 Cells
Average voltage changes 2.5 minutes after addition of EGF to serum deprived A431 cells were determined at different EGF concentrations, and these means were fit to a four parameter logistic function in MATLAB (MathWorks, Natick, MA).
Goldman-Hodgkin-Katz Estimation of Vmem in Different Imaging Buffers
If intracellular and extracellular concentrations, as well as relative permeabilities, of all ionic species are known, the Goldman-Hodgkin-Katz (GHK) equation (eqn. 1) can be used to calculate the resting membrane potential of a cell 32. In practice, the intracellular ion concentrations [X]in and relative permeabilities Px are difficult to determine. To obtain a range of reasonable Vmem values in systems where these concentrations and relative permeabilities are not known, we calculated possible Vmem using the “standard” parameters derived from the work of Hodgkin and Katz 32, as well as a value above and a value below each “standard” point. The values evaluated were the following: Pk 1; PNa 0.01, 0.05, 0.2; Pcl 0.2, 0.45, 0.9; [K+]in 90, 150, 200 mM; [Na+]in 5, 15, 50 mM; [C1−]in 2, 10, 35 mM. Extracellular ion concentrations [X]out were known (see Materials). In eqn. 1, R is the universal gas constant, T is the temperature (293 K for this experiment), and F is Faraday’s constant.
Fluorescence Lifetime Data Acquisition
Fluorescence lifetime imaging was conducted on a LSM 510 inverted scanning confocal microscope (Carl Zeiss AG, Oberkochen, Germany) equipped with an SPC-150 or SPC-150N single photon counting card (Becker & Hickl GmbH, Berlin, Germany) (Scheme S1). 80 MHz pulsed excitation was supplied by a Ti:Sapphire laser (MaiTai HP; SpectraPhysics, Santa Clara, CA) tuned to 958 nm and frequency-doubled to 479 nm. The laser was cooled by a recirculating water chiller (Neslab KMC100). Excitation light was directed into the microscope with a series of silver mirrors (Thorlabs, Newton, NJ or Newport Corporation, Irvine, CA).
Excitation light power at the sample was controlled with a neutral density (ND) wheel and a polarizer (P) followed by a polarizing beamsplitter (BS). Light was titrated such that VoltageFluor lifetime did not drift during the experiment, no phototoxicity was visible, and photon pile-up was not visible on the detector. For recordings at high VoltageFluor concentrations (Fig. 1-supplement 3, Fig. 2-supplement 5), reduced power was used to avoid saturating the detector. For optical voltage determinations using 50 or 100 nM VoltageFluor, typical average power at the sample was 5 μW.
Fluorescence emission was collected through a 40x oil immersion objective (Zeiss) coated with immersion oil (Immersol 518F, Zeiss). Emitted photons were detected with a hybrid detector, HPM-100-40 (Becker & Hickl), based on a Hamamatsu R10467 GaAsP hybrid photomultiplier tube. Detector dark counts were kept below 1000 per second during acquisition. Emission light was collected through a 550/49 bandpass filter (Semrock, Rochester, NY) after passing through a 488 LP dichroic mirror (Zeiss). The reference photons for determination of photon arrival times were detected with a PHD-400-N high speed photodiode (Becker & Hickl). Data were acquired with 256 time bins in the analog-to-digital-converter and either 64×64 or 256×256 pixels of spatial resolution (see discussion of pixel size below).
Routine evaluation of the proper functioning of the lifetime recording setup was performed by measurement of three standards (Fig. 1-supplement 2): 2 μM fluorescein in 0.1 N NaOH, 1 mg/mL erythrosin B in water (pH 7), and the instrument response function (IRF). The IRF was determined from a solution of 500 μM fluorescein and 12.2 M sodium iodide in 0.1 N NaOH. Because of the high concentration of iodide quencher, the IRF solution has a lifetime shorter than the detector response time, allowing approximation of the instrument response function under identical excitation and emission conditions as data acquisition 62.
Fluorescence Lifetime Data Processing and Conversion to Voltage
IRF Deconvolution
Signal from photons detected in a TCSPC apparatus are convolved with the instrument response (IRF). IRFs can be approximated by the SPCImage fitting software, but consistency of lifetime fits on VF-FLIM datasets was improved by using a measured IRF. Measured IRFs were incorporated by the iterative reconvolution method using SPCImage analysis software 63.
VoltageFluor Lifetime Fitting Model
All VoltageFluor lifetime data were fit using SPCImage (Becker & Hickl), which solves the nonlinear least squares problem using the Levenberg-Marquadt algorithm. VF2.1.Cl lifetime data were fit to a sum of two exponential decay components (eqn. 2). Attempts to fit the VF2.1.Cl data with a single exponential decay (eqn. 3) were unsatisfactory.
The fluorescence lifetime of VF2.0.Cl was adequately described by a single exponential decay for almost all data (eqn. 3). A second exponential component was necessary to fit data at VF2.0.Cl concentrations above 500 nM, likely attributable to the concentration-dependent decrease in lifetime that was observed high VF concentrations.
For all data fit with the two component model, the weighted average of the two lifetimes, τm (eqn. 4), was used in subsequent analysis.
All lifetime images are represented as an overlay of photon count (pixel intensity) and weighted average lifetime (pixel color) throughout the text (τm + PC, Scheme S2). Pixels with insufficient signal to fit a fluorescence decay are shown in black. The photon counts, as well as the lifetimes, in image sequences on the same set of cells are scaled across the same range.
Additional Fit Parameters for VoltageFluor Lifetimes
Pixels with photon counts below 300 (VF2.1.Cl) or 150 (VF2.0.Cl) photons at the peak of the decay (time bin with the most signal) were omitted from analysis to ensure reproducible fits. Because the lifetime of VFs does not fully decay to baseline in a single 12.5 ns laser cycle, the incomplete multiexponentials fitting option was used, allowing the model to attribute some signal early in the decay to the previous laser cycle. Out of 256 time bins from the analog-to-digital converter (ADC), only data from time bins 23 to 240 were used in the final fit. The offset parameter (detector dark counts per ADC time bin per pixel) was set to zero. The number of iterations for the fit in SPCImage was increased to 20 to obtain converged fits. Shift between the IRF and the decay trace was fixed to 0.5 (in units of ADC time bins), which consistently gave lifetimes of standards erythrosin B (1 mg/mL in H2O) 64 and fluorescein (2 μM in 0.1 N NaOH, H2O) 27 closest to reported values (Fig. 1 – supplement 2).
Effective Pixel Size
To obtain sufficient photons but keep excitation light power minimal, binning between neighboring pixels was employed during fitting. This procedure effectively takes the lifetime as a spatial moving average across the image by including adjacent pixels in the decay for a given pixel.
Pixel Sizes. For each recording type, the width of each pixel at acquisition is reported, as well as the width of the area included in the binned lifetime signal during fitting. All pixels are square.
Determination of Regions of Interest
Images were divided into cell groups, with each cell group as a single region of interest (ROI). ROIs were determined from photon count images, either manually from the cell morphology in ImageJ or automatically by sharpening and then thresholding the signal intensity with custom MATLAB code. Regions of images that were partially out of the optical section or contained punctate debris were omitted. Sample ROIs are shown in Scheme S2.
For cells that adjoin other cells, attribution of a membrane region to one cell versus the other is not possible. As such, we chose to interpret each cell group as an independent sample (‘n’) instead of extracting Vmem values for individual cells. Adjacent cells in a group are electrically coupled to varying degrees, and their resting membrane potentials are therefore not independent 31. While this approach did not fully utilize the spatial resolution of VF-FLIM, it prevented overestimation of biological sample size for the effect in question.
Conversion of Lifetime to Transmembrane Potential
The mean τm across all pixels in an ROI was used as the lifetime for that ROI. Lifetime values were mapped to transmembrane potential via the lifetime-Vmem standard curves determined with whole-cell voltage-clamp electrophysiology. For electrophysiology measurements, the relationship between the weighted average lifetime (eqn. 4) and membrane potential for each patched cell was determined by linear regression, yielding a sensitivity (m, ps/mV) and a 0 mV lifetime (b, ps) for each cell (eqn. 5). The average sensitivity and 0 mV point across all cells of a given type were used to convert subsequent lifetime measurements (τ) to Vmem (Figure 2-supplement 3, eqn. 6). For quantifying changes in voltage (ΔVmem) from changes in lifetime (Δτ), only the average sensitivity is necessary (eqn. 7).
Where standard error of the mean of a voltage determination (δVmem) is given, error was propagated to include the standard errors of the slope (δm) and y-intercept (δb) of the voltage calibration, as well as the standard error of the lifetime measurements (δτ) in the condition of interest (eqn. 8). For error in a voltage change (δΔVmem), only error in the calibration slope was included in the propagated error (eqn. 9). Where standard deviation of VF-FLIM derived Vmem values is shown, a similar error propagation procedure was applied, using the standard deviation of the average sensitivity and 0 mV lifetime for that cell line.
Resolution of VF-FLIM Voltage Determination
The intrinsic nature of fluorescence lifetime introduces a point of reference into the voltage measurement, from which a single lifetime image can be interpreted as resting membrane potential. The reproducibility of this reference point (reported here as the 0 mV lifetime) over time and across cells determines the accuracy of optical Vmem measurements. Because the sensitivities exhibited little variability within each cell type, the slope parameter contributes very little to the overall error.
The amount of voltage-independent noise in VF-FLIM can be estimated from lifetime-Vmem calibration data. We report resolution as the root-mean-square deviation (RMSD) of the optically calculated voltage (VFLIM) from the voltage set by whole-cell voltage clamp (Vephys). The RMSD of n measurements (eqn. 10) can be determined from the variance σ2 (eqn. 11) and the bias (eqn. 12) of the estimator (in this case, VF-FLIM) relative to the “true” value (in this case, electrophysiology).
The voltage-independent variations in lifetime are much larger between cells than within a cell. Therefore, the error in tracking the magnitude of voltage changes on an individual cell (“intracell” comparisons) is much lower than the error in making a comparison of absolute Vmem between two cells (“inter-cell” comparisons). We can therefore determine an “intra-cell” RMSD and an “inter-cell” RMSD to reflect the voltage resolution of these two types of measurements. To calculate “intra-cell” error, we look at the RMSD between Vephys and VFLIM using the τfl-Vmem relationship for that specific cell. Phrased another way, we are looking at the amount of error that would be expected in estimating a new Vmem for a cell based on a previous, optically-determined potential at that cell (i.e. changes in voltage). By averaging these “intra cell” RMSD values across all cells of a given type, we estimate the single-trial resolution for quantifying voltage changes is at or below 5 mV (Fig. 2-supplement 3).
The error in the absolute membrane potential determination (“inter-cell”) is calculated here as the RMSD between the y-intercept (0 mV lifetime) of all of the individual cells’ lifetime-voltage relationships and the 0 mV value for the averaged calibration for all cells of a given type. This metric addresses how well the lifetime-Vmem relationship for a given cell type is likely to represent an individual cell’s lifetime-Vmem relationship. This “inter cell” RMSD ranged from 11 to 24 mV for the tested cell lines (Fig. 2-supplement 3). Because of the improved throughput of VF-FLIM, much smaller errors for a population value of Vmem can be obtained by and averaging Vmem recordings from multiple cells.
This method of calculating error assumes that the electrophysiology measurement is perfectly accurate and precise. Realistically, it is likely that some of the variation seen is due to the quality of the voltage clamp. As a result, these RMSD values provide a conservative upper bound for the voltage errors in VF-FLIM.
Analysis of CAESR Lifetimes
For sample images of CAESR in HEK293T (Fig. 1-supplement 5), fluorescence decays were fit using SPCImage to a biexponential decay model as described for VF2.1.Cl above, using a peak photon threshold of 150 and a bin of 2 (binned pixel width of 5 μm). To better match the studies by Cohen and co-workers 21, which isolated the membrane fluorescence from cytosolic fluorescence by directing the laser path, the lifetime-voltage relationships were not determined with these square-binned images. Instead, membranes were manually identified, and the fluorescence decays from all membrane pixels were summed together before fitting once per cell. (This is in contrast to the processing of VoltageFluor data, where the superior signal to noise and localization enables fitting and analysis of the lifetime on a pixel by pixel basis). This “one fit per membrane” analysis of CAESR was performed in custom MATLAB code implementing a Nelder-Meade algorithm, in which CAESR data were fit to a biexponential model with the offset fixed to 0 and the color shift as a free parameter.
Acknowledgments
We thank Holly Aaron and Vadim Degtyar for expert technical assistance and training in the use of FLIM, Prof. John Kuriyan and Dr. Sean Peterson for helpful discussions, and members of the Miller lab for providing VF dyes. FLIM experiments were performed at the CRL Molecular Imaging Center, supported by NSF DBI-0116016. Cell lines were from the UCB Cell Culture Facility. FCK-QuasAr2-Citrine was a gift from Adam Cohen (Addgene plasmid # 59172). JLD was supported by an NSF Graduate Research Fellowship. EWM acknowledges support from the Sloan Foundation (FG-2016-6359), March of Dimes (5-FY16-65), and the NIH (R35GM119855).
References
- (1).↵
- (2).↵
- (3).↵
- (4).↵
- (5).↵
- (6).↵
- (7).
- (8).↵
- (9).↵
- (10).↵
- (11).↵
- (12).↵
- (13).↵
- (14).↵
- (15).↵
- (16).↵
- (17).↵
- (18).↵
- (19).↵
- (20).↵
- (21).↵
- (22).↵
- (23).↵
- (24).↵
- (25).↵
- (26).↵
- (27).↵
- (28).↵
- (29).↵
- (30).↵
- (31).↵
- (32).↵
- (33).↵
- (34).↵
- (35).↵
- (36).↵
- (37).↵
- (38).↵
- (39).↵
- (40).↵
- (41).↵
- (42).↵
- (43).↵
- (44).↵
- (45).↵
- (46).↵
- (47).↵
- (48).
- (49).↵
- (50).↵
- (51).↵
- (52).↵
- (53).↵
- (54).↵
- (55).↵
- (56).↵
- (57).↵
- (58).↵
- (59).↵
- (60).↵
- (61).↵
- (62).↵
- (63).↵
- (64).↵
- (65).↵
- (66).
- (67).
- (68).
- (69).
- (70).
- (71).
- (72).
- (73).
- (74).
- (75).
- (76).