Research ArticleCell Biology

Fluorescent Ca2+ indicators directly inhibit the Na,K-ATPase and disrupt cellular functions

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Sci. Signal.  30 Jan 2018:
Vol. 11, Issue 515, eaal2039
DOI: 10.1126/scisignal.aal2039

The dark side of tracking Ca2+ in cells

Ca2+ signaling events in many different cell types are tracked with fluorescent Ca2+ indicators, such as Fluo-4, Rhod-2, and Fura-2, and can be inhibited with the Ca2+ chelator BAPTA. Smith et al. found that these commonly used reagents inhibited the Na,K-ATPase, a membrane protein that exchanges intracellular Na+ for extracellular K+ and thus helps set the resting membrane potential and regulate cellular volume. This inhibition, which was accompanied by reduced cell viability, decreased glucose uptake, and cell swelling, occurred in multiple cell types, including neurons, astrocytes, and cardiomyocytes, and in mice when Rhod-2 or Fluo-4 was microdialyzed into the CNS. However, a genetically encoded Ca2+ indicator did not inhibit the Na,K-ATPase. These results suggest that it may be necessary to use these reagents with caution or rely on genetically encoded indicators to prevent cellular toxicity from affecting experimental outcomes.


Fluorescent Ca2+ indicators have been essential for the analysis of Ca2+ signaling events in various cell types. We showed that chemical Ca2+ indicators, but not a genetically encoded Ca2+ indicator, potently suppressed the activity of Na+- and K+-dependent adenosine triphosphatase (Na,K-ATPase), independently of their Ca2+ chelating activity. Loading of commonly used Ca2+ indicators, including Fluo-4 acetoxymethyl (AM), Rhod-2 AM, and Fura-2 AM, and of the Ca2+ chelator BAPTA AM into cultured mouse or human neurons, astrocytes, cardiomyocytes, or kidney proximal tubule epithelial cells suppressed Na,K-ATPase activity by 30 to 80%. Ca2+ indicators also suppressed the agonist-induced activation of the Na,K-ATPase, altered metabolic status, and caused a dose-dependent loss of cell viability. Loading of Ca2+ indicators into mice, which is carried out for two-photon imaging, markedly altered brain extracellular concentrations of K+ and ATP. These results suggest that a critical review of data obtained with chemical Ca2+ indicators may be necessary.


The use of fluorescent Ca2+ indicators has provided valuable insight into the dynamics of intracellular Ca2+ signaling (1). Fluorescent Ca2+ indicators were originally developed to take advantage of the fluorescence spectrum shift of the Ca2+ selective compound 1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) that occurs upon Ca2+ binding (2). Multiple Ca2+ indicators have been introduced since Roger Tsien’s original publication in 1979, and more than 25,700 publications using Ca2+ indicators are listed in PubMed (fig. S1, A to D). However, despite widespread use, the potential possible side effects of Ca2+ indicator dyes have not been systematically investigated. Various genetically encoded Ca2+ indicators (GECIs) have been developed, including the genetically encoded Ca2+ indicator abbreviated as “GCaMP” [a fusion product of enhanced green fluorescent protein (GFP), calmodulin, and the M13 sequence from myosin light chain kinase]. After several generations of modifications to increase fluorescence output and optimize the signal-to-noise ratio, GECIs and GCaMPs have become widely used tools in multiple domains of research (3, 4). However, Ca2+ indicator dyes are also still widely used (fig. S1A), and there is a lack of comparative analysis of the effect of chemical and genetically encoded Ca2+ indicators on cellular functions. However, a few reports have shown that Ca2+ indicators can alter cellular functions, such as Ca2+-induced Ca2+ release from endoplasmic reticulum in cerebellar granule neurons and human embryonic kidney (HEK) 293 cells (5), field excitatory postsynaptic potentials in the brain hippocampal slice (6), the activity of K+ channels expressed in HEK293 cells (7), and K+ currents in brain slices (8).

Here, we analyzed the effects of three Ca2+ indicators (Rhod-2, Fluo-4, and Fura-2), the Ca2+ chelator BAPTA, and GCaMP3 on basic cellular functions. As one of the several end points, we assessed the effects of Ca2+ indicators on the activity of the Na+- and K+-dependent adenosine triphosphatase (Na,K-ATPase), a membrane-bound protein, which consumes ~40% of cellular energy supply (9, 10) and is essential for many aspects of cellular membrane functions (11, 12). We found that all three indicators and BAPTA inhibited Na,K-ATPase activity in both intact primary cells and membrane preparations of mouse and human brain astrocytes, mouse heart cardiomyocytes, and human neurons and kidney proximal tubule epithelial cells. Further analysis focused on the characterization of the effects of Ca2+ indicators on cellular functions in the central nervous system (CNS) demonstrated that Ca2+ indicators and BAPTA loading also significantly altered neuronal and astrocytic energy metabolism, as detected by uptake of [3H]-2-deoxyglucose ([3H]-2-DG) and lactate release. Loading of Rhod-2 AM or Fluo-4 AM in the cerebral cortex of awake freely behaving mice induced a marked increase in extracellular K+ and adenosine triphosphate (ATP). In contrast, the genetically encoded Ca2+ indicator GCaMP3 had minimal adverse effects. Together, these observations raise concerns regarding the use of Ca2+ indicator dyes and suggest that genetically encoded Ca2+ indicators interfere to a lesser extent with basic neuronal-glial cell signaling interactions. Our observations, which are focused on analysis of intact brain and cultured neurons and astrocytes, suggest a reassessment of previous conclusions based on loading of chemical Ca2+ indicators due to their cytotoxicity and effect on basic cellular parameters.


Ca2+ indicators suppress Na,K-ATPase–mediated ion uptake independently of cell type

We first assessed the degree to which two of the most commonly used single-wavelength Ca2+ indicators, Fluo-4 and Rhod-2, or of the Ca2+ chelator BAPTA acetoxymethyl (AM) ester (fig. S1A) affected ouabain-sensitive 86RB+ uptake when loaded into the cytosol of cultured mouse astrocytes. Fluo-4 AM, Rhod-2 AM, and BAPTA AM were added to the medium at commonly used concentrations (Fig. 1, A and B, and fig. S1B). Ca2+ indicator dyes are often chemically derivatized with an AM, which is Ca2+-insensitive and nonfluorescent, but can readily permeate cellular membranes. The AM ester is then cleaved by endogenous cytosolic esterases, which leads to an accumulation of a membrane-impermeable Ca2+ indicator (Fig. 1A) (13). Quantification of the intracellular concentrations of Fluo-4, Rhod-2, and BAPTA showed that all were present in the cytosol at considerably higher concentrations than those added to the medium (~100-fold) (Fig. 1B). Similarly, the cytosolic concentration of the Ca2+ indicator Quin2 was several hundred times higher than the loading concentration (14).

Fig. 1 Suppression of ouabain-sensitive 86Rb+ uptake by Ca2+ indicators in cultured mouse astrocytes.

(A) Acetoxymethyl (AM) ester derivatives of Ca2+ indicators (Ca2+-insensitive and nonfluorescent), which cross cell membranes noninvasively, are cleaved by esterases in the cells. The ionized active form of Ca2+ indicators (which is Ca2+-sensitive) remains trapped within the cells. (B) Intracellular Fluo-4, Rhod-2, and BAPTA concentrations after the loading of various concentrations of AM indicators (n = 4 wells for each treatment). (C) Ouabain (1 mM)–sensitive 86Rb+ uptake without or with preincubation of Fluo-4 AM (1 to 9 μM), Rhod-2 AM (1 to 9 μM), or BAPTA AM (5 to 40 μM). The data were expressed as percent change from control and plotted against intracellular concentrations (n = 4 to 6 wells for each treatment). **P < 0.01, ****P < 0.0001 compared to vehicle control (0.2% DMSO). Means ± SEM are shown.

The effect of the Ca2+ indicators on the Na,K-ATPase was assessed by quantifying ouabain-sensitive uptake of a commonly used K+ analog, rubidium-86 (86Rb+), in cultured mouse astrocytes (15, 16). Both Ca2+ indicators and BAPTA AM, but not the solvent dimethyl sulfoxide (DMSO), effectively suppressed the ouabain-sensitive fraction of 86Rb+ uptake by astrocytes in a dose-dependent manner (Fig. 1C and fig. S2).

We next tested whether the suppression of Na,K-ATPase activity caused by Ca2+ indicators could be observed in different cell types, including cultures of human astrocytes, mouse cardiomyocytes, human renal proximal tubule epithelial cells, and mouse neurons (Fig. 2A and movie S1). We also expanded the analysis to other BAPTA-based Ca2+ indicators because of the similarities in their structures (fig. S1D). Fura-2 is the most widely used ratiometric dye (fig. S1A) (17). All cell types analyzed exhibited ouabain-sensitive 86Rb+ uptake (Fig. 2, B to F). Pretreatment with Fluo-4 AM, Rhod-2 AM, Fura-2 AM, or BAPTA AM significantly reduced 86Rb+ uptake in mouse astrocytes (Fig. 2B). 86Rb+ uptake was not further suppressed upon co-application of ouabain (fig. S3), suggesting that Ca2+ indicators do not affect ouabain-insensitive 86Rb+ uptake. Ca2+ indicators and BAPTA AM also inhibited 86Rb+ uptake in human astrocytes (Fig. 2C), mouse cardiomyocytes (Fig. 2D), human proximal tubular epithelial cells (Fig. 2E), mouse neurons (Fig. 2F), and rat astrocytes (fig. S4A). Cultured human astrocytes and mouse neurons were the most sensitive to loading with Ca2+ indicators and displayed the greatest decrease in 86Rb+ uptake (Fig. 2, C and F).

Fig. 2 Ca2+ indicators inhibited Na,K-ATPase in four different types of cultured cells.

(A) Schematic diagram outlining the measurement of 86Rb+ uptake by cells and Na,K-ATPase activity in membrane preparation. (B to K) Ca2+ indicators (Fluo-4 AM, Rhod-2 AM, and Fura-2 AM) or BAPTA AM was preloaded for 30 min in cultured mouse astrocytes (B and G; n = 8 to 27 wells for each treatment), human astrocytes (C and H; n = 5 to 12 wells for each treatment), mouse cardiomyocytes (D and I; n = 6 to 12 wells for each treatment), human proximal tubule epithelial cells (E and J; n = 5 to 12 wells for each treatment), or mouse neurons (F and K; n = 5 wells for each treatment), and then 86Rb+ uptake (B to F) or Na,K-ATPase activity (G to K) was measured. Ouabain (1 mM) was used to identify the portion of 86Rb+ uptake or ATP hydrolysis mediated by the Na,K-ATPase. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 compared to vehicle control (0.2% DMSO). Means ± SEM are shown.

Ca2+ indicators decrease ATP hydrolysis mediated by Na,K-ATPase

Because the Na,K-ATPase is sensitive to changes in Ca2+ concentrations (11, 18), we next tested whether Ca2+ indicators and BAPTA AM reduced the activity of the Na,K-ATPase by altering intracellular Ca2+ concentrations. To control the Ca2+ concentration, we used a membrane preparation to eliminate cellular control of cytosolic Ca2+. Quantification of ouabain-sensitive ATP hydrolysis was used as an indirect measurement of the Na,K-ATPase activity (1921). Preloading of cultured mouse astrocytes with Ca2+ indicators or BAPTA AM significantly inhibited ouabain-sensitive ATP hydrolysis (Fig. 2G). Similarly, human astrocytes (Fig. 2H), mouse cardiomyocytes (Fig. 2I), human proximal tubule epithelial cells (Fig. 2J), mouse neurons (Fig. 2K), and rat astrocytes (fig. S4B) also exhibited suppression of ouabain-sensitive ATP hydrolysis. Thus, observations based on membrane preparations were consistent with the findings in intact cells. However, Fluo-4 AM did not significantly affect kidney cells, cardiomyocytes, and neurons at 2 μM, which is within the recommended usage range (<5 μM).

Ca2+ indicators directly suppress Na,K-ATPase activity independently of Ca2+ chelation

We next sought to evaluate whether the effect of Ca2+ indicators was attributed to AM loading by using a cell-impermeant form of BAPTA or Rhod-2 (Fig. 3A). Addition of BAPTA (100 μM) or Rhod-2 (10 μM) inhibited ouabain-sensitive ATP hydrolysis in membrane preparations derived from cultured mouse or human astrocytes, with a potency that was directly comparable to intracellular loading with BAPTA AM or Rhod-2 AM (Fig. 3, B and C). This result suggests that suppression of the Na,K-ATPase is a direct effect of the Ca2+ indicators and not a consequence of the preloading of the AM version of the Ca2+ indicators. High levels of Ca2+ can inhibit the activity of Na,K-ATPase (11). However, it is unlikely that Ca2+ plays a role in Ca2+ indicator–mediated suppression of the Na,K-ATPase activity, because ouabain-sensitive ATP hydrolysis was inhibited by BAPTA, even in the absence of Ca2+ (Fig. 3D). Moreover, 5,5′-dibromo BAPTA, which has a ~15-fold lower affinity for Ca2+ than its parent compound BAPTA [dissociation constant (Kd) = 1.6 μM compared to 110 nM], also potently suppressed the Na,K-ATPase (Fig. 3D).

Fig. 3 Ca2+ indicators directly inhibited Na,K-ATPase and diminished the GPCR-mediated enhancement of K+ uptake.

(A) Schematic diagram outlining the ATPase assay in membrane preparations using salt and AM forms of BAPTA or Rhod-2. (B) Effect of BAPTA tetrapotassium salt (100 μM) to membrane preparations on Na,K-ATPase activity, compared to the preincubation of BAPTA AM (20 μM) in cultured mouse astrocytes (n = 8 to 14 wells for each treatment). (C) Effect of Rhod-2 tripotassium salt (10 μM) on ouabain-sensitive ATP hydrolysis in membrane preparation compared to the preincubation of Rhod-2 AM (2 μM) in cultured human astrocyte (n = 8 wells for each treatment). (D) Na,K-ATPase activity measurement in the absence or presence of BAPTA tetrapotassium salt (100 μM) or low–Ca2+ affinity 5,5′-dibromo BAPTA tetrapotassium salt (100 μM) in the zero Ca2+ assay buffer in cultured mouse astrocytes (n = 8 to 14 wells for each treatment). *P < 0.05, **P < 0.01, ****P < 0.0001 compared to vehicle control (B and D, 0.1% DMSO; C, 0.04% DMSO). Means ± SEM are shown.

To further test whether Ca2+ indicators directly affected Na,K-ATPase activity, we next evaluated Na,K-ATPase current in hippocampal pyramidal neurons in the presence and absence of Ca2+ indicators (Fig. 4A). Using 0.5 mM strophanthidin, an inhibitor of Na,K-ATPase pump activity (22), we could directly measure Na,K-ATPase current (Fig. 4, B and C). Fluo-4(5 μM), Rhod-2 (5 μM), Fura-2 (5 μM), and BAPTA (20 μM) all significantly inhibited Na,K-ATPase current in hippocampal pyramidal neurons (Fig. 4, D to H). Together, these observations suggest that inhibition of the Na,K-ATPase is not attributable to the Ca2+ chelating effects of Ca2+ indicators but rather to a direct inhibitory effect of the Ca2+ indicators on the Na,K-ATPase or its associated proteins.

Fig. 4 Ca2+ indicators inhibited Na,K-ATPase current in pyramidal neurons of hippocampal CA1 region.

(A) Schematic diagram outlining the measurement of Na,K-ATPase current in a pyramidal neuron. (B) Na,K-ATPase current was measured by the application of 0.5 mM strophanthidin for 2 min. (C to G) Baseline recording of the holding current using vehicle without strophanthidin (B) (88). The membrane-impermeable Ca2+ indicators Fluo-4 (D), Rhod-2 (E), and Fura-2 (F) or the membrane-impermeable BAPTA (G) was included in the recording pipette to measure their effect on Na,K-ATPase current. Ten minutes after breaking into the neuron, Ca2+ indicator loading was detectable [inset in (D); scale bar, 10 μm]. (H) Na,K-ATPase current density was plotted and analyzed using one-way analysis of variance (ANOVA) (n = 4 to 5 mice; **P < 0.01, ***P < 0.001, ****P < 0.0001). Means ± SEM are shown.

Ca2+ indicator loading leads to cell swelling and dose-dependent loss of cell viability

Blockade of the Na,K-ATPase by ouabain leads to death of cultured rat astrocytes (23). Because Ca2+ indicators inhibited the Na,K-ATPase, we next asked whether Ca2+ indicators also compromise the viability of astrocytes. Cell viability was quantified by cell counting and by leakage of intracellular constituents, using lactate dehydrogenase (LDH) as a proxy. In agreement with previous findings (23), the addition of ouabain for 24 hours decreased the number of viable cells in a dose-dependent manner (Fig. 5A). We next performed this same assessment of cell viability 24 hours after loading of the Ca2+ indicators or BAPTA AM and washing to determine whether, similar to ouabain, there was a latent cytotoxic effect of Ca2+ indicators or BAPTA AM loading. Viable cell numbers were decreased by loading of Ca2+ indicators and BAPTA AM for 30 min at higher concentrations (Fig. 5A) or for a longer loading period (2 hours), which is used in many studies (Fig. 5B) (2428). This reduction on cell viability was not triggered by application of DMSO, a vehicle used for Ca2+ indicators. We also examined cell viability using two alternative markers of cell injury: leakage of the intracellular constituent LDH and cell volume changes (shrinkage or swelling) (29). Loading of Ca2+ indicators and BAPTA AM for 2 hours induced significant LDH release (Fig. 5C). In particular, Rhod-2 AM and BAPTA AM were poorly tolerated, and their adverse effect on cell viability was comparable to that of ouabain (Fig. 5, A to D). In neuronal cultures, loading of Ca2+ indicators or BAPTA also had marked cytotoxic effects detected as a significant increase in LDH release (fig. S5A).

Fig. 5 Ca2+ indicators induced swelling and cell death of mouse astrocytes.

(A) Assessment of viable cell numbers by counting. Cultured mouse astrocytes were treated with increasing concentrations of ouabain (0 to 5 mM), DMSO (0 to 0.2%), Ca2+ indicators (1 to 10 μM in 0.2% DMSO), and BAPTA AM (5 to 40 μM in 0.2% DMSO) for 30 min, and cell viability was examined at 24 hours (n = 8 wells for each treatment). *P < 0.05, **P < 0.01 compared to no-treatment control. (B) Same as in (A), except the loading time was extended to 2 hours (n = 5 to 6 wells for each treatment). *P < 0.05, ***P < 0.001 compared to no-treatment control. (C) LDH release was examined immediately after 2 hours of treatment with the indicators (n = 6 wells for each treatment). *P < 0.05, **P < 0.001, ****P < 0.0001 compared to no-treatment control. (D) Relative increases in the diameter of viable cultured mouse astrocytes at 24 hours induced by treatment of various concentrations of ouabain, DMSO, Ca2+ indicators, and BAPTA AM for 2 hours (n = 4 wells for each treatment). (E) Effect of loading of BAPTA AM (20 μM) on the ability of TFLLR-NH2 (30 μM) and FMRF (15 μM) to enhance 86Rb+ uptake in cultured wild-type or MrgA1+/− mouse astrocytes, respectively (n = 4 to 38 wells for each treatment). n.s. (not significant), *P < 0.05, **P < 0.01 compared to (−) BAPTA AM of control group (0.1% DMSO). (F) Effect of preloading Rhod-2 AM (4.5 μM) and BAPTA AM (20 μM) on the ability of ATP (100 μM) to increase 86Rb+ uptake in cultured mouse astrocytes (n = 4 to 38 wells for each treatment). n.s., *P < 0.05 compared to (−) ATP of each group, #P < 0.05 compared to (+) ATP of control group (0.1% DMSO). Means ± SEM are shown.

The Na,K-ATPase, which exports three Na+ ions in exchange for import of two K+ ions, is an important determinant of cellular volume. Thus, as an independent assay of inhibition of the Na,K-ATPase, we also assessed cell volume at 24 hours after Ca2+ or BAPTA application. Ouabain, as well as the Ca2+ indicators and BAPTA AM, but not DMSO, increased cell volume in a dose-dependent manner (Fig. 5D and fig. S5B).

Ca2+ indicators impair agonist-induced activation of Na,K-ATPase

We have previously found that activation of Gq-linked G protein (guanine nucleotide–binding protein)–coupled receptors (GPCRs), including the protease-activated receptor-1 (PAR-1)–selective agonist Thr-Phe-Leu-Arg-NH2 (TFLLR-NH2), can stimulate Na,K-ATPase activity in astrocytes through the production of inositol 1,4,5-trisphosphate (IP3) (30). Here, we found that preloading of BAPTA AM suppressed the ability of TFLLR-NH2 to increase 86Rb+ uptake (Fig. 5E). We also assessed the effect of Ca2+ indicators on the Gq-coupled receptor MrgA1 using astrocyte cultures from transgenic mice expressing MrgA1 under the inducible control of the human GFAP (which encodes glial fibrillary acidic protein) promoter (31). As expected, the MrgA1 agonist peptide Phe-Met-Arg-Phe amide (FMRF) increased 86Rb+ uptake, and similar to the PAR-1 agonist, BAPTA AM loading also inhibited FMRF-induced 86Rb+ uptake (Fig. 5E). Finally, we assessed the effect of Ca2+ indicators on purinergic GPCRs. In cultured mouse astrocytes, the purinergic agonist ATP enhanced ouabain-sensitive 86Rb+ uptake (Fig. 5F), which was suppressed by the application of Rhod-2 AM or BAPTA AM (Fig. 5F). These results extended the finding that Ca2+ indicators inhibit the Na,K-ATPase activity by demonstrating that stimulation of Na,K-ATPase activity by various GPCRs was also disrupted after loading of Ca2+ indicators and BAPTA AM.

Ca2+ indicators alter energy metabolism and Na,K-ATPase activity in vitro and in vivo

To assess whether Ca2+ indicators also affect cellular energy metabolism, we next quantified [3H]-2-DG uptake in cultured mouse astrocytes and neurons (Fig. 6A). Fluo-4 AM, Fura-2 AM, and BAPTA AM all significantly reduced [3H]-2-DG uptake in cultured mouse astrocytes, whereas Fluo-4 AM and BAPTA AM reduced [3H]-2-DG uptake in cultured mouse neurons. Unexpectedly, however, Rhod-2 AM increased both astrocytic and neuronal [3H]-2-DG uptake (Fig. 6B). Rhod-2 is a derivative of Rhodamine 123 that inhibits the mitochondrial electron transport chain, thereby slowing respiration (Fig. 6B) (32, 33). Thus, suppression of mitochondrial ATP synthesis could enhance glycolysis to fulfill cellular energy demands. In agreement with this hypothesis, Rhod-2 AM induced an increase in [3H]-2-DG uptake that was inhibited by iodoacetate, an inhibitor of glycolysis (Fig. 6C). Iodoacetate alone decreased [3H]-2-DG uptake, whereas sodium cyanide (NaCN), an inhibitor of mitochondrial oxidative metabolism (cytochrome c oxidase), enhanced [3H]-2-DG uptake in both astrocytes and neurons (Fig. 6C). As an alternative strategy to test whether Rhod-2 AM increased glycolysis, we measured lactate production in the same cultures. As expected, both the inhibition of oxidative energy metabolism by NaCN and the inhibition of glycolysis by iodoacetate potently limited lactate production in mouse cultured astrocytes (Fig. 6D). Rhod-2 AM loading was also associated with a significant increase in lactate production that was inhibited by the co-application of iodoacetate (Fig. 6D). Moreover, Rhod-2 AM loading caused morphological alterations in mitochondria (Fig. 6E), a finding that is supported by a previous study (34). These observations suggest that Rhod-2 AM loading is linked to an increase in glycolysis. In contrast, Fluo-4 AM, Fura-2 AM, and BAPTA AM all reduced glucose utilization, consistent with the suppression of the Na,K-ATPase activity.

Fig. 6 Ca2+ indicators altered glucose uptake and lactate release of cultured mouse astrocytes and neurons.

(A) Effect of loading Fluo-4 AM, Fura-2 AM, and BAPTA AM on [3H]-2-deoxyglucose ([3H]-2-DG) uptake by mouse astrocyte cultures and neuronal cultures (n = 4 to 12 wells for each treatment). (B) Effect of Rhod-2 AM on [3H]-2-DG uptake by mouse astrocyte and neuronal cultures (n = 4 to 8 wells for each treatment). *P < 0.05, **P < 0.01, ***P < 0.001 compared to vehicle control (0.2% DMSO; A and B). (C) Effect of mitochondrial oxidative metabolism inhibition with sodium cyanide (NaCN; 100 μM; n = 8 wells for each treatment) or glycolysis inhibition with iodoacetate (IAA; 300 μM; n = 8 to 16 wells for each treatment) on [3H]-2-DG uptake by cultured mouse astrocytes and neurons in the presence or absence of Rhod-2 AM (2 μM). (D) Effect of inhibition of mitochondrial oxidative metabolism by Rhod-2 AM (2 μM) and NaCN (100 μM) or IAA (300 μM) on lactate release by cultured mouse astrocytes (n = 4 to 8 wells for each treatment). *P < 0.05, **P < 0.01, ****P < 0.001 compared to vehicle control (0.04% DMSO); #P < 0.01, ####P < 0.01 compared to (+) Rhod-2 AM of control group (C and D). (E) Images of astrocytes loaded with MitoTracker (20 nM) (top) and Rhod-2 AM (2 and 10 μM) (bottom). Scale bar, 20 μm. White arrows correspond to mitochondrial morphological changes. Means ± SEM are shown.

Because our in vitro observations raised concern that Ca2+ indicator loading adversely affects cellular functions, we also used microdialysis in freely moving awake mice to evaluate the effect of Rhod-2 AM and Fluo-4 AM in the intact CNS (Fig. 7A). These experiments are relevant for in vivo two-photon imaging experiments, which frequently use Ca2+ indicators to analyze cortical Ca2+ dynamics in response to stimulation (3537). Rhod-2 AM (5 μM) was delivered by reverse microdialysis, and vibratome sections were prepared immediately after the experiments to confirm conversion of Rhod-2 AM to Rhod-2 based on its fluorescence signal (Fig. 7B). The diameter of tissue with Rhod-2 uptake was ~400 μm or comparable to the spatial extent of tissue loading after bolus injection (35). Microdialysates collected before, during, and after loading of Rhod-2 AM, Fluo-4 AM, or DMSO were used for measurement of extracellular K+ concentration, as an indicator of Na,K-ATPase activity (Fig. 7C). Compared to DMSO control, Rhod-2 and Fluo-4 AM treatment induced a marked increase in K+ concentration that lasted for 45 min for both Rhod-2 AM and Fluo-4 AM before gradually returning to baseline levels (Fig. 7C). The extracellular concentration of ATP also increased at 1 to 2 hours after Rhod-2 or Fluo-4 AM application (Fig. 7D). To examine whether this increase was due to leakage of ATP from dying cells, we evaluated the glycerol concentrations, a marker of cell membrane damage (38). Application of Rhod-2 and Fluo-4 AM did not affect glycerol concentration (Fig. 7D), suggesting that the ATP increase observed after Rhod-2 AM infusion is not a result of cellular damage.

Fig. 7 Ca2+ indicator disrupts astrocyte K+ uptake and tissue environments in vivo.

(A) Schematic diagram outlining the application of Ca2+ indicator and sample collection using microdialysis in freely moving mice. After overnight probe equilibration, samples were collected every 15 min before, during, and after infusion of artificial cerebrospinal fluid (aCSF) containing Fluo-4 AM, Rhod-2 AM, or vehicle. (B) Image of acutely prepared coronal brain section confirming the delivery of Rhod-2 AM into tissue. Dashed circle indicates position of microdialysis probe. Scale bar, 500 μm. (C) Comparison of extracellular K+ measurements with Fluo-4 AM, Rhod-2 AM, or control application in vivo (n = 6 to 8 mice for each treatment; *P < 0.05, **P < 0.01 compared to control group). BL, baseline. (D) Comparison of ATP and glycerol measurements with Fluo-4 AM, Rhod-2 AM, or control application in vivo (n = 6 to 8 mice for each treatment; n.s., *P < 0.05, **P < 0.01, ***P < 0.001 compared to control group). Means ± SEM are shown.

GCaMP expression in astrocytes does not alter Na,K-ATPase–mediated K+ uptake

Genetically encoded Ca2+ indicators have been developed as alternative tools for the analysis of Ca2+ signaling (39). To assess whether genetically encoded Ca2+ indicators also affect basic cellular functions, we prepared astrocytes from double-transgenic GFAP-Cre and floxed GCaMP3 mice, in which GCaMP3 is expressed under the control of the GFAP promoter (GFAP-GCaMP3) (Fig. 8A). Astrocytes expressing GCaMP3 (AdV-GCaMP3) or GFP (AdV-GFP control) showed similar amounts of Na,K-ATPase–mediated 86Rb+ uptake (Fig. 8B). Conversely, Rhod-2 AM loading (4 μM) consistently decreased Na,K-ATPase–mediated 86Rb+ uptake in all cell types (fig. S6, A and B). However, it is possible that the cells compensated for GCaMP3 expression because the transgene is expressed during development. To assess this possibility, we next prepared cultured astrocytes from GCaMP3fl transgenic mice and introduced GCaMP3 by transfection with an adenovirus bearing Cre recombinase under the control of the CMV promoter (AdV-GCaMP3). At 6 days after transduction, GCaMP3 fluorescence and Ca2+ responses to pharmacological stimulation with ATP were confirmed (fig. S6, C and D, and movies S2 to S5). Analysis of Ca2+ transients in response to application of 5 μM ATP revealed that AdV-GCaMP3 astrocytes exhibited long-lasting patterns of rhythmic oscillations. GFAP-GCaMP3 astrocytes also exhibited both shorter (~60 s) and longer (~120 s) patterns of oscillation, whereas Rhod-2 AM (2.25 and 4.5 μM)–loaded cells showed small, fluctuated Ca2+ signals but no oscillations (fig. S6, C and D). For this set of experiments, we used a lower concentration of ATP (5 μM). GCaMP3 showed a greater response than Rhod-2, suggesting that GCaMP3 could detect intracellular Ca2+ changes triggered by low doses of GPCR agonists. Although GFAP-GCaMP3 expression did not affect ouabain-sensitive 86Rb+ uptake, total 86Rb+ uptake was slightly inhibited in GFAP-GCaMP3–expressing astrocytes (fig. S6E), suggesting that ouabain-insensitive K+ uptake through other ion exchangers might be altered. In contrast, in transfected astrocytes, AdV-GCaMP3 expression did not affect either ouabain-sensitive or ouabain-insensitive 86Rb+ uptake (Fig. 8B and fig. S6E). As expected, Rhod-2 AM loading suppressed ouabain-sensitive 86Rb+ uptake in astrocytes expressing GFAP-GCaMP3, AdV-GCaMP3, or AdV-GFP (Fig. 8B). Consistent with previous findings, few spontaneous Ca2+ events at similar frequencies were recorded with Rhod-2 and Fluo-4 AM (40). Imaging of baseline Ca2+ activity in astrocytes expressing GCaMP3 revealed frequent spontaneous Ca2+ spikes (Fig. 8C). However, after loading GCaMP3-expressing astrocytes with Rhod-2 AM, we observed a low number of Ca2+ events for both GCaMP3 and Rhod-2 (Fig. 8C). Ca2+ binding to Rhod-2 or Fluo-4 results in an ~100-fold change in peak emission intensity, whereas GCaMP3 shows ~12-fold change in peak emission intensity upon Ca2+ binding (13, 41). To further test whether Rhod-2 reduced the number of spontaneous Ca2+ events in Adv-GCaMP3–expressing cells by mechanisms other than Ca2+ chelation, we loaded mouse astrocytes expressing Adv-GCaMP3 with 5,5′-dibromo BAPTA at a low concentration (1.6 mM). 5,5′-Dibromo BAPTA has a high Kd for Ca2+ binding (1.6 μM), thus rendering Ca2+ chelation during physiological conditions negligible. We found that 5,5′-dibromo BAPTA significantly reduced spontaneous Ca2+ events compared to controls (Fig. 8D). This observation showed that a Ca2+ buffer with negligible ability to bind Ca2+ suppressed Ca2+ signaling events in mouse astrocytes, thus suggesting that Rhod-2 AM and Fluo-4 AM may also reduce spontaneous Ca2+ signaling partially by mechanisms other than Ca2+ chelation (Fig. 8, C and D, and fig. S6, F and G).

Fig. 8 Expression of GCaMP3 does not affect K+ uptake of mouse astrocytes.

(A) Representative images of astrocytes loaded with Rhod-2 AM (4.5 μM) or prepared from GFAP-Cre:GCaMP3fl transgenic mice (GFAP-GCaMP3). Scale bar, 100 μm. (B) Ouabain-sensitive 86Rb+ uptake in mouse astrocytes expressing GFAP-GCaMP3 or in astrocytes from GCaMP3fl transgenic mice induced to express GCaMP3 by Cre recombinase adenovirus in the presence or absence of Rhod-2 AM (4 μM). n = 6 to 23 wells for each treatment; *P < 0.05, **P < 0.01, ****P < 0.0001 compared to (−) Rhod-2 AM of control group (0.1% DMSO). (C) A number of spontaneous Ca2+ events in cultured mouse astrocytes were detected by Fluo-4 AM (2 and 4 μM), Rhod-2 AM (2.25 and 4.5 μM), or GCaMP3. In groups combining Ca2+ imaging of GCaMP3 with Rhod-2 AM, the number of Ca2+ events detected with Rhod-2 (left, orange channel) and GCaMP3 (right, green channel) was counted. n = 4 to 6 wells for each treatment; ***P < 0.001 compared to GFAP-GCaMP3 control group (0.1% DMSO). (D) Histogram comparing the number of spontaneous Ca2+ events in Adv-GCaMP3–expressing mouse astrocytes in the presence and absence of 5,5′-dibromo BAPTA [P = 0.0238, unpaired t test (Mann-Whitney test); n = 4 to 8 wells for each treatment]. Means ± SEM are shown.


Commercially available Ca2+ indicators have revolutionized our understanding of intracellular Ca2+ signaling over the past three decades (13). Because all the Ca2+ indicators are derivatized from the Ca2+ chelator BAPTA (fig. S1), it is generally acknowledged that they, to some extent, buffer intracellular Ca2+ and potentially interfere with basic cellular processes (13, 42). However, it was surprising that loading cultured cells with Ca2+ indicators suppressed the activity of the Na,K-ATPase (Figs. 2 and 3 and fig. S4). Fura-2, Fluo-4, Rhod-2, or BAPTA AM suppressed the Na,K-ATPase activity in five different types of cultured primary cells harvested from brain, heart, and kidney from either mouse, rat, or human (Fig. 2 and fig. S4). Loading of the same Ca2+ indicators or BAPTA AM in the frequently used higher concentrations also resulted in loss of cell viability (Fig. 5 and fig. S1B). Although most of our study was performed on cells from the CNS, the consistency in observations across preparations of different cell types and species suggests that the adverse effects of Ca2+ indicators are a general phenomenon. In vivo, we found that the Ca2+ indicators caused a transient but marked increase in extracellular K+ (from 4 mM up to 10 mM) after cortical loading of Fluo-4 AM or Rhod-2 AM, which was also accompanied by a significant increase in extracellular ATP (Fig. 7). Extracellular K+ concentration determines membrane potential and is tightly maintained within a range of 3 to 4 mM (43). Disruption of K+ homeostasis is likely to affect neuronal excitability because even minor increases in extracellular K+ concentration will increase neuronal excitability by narrowing the gap between the resting membrane potential and the threshold for activation of voltage-gated Na+ channels (30, 44, 45). Thus, Ca2+ indicators may alter neuronal activity by inhibiting K+ uptake. Moreover, our data suggest that this would occur within time frames used for in vivo imaging of Ca2+ signaling, which typically starts after a 1-hour loading period of Ca2+ indicators (3537). As expected, toxicity was most pronounced when high loading concentrations were used (Fig. 5). This finding is particularly relevant given that a search of the literature showed that high loading concentrations are still used (fig. S1A).

A key question is whether Ca2+ indicators and BAPTA reduce the Na,K-ATPase activity by chelation of cytosolic Ca2+. Ca2+ ions decrease, rather than increase, the activity of Na,K-ATPase (11, 18, 46). Moreover, several observations reported here suggest that cytosolic Ca2+ was not the main driver of the Ca2+ indicator–mediated suppression of the Na,K-ATPase. For example, the potency by which the Ca2+ indicator and BAPTA induced the inhibition of ouabain-sensitive 86Rb+ uptake (Rhod-2 AM > Fluo-4 AM > BAPTA AM; Fig. 1C) was not a function of their Kd (BAPTA = Fluo-4 < Rhod-2; fig. S1C). In addition, the rank order of inhibition was not a function of their intracellular accumulation (BAPTA > Fluo-4 > Rhod-2; Fig. 1, B and C). Furthermore, 5,5′-dibromo BAPTA suppressed the Na,K-ATPase with a potency comparable to BAPTA (Fig. 3D), although it has a lower Ca2+ affinity than BAPTA (fig. S1B). Moreover, a 0 Ca2+ concentration in cell-free ATPase assay buffer increased Na,K-ATPase–mediated ATP hydrolysis (Fig. 3D). Finally, GCaMP3 expression did not affect ouabain-sensitive 86Rb+ uptake, although GCaMP3 can chelate Ca2+ with a Kd of 345 nM, which is identical to that of Fluo-4 (Kd = 345 nM) (Fig. 8B) (4). An alternative explanation is that the Ca2+ indicators and BAPTA reduced Na,K-ATPase activity by binding to the Na,K-ATPase directly or interacting with associated proteins. Only 15 to 35% of the diffusible Ca2+ indicators are free and in an unbound state in the cytosol, whereas the rest is bound to various cytosolic proteins or trapped in organelles (47, 48). The Na,K-ATPase interacts with many membrane and cytosolic proteins, including Src and calmodulin, to form a protein complex (18, 49, 50). Powis and Wattus have proposed that Ca2+-bound calmodulin can stimulate the Na,K-ATPase (51). BAPTA directly interacts with several different Ca2+-binding proteins (calmodulin, parvalbumin, and trypsin), and the formation of BAPTA-protein complex occurs regardless whether these proteins are in their Ca2+-free or Ca2+-bound forms (52). The membrane preparations in this study were prepared by hypotonic shock, followed by several washes to restore osmolarities. This procedure eliminates most cytoplasmic but not all membrane-bound proteins (53). Therefore, our analysis cannot define whether Ca2+ indicators and BAPTA reduced pump activity by directly interacting with the Na,K-ATPase or interacting with other membrane-bound modulatory proteins. This limitation is shared by many previous studies and should be subjected to further scrutiny to avoid misinterpretation of past results. In addition to the effect of Ca2+ indicators on the Na,K-ATPase reported here, the BAPTA-based Ca2+ chelators, diazo-2 and dibromo BAPTA, can activate K+ channels in rat hippocampal CA1 neurons (54). However, it is unlikely that formaldehyde generated by hydrolysis of AM form of the Ca2+ indicator played an important role in the toxicity observed. For example, the potency of 86Rb+ uptake inhibition (Rhod-2 AM > Fluo-4 AM > BAPTA AM) did not correlate with the intracellular concentration of the hydrolyzed indicators (BAPTA > Fluo-4 > Rhod-2; Fig. 1B), which, in a ratio of 1:4, reflects the formation of the hydrolysis product formaldehyde (55). Changes in [3H]-2-DG uptake were inversely correlated with the intracellular concentration of indicators, because Fluo-4 AM and Fura-2 AM were present at higher concentrations than Rhod-2 AM but decreased uptake (Fig. 6, A and B). Furthermore, the salt form of Rhod-2 and BAPTA (without the AM moiety) directly inhibited the Na,K-ATPase activity (Figs. 3, B and C, and 4, D to H). Thus, the Ca2+ indicator itself rather than formaldehyde produced by AM hydrolysis was the trigger for the deleterious effects on Na,K-ATPase function and metabolic status in our studies. Another important issue is the possible toxicity of the organic solvent DMSO, which has multiple pharmacological and pathological effects (56). However, our analysis showed that DMSO itself did not significantly affect 86Rb+ uptake at the concentrations that are typically used in Ca2+ imaging experiments (fig. S2). Long-term treatment of DMSO also did not significantly affect the number of viable cells, LDH release, or cell volume (Fig. 5, A to D).

Tsien’s group, who also engineered the original Ca2+ indicator dyes, developed the genetically encoded Ca2+ indicator Cameleon (57), which has led to the generation of other types of genetically encoded Ca2+ indicators, including GCaMPs (58). GCaMPs are the most used of these indicators because of their high fluorescence, rapid kinetics, and thermal stability and because they have a suitable Kd range for measuring cytosolic Ca2+ levels (59). Here, we showed that GCaMP3 did not inhibit the activity of the Na,K-ATPase (Fig. 8B). These findings suggest that GCaMPs or potentially other genetically encoded Ca2+ indicators may be more suitable than chemical Ca2+ indicators for future studies and reassessing previous observations.

In summary, our analysis found that BAPTA-derived Ca2+ indicators potently inhibited the Na,K-ATPase activity and suppressed spontaneous Ca2+ signaling. In contrast, the genetically encoded Ca2+ indicator GCaMP3 interfered minimally with Na,K-ATPase activity. The Na,K-ATPase regulates physiological processes at multiple levels ranging from single-cell Na+-coupled glutamate uptake (60) to glucose utilization (61), cell adhesion (62), cellular excitability, and synaptic transmission (30, 63), as well as cardiac (64) and skeletal muscle contraction (65) and blood pressure (66). Given the many roles of the Na,K-ATPase in physiological and pathological processes, it is necessary to consider the consequences of Na,K-ATPase inhibition in experiments using Ca2+ indicators or BAPTA. Our study highlights limitations of studies that used Ca2+ indicators in cellular physiology and suggests that a reevaluation of previous key observations, potentially with genetically encoded Ca2+ indicators, is warranted.


Cell culture

Mouse cortical astrocyte cultures were prepared from 0- to 2-day-old C57BL/6 (Charles River Laboratories) mouse pups, as previously described (67). For some experiments, we used MrgA1+/− transgenic and littermate control MrgA1−/− mice (courtesy of K. McCarthy, University of North Carolina at Chapel Hill) (31). We also used the GCaMP3 transgenic mouse, which is based on the flexible Cre/lox system (GCaMP3fl) (the Jackson Laboratory). GCaMP3fl mice were crossed with transgenic mice expressing Cre recombinase under the control of GFAP promoter. Astrocytes were prepared from GCaMP3fl and GFAP-Cre:GCaMP3fl double-positive (GFAP-GCaMP3) mice. Briefly, dissociated cells were plated on culture flasks and maintained in Dulbecco’s modified Eagle’s medium (DMEM)/F12 containing 10% fetal bovine serum (FBS), penicillin (100 U/ml), streptomycin (100 μg/ml), and amphotericin B (25 μg/ml). Astrocytes were subcultured on the plates for subsequent assays.

Mouse neocortical neurons were prepared from 0- to 3-day-old C57BL/6 mouse pups, as previously described (68), with slight modifications. Briefly, cerebral cortices were dissected and meninges were removed. The tissues were minced and then suspended in Hibernate A solution minus calcium (BrainBits LLC) containing papain (Worthington) at 37°C for 30 min. The suspension was then triturated with a 0.23 m siliconized pipette. The cell suspension was then separated by density gradient centrifugation. The neuronal cells were then seeded onto poly-d-lysine–coated 24-well plates and maintained in Neurobasal A medium (Invitrogen) with 0.5 ml of B27, 0.5 mM GlutaMAX, gentamicin (10 mg/ml), and human fibroblast growth factor 2 (FGF2) (5 ng/ml) at 37°C in an incubator containing humidified air and 5% CO2. Immunohistochemistry showed that more than 90% cells in the cultures [identified by DAPI (4′,6-diamidino-2-phenylindole)–stained nuclei] were MAP2-positive. Neuronal cultures were used 5 to 7 days after plating.

Human cortical astrocyte cultures were derived from temporal neocortex, as previously described (69), under protocols approved by the University of Rochester–Strong Memorial Hospital Research Subjects Review Board. Briefly, the forebrain tissue samples were collected and washed two to three times with sterile Hanks’ balanced salt solution with Ca2+/Mg2+ (HBSS+/+). The cortical plate region (CTX) of the fetal forebrain was dissected and separated from the ventricular zone/subventricular zone portion. The CTX was then dissociated with papain, as previously described (70). The cells were resuspended at 2 × 106 to 4 × 106 cells/ml in DMEM/F12 supplemented with N1, 0.5% plasma-derived FBS, and basic FGF (10 ng/ml) and plated in suspension culture dishes. The day after dissociation, cortical cells were recovered and subjected to magnetic-activated cell sorting (MACS) for purification of the bipotential glial progenitor cell (GPC) population (PSA-NCAM/A2B5+), as previously described (71). The purified GPCs were cultured in DMEM/F12 supplemented with N1 and 5% FBS to further differentiate them. To prepare culture dishes for ATPase assays or 86Rb+ uptake measurement, the fetal cortical astrocytes were passaged with papain into single cells and then plated onto poly-l-ornithine/laminin–coated 24-well plates (50,000 cells per well).

Mouse cardiomyocyte cultures were prepared from 0- to 2-day-old C57BL/6 mouse pups (Charles River Laboratories), as previously described (72). To prepare the cardiogel matrix–coated plates, cardiac fibroblasts were obtained from whole hearts of 0- to 5-day-old C57BL/6 mouse pups. The cells were cultured on dishes precoated with 1% gelatin for 5 days and reseeded onto the plates. When the cultured fibroblasts reached confluency (3 to 4 days), the extraction buffer (0.5% Triton X-100 and 20 mM NH4OH in HBSS) was added to the plate to lyse the cells, and the cellular debris were washed off with chilled HBSS. For cardiomyocytes, the heart ventricles were excised, and the auricles were removed and discarded. The cardiomyocytes were dispersed by incubating with 0.5% trypsin-EDTA (4 × 5 min, 37°C) and purified by the differential attachment of nonmyocardial cells to the 1% gelatin–precoated dishes (twice for 1.5 hours each time). The nonadhesive cells (cardiomyocytes) were transferred into the cardiogel matrix–coated plates and cultured in the DMEM/F12 medium supplemented with 20% FBS, 5% horse serum, 2 mM glutamine, 0.1 mM nonessential amino acids, 3 mM sodium pyruvate, bovine insulin (1 μg/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), and amphotericin B (25 μg/ml). Beating cardiomyocytes were present on the third day in culture (movie S1).

Human renal proximal tubule epithelial cells were obtained from Lifeline Cell Technology and cultured according to the manufacturer’s protocol with the RenaLife basal medium containing 0.5% FBS, recombinant human insulin (5 μg/ml), hydrocortisone hemisuccinate (0.1 μg/ml), 2.4 mM l-alanyl-l-glutamine, 1 μM epinephrine, transferrin (5 μg/ml), 10 nM triiodothyronine, recombinant human epidermal growth factor (10 ng/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), and amphotericin B (25 μg/ml). Cultured cells were incubated in 5% CO2/95% air at 37°C. Animal procedures were approved by the Institutional Animal Care and Use Committee of University of Rochester.

Titration of adenoviral infection of GCaMP3fl astrocytes

To induce GCaMP3 expression in cortical mouse astrocyte cultures prepared from GCaMP3fl mice, cells were infected with an adenovirus bearing Cre recombinase under the control of the constitutive cytomegalovirus (CMV) promoter (courtesy of A. N. Economides, Regeneron Pharmaceuticals, Tarrytown, NY) for 5 hours, with 10, 50, or 100 multiplicities of infection (MOIs) (73). At day 6 after transduction, GCaMP3 expression was confirmed under the microscope, and cells were used for Ca2+ imaging and 86Rb+ uptake experiments (AdV-GCaMP3). Cytotoxicity was not observed with 10 to 100 MOIs. For control experiments, cells were infected with an adenovirus bearing GFP under the control of the CMV promoter (AdV-GFP control) (74).

Ca2+ imaging

Ca2+ imaging was performed as previously described (30). Astrocytes without or with GCaMP3 expression (GFAP-GCaMP3 and AdV-GCaMP3) were loaded with Fluo-4 AM (2 or 4 μM), Rhod-2 AM (2.25 or 4.5 μM), or DMSO (0.1%) for 30 min at 37°C, followed by washing with DMEM/F12 medium for the intracellular Ca2+ imaging. AM indicators were initially dissolved in DMSO and then introduced to the culture medium. Images were taken using a confocal microscope (FV500, Olympus) at 1.2 s per frame with 488-nm laser for GCaMP and Fluo-4 or 532-nm laser for Rhod-2. Cells were kept at 37°C with a stage heater during the imaging. For stimulation with 5 μM ATP, 15 visually identifiable cell bodies were randomly selected in each image. Circular regions of interest (ROIs) were used, and the fluorescence time course of each cell was measured by averaging all pixels within the ROI. The frequency of spontaneous Ca2+ events was obtained by counting the total numbers of Ca2+ events for a period of 5 min in a field spanning 1273 μm × 1273 μm of each image.

86Rb+ uptake measurement

86Rb+ uptake measurement was conducted as previously described, with a slight modification (30). We used DMSO to initially dissolve membrane-permeable AM-linked Ca2+ indicators, and therefore, DMSO was used as a vehicle control for experiments. In each set of experiments, all groups received the same amount of DMSO, except in fig. S2, in which the effect of DMSO was examined. Cells were loaded with various concentrations of Fluo-4 AM (0 to 10 μM), Rhod-2 AM (0 to 10 μM), Fura-2 AM (0 to 8 μM), BAPTA AM (0 to 40 μM), or identical amount of DMSO (0 to 0.2%) for 20 min and further incubated in the presence or absence of ouabain (1 mM) for 10 min (30-min total incubation with AM indicators at 37°C). The potassium analog 86Rb+ was added to each well for 15 min (1 μCi; PerkinElmer), and the 86Rb+ uptake was quantified by liquid scintillation counting (Beckman Coulter). Data were normalized to cell numbers for comparison of cultures prepared from different transgenic mice. Ouabain-sensitive 86Rb+ uptake was calculated by subtracting 86Rb+ uptake during the same conditions but in the presence of ouabain.

[3H]-2-DG uptake and lactate release measurement

Glucose uptake was determined by using 2-deoxyglucose labeled with tritium (75). Cells were loaded with various concentrations of Ca2+ indicators, BAPTA AM, or identical amount of DMSO, as described in the 86Rb+ uptake measurement section in the presence or absence of iodoacetate (300 μM) and sodium cyanide (100 μM). The supernatants (100 μl) were collected for lactate release measurement after a 30-min incubation period at 37°C, followed by the addition of the glucose analog [3H]-2-DG to each well and incubation for 15 min (1 μCi; PerkinElmer). [3H]-2-DG uptake and lactate release can therefore not directly be compared. [3H]-2-DG uptake was quantified by liquid scintillation counting (Beckman Coulter). [3H]-2-DG uptake was expressed as percentage of control [3H]-2-DG uptake. The concentrations of lactate in samples were measured by using lactate assay kit (Abcam Inc.).

Neuronal lactate release measurement

Cells were incubated with ouabain (1 mM), Ca2+ indicators (2 to 5 μM in 0.2% DMSO), or BAPTA AM (5 to 10 μM in 0.2% DMSO) for 30 min at 37°C and then washed four times. The supernatant was collected 6 and 24 hours after incubation. The concentrations of lactate were measured by using LDH Colorimetric Assay kit (Abcam Inc.).

ATPase assay

Na,K-ATPase–mediated ATP hydrolysis was measured as reported previously, with a slight modification (19, 21, 76). After the loading of Ca2+ indicators, BAPTA AM, or DMSO, as described in 86Rb+ uptake measurement, the cells were washed five times with tris-buffered water (15 mM tris-HCl, pH 7.4) and lysed for 4 to 12 hours at 4°C. The cell ghosts were washed and collected from the culture plate by using cell lifter (Corning) in assay buffer containing 100 mM NaCl, 15 mM KCl, 5 mM MgCl2, 500 nM CaCl2, 50 mM tris, and protease inhibitor cocktail (100×) (pH 7.4). Ca2+ concentration in the assay buffer was modified (0 or 500 nM), where indicated. The collected cell ghosts were divided into two groups and used for ATP hydrolysis and protein measurement. For ATP hydrolysis measurements, the membrane preparation was preincubated for 10 min at 37°C. For some experiments, Rhod-2 tripotassium salt (10 μM), BAPTA tetrapotassium salt (100 μM), or 5,5′-dibromo BAPTA tetrapotassium salt (100 μM) was added to membrane preparations. Vanadate-free ATP (1 mM) was used as the substrate (77, 78), and the liberated inorganic phosphate during 10-min incubation at 37°C in the presence or absence of ouabain (1 mM) was quantified by using malachite green phosphate detection kit (R&D Systems). Ouabain-sensitive ATPase activity was calculated and expressed as nanomoles of inorganic phosphate released per milligram protein per minute. The protein concentration in the samples was measured by the bicinchoninic acid assay (BCA) protein assay (79).

Intracellular Fluo-4, Rhod-2, and BAPTA concentration measurements

After the incubation period with Fluo-4 AM, Rhod-2 AM, BAPTA AM, or DMSO, cells were gently washed four times with assay buffer and then collected in assay buffer by using a cell lifter. For Fluo-4 and Rhod-2 sample collection, we used the assay buffer supplemented with 100 μM Ca2+ to maximize the fluorescent intensity. For BAPTA sample collection, we used the assay buffer without CaCl2 and supplemented with 1 mM EGTA to maximize the optical density. In pilot studies, we confirmed that EGTA itself (1 mM) did not show detectable absorption at 260 nm but effectively maximized the absorption of BAPTA by replacing the Ca2+ bound to BAPTA. Cells were dissolved by sonication and centrifuged at 12,000 rpm for 5 min. The supernatant was used for measurement. The fluorescence readings (Fluo-4: excitation, 485 nm; emission, 535 nm; Rhod-2: excitation, 530 nm; emission, 620 nm) were performed using a luminometer (Victor2, Wallac, PerkinElmer), and absorption at 260 nm was detected by using a spectrophotometer (SpectraMax, Molecular Devices). The standards were generated from DMSO-treated control cells, as described above, and supplemented with various concentrations of Fluo-4 (10 to 1000 nM), Rhod-2 (10 to 1000 nM), or BAPTA (0.5 to 32 μM) salts. The intracellular concentrations were calculated from the total cell number, and cell volume was measured by using a Scepter 2.0 cell counter (EMD Millipore Corp.).

LDH assay

LDH released into the medium was measured as a marker of dead cells (80). Various concentrations of ouabain (0 to 5 mM), Ca2+ indicators (1 to 10 μM in 0.2% DMSO), BAPTA AM (5 to 40 μM in 0.2% DMSO), or DMSO (0.02 to 0.2%) were incubated with cells for 2 hours at 37°C, and the culture supernatant was collected for LDH assay. LDH concentrations in the samples were detected by using the LDH Cytotoxicity Assay kit II (Abcam Inc.).

Cell volume and viability assessment

To count viable cells, we used the Scepter 2.0 cell counter to rapidly count cell numbers and diameters. The cells were incubated with ouabain (0 to 5 mM), Ca2+ indicators (1 to 10 μM in 0.2% DMSO), BAPTA AM (5 to 40 μM in 0.2% DMSO), or DMSO (0.02 to 0.2%) for 30 min or 2 hours at 37°C. After washing with serum-free medium at 30 min or 2 hours, the cells were further incubated for a total period of 24 hours and directly sampled from the plate into the tip attached to the cell counter. A debris/dead cell fraction appeared in the smaller diameter range (>11 μm), which was excluded from the viable cell number. In fig. S4, cells were washed four times in phosphate-buffered saline and centrifuged. The cells were then resuspended in various concentrations of Ca2+ indicator buffers and incubated for 30 min at room temperature. The samples were analyzed using ImageStream (81).

In vivo microdialysis and measurement of K+, ATP, and glycerol

The effects of Rhod-2 and Fluo-4 AM loading in the intact brain were evaluated by using microdialysis in freely moving awake mice. This approach permits direct monitoring of the changes before, during, and after loading of Rhod-2 and Fluo-4 AM by collection of interstitial fluid. For surgical implantation of microdialysis guide cannula, 8- to 12-week-old C57BL/6 mice (Charles River Laboratories) were anesthetized with a mixture of ketamine [60 mg/kg, intraperitoneally (i.p.)] and xylazine (10 mg/kg, i.p.). The guide cannula was stereotactically implanted in the somatosensory cortex, as previously described (82). After surgery, animals were given a recovery period (24 hours). Microdialysis probes (2 mm membrane length; 30-kDa cutoff; BASi) were carefully inserted, and the probe tip was positioned at the following coordinate: anterior-posterior, −1.5 mm; medial-lateral, +3.96 mm (probe angle, 35°); dorsal-ventral, −2.0 mm (probe angle, 45°). After overnight equilibration with aCSF (145 mM NaCl, 4 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 1.25 mM NaH2PO4, 15 mM Hepes, pH 7.4), the microdialysates were collected every 15 min (flow rate at 2 μl/min). Rhod-2 and Fluo-4 AM (5 μM) were prepared in 0.1% Pluronic F-127, 0.1% DMSO, and aCSF and filtered with 0.2-μm-pore syringe filters (Nalge Nunc International). In two-photon imaging experiments, Ca2+ indicators are usually loaded into tissue with multicell bolus loading or surface application. For multicell bolus loading, high concentrations of Ca2+ indicators in a small volume are pressure-ejected intracortically from the micropipette positioned at 150 to 750 μm below the pia (approximately 0.4 to 1.0 fmol), followed by a 1- to 1.5-hour waiting period (35, 36, 83). For surface application, which was also used in our previous studies (82, 84), 18 to 45 nmol of Ca2+ indicators are generally applied onto the exposed surface of cortical tissue and incubated for 45 to 60 min to stain astrocytes in the superficial layer (37, 83, 85). Here, we delivered 20 to 38 pmol of the Ca2+ indicator into the cortex for a period of 1 hour. The aCSF containing Pluronic F-127 and DMSO was used as control. The collected samples were divided into two aliquots, 12 μl for the ATP and glycerol measurement and 18 μl for the K+ measurement, and were immediately stored at −80°C.

To measure K+ changes in the microdialysates, ion-sensitive microelectrodes were made as previously described (30, 86). Briefly, pipettes were silanized by dimethyldichlorosilane (Silanization Solution I, Sigma-Aldrich) and loaded with a 100- to 150-μm column of K+ exchanger (potassium ionophore I—cocktail B, Sigma-Aldrich). The electrodes were calibrated before the experiments with 2 to 20 mM K+ in aCSF and were recalibrated at the end of each experiment. The baseline was monitored by randomly running standard during measurement of samples. The ATP concentrations in the microdialysates were measured by using a bioluminescent ATP assay mix (87) and a Victor 2 plate reader (Wallac). The concentration of glycerol in the samples was detected by using Free Glycerol Detection, Glucose Assay, and Lactate Assay kits (Abcam Inc.), respectively, following the manufacturer’s instructions.

Recording of Na,K-ATPase current on hippocampal slice

Coronal hippocampal brain slices (300 μm) were prepared from 8- to 12-week-old mice. Briefly, hippocampal slices were cut using a Leica VT1000s (Leica Biosystems) in ice-cold slicing buffer (127 mM NaCl, 26 mM NaHCO3, 1.2 mM KH2PO4, 1.9 mM KCl, 1.1 mM CaCl2, 2 mM MgSO4, and 10 mM d-glucose) bubbled with 95% O2 and 5% CO2. Slices were then transferred to a holding chamber containing oxygenated aCSF (127 mM NaCl, 26 mM NaHCO3, 1.2 mM KH2PO4, 1.9 mM KCl, 2.2 mM CaCl2, 1 mM MgSO4, and 10 mM d-glucose) for 30 min at 34°C and for another 30 min at room temperature for recovery and then transferred to a submersion recording chamber continually perfused with oxygenated aCSF containing 1 μM tetrodotoxin (Sigma-Aldrich) at room temperature (rate, 2 ml/min). Slices were equilibrated for at least 15 min before each recording.

Whole-cell recordings from pyramidal neurons in hippocampal CA1 region were made using a MultiClamp 700B amplifier (Axon Instruments), sampled at 10 kHz, digitized by a Digidata 1440A, and later analyzed offline by Clampfit (Axon software). Recording pipettes with resistances ranging between 3 and 6 megohms were pulled using standard borosilicate capillaries by a two-step electrode puller (PC-10, Narishige) and were filled with a K-gluconate–based patch solution (125 mM K-gluconate, 20 mM KCl, 10 mM NaCl, 2 mM Mg-ATP, 0.3 mM Na-GTP, 2.5 mM QX314, 10 mM Hepes, pH 7.3 adjusted with KOH). Because Na,K-ATPase constantly moves three Na+ ions out of the cell and two K+ ions into the cell, the pump generates a constant outward current. By inhibiting this constant outward current with strophanthidin (0.5 mM; Sigma-Aldrich), the holding current is decreased, and such inward move of the holding current recorded at −70 mV is measured as the current carried by Na,K-ATPase. Membrane-impermeable calcium indicators were added into patch solution to test their effect on the current carried by Na,K-ATPase.


Values are expressed as means ± SEM. All data represent four or more independent experiments, and in vitro results were obtained from separate cultures. Normality of the data was evaluated with Shapiro-Wilk test, with α = 0.05. Differences between two means were assessed by Student’s t test, and differences among multiple means were assessed by one-way ANOVA with Tukey-Kramer post hoc test or two-way ANOVA with Bonferroni post hoc test. For data that failed normality test, differences were assessed with Kruskal-Wallis and Mann-Whitney post hoc tests. The null hypothesis was rejected when P < 0.05.


Fig. S1. Structures and characters of Ca2+ indicator derivatives used in publications listed in PubMed.

Fig. S2. DMSO does not affect 86Rb+ uptake by astrocytes.

Fig. S3. Ca2+ indicators do not affect ouabain-insensitive 86Rb+ uptake.

Fig. S4. Ca2+ indicators inhibit Na,K-ATPase–mediated ATP hydrolysis and 86Rb+ uptake in rat astrocyte cultures.

Fig. S5. Ca2+ indicators induce lactate release in neurons and increase cell volume in neurons and astrocytes.

Fig. S6. Quantification of spontaneous and pharmacologically evoked Ca2+ signals and 86Rb+ uptake in GCaMP3-expressing astrocytes.

Movie S1. Beating cardiomyocytes in culture prepared from mouse (2× frame rate).

Movie S2. ATP-induced Ca2+ response in astrocytes expressing GFAP-GCaMP3.

Movie S3. ATP-induced Ca2+ response in astrocytes expressing AdV-GCaMP3.

Movie S4. ATP-induced Ca2+ response in astrocytes loaded with Rhod-2 AM.

Movie S5. Spontaneous Ca2+ response in astrocytes expressing GFAP-GCaMP3.


Acknowledgments: We thank V. Gallo for valuable comments on the manuscript. We thank K. McCarthy for sharing transgenic mice. Some microscopic analysis was carried out at the Children’s Research Institute (CRI) Light Microscopy and Image Analysis Core supported by CRI and DC-IDDRC grant U54HD090257 by the National Institute of Child Health and Human Development. Funding: This work was supported by NIH grants (NS078167 and NS078304 to M.N.) and F31 National Research Service Award and T32 Postdoctoral Training Grant (F31NS073390 and 5T32HD046388 to N.A.S.). Author contributions: N.A.S., Y.L., and D.C.-M. performed the experiments. N.A.S. and Y.L. analyzed data. N.A.S. and M.N. designed the experiments. A.B. provided adenoviruses. N.A.S., B.T.K., and M.N. wrote the paper. Competing interests: The authors declare that they have no competing interests.
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