Genetically encoded sensors based on fluorescence resonance energy transfer (FRET) are powerful tools for quantifying and visualizing analytes in living cells, and when targeted to organelles have the potential to define distribution of analytes in different parts of the cell. However, quantitative estimates of analyte distribution require rigorous and systematic analysis of sensor functionality in different locations. In this work, we establish methods to critically evaluate sensor performance in different organelles and carry out a side-by-side comparison of three different genetically encoded sensor platforms for quantifying cellular zinc ions (Zn2+). Calibration conditions are optimized for high dynamic range and stable FRET signals. Using a combination of single-cell microscopy and a novel microfluidic platform capable of screening thousands of cells in a few hours, we observe differential performance of these sensors in the cytosol compared to the ER of HeLa cells, and identify the formation of oxidative oligomers of the sensors in the ER. Finally, we use new methodology to re-evaluate the binding parameters of these sensors both in the test tube and in living cells. Ultimately, we demonstrate that sensor responses can be affected by different cellular environments, and provide a framework for evaluating future generations of organelle-targeted sensors.

Fluorescent biosensors are important measurement tools for in vivo quantification of pH, concentrations of metal ions and other analytes, and physical parameters such as membrane potential. Both the development of these sensors and their implementation in examining cellular heterogeneity requires technology for measuring and sorting cells based on the fluorescence levels before and after chemical or physical perturbations. We developed a droplet microfluidic platform for the screening and separation of cell populations on the basis of the in vivo response of expressed fluorescence-based biosensors after addition of an exogenous analyte. We demonstrate the capability to resolve the responses of two genetically encoded Zn2+ sensors at a range of time points spanning several seconds and subsequently sort a mixed-cell population of varying ratios with high accuracy.

The bacterial pathogen Salmonella uses sophisticated type III secretion systems (T3SS) to translocate and deliver bacterial effector proteins into host cells to establish infection. Monitoring these important virulence determinants in the context of live infections is a key step in defining the dynamic interface between the host and pathogen. Here, we provide a modular labeling platform based on fluorescence complementation with split-GFP that permits facile tagging of new Salmonella effector proteins. We demonstrate enhancement of split-GFP complementation signals by manipulating the promoter or by multimerizing the fluorescent tag and visualize three effector proteins, SseF, SseG, and SlrP, that have never before been visualized over time during infection of live cells. Using this platform, we developed a methodology for visualizing effector proteins in primary macrophage cells for the first time and reveal distinct differences in the effector-defined intracellular niche between primary macrophage and commonly used HeLa and RAW cell lines.Keywords: bacterial effector proteins; host−pathogen interface; live cell imaging; Salmonella; split-GFP;

Various fluorescent probes have been developed to reveal the biological functions of intracellular labile Zn2+. Here, we present Green Zinc Probe (GZnP), a novel genetically encoded Zn2+ sensor design based on a single fluorescent protein (single-FP). The GZnP sensor is generated by attaching two zinc fingers (ZF) of the transcription factor Zap1 (ZF1 and ZF2) to the two ends of a circularly permuted green fluorescent protein (cpGFP). Formation of ZF folds induces interaction between the two ZFs, which induces a change in the cpGFP conformation, leading to an increase in fluorescence. A small sensor library is created to include mutations in the ZFs, cpGFP and linkers between ZF and cpGFP to improve signal stability, sensor brightness and dynamic range based on rational protein engineering, and computational design by Rosetta. Using a cell-based library screen, we identify sensor GZnP1, which demonstrates a stable maximum signal, decent brightness (QY = 0.42 at apo state), as well as specific and sensitive response to Zn2+ in HeLa cells (Fmax/Fmin = 2.6, Kd = 58 pM, pH 7.4). The subcellular localizing sensors mito-GZnP1 (in mitochondria matrix) and Lck-GZnP1 (on plasma membrane) display sensitivity to Zn2+ (Fmax/Fmin = 2.2). This sensor design provides freedom to be used in combination with other optical indicators and optogenetic tools for simultaneous imaging and advancing our understanding of cellular Zn2+ function.

There is a critical need for high-speed multiparameter photophysical measurements of large libraries of fluorescent probe variants for imaging and biosensor development. We present a microfluidic flow cytometer that rapidly assays 104–105 member cell-based fluorophore libraries, simultaneously measuring fluorescence lifetime and photobleaching. Together, these photophysical characteristics determine imaging performance. We demonstrate the ability to resolve the diverse photophysical characteristics of different library types and the ability to identify rare populations.

Fluorescent proteins offer exceptional labeling specificity in living cells and organisms. Unfortunately, their photophysical properties remain far from ideal for long-term imaging of low-abundance cellular constituents, in large part because of their poor photostability. Despite widespread engineering efforts, improving the photostability of fluorescent proteins remains challenging due to lack of appropriate high-throughput selection methods. Here, we use molecular dynamics guided mutagenesis in conjunction with a recently developed microfluidic-based platform, which sorts cells based on their fluorescence photostability, to identify red fluorescent proteins with decreased photobleaching from a HeLa cell-based library. The identified mutant, named Kriek, has 2.5- and 4-fold higher photostability than its progenitor, mCherry, under widefield and confocal illumination, respectively. Furthermore, the results provide insight into mechanisms for enhancing photostability and their connections with other photophysical processes, thereby providing direction for ongoing development of fluorescent proteins with improved single-molecule and low-copy imaging capabilities.

Fluorescent sensors are powerful tools for visualizing and quantifying molecules and ions in living cells. A variety of small molecule and genetically encoded sensors have been developed for studying intracellular Zn2+ homeostasis and signaling, but no direct comparisons exist, making it challenging for researchers to identify the appropriate sensor for a given application. Here we directly compare the widely used small molecule probe FluoZin-3 and a genetically encoded sensor, ZapCY2. We demonstrate that, in contrast to FluoZin-3, ZapCY2 exhibits a well-defined cytosolic localization, provides estimates of Zn2+ concentration with little variability, does not perturb cytosolic Zn2+ levels, and exhibits rapid Zn2+ response dynamics. ZapCY2 was used to measure Zn2+ concentrations in 5 different cell types, revealing higher cytosolic Zn2+ levels in prostate cancer cells compared to normal prostate cells (although the total zinc is reduced in prostate cancer cells), suggesting distinct regulatory mechanisms.

Fluorescence resonance energy transfer (FRET)-based genetically encoded metal-ion sensors are important tools for studying metal-ion dynamics in live cells. We present a time-resolved microfluidic flow cytometer capable of characterizing the FRET-based dynamic response of metal-ion sensors in mammalian cells at a throughput of 15 cells/s with a time window encompassing a few milliseconds to a few seconds after mixing of cells with exogenous ligands. We have used the instrument to examine the cellular heterogeneity of Zn2+ and Ca2+ sensor FRET response amplitudes and demonstrated that the cluster maps of the Zn2+ sensor FRET changes resolve multiple subpopulations. We have also measured the in vivo sensor response kinetics induced by changes in Zn2+ and Ca2+ concentrations. We observed an ∼30 fold difference between the extracellular and intracellular sensors.

Traditional flow cytometers are capable of rapid cellular assays on the basis of fluorescence intensity and light scatter. Microfluidic flow cytometers have largely followed the same path of technological development as their traditional counterparts; however, the significantly smaller transport distance and resulting lower cell speeds in microchannels provides for the opportunity to detect novel spectroscopic signatures based on multiple, nontemporally coincident excitation beams. Here, we characterize the design and operation of a cytometer with a three-beam, probe/bleach/probe geometry, employing HeLa suspension cells expressing fluorescent proteins. The data collection rate exceeds 20 cells/s under a range of beam intensities (5 kW to 179 kW/cm2). The measured percent photobleaching (ratio of fluorescence intensities excited by the first and third beams: Sbeam3/Sbeam1) partially resolves a mixture of four red fluorescent proteins in mixed samples. Photokinetic simulations are presented and demonstrate that the percent photobleaching reflects a combination of the reversible and irreversible photobleaching kinetics. By introducing a photobleaching optical signature, which complements traditional fluorescence intensity-based detection, this method adds another dimension to multichannel fluorescence cytometry and provides a means for flow-cytometry-based screening of directed libraries of fluorescent protein photobleaching.

Zinc (Zn2+) homeostasis plays a vital role in cell function, and the dysregulation of intracellular Zn2+ is associated with mitochondrial dysfunction. Few tools exist to quantitatively monitor the buffered, free Zn2+ concentration in mitochondria of living cells ([Zn2+]mito). We have validated three high dynamic range, ratiometric, genetically encoded, fluorescent Zn2+ sensors that we have successfully used to precisely measure and monitor [Zn2+]mito in several cell types. Using one of these sensors, called mito-ZapCY1, we report observations that free Zn2+ is buffered at concentrations about 3 orders of magnitude lower in mitochondria than in the cytosol and that HeLa cells expressing mito-ZapCY1 have an average [Zn2+]mito of 0.14 pM, which differs significantly from other cell types. These optimized mitochondrial Zn2+ sensors could improve our understanding of the relationship between Zn2+ homeostasis and mitochondrial function.

Zn2+ plays essential roles in biology, and cells have adopted exquisite mechanisms for regulating steady-state Zn2+ levels. Although much is known about total Zn2+ in cells, very little is known about its subcellular distribution. Yet defining the location of Zn2+ and how it changes with signaling events is essential for elucidating how cells regulate this essential ion. Here we create fluorescent sensors genetically targeted to the endoplasmic reticulum (ER) and Golgi to monitor steady-state Zn2+ levels as well as flux of Zn2+ into and out of these organelles. These studies reveal that ER and Golgi contain a concentration of free Zn2+ that is 100 times lower than the cytosol. Both organelles take up Zn2+ when cytosolic levels are elevated, suggesting that the ER and Golgi can sequester elevated cytosolic Zn2+ and thus have the potential to play a role in influencing Zn2+ toxicity. ER Zn2+ homeostasis is perturbed by small molecule antagonists of Ca2+ homeostasis and ER Zn2+ is released upon elevation of cytosolic Ca2+ pointing to potential exchange of these two ions across the ER. This study provides direct evidence that Ca2+ signaling can influence Zn2+ homeostasis and vice versa, that Zn2+ dynamics may modulate Ca2+ signaling.

Long regarded as a toxic byproduct, hydrogen peroxide is increasingly recognized as an important cellular signal. Efforts at defining the spatiotemporal nature of hydrogen peroxide production recently got a boost by the development of a series of organelle-targeted fluorescent probes by Srikun et al. (2010).

The ER plays a fundamental role in storing cellular Ca2+, generating Ca2+ signals, and modulating Ca2+ in both the cytosol and mitochondria. Genetically encoded Ca2+ sensors can be explicitly targeted to the ER to directly define Ca2+ levels and monitor fluxes of Ca2+ within this organelle. In this study we use an ER-targeted Ca2+ sensor to define both the level and dynamics of ER Ca2+ in cells expressing mutant presenilin proteins. Growing evidence suggests the enigmatic presenilin-1 plays a role in regulating ER Ca2+. Presenilin-1 was initially identified in a screen for genetic causes of inherited familial Alzheimer's disease (fAD). The connection between presenilin-1, calcium regulation, and Alzheimer's disease may provide the key to understanding the long-observed, but poorly understood, link between Alzheimer's disease and Ca2+ dysregulation. In this study we examined seven fAD-causing mutations in presenilin-1 to define how they influence ER Ca2+ levels and dynamics. We observed that some, but not all, mutations in PS1 decrease the level of Ca2+ within the ER and this difference depends on the enzymatic activity of PS1. Two mutations tested altered the kinetics of Ca2+ release from the ER upon ATP stimulation, resulting in faster spiking. Combined, these results indicate that mutations in PS1 can alter the balance of Ca2+ in cells and have the potential to influence the nature of Ca2+ signals.

Fluorescent indicators for calcium are incredibly powerful because they enable researchers to watch the movement of calcium ions in real time in living cells. The popular small molecule indicator Indo-1 has now been targeted to a defined location, namely, the nucleus of muscle cells, using SNAP-tag technology. This combination of a chemical probe with genetic targeting expands the available options for measuring local calcium events in cells.

Gram-negative pathogenic bacteria such as Salmonella utilize the type III secretion system to inject bacterial effector proteins into a host cell. Upon entry, these effectors bind mammalian cell proteins, hijack cellular signaling pathways, and redirect cellular function, thus enabling bacterial infection. In this study we use the FlAsH/tetracysteine labeling system to fluorescently tag specific effectors in Salmonella to observe real-time secretion of these proteins into a mammalian host cell. The tetracysteine tag is genomically incorporated, thus preserving endogenous control of bacterial effectors. We demonstrate that two effectors, SopE2 and SptP, exhibit different secretion kinetics, as well as different rates of degradation within the host cell. These proteins respectively activate and suppress GTPase Cdc42, suggesting that there is a temporal hierarchy for effector delivery and persistence within the cell that is directly related to effector function.

Over the past 20 years, protein engineering has been extensively used to improve and modify the fundamental properties of fluorescent proteins (FPs) with the goal of adapting them for a fantastic range of applications. FPs have been modified by a combination of rational design, structure-based mutagenesis, and countless cycles of directed evolution (gene diversification followed by selection of clones with desired properties) that have collectively pushed the properties to photophysical and biochemical extremes. In this review, we provide both a summary of the progress that has been made during the past two decades, and a broad overview of the current state of FP development and applications in mammalian systems.

Fluorescent proteins (FPs) are powerful tools that permit real-time visualization of cellular processes. The utility of a given FP for a specific experiment depends strongly on its effective brightness and overall photostability. However, the brightness of FPs is limited by dark-state conversion (DSC) and irreversible photobleaching, which occur on different timescales. Here, we present in vivo ensemble assays for measuring DSC and irreversible photobleaching under continuous and pulsed illumination. An analysis of closely related red FPs reveals that DSC and irreversible photobleaching are not always connected by the same mechanistic pathway. DSC occurs out of the first-excited singlet state, and its magnitude depends predominantly on the kinetics for recovery out of the dark state. The experimental results can be replicated through kinetic simulations of a four-state model of the electronic states. The methodology presented here allows light-driven dynamics to be studied at the ensemble level over six orders of magnitude in time (microsecond to second timescales).