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A method for estimating intracellular ion concentration using optical nanosensors and ratiometric imaging

Scientific Reportsvolume 7, Article number: 10819 (2017) | Download Citation


Optical nanoparticle (NP)-based sensors have been widely implemented as tools for detection of targeted ions and biomolecules. The NP sensing platform offer a modular design that can incorporate different sensing components for greater target specificity and the ability to tune the dynamic range, as well as encapsulation of multiple dyes to generate a ratiometric signal with varying spectra. Despite these advantages, demonstrating quantitative ion imaging for intracellular measurement still possess a major challenge. Here, we describe fundamentals that enable intracellular validation of this approach using ion-selective nanosensors for investigating calcium (Ca2+) as a model ion. While conventional indicators can improve individual aspects of indicator performance such as Kd, wavelength, and ratiometric measurements, the use of NP sensors can achieve combined benefits of addressing these issues simultaneously. The nanosensor incorporates highly calcium-selective ionophores and two fluorescence indicators that act as signal transducers to facilitate quantitative ratiometric imaging. For intracellular Ca2+ application, the sensors are fine-tuned to physiological sensing range, and live-cell imaging and quantification are demonstrated in HeLa cells loaded with nanosensors and their responsiveness to carbachol-evoked store release (~400 nM). The current nanosensor design thus provides a promising sensing platform for real-time detection and optical determination of intracellular ions.


Optical imaging and fluorescent probes provide key tools for exploring the spatiotemporal dynamics of ions in living systems. In particular, real-time detection of ion dynamics in the cellular environment requires designing optical probes with high target specificity and sensitivity. Major progress has been achieved with the advent of fluorescent calcium (Ca2+) indicators1, 2, luminescent metal-complexes3, 4, quantum dots5, 6, and genetically encoded indicators7, 8 which continue to provide powerful tools for the imaging of ions. However, when applied to intracellular measurement, accurate determination of the intracellular concentration with these indicators can be complicated by a non-optimal dissociation constant (Kd), low signal, undesired compartmentalization, and interference with cellular components potentially leading to variability and inaccurate measurements. A number of fluorophores are commercially available to achieve quantitative ion imaging as demonstrated by single wavelength dyes in Ca2+ recordings9, 10, as well as other approaches including fluorescence resonance energy transfer (FRET)-based sensors11, 12. The accuracy of optical quantification is often improved by ratiometric methods which eliminates interference caused by fluctuations in excitation intensity, fluorophore concentration, and light scattering due to specimen thickness. Traditionally, quantitative Ca2+ imaging has been achieved with ratiometric indicator dyes such as Fura-22, 13 and Indo-114, and fluorescence lifetime imaging (FLIM) methods15. However, ratiometric imaging in practice is limited to only a few indicators in the UV spectrum associated with excitation light-induced phototoxicity, higher tissue scattering, and most importantly, the requirement of structural modification to alter its physiochemical properties such as the sensing range for desired biological applications.

Nanoparticle (NP)-based sensors16,17,18,19,20,21,22 provides an efficient and promising approach for developing robust imaging probes for quantitative ion detection. For example, while molecular probes can improve individual aspects of indicator performance such as Kd, wavelength, and ratiometric measurements, the use of NP-based sensors can achieve combined benefits of addressing these issues simultaneously. By incorporating multiple agents in the NP matrix, the advantages of these classes of sensors include: (i) brighter and photostable probes by integrating a high number of dyes; (ii) ability to fine-tune the sensor dynamic range; (iii) greater selectivity and range of analyte detection; (iv) adjustable emission spectra and ratiometric response; (v) reduced effects from light-induced cell damage23 by integrating longer wavelength dyes; and (vi) low cytoplasmic mobility24,25,26,27 to prevent dye diffusion, and protection from intracellular organelle sequestration28, 29. Indeed, the recent advances in NP-based sensors from various nanomaterials have been widely implemented in cellular tracking and imaging applications16,17,18,19,20,21,22. For example, there are several nano- and micro-particle sensors for detection of ions reported for sodium30,31,32, potassium33,34,35,36,37, calcium38, 39, magnesium40, 41, zinc42, silver43, and chloride44. Other efforts include nanosensors termed PEBBLEs (probes encapsulated by biologically localized embedding) developed by Kopelman and co-workers28, 45. These sensors incorporate molecular indicators such as rhod-2 into a polymeric matrix, however, the flexibility to tune the dynamic range remains challenging with this mechanism46. While the development of NP-based sensors has been an increasingly growing field of research, the application for measurement of intracellular ions, and detailed procedure for quantitative ratiometric ion imaging has not been fully established in these sensors.

To address this issue, we present a ratiometric sensor termed calcium-optode nanosensor (opCaNS) for quantitative fluorescence imaging of ions in living cells, and establish a procedure for estimating ionic concentration using ratiometric fluorescent signal. Calcium is selected as a model target ion to reliably assess our sensor characteristics to the extensive work established on the local control of Ca2+ signaling, as widely exemplified by cellular processes ranging from gene expression, synaptic transmission, hormone secretion to cell survival and death47,48,49,50.

In the nanosensor design, the ion-selective optodes (ISOs)21, 46, 51, an optical counterpart to ion-selective electrodes (ISEs), were applied and miniaturized to nanoscale for intracellular investigation. In the ISO sensing scheme, three reagents are embedded within the NP matrix: a highly selective, optically silent ion carrier (ionophore)21, 46, 51 that is capable of reversibly binding ions; a fluorescent pH indicator dye (chromoionophore) which acts as a reporter to yield a fluorescence readout based on its protonation degree; and a lipophilic charge-carrying molecule (ion exchanger) to maintain electroneutrality within the NP. Hence, ion-specific optical change is facilitated by binding of calcium ions by the ionophores, which causes displacement of protons of the pH-indicator with a quantifiable fluorescence signal. Furthermore, we have incorporated a second pH-indicator [octadecyl rhodamine (R18)] and term it as reference dye, whereby fluorescence intensity moves in opposite direction to pH changes in comparison to the chromoionophore to achieve a highly sensitive sensor for ratiometric measurement of intracellular Ca2+. Most importantly, the sensing range of the probe is easily modified and tuned by changing the ratio of components, which provides flexibility to the user for specific imaging applications.

Overall, the opCaNS provides a sensing platform suitable for intracellular Ca2+ measurement using ratiometric approach with sensor characteristics tuned to exhibit physiological sensing range, high selectivity, fast response time, and a reversible profile. Herein, we demonstrate the performance of this class of ion-selective nanosensors for cellular applications, achieving real-time detection and quantitative ion measurements by live-cell imaging for the first time.


Principle and characterization of ratiometric nanosensor

To achieve nanosensors for real-time detection and measurement of intracellular free calcium ions ([Ca2+]i), we applied ISOs as the principle sensing scheme, and further developed into miniaturized nanoscale sensors for ratiometric fluorescence imaging. In this sensing platform, the nanosensors are fabricated by co-immobilizing hydrophobic optode components comprised of calcium ionophore (I), a fluorescent chromoionophore (C), ion exchanger (R), and reference fluorophore (Ref) within a biocompatible plasticizer nano-emulsion matrix, and stabilized by a biocompatible polyethylene glycol (PEG)-lipid as surfactant (Fig. 1). The hydrodynamic diameter of nanosensors measured 67 nm (polydispersity index 0.16) (Fig. 2a) with zeta-potential of −29.9 ± 1.4 mV, indicating nanoparticles that are stable in aqueous solution. Mechanistically, the highly selective calcium ionophore acts as a recognition element that extracts Ca2+ into the sensor, which causes the pH-sensitive fluorophore (chromoionophore) to deprotonate to maintain charge neutrality within the sensor with an accompanying change in fluorescence. In this scheme, the opCaNS respond to Ca2+ in accordance with optode function (Eqs 13)21, 46, 52. The ion-exchange optode mechanism in a system at equilibrium is described by:

$${M}^{z+}(aq)+n{\rm{I}}(org)+z{{\rm{CH}}}^{+}(org)\leftrightarrow M{{\rm{I}}}_{n}^{z+}(org)+z{{\rm{C}}}^{-}(org)+z{{\rm{H}}}^{+}(aq)$$

where, M, I, and C are the ion of interest, ionophore, and chromoionophore, respectively, and aq and org denote an ion, M, in aqueous or organic phase of the optode component, respectively. The exchange constant to describe the partitioning between the two phases in the ion-exchange system is then given by:

$${K}_{exch}^{M{{\rm{I}}}_{n}}={(\frac{{\alpha }_{H}[{\rm{C}}]}{[{{\rm{CH}}}^{+}]})}^{z}\frac{[M{{\rm{I}}}_{n}^{z+}]}{{\alpha }_{M}{[{\rm{I}}]}^{n}}={(\frac{{K}_{\alpha }}{{K}_{H}})}^{z}{k}_{{\rm{M}}}{\beta }_{{\rm{M}}{I}_{n}}$$

where, αH and αM denote the activity of a proton, H, or the ion, M, in the sensor. The optical activity reflects the concentration of respective complexes within the organic phase of the optode component by maintaining electroneutrality46. Under these conditions, the relation between the deprotonation ratio (α) of the nanosensor and the ion activity is:

$${\alpha }_{{\rm{I}}}={(z{K}_{exch}^{M{{\rm{I}}}_{n}})}^{-1}{(\frac{1-\alpha }{\alpha }{\alpha }_{H})}^{z}\times \frac{{R}_{T}^{-}-\alpha {C}_{T}}{{\{{I}_{T}-({R}_{T}^{-}-\alpha {C}_{T})(n/z)\}}^{n}}$$
Figure 1
Figure 1

Calcium nanosensor working principle. The sensing components (chromoionophore, C; ionophore, I; ion exchanger, R) embedded in an organic hydrophobic sensor matrix. The recognition of target ion by selective ionophore-mediated recruitment results in an optical signal output as determined by the protonation degree of the chromoionophore. The overall electroneutrality of the sensor is maintained by ion exchanger. Ratiometric responses are obtained by incorporation of additional reference dye (Ref). The surface is covered with polyethylene glycol (PEG)-lipid molecules to offer particle stabilization. On the bottom are chemical structures and abbreviations of the main sensor components used in this work.

Figure 2
Figure 2

Sensor characterization and calibration. (a) Size distribution of nanosensors as measured by dynamic light scattering show an effective average diameter of 67 nm. (b,c) Calibration of the calcium nanosensors performed in solution. (b) Sensor response can be fine-tuned by altering amounts of sensing components. (c) For intracellular applications, the sensor response is tuned to a characteristic Ca2+ concentration derived from Hill fit (defined here as, EC50) at nanomolar range (105 ± 19 nM), with a corresponding linear range between 38 to 600 nM (inset). (d) Characterization of nanosensors in response to Ca2+ (blue) demonstrate 4 orders of magnitude selectivity over Mg2+ (purple). (e) Two-photon Ca2+ uncaging experiment using opCaNS. Nanosensors (n = 10) respond instantaneously to elevations of local Ca2+ levels by DMNP uncaging, as indicated by an increase of fluorescence levels from the base line (arrow). (f) Reversible response over repeated cycles between exchanged Ca2+ concentrations (100 nM and 1350 nM) characterized by confocal microscopy (n = 3). Measurements are taken in triplicate from three individual sets of nanosensors (n = 9), unless specified otherwise. Data is shown as mean ± standard deviation (SD).

For sensor calibration, the emission intensity ratios of two fluorophores (CHIII:R18, Eqs 46, see Methods) in response to Ca2+ was used to plot a calibration curve with a Hill fit to determine effective Ca2+ concentration at half-maximal sensor response (defined here as EC50). The emission spectra for individual fluorophores are shown in Supplementary Figure S1. The sensing range can be fine-tuned (Fig. 2b) by altering the amount of the individual sensing components (CaI-II, CHIII, NaTFPB, see Methods) as previously demonstrated53.

For intracellular applications, the ratiometric response of nanosensor can be tuned to exhibit EC50 at 105 ± 19 nM in solution (Fig. 2c), which is close to 100 nM intracellular calcium basal level48. There were batch-to-batch differences in sensor response and calibrations were routinely performed to ensure the sensors maintain optimal responsiveness for subsequent cellular studies. An index of nanosensor sensitivity, as determined by the slope of linear portion of the normalized response at EC50 (Fig. 2c, inset), measured a decrease of 80% between 38 to 600 nM.

For sensor selectivity, the sensor responded to Ca2+ with high selectivity over the primary intracellular interfering ion, magnesium (Mg2+) (Fig. 2d). The opCaNS demonstrated EC50 for magnesium >1 mM, which produces a log selectivity coefficient of 4 (Eq. 7, zero background ion). The physiological range of ionized Mg2+ concentration is known to be 0.5–1 mM54, suggesting there is minimal interference of sensor response by Mg2+.

It is critical to determine the sensor response time to assess the ability of nanosensors to capture ionic transients which occur on the millisecond-to-second time scale47,48,49,50. Response time of the nanosensor was assessed by a two-photon (2P) Ca2+ uncaging experiment (Fig. 2e). We first demonstrated that the opCaNS is 2P-excitable with maximum fluorescence observed at excitation wavelengths centered at 875 nm (Supplementary Figure S2). Subsequent uncaging data shows that nanosensors responded instantaneously to elevations of local Ca2+ levels when imaged at a frame-rate of 6 Hz, as indicated by an increase of fluorescence from baseline. This suggests the capability of nanosensors to capture intracellular Ca2+ signals by store-release mechanisms (see next section). It is worth noting that the measured response time of the nanosensors here is limited by the imaging frame-rate, and the temporal resolution can be significantly improved by other imaging methods31.

Reversibility is also key to monitoring Ca2+ release from the intracellular store due to its rise and decay characteristics47,48,49,50. The ratiometric response of the opCaNS exhibited a clear reversibility profile (Fig. 2f) over repeated cycles of alternating Ca2+ concentrations (100 and 1350 nM), which is indicative of excellent cycling capability of the nanosensors. The reversibility profile for individual fluorophores (CHIII and R18) is shown in Supplementary Figure S3.

Collectively, the results show applicable sensor characteristics for reporting local cytoplasmic Ca2+ concentration by exhibiting nanomolar response range, high selectivity, fast response time, and reversible response.

Sensing intracellular calcium signals

To demonstrate the capabilities of our nanosensors for intracellular applications, we examined carbachol-induced Ca2+ store release in HeLa cells (n = 3) which acts on inositol 1,4,5-triphosphate receptors (IP3Rs) on the endoplasmic reticulum (ER) membrane55,56,57. First, nanosensors were loaded into HeLa cells using pressure controlled microinjection (Fig. 3a, inset). Although nanosensors have previously been introduced to intracellular space through endocytosis39, 58, the microinjection technique ensures nanoparticle delivery to the cell cytosol and prevents encapsulation by endosomes31, 59,60,61. Each injection was about one tenth of total volume correlating to ~102−103 nanosensors (picomolar) per cell (Supplementary Figure S4), which is an amount that provided robust signal-to-noise ratio for imaging without inflicting damage to the cells.

Figure 3
Figure 3

Intracellular Ca2+ detection by carbachol-mediated calcium store release. (a) Time-lapse confocal imaging of nanosensor-loaded cells (inset) reveal an initial decrease in fluorescence intensity ratio followed by an increase to the base level, indicating a rise in intracellular Ca2+ level followed by cytoplasmic depletion. (b) Conversion of fluorescence signal to Ca2+ levels indicate cytosolic elevations reach up to ~400 nM. (c) In situ cell calibration (green; EC50 = 110 ± 42 nM) was performed by superfusing varying concentrations of Ca2+ in presence of ionomycin to allow Ca2+ levels to be equilibrated across the cell membrane. This calibration is compared with the result (EC50 = 54 ± 14 nM) obtained from in-solution calibration experiment using identical nanosensors (blue). CCh: carbachol. Scale bar: 10 µm.

Time-lapse confocal imaging (Fig. 3a) revealed an initial short-lived decrease followed by a slower increase in sensor intensity ratio. This indicates an elevation of Ca2+ levels followed by a slow decay of cytoplasmic Ca2+ to its basal level. When the intensity ratios are converted to Ca2+ concentration, the cytosolic elevations reached up to ~400 nM (Fig. 3b). The empirical conversion involved performing in situ cell calibration (Fig. 3c) and curve fitted by the Hill equation (Eq. 8) to determine the EC50 after the carbachol experiment on the same cell. Briefly, in situ cell calibrations were performed by superfusing nanosensor-loaded HeLa cells with varying concentrations of Ca2+ using calibration buffer in presence of 5 µM ionomycin, which serve as ion carriers to facilitate the equilibrium of Ca2+ levels across the cell membrane. Although measurement of EC50 in solution can serve as an index of sensor performance, the behavior can change when loaded in living cells62, 63. As shown in Fig. 3c, the EC50 of in situ cell calibration was determined to be 110 ± 42 nM. In comparison, the EC50 of the same batch of nanosensor in solution was 54 ± 14 nM. We note that the differences between in situ and in solution calibrations in our sensor response (EC50) is less or comparable to those observed in previous studies using dye indicators and PEBBLE NP-sensors (Table S1), suggesting the validity of nanosensors for intracellular studies. The in situ calibration curve defined by optode function, α, is also provided in Supplementary Figure S5. The recovery of Ca2+ signal to baseline in our sensor carbachol experiment suggests that reversible characteristics of the nanosensors remained intact. Also, comparable signal amplitudes were observed from Ca2+ indicator, Fluo-464 (Supplementary Figure S6), suggesting that taken together, we demonstrate the use of optode-based nanosensors as a sensing platform for monitoring and quantitative imaging of intracellular ion dynamics.


The main advantage of our sensor platform is the flexibility to tune the sensor to a given response range and selectivity for various applications by altering the sensing components. While molecular probes can improve individual aspects of indicator performance such as Kd, wavelength, and ratiometric measurements, NP-based sensors benefit from addressing these issues simultaneously. For example, sensor sensitivity can be tuned to a dynamic range of interest which can mitigate issues with indicator buffering and affinities that are either lower or higher relative to the magnitude of the Ca2+ signal - which often leads to indicator saturation and causing erroneous estimation of the magnitude of the signal65, 66. Also, the current sensing scheme utilizes a recognition agent that is separate from an optical signal transducer. As a result, a wide range of highly selective ionophores22, 31, 32 applied for potentiometric biosensing51, 67,68,69,70,71,72 can be explored for optical detection of other physiologically important ions that lack such specific probes. In addition, the current nanosensor platform permits selection of fluorophores in the visible and infrared range to permit ratiometric imaging where possible photodamaging effects induced by UV light excitation of common ratiometric indicators such as Fura-22, 13 and Indo-114, can be minimized. While the damage imposed on cells was reduced during microinjection procedure where possible, microinjection is an invasive method which may perturb the normal physiology and delay restoration of elevated Ca2+ levels. This can be improved by further sensor miniaturization through bottom-up fabrication approaches22, 73 and sensor delivery through facilitated uptake methods such as functionalizing nanosensors to cell penetrating peptides (CPPs)73,74,75 in order to minimize physical perturbation of living cells. Lastly, we find that the nanosensors are dynamically fluorescent under 2P excitation, demonstrating their potential use in vivo.

Similar to the pH dependent nature of conventional dye indicators76, 77, our nanosensor response is also subject to pH changes due to the encapsulation of pH-sensitive dyes. This is illustrated in Supplementary Figure S7, based on theoretical optode function (Eq. 3). However, the intracellular environment is governed and protected by a buffered system to maintain pH homeostasis near pH 7.278, and the effect of pH is minimized. Also, the CHIII (ETH 5350) used in this work has a pKa of >1021, which is highly protonated under standard physiological conditions, and as such, the predominant sensor response is dictated by the changes in Ca2+ concentrations. Currently, there are developments of other efforts to circumvent such pH dependency52, 79.

This work expands on the continual development of optode-based nanosensors and highlights its importance as tools for monitoring physiological analytes. The current work introduces a ratiometric method for demonstrating quantitative determination of intracellular Ca2+ dynamics using these sensors which has not been described previously. The current sensor platform combines the ease of synthesis and the flexibility to allow incorporation of different sensing components. As such, the sensor response can be tuned for specific dynamic range, selectivity, and wavelengths. In future work, detection of other physiologically important ions such as sodium, potassium, and chloride will be explored for intracellular signaling applications.



Bis(2-ethylhexyl) sebacate (DOS), calcium ionophore II (CaI II; N,N,N′,N′-Tetra[cyclohexyl]diglycolic acid diamide), chromoionophore III (CHIII; 9-(Diethylamino)-5-[(2-octyldecyl)imino]benzo[a]phenoxazine, ETH 5350), dichloromethane (DCM), ethylene glycol tetraacetic acid (EGTA), 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), magnesium chloride (MgCl2), sodium tetrakis-[3,5-bis(trifluoromethyl)phenyl]-borate (NaTFPB), and tetrahydrofuran (THF) were obtained from Sigma-Aldrich. 1-(4,5-dimethoxy-2-nitrophenyl)-1,2-diaminoethane-N,N,N′,N′-tetraacetic acid (DMNP-EDTA), octadecyl rhodamine B chloride (R18), ionomycin, calcium calibration buffer kit (C3008MP), Fluo-4 AM, and all cell culture media and supplements were obtained from Thermo Fisher. Tris-base was obtained from Fisher BioReagents. 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-550] (ammonium salt) (DSPE-mPEG550) was obtained from Avanti Polar Lipids, Inc.


Fluorescent spectra for routine nanosensor calibration in solution and selectivity experiments were measured with Spectramax M3 plate reader (Molecular Devices). Sensor reversibility test and cell imaging were performed with Zeiss LSM700 inverted confocal laser scanning microscope (Carl Zeiss). Hydrodynamic diameter and zeta-potential measurements were determined using a 90 Plus particle size analyzer (Brookhaven Instruments Corporation). Ca2+ uncaging/imaging experiments were carried out on a two-photon microscope (Bruker) equipped with two Ti:Sa lasers (SpectraPhysics) and separate beam paths/galvanometer mirrors for independent imaging and uncaging. Laser power was controlled independently for each beam path using two Pockels cells (Conoptics).

Nanosensor Fabrication

To prepare the opCaNS, the calcium-selective optode sensing components comprising CaI-II (2 μmol, 2 mg), CHIII (878 nmol, 0.5 mg), NaTFPB (1.12 μmol, 1 mg), and R18 (123 nmol, 0.09 mg) were first reconstituted separately in 300 μL THF. DOS (428 μmol, 200 μL) was then added to the optode sensing components and vortexed briefly for 30 s to form a homogeneous solution. Just prior to nanosensor fabrication, DSPE-mPEG550 (250 μg) was dried in a glass scintillation vial and rehydrated in HEPES buffer (130 mM KCl; 10 mM NaCl; 1 mM MgCl2, 10 mM HEPES; pH adjusted to 7.2 with tris base). In a typical opCaNS fabrication, 50 μL of polymer-free optode was added to 70 μL DCM, and this mixture was sonicated with the 4 mL of HEPES buffer containing DSPE-mPEG550 at 10% intensity for 1 min (Branson digital sonifier S-450D; 1/8′′ diameter tip). After sonication, the organic solvents were removed using a rotavap (Buhle) for 15 min at room temperature, and the resulting emulsion was filtered with a 100 nm syringe filter (Millipore).

Particle Size and Zeta-Potential Measurement

Hydrodynamic diameters were measured by dynamic light scattering (DLS) based on the intensity of scattered 640 nm light at a fixed angle of 90°. All particle size measurements were taken in triplicates with sample solutions diluted in 10 mM HEPES buffer, pH 7.2 adjusted to a detector count rate of 150–450 kcps. Zeta-potentials were measured in PALS mode collected over 25 cycles each 10 runs per cycle. Size and zeta-potential measurements were analyzed from three separate batches of nanosensors each from three discrete optode stock preparations.

Fluorescent Response to Calcium

Nanosensors were calibrated in solution for fluorescent response to a series of Ca2+ EGTA buffers with controlled concentrations of free calcium ion (Ca2+ calibration buffer). In order to determine the fraction protonated during measurements in Ca2+, acid and base standards were used as endpoints for fully protonated or deprotonated chromoionophore conditions, respectively.

The fluorescence intensities for CHIII (λEX: 639 nm, λEM: 670 nm) and R18 (λEX: 555 nm, λEM: 575 nm) were measured with plate reader in bottom read mode through clear-bottom 96-well plates (emission spectra of the fluorophores shown in Figure S1). The emission fluorescence intensity ratio, R, of two fluorophores (CHIII:R18) was calculated as:


Traditionally, optode data is converted to a normalized value termed α, by ratios of 670 nm/575 nm defined as:

$$\alpha =\frac{R-{R}_{D}}{{R}_{P}-{R}_{D}}$$

where, R is the fluorescence ratio measured experimentally, and R P and R D is the ratio of fully protonated and deprotonated state of the chromoionophore, respectively. The effective Ca2+ concentration at half-maximal sensor response (defined here as, EC50), for the nanosensors was calculated from Ca2+ concentrations and the corresponding values of alpha, α, was fitted according to the dose-response (Hill) equation:

$$R(\alpha )={R}_{\min }+\frac{{R}_{\max }-{R}_{\min }}{1+{10}^{(LogE{C}_{50}-LogC)p}}$$

where R(α) is intensity ratio of measured response, C, denotes Ca2+ concentration at which a ratio was measured, and R min, R max, and EC50 are parameters obtained from the fitted sigmoidal curve defined by the Hill equation.

Selectivity for Calcium over Magnesium

Nanosensor selectivity was determined by measuring the ratiometric fluorescent response (CHIII:R18) to Ca2+ and Mg2+ as described above, and fitting the calibration data to the equation comparable to the Nicolskii-Eisenman model with a fixed interfering ion51, 80. The selectivity coefficient for an optical sensor is defined as:


where, \(E{C}_{50}^{i}\) and \(E{C}_{50}^{j}\) are the constants for the interfering ion (Mg2+) and primary ion (Ca2+), respectively.

Confocal Microscope Settings

For reversibility and cell imaging experiments (see below), confocal laser scanning microscope (Zeiss LSM 700, Carl Zeiss) was used to measure fluorescence intensity ratios (CHIII:R18), as described above. Briefly, simultaneous images were acquired from dual excitation (λEX: 555 and 639 nm) and peak intensities were recorded at two emission channels (R18 λEM: 575 nm and CHII λEM: 670 nm) using diffraction grating set to split the emission at 620 nm. The 40× oil-immersion objective lens (NA = 1.3) was used with the pinhole aperture setting at 2 µm which remained constant throughout the entire experiment.

Sensor Reversibility

For reversibility experiments, nanosensors were firstly bound to the surface of a neutravidin functionalized rectangular glass flowchamber (0.1× 2 mm ID) as described by Reinhard et al.81, and imaged on the confocal microscope. Simultaneous images from both color channels were acquired every 10 s intervals with alternating solutions of 100 or 1350 nM calcium buffers exchanged every 2 min for five exchange cycles. Images were analyzed using Matlab with identical regions of interest defined on both channels.

Two-Photon Uncaging and Imaging

Calcium caging agent (DMNP, 1.05 mM) and opCaNS were loaded into 20 μL glass capillaries (Drummond) with 1 mM Ca2+, and the ends of the capillary were sealed with wax. Imaging was carried out at 875 nm excitation wavelength through a 20 ×/0.45 N.A. objective (Nikon). Images were collected via photomultiplier tube (PMT, Hamamatsu) after a 595/50 emission filter (Semrock), at a frame-rate of 6 Hz. For DMNP uncaging, the uncaging beam (730 nm) dwelled on the target for 500 ms. Images were analyzed using ImageJ to extract raw data at the site of uncaging pulse (20 × 20 pixels).

Cell Culture

HeLa cells (Sigma, 93021013) were grown in Eagle’s Minimum Essential Medium (EMEM) supplemented with 10% fetal bovine serum, 2 mM glutamine, 1% non-essential amino acids (NEAA), penicillin (50 U/ml), and streptomycin (50 μg/ml) at 37 °C in a humidified atmosphere containing 5% CO2. The cells were plated on poly-d-lysine coated 50 mm glass bottom dishes at a density of 4–8 × 104 cells. For Ca2+ imaging studies, cells were cultured in serum-free medium for 12 h, washed with PBS, and subsequently imaged using Ca2+-free Hank’s Balanced Salt Solution (HBSS; Thermo Fisher, 14175-079) with 10 mM HEPES, adjusted to pH 7.2 (Ca2+-free imaging buffer).

Delivery and in situ Cell Calibration

Nanosensors were delivered to cytosolic compartment of HeLa cells using a microinjection technique (Eppendorf InjectMen 4) with 1.0 mm/0.5 mm (OD/ID) borosilicate glass tubes pulled to a tip size 0.1–0.5 µm using P-97 micropipette puller (Sutter Instrument). For in situ cell calibration, nanosensor loaded cells were initially incubated with Ca2+-free imaging buffer containing 5 µM ionomycin for 15 min to equilibrate Ca2+ levels within the cytosolic environment to the extracellular space. Ca2+ calibration buffer with incremental levels of Ca2+ were subsequently added and fluorescence images from the confocal microscope were acquired after 10 min incubation at each calcium concentration. The values of alpha, α, and the EC50 was determined based on the same methods used to calibrate sensors in solution. There were batch-to-batch differences in sensor response and consequently, in solution and in situ cell calibrations were routinely performed to ensure the sensors maintain responsiveness to the intracellular Ca2+ levels.

Carbachol-Induced Intracellular Calcium Measurement

Single cell determination of the intracellular calcium concentration ([Ca2+]i) was carried out in opCaNS-loaded HeLa cells. Carbachol (1 mM) was used to invoke changes in [Ca2+]i by acting on IP3R on the endoplasmic reticulum (ER) membrane. Fluorescence confocal images were acquired at 1 s interval for 10 min after addition of carbachol to the 50 mm culture dish. Fluorescence intensities in the regions of interest (ROI) on each channel (575 nm and 670 nm) were first background subtracted by the intensities of nearby area that were void of cells. For individual regions of interest (ROI) in the cytosolic cellular region we subsequently used the corrected fluorescence intensities on both channels to calculate intensity ratio R (F 670 /F 575) normalized by the photobleaching profile. Changes in nanosensor fluorescence ratio were expressed as changes in Ca2+ concentration based on in situ calibrations as described above. Conversion of fluorescence ratio to Ca2+ concentration, C, involved empirically deriving from acquired fluorescence ratio using the dose-response (Hill) equation (Eq. 6) and plotted against time. Image processing and data analysis were performed using Matlab and Origin.

$$C=E{C}_{50}\sqrt[p]{\frac{R-{R}_{\min }}{{R}_{\max }-R}}$$

Data and Image Analysis

Data and image analysis was performed using Matlab and images were constructed in Origin 8 (OriginLab). Numerical values are represented as a mean ± standard deviation, and graphical representations contain error bars for the standard deviation values.

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  1. 1.

    Tsien, R. Y. Fluorescent indicators of ion concentrations. Methods Cell Biol 30, 127–156 (1989).

  2. 2.

    Grynkiewicz, G., Poenie, M. & Tsien, R. Y. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260, 3440–3450 (1985).

  3. 3.

    Demas, J. N. & DeGraff, B. A. Applications of luminescent transition platinum group metal complexes to sensor technology and molecular probes. Coordin Chem Rev 211, 317–351 (2001).

  4. 4.

    Bunzli, J. C. & Piguet, C. Taking advantage of luminescent lanthanide ions. Chem Soc Rev 34, 1048–1077 (2005).

  5. 5.

    Medintz, I. L., Uyeda, H. T., Goldman, E. R. & Mattoussi, H. Quantum dot bioconjugates for imaging, labelling and sensing. Nat Mater 4, 435–446 (2005).

  6. 6.

    Michalet, X. et al. Quantum dots for live cells, in vivo imaging, and diagnostics. Science 307, 538–544 (2005).

  7. 7.

    Looger, L. L. & Griesbeck, O. Genetically encoded neural activity indicators. Curr Opin Neurobiol 22, 18–23 (2012).

  8. 8.

    Badura, A., Sun, X. R., Giovannucci, A., Lynch, L. A. & Wang, S. S. Fast calcium sensor proteins for monitoring neural activity. Neurophotonics 1, 025008 (2014).

  9. 9.

    Maravall, M., Mainen, Z. F., Sabatini, B. L. & Svoboda, K. Estimating intracellular calcium concentrations and buffering without wavelength ratioing. Biophys J 78, 2655–2667 (2000).

  10. 10.

    Sabatini, B. L., Oertner, T. G. & Svoboda, K. The life cycle of Ca(2+) ions in dendritic spines. Neuron 33, 439–452 (2002).

  11. 11.

    Mank, M. et al. A FRET-based calcium biosensor with fast signal kinetics and high fluorescence change. Biophys J 90, 1790–1796 (2006).

  12. 12.

    Miyawaki, A. Development of probes for cellular functions using fluorescent proteins and fluorescence resonance energy transfer. Annu Rev Biochem 80, 357–373 (2011).

  13. 13.

    Nitschke, R. et al. A modified confocal laser scanning microscope allows fast ultraviolet ratio imaging of intracellular Ca2+ activity using Fura-2. Pflugers Arch 433, 653–663 (1997).

  14. 14.

    Kuba, K., Hua, S. Y. & Nohmi, M. Spatial and dynamic changes in intracellular Ca2+ measured by confocal laser-scanning microscopy in bullfrog sympathetic ganglion cells. Neurosci Res 10, 245–259 (1991).

  15. 15.

    Lakowicz, J. R. et al. Fluorescence lifetime imaging of intracellular calcium in COS cells using quin-2. Cell Calcium 15, 7–27 (1994).

  16. 16.

    Howes, P. D., Chandrawati, R. & Stevens, M. M. Bionanotechnology. Colloidal nanoparticles as advanced biological sensors. Science 346, 1247390 (2014).

  17. 17.

    Ruedas-Rama, M. J., Walters, J. D., Orte, A. & Hall, E. A. Fluorescent nanoparticles for intracellular sensing: a review. Anal Chim Acta 751, 1–23 (2012).

  18. 18.

    Wolfbeis, O. S. An overview of nanoparticles commonly used in fluorescent bioimaging. Chem Soc Rev 44, 4743–4768 (2015).

  19. 19.

    de Aberasturi, D. J. et al. Optical sensing of small ions with colloidal nanoparticles. Chem Mater 24, 738–745 (2012).

  20. 20.

    Bau, L., Tecilla, P. & Mancin, F. Sensing with fluorescent nanoparticles. Nanoscale 3, 121–133 (2011).

  21. 21.

    Mistlberger, G., Crespo, G. A. & Bakker, E. Ionophore-based optical sensors. Annu Rev Anal Chem 7, 483–512 (2014).

  22. 22.

    Wang, X. D. et al. Ultra-small, highly stable, and sensitive dual nanosensors for imaging intracellular oxygen and pH in cytosol. J Am Chem Soc 134, 17011–17014 (2012).

  23. 23.

    Magidson, V. & Khodjakov, A. Circumventing photodamage in live-cell microscopy. Methods Cell Biol 114, 545–560 (2013).

  24. 24.

    Delehanty, J. B. et al. Spatiotemporal multicolor labeling of individual cells using peptide-functionalized quantum dots and mixed delivery techniques. J Am Chem Soc 133, 10482–10489 (2011).

  25. 25.

    Yum, K., Na, S., Xiang, Y., Wang, N. & Yu, M. F. Mechanochemical delivery and dynamic tracking of fluorescent quantum dots in the cytoplasm and nucleus of living cells. Nano Lett 9, 2193–2198 (2009).

  26. 26.

    Grunwald, D., Hoekstra, A., Dange, T., Buschmann, V. & Kubitscheck, U. Direct observation of single protein molecules in aqueous solution. Chemphyschem 7, 812–815 (2006).

  27. 27.

    Courty, S., Luccardini, C., Bellaiche, Y., Cappello, G. & Dahan, M. Tracking individual kinesin motors in living cells using single quantum-dot imaging. Nano Lett 6, 1491–1495 (2006).

  28. 28.

    Clark, H. A., Hoyer, M., Philbert, M. A. & Kopelman, R. Optical nanosensors for chemical analysis inside single living cells. 1. Fabrication, characterization, and methods for intracellular delivery of PEBBLE sensors. Anal Chem 71, 4831–4836 (1999).

  29. 29.

    Monteith, G. R. & Blaustein, M. P. Heterogeneity of mitochondrial matrix free Ca2+: resolution of Ca2+ dynamics in individual mitochondria in situ. Am J Physiol 276, C1193–1204 (1999).

  30. 30.

    Ruckh, T. T., Mehta, A. A., Dubach, J. M. & Clark, H. A. Polymer-free optode nanosensors for dynamic, reversible, and ratiometric sodium imaging in the physiological range. Sci Rep 3, 3366 (2013).

  31. 31.

    Dubach, J. M., Das, S., Rosenzweig, A. & Clark, H. A. Visualizing sodium dynamics in isolated cardiomyocytes using fluorescent nanosensors. Proc Natl Acad Sci U S A 106, 16145–16150 (2009).

  32. 32.

    Tsagkatakis, L., Peper, S., Retter, R., Bell, M. & Bakker, E. Monodisperse plasticized poly(vinyl chloride) fluorescent microspheres for selective ionophore-based sensing and extraction. Anal Chem 73, 6083–6087 (2001).

  33. 33.

    Sahari, A., Ruckh, T. T., Hutchings, R. & Clark, H. A. Development of an optical nanosensor incorporating a pH-sensitive quencher dye for potassium imaging. Anal Chem 87, 10684–10687 (2015).

  34. 34.

    Brasuel, M., Kopelman, R., Miller, T. J., Tjalkens, R. & Philbert, M. A. Fluorescent nanosensors for intracellular chemical analysis: decyl methacrylate liquid polymer matrix and ion-exchange-based potassium PEBBLE sensors with real-time application to viable rat C6 glioma cells. Anal Chem 73, 2221–2228 (2001).

  35. 35.

    Ruedas-Rama, M. J. & Hall, E. A. K+-selective nanospheres: maximising response range and minimising response time. Analyst 131, 1282–1291 (2006).

  36. 36.

    Ruckh, T. T. et al. Ion-switchable quantum dot Forster resonance energy transfer rates in ratiometric potassium sensors. ACS Nano 10, 4020–4030 (2016).

  37. 37.

    Xie, X., Crespo, G. A., Zhai, J., Szilagyi, I. & Bakker, E. Potassium-selective optical microsensors based on surface modified polystyrene microspheres. Chem Commun 50, 4592–4595 (2014).

  38. 38.

    Kisiel, A., Klucinska, K., Gniadek, M., Maksymiuk, K. & Michalska, A. Optimizing calcium selective fluorimetric nanospheres. Talanta 144, 398–403 (2015).

  39. 39.

    Yang, C. Y., Qin, Y., Jiang, D. C. & Chen, H. Y. Continuous fluorescence imaging of intracellular calcium by use of ion-selective nanospheres with adjustable spectra. ACS Appl. Mater. Interfaces 8, 19892–19898 (2016).

  40. 40.

    Martin-Orozco, N. et al. Visualization of vacuolar acidification-induced transcription of genes of pathogens inside macrophages. Mol Biol Cell 17, 498–510 (2006).

  41. 41.

    Park, E. J., Brasuel, M., Behrend, C., Philbert, M. A. & Kopelman, R. Ratiometric optical PEBBLE nanosensors for real-time magnesium ion concentrations inside viable cells. Anal Chem 75, 3784–3791 (2003).

  42. 42.

    Sumner, J. P., Aylott, J. W., Monson, E. & Kopelman, R. A fluorescent PEBBLE nanosensor for intracellular free zinc. Analyst 127, 11–16 (2002).

  43. 43.

    Jagerszki, G., Grun, A., Bitter, I., Toth, K. & Gyurcsanyi, R. E. Ionophore-gold nanoparticle conjugates for Ag(+)-selective sensors with nanomolar detection limit. Chem Commun 46, 607–609 (2010).

  44. 44.

    Brasuel, M. G., Miller, T. J., Kopelman, R. & Philbert, M. A. Liquid polymer nano-PEBBLEs for Cl- analysis and biological applications. Analyst 128, 1262–1267 (2003).

  45. 45.

    Si, D., Epstein, T., Lee, Y. E. & Kopelman, R. Nanoparticle PEBBLE sensors for quantitative nanomolar imaging of intracellular free calcium ions. Anal Chem 84, 978–986 (2012).

  46. 46.

    Xie, X. & Bakker, E. Ion selective optodes: from the bulk to the nanoscale. Anal Bioanal Chem 407, 3899–3910 (2015).

  47. 47.

    Berridge, M. J., Lipp, P. & Bootman, M. D. The versatility and universality of calcium signalling. Nat Rev Mol Cell Biol 1, 11–21 (2000).

  48. 48.

    Clapham, D. E. Calcium signaling. Cell 131, 1047–1058 (2007).

  49. 49.

    Neher, E. Vesicle pools and Ca2+ microdomains: new tools for understanding their roles in neurotransmitter release. Neuron 20, 389–399 (1998).

  50. 50.

    Ross, W. N. Understanding calcium waves and sparks in central neurons. Nat Rev Neurosci 13, 157–168 (2012).

  51. 51.

    Bakker, E., Buhlmann, P. & Pretsch, E. Carrier-based ion-selective electrodes and bulk optodes. 1. General characteristics. Chem Rev 97, 3083–3132 (1997).

  52. 52.

    Xie, X., Zhai, J., Crespo, G. A. & Bakker, E. Ionophore-based ion-selective optical nanosensors operating in exhaustive sensing mode. Anal Chem 86, 8770–8775 (2014).

  53. 53.

    Dubach, J. M., Balaconis, M. K. & Clark, H. A. Fluorescent nanoparticles for the measurement of ion concentration in biological systems. J Vis Exp, (2011).

  54. 54.

    Romani, A. Regulation of magnesium homeostasis and transport in mammalian cells. Arch Biochem Biophys 458, 90–102 (2007).

  55. 55.

    Berridge, M. J. Neuronal calcium signaling. Neuron 21, 13–26 (1998).

  56. 56.

    Bezprozvanny, I., Watras, J. & Ehrlich, B. E. Bell-shaped calcium-response curves of Ins(1,4,5)P3- and calcium-gated channels from endoplasmic reticulum of cerebellum. Nature 351, 751–754 (1991).

  57. 57.

    Sanderson, M. J., Smith, I., Parker, I. & Bootman, M. D. Fluorescence microscopy. Cold Spring Harb Protoc 2014, pdb top071795, (2014).

  58. 58.

    Peng, H. S., Stolwijk, J. A., Sun, L. N., Wegener, J. & Wolfbeis, O. S. A nanogel for ratiometric fluorescent sensing of intracellular pH values. Angew Chem Int Ed 49, 4246–4249 (2010).

  59. 59.

    Candeloro, P. et al. Nanoparticle microinjection and Raman spectroscopy as tools for nanotoxicology studies. Analyst 136, 4402–4408 (2011).

  60. 60.

    McDougall, C., Stevenson, D. J., Brown, C. T., Gunn-Moore, F. & Dholakia, K. Targeted optical injection of gold nanoparticles into single mammalian cells. J Biophotonics 2, 736–743 (2009).

  61. 61.

    Feldherr, C. M., Lanford, R. E. & Akin, D. Signal-mediated nuclear transport in simian virus 40-transformed cells is regulated by large tumor antigen. Proc Natl Acad Sci U S A 89, 11002–11005 (1992).

  62. 62.

    Du, C., MacGowan, G. A., Farkas, D. L. & Koretsky, A. P. Calibration of the calcium dissociation constant of Rhod(2) in the perfused mouse heart using manganese quenching. Cell Calcium 29, 217–227 (2001).

  63. 63.

    Takahashi, A., Camacho, P., Lechleiter, J. D. & Herman, B. Measurement of intracellular calcium. Physiol Rev 79, 1089–1125 (1999).

  64. 64.

    Gee, K. R. et al. Chemical and physiological characterization of fluo-4 Ca(2+)-indicator dyes. Cell Calcium 27, 97–106 (2000).

  65. 65.

    Sinha, S. R., Wu, L. G. & Saggau, P. Presynaptic calcium dynamics and transmitter release evoked by single action potentials at mammalian central synapses. Biophys J 72, 637–651 (1997).

  66. 66.

    Regehr, W. G. & Atluri, P. P. Calcium transients in cerebellar granule cell presynaptic terminals. Biophys J 68, 2156–2170 (1995).

  67. 67.

    Carey, J. L. 3rd, Whitcomb, D. R., Chen, S., Penn, R. L. & Buhlmann, P. Potentiometric in situ monitoring of anions in the synthesis of copper and silver nanoparticles using the polyol process. ACS Nano 9, 12104–12114 (2015).

  68. 68.

    Buhlmann, P., Pretsch, E. & Bakker, E. Carrier-based ion-selective electrodes and bulk optodes. 2. Ionophores for potentiometric and optical sensors. Chem Rev 98, 1593–1688 (1998).

  69. 69.

    Chumbimuni-Torres, K. Y. et al. Potentiometric biosensing of proteins with ultrasensitive ion-selective microelectrodes and nanoparticle labels. J Am Chem Soc 128, 13676–13677 (2006).

  70. 70.

    Chumbimuni-Torres, K. Y. & Wang, J. Nanoparticle-induced potentiometric biosensing of NADH at copper ion-selective electrodes. Analyst 134, 1614–1617 (2009).

  71. 71.

    Uhlig, A., Lindner, E., Teutloff, C., Schnakenberg, U. & Hintsche, R. Miniaturized ion-selective chip electrode for sensor application. Anal Chem 69, 4032–4038 (1997).

  72. 72.

    Lindner, E. & Buck, R. P. Microfabricated potentiometric electrodes and their in vivo applications. Anal Chem 72, 336A–345A (2000).

  73. 73.

    Zamaleeva, A. I. et al. Cell-penetrating nanobiosensors for pointillistic intracellular Ca2+ -transient detection. Nano Lett 14, 2994–3001 (2014).

  74. 74.

    Schwarze, S. R., Ho, A., Vocero-Akbani, A. & Dowdy, S. F. In vivo protein transduction: delivery of a biologically active protein into the mouse. Science 285, 1569–1572 (1999).

  75. 75.

    Boeneman, K. et al. Selecting improved peptidyl motifs for cytosolic delivery of disparate protein and nanoparticle materials. ACS Nano 7, 3778–3796 (2013).

  76. 76.

    Larsson, D., Larsson, B., Lundgren, T. & Sundell, K. The effect of pH and temperature on the dissociation constant for fura-2 and their effects on [Ca(2+)](i) in enterocytes from a poikilothermic animal, Atlantic cod (Gadus morhua). Anal Biochem 273, 60–65 (1999).

  77. 77.

    Lattanzio, F. A. Jr. The effects of pH and temperature on fluorescent calcium indicators as determined with Chelex-100 and EDTA buffer systems. Biochem Biophys Res Commun 171, 102–108 (1990).

  78. 78.

    Casey, J. R., Grinstein, S. & Orlowski, J. Sensors and regulators of intracellular pH. Nat Rev Mol Cell Biol 11, 50–61 (2010).

  79. 79.

    Xie, X., Zhai, J. & Bakker, E. pH independent nano-optode sensors based on exhaustive ion-selective nanospheres. Anal Chem 86, 2853–2856 (2014).

  80. 80.

    Bakker, E. & Simon, W. Selectivity of ion-sensitive bulk optodes. Anal Chem 64, 1805–1812 (1992).

  81. 81.

    Sonnichsen, C., Reinhard, B. M., Liphardt, J. & Alivisatos, A. P. A molecular ruler based on plasmon coupling of single gold and silver nanoparticles. Nat Biotechnol 23, 741–745 (2005).

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This work was supported by grant R01 NS081641 from the National Institutes of Health (NIH) (H.A.C.) and the UCSF Program for Breakthrough Biomedical Research funded in part by the Sandler Foundation (K.E.P.).

Author information

Author notes

  1. Guoxin Rong and Eric H. Kim contributed equally to this work.


  1. Department of Bioengineering, Northeastern University, Boston, MA, 02115, United States

    • Guoxin Rong
    • , Eric H. Kim
    •  & Heather A. Clark
  2. Department of Biochemistry & Biophysics and Kavli Institute for Fundamental Neuroscience, University of California, San Francisco, San Francisco, CA, 94143, United States

    • Kira E. Poskanzer
  3. Department of Chemistry and Chemical Biology, Northeastern University, Boston, MA, 02115, United States

    • Heather A. Clark


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G.R. and E.H.K. designed and performed experiments, analyzed data, and wrote the manuscript. K.E.P. designed and performed two-photon experiments, analyzed data, and revised the manuscript. H.A.C. involved in the experimental design, data interpretation, supervision of the project, and revising the manuscript.

Competing Interests

The authors declare that they have no competing interests.

Corresponding author

Correspondence to Heather A. Clark.

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