H9C2 cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) (Gibco Laboratories, Gaithersburg, MD) supplemented with 10% (v/v) Fetal Bovine Serum (FBS) (Gibco Laboratories), 2 mmol/L L-Glutamine (Gibco Laboratories), and 1X Penicillin-Streptomycin (P/S) (100 U/mL of penicillin and 100 μg/mL of streptomycin) Solution (EMD Millipore, Burlington, MA). AC16 cells were cultured DMEM/F12 (Sigma, St Louis, MO) containing 2 mmol/L L-Glutamine, 12.5% FBS, and 1X P/S Solution. Cells were maintained in 5% CO₂ at 37°C with a confluence between 10% and 80%. Induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) were reprogrammed from human cardiac fibroblasts, following the procedure outlined in Zhao et al. (Zha et al., 2018). They were cultured on Matrigel Membrane Matrix in mTeSR™ medium until reaching 75% confluence. Cardiomyocyte differentiation was achieved using a small molecule-based protocol, as previously described (Ernst et al., 2021). Beating iPSC-CMs typically emerged 9–12 days after the initiation of differentiation. To purify the iPSC-CM cultures and eliminate non-cardiomyocytes, a glucose-free medium containing lactate was employed. Following purification, iPSC-CMs were maintained in RPMI 1640 supplemented with B27 Supplement and 1X P/S. All iPSC-CMs utilized in experiments were within 3 weeks of differentiation.

Mouse cardiomyocytes were isolated from 8-week-old C57BL/6 mice via Langendorff perfusion, as previously described (Goh et al., 2019; Kim et al., 2020). Briefly, following thoracotomy, hearts were quickly excised, mounted on a Langendorff apparatus, and perfused with Tyrode’s solution containing (in mmol/L) 113 NaCl, 4.7 KCl, 0.6 KH₂PO₄, 0.6 Na₂HPO₄, 1.2 MgSO₄, 10 Na-HEPES, 12 NaHCO₃, 10 KHCO₃, 0.032 phenol red, 30 taurine, 10 2,3-Butanedione monoxime, and 5.5 glucose at 37°C for 3–5 min. Thereafter, the heart was digested with collagenase II (Worthington Biochemicals, Lakewood, NJ) for 10–12 min. The digested heart was pulled apart with forceps into tiny pieces and then piped in digestion solution several times to further break up cell clusters. The cell solution was filtered through a 100 μm cell strainer to remove any undigested tissue. Next, cardiomyocytes were introduced to grade-elevating concentrations of CaCl₂ (in μmol/L: 12.5, 100, 400, and 900) to establish Ca²⁺ tolerance. Finally, isolated cells were stored in a high K⁺ solution (in mmol/L: 120 Glutamate, 25 KCl, 1 MgCl₂, 10 HEPES, 1 EGTA, and pH 7.5 with KOH) temporarily before fluorescent dye loading and experiments.

Determination of MitoView 633 excitation and emission spectra was performed using a Spectramax i3x microplate reader (Molecular Devices, San Jose, CA). Cells were loaded with MitoView 633 of a desired concentration. Fifteen minutes later, the dye-containing medium was replaced with a colorless culture medium titrated to the indicated pH before spectral measurement. The temperature was adjusted using Spectramax’s internal temperature control. Excitation spectra were determined by measuring 720 nm emission using excitation light from 500–700 nm in 10 or 2 nm increments. Emission spectra were determined by measuring emission from 600–800 nm in 10 or 2 nm increments using excitation light at 580 nm. All spectra (absorption and emission) were normalized to their maximum values.

The Cellular ATP content was quantified employing the CellTiter-Glo Luminescent Viability Assay (Promega, Madison, WI). In brief, an equivalent number of cells were seeded into 96-well plates on Day 0. On Day 1, cells were exposed to Mitoview 633 at varying concentrations (ranging from 0 to 200 nmol/L) for a 30-min incubation period. Following incubation in darkness, cells were rinsed and resuspended in 100 μL of phenol red-free DMEM. Simultaneously, an ATP standard curve was generated using ATP disodium salt (Sigma Cat. # A7699) within the same plate on which stained cells were seeded. Subsequently, 100 μL of CellTiter-Glo reagent was introduced into each well, followed by a brief 2–3-min agitation of cellular contents on an orbital shaker. The plate was then allowed to incubate for 10 min to stabilize the luminescent signal, after which luminescence was measured using a SpectraMax microplate reader.

Cells (adult mouse cardiomyocytes or iPSC-CMs) were loaded with MitoView 633 (25 nmol/L for 15 min at 37°C), MitoTracker Green (250 nM for 30 min at 37°C), or TMRM (20 nmol/L for 15 min at 37°C). The glass-bottom dish containing dye-loaded cells was equilibrated at 37°C (or specified temperature) with unrestricted access to atmospheric oxygen on the stage of an Olympus FV1000 confocal microscope (Olympus, Center Valley, PA). The 488 nm argon laser, 543 nm HeNe laser, and 635 nm LD laser lines were used to image MitoTracker Green, TMRM, and MitoView 633, respectively. The emission filters used to collect images were 505–605 nm, 560–620 nm, and 655–755 nm, respectively. Colocalization of Mitotracker Green and MitoView 633 was analyzed using the previously described method (Kim et al., 2020).

To access the performance of MitoView 633 in measuring stress-induced ΔΨ_m depolarization, cells loaded with MitoView 633 alone or in conjunction with TMRM were deposited onto a cover glass in a microincubation system (Harvard Apparatus, Cambridge, MA). The chamber was held at 37°C and perfused with Tyrode’s solution containing (in mmol/L): 137 NaCl, 2.7 KCl, 1 MgCl₂, 20 HEPES, 4.5 glucose, 1.8 CaCl₂, pH 7.4, at a flow rate of 1 mL/minute. Mitochondrial depolarization was induced by introducing FCCP into the perfusate at a final concentration of either 1 or 10 μmol/L. The 543 nm HeNe laser and 635 nm LD laser lines were used to image TMRM and MitoView 633, respectively.

To simultaneously measure ΔΨ_m, ROS levels, and cytosolic Ca²⁺, isolated cardiomyocytes were loaded with MitoSOX Red (2 μmol/L), MitoView 633 (25 nmol/L), and Fluo4-AM (5 μmol/L for 30 min, followed by a 30-min wash out to allow for de-esterification). The laser flash method described previously (Aon et al., 2003; Zhou et al., 2011; Goh et al., 2016) was adopted to elicit local oxidative stress and mitochondrial depolarization. In brief, a high-intensity laser flash (453 nm, ∼25 mW, 100 ms duration) was applied to a small (∼100 μm³) region within the cell volume. After photoactivation, Fluo4-AM, MitoSOX Red, and MitoView 633 were concurrently imaged using the 488 nm argon laser, 543 nm HeNe laser, and 635 nm LD laser, respectively with x60 oil immersion lens (numerical aperture = 1.4). For fast time resolution of Ca²⁺ sparks, the linescan mode of the confocal microscope was used. The linescan rate was 2.0 ms per line; and 512 × 512 pixel xy images were acquired every 3.7 s, as previously described (Zhou et al., 2011).

Images were processed offline using ImageJ software (Wayne Rasband, NIH) or Matlab (The MathWorks, Inc., Natick, MA).

Comparisons were performed using paired or unpaired 2-tailed Student’s t-test. Data were considered significantly different at p < 0.05. Results are presented as mean ± SEM.

We first analyzed the spectrum of MitoView 633 in H9C2 cells using the Spectramax i3x microplate reader. The results showed that MitoView 633 excitation and emission spectrum peaked at 622 nm and 648 nm, with the full width at half maximum (FWHM) of 20 nm and 25 nm, respectively, at 37°C and pH 7.4 in colorless culture medium (Figure 1A). These data were consistent with the specifications provided by the manufacturer.

To investigate the influence of extracellular pH on the fluorescent spectra of MitoView 633, we conducted measurements in H9C2 cells at 37°C, using a colorless culture medium adjusted to pH levels of 6.6, 7.0, or 7.4. At a neutral pH of 7.4, both spectra exhibited a minor blue shift in the peak wavelength, approximately 5 nm, along with a slight reduction in bandwidth, resulting in a 5 nm narrower full width at half maximum (FWHM) when compared to the neutral pH of 7.0 (Figures 1B, C; Table 1). Conversely, at a lower pH of 6.6, we observed an approximate 5 nm red shift in the peak wavelength compared to the neutral pH. The bandwidth at pH 6.6 showed a more pronounced narrowing, resulting in an FWHM 10 nm narrower than that at neutral pH (Table 1). It's important to note that changes in extracellular pH may not accurately reflect intracellular pH, especially in the vicinity of mitochondria or within the mitochondrial matrix. We proceeded to evaluate the impact of pH on MitoView 633 spectra in a cell-free medium. As depicted in Figures 2A, B, alterations in pH had minimal influence on both the excitation and emission spectra of MitoView 633, unlike the observations in cells. Given that the loading of MitoView 633 into the mitochondrial matrix relies on the potential gradient across the membrane, we further examined the MitoView 633 spectra under varying dye concentrations in the cell-free medium. As anticipated, increasing concentrations of MitoView 633 resulted in more pronounced fluorescence signals (Figures 2C, D). Interestingly, it became apparent that dye concentration also exerted an impact on the spectra. With increasing concentration from 1 μmol/L to 25 μmlo/L, the peak excitation wavelength shifted slightly from 624 nm to >626 nm, while the peak emission wavelength shifted significantly from 641 nm to 651 nm (Figures 2E, F). The observed red shift in spectra induced by elevated dye concentration mirrored that caused by reduced pH, suggesting a connection between extracellular pH (or plasma membrane potential) and the accumulation of cationic MitoView 633 within mitochondria.

Temperature dependence was measured in H9C2 cells at neutral pH using Spectramax’s heating capabilities to adjust the culture temperature from 25°C up to 37°C. Neither excitation (Figure 1D) nor emission (Figure 1E) spectra exhibited a significant red- or blue shift across the entire range measured. A small difference in bandwidth was observed between the two most extreme temperatures, amounting to an about 5 nm narrower FWHM at 25°C as compared to 37°C (Table 2). These data indicate MitoView 633’s fluorescent spectral stability across the physiologically relevant temperature and pH ranges. MitoView 633’s spectra stability is a very useful characteristic; it is highly likely that local intracellular variances in either property could not reach such a magnitude without first causing significant off-target physiological effects.

We next examined the subcellular staining and distribution of MitoView 633 across various cell types. Confocal imaging unveiled that MitoView 633 consistently displayed specific staining within the mitochondria of adult cardiomyocytes. This was confirmed by the colocalization observed with MitoTracker Green, a well-established mitochondrial marker (Figure 3A). To enhance clarity, simultaneous line scan imaging was performed, demonstrating a nearly perfect overlap of the MitoView 633 (red line) and MitoTracker Green (green line) fluorophore peaks (Pearson correlation coefficient, PCC = 0.93 ± 0.04), providing unequivocal evidence of MitoView 633’s mitochondrial localization. The mitochondrial-specific staining of MitoView 633 was also consistently observed in all other tested cell types, including hiPSC-CMs (Figure 3B, PCC = 0.97 ± 0.18), as well as myoblast cells H9C2, triple-negative breast cancer cells, and neurons (data not shown). These findings collectively indicate that MitoView 633 possesses the versatility to serve as a universal far-red spectrum mitochondrial indicator.

Photobleaching represents a phenomenon where a fluorophore experiences a permanent loss of its ability to fluoresce due to photon-induced chemical damage and covalent modifications (BERNAS et al., 2004). This occurrence leads to a reduction in signal-to-noise ratio and image resolution, thereby limiting the utility of repeated scanning, often necessary in life science research. To investigate the potential photobleaching effect of MitoView 633, cardiomyocytes were loaded with various concentrations of MitoView 633 and subjected to repeated XY scanning without external stress. Confocal imaging results revealed that at a concentration of 10 nmol/L, the fluorescence intensity of MitoView 633 remained relatively stable throughout the entire scanning period (80 scans, approximately 250 s) (Figure 3C). The slope coefficient of −8.17 ± 0.37 × 10⁻⁵ indicated no significant change. At a concentration of 25 nmol/L, the fluorescence of MitoView 633 showed minimal variation (<2%) during the initial 10 scans but experienced a decrease of approximately 9% after 80 repeated scans (slope coefficient: −1.32 ± 0.42 × 10⁻⁴). At 50 nmol/L, the photobleaching effect became more noticeable, resulting in an approximately 15% reduction in fluorescent intensity by the end of repeated scans (slope coefficient: −1.96 ± 0.45 × 10⁻³). These observations emphasize the need for careful consideration of MitoView 633 concentration to mitigate significant photobleaching. It's noteworthy that the extent of photobleaching appears to be influenced by cell type. As illustrated in Figure 3D, the fluorescence of MitoView 633 in H9C2 cells decreased by nearly 18% after 40 scans (slope coefficient: −4.42 ± 0.23 × 10⁻³), whereas in cardiomyocytes, the reduction was less than 9% (slope coefficient: −2.33 ± 0.16 × 10⁻³). Consequently, thorough characterization of this fluorescent probe is crucial when applying it to new cell types.

We next examined the effect of temperature or pH on MitoView 633 fluorescence and compared it with TMRM. A microincubation system was used to control the temperature of cardiomyocytes loaded with MitoView 633 or TMRM. MitoView 633 and TMRM were recorded with confocal microscopy using the same imaging conditions as the temperature was increased incrementally from room temperature (25°C) up to 37°C, allowing time for the temperature to equilibrate across the incubation system. Whereas TMRM fluorescent intensity increased nearly linearly as temperature elevated, MitoView 633 fluorescence showed no significant change when temperature increased from 28°C to 37°C, and only ∼ 10% when increased from 25°C to 37°C (Figure 3E). This effect was approximately one-third of that observed with TMRM. The temperature changes similarly affected the peak intensity of these two dyes in cell free solution. Both the peak excitation and emission intensities of TMRM increased ∼6.7 folds when the temperature was elevated from 25°C to 37°C (Table 3). On the contrary, the peak intensities of MitoView 633 only changed about 4% with the same temperature fluctuations. This enhanced stability of MitoView 633 across the physiologically relevant temperature range could facilitate more sensitive measurements of mitochondrial ΔΨ_m at lower temperatures and potentially minimize confounding factors in scenarios where significant temperature fluctuations are encountered during recording. To assess sensitivity to pH change, cells loaded with either TMRM or MitoView 633 were immersed in Tyrode’s solution adjusted to a neutral pH of 7.4, followed by imaging with confocal microscopy. Subsequently, the chamber solution was switched to one with progressively lower (6.6) or higher (7.4) pH levels, allowing adequate time for cellular equilibration to the new pH before imaging. The data indicated that changes in pH exerted similar effects on TMRM and MitoView 633 fluorescence: the fluorescence intensity of both dyes decreased when pH was shifted away from neutrality, either towards acidity or alkalinity (Figure 3F). These observations underscore the importance of exercising caution when measuring ΔΨ_m with either MitoView 633 or TMRM under conditions where pH alterations may occur.

We next investigated the potential impact of MitoView 633 on cellular bioenergetics. We exposed isolated adult cardiomyocytes to varying concentrations (ranging from 0 to 200 nmol/L) of MitoView 633 and subsequently measured cellular ATP content using the Cell Titer-Glo Luminescent Viability Assay. The results, as depicted in Figure 3G, revealed no significant difference in ATP levels among the tested groups, even at the higher concentration of 200 nmol/L. These findings underscore the notion that MitoView 633 staining exerts minimal influence on the cellular respiratory chain and overall energetics. Furthermore, we extended our investigation to AC16 cells, a proliferating human cardiomyocyte cell line derived from the fusion of primary cells from adult human ventricular heart tissues (Davidson et al., 2005). As an immortalized cardiomyocyte cell line, AC16 cells are more glycolytic, and the energy requirements and other physiological responses (e.g., relative stress tolerance) are different from adult cardiomyocytes that have preserved myofibrils, t-tubules, and gap junctions. Hence, we also assessed the effects of MitoView 633 staining on AC16 cells. In line with our observations in adult cardiomyocytes, the loading of MitoView 633 did not induce significant alterations in intracellular ATP concentrations within AC16 cells (Figure 3H). Altogether, our data indicate that MitoView 633 can be effectively employed as an indicator for ΔΨ_m measurement in live cell imaging without evident effect on overall cellular energetics. It should be noted that previous studies revealing that TMRM or TMRE moderately inhibits mitochondrial respiration were performed with isolated mitochondria; in intact cells, we found that TMRM loading, at a concentration below 100 nmol/L, had a very small effect on cellular ATP (less than 4%).

FCCP is a potent uncoupler of mitochondrial oxidative phosphorylation, which disrupts ATP synthesis by transporting protons across the mitochondrial inner membrane, interfering with the proton gradient (Perry et al., 2011). Our subsequent investigation focused on assessing how MitoView 633 responds to FCCP treatment and comparing its dynamics with that of TMRM. To accomplish this, isolated adult cardiomyocytes were co-stained with both TMRM and MitoView 633 (Figure 4A) and subjected to FCCP (10 μmol/L) perfusion.

Simultaneous confocal imaging of TMRM and MitoView 633 revealed a similar decline in their fluorescence following FCCP treatment, with a reduction of approximately 70% occurring over a span of about 6 min (Figure 4B). In a separate experiment, cardiomyocytes loaded solely with MitoView 633 were perfused with varying concentrations of FCCP. The confocal recordings demonstrated that while MitoView 633 fluorescence intensity dropped by 70% with 10 μmol/L FCCP, consistent with data shown in Figure 3B, its fluorescence diminished only 25% when the FCCP concentration was reduced to 1 μmol/L (Figure 4C). The concentration-dependent responses of FCCP on cardiomyocyte ΔΨ_m measured via MitoView 633 are consistent with those measured by TMRM (Zhao et al., 2013). Interestingly, our data unveiled a cell-type-dependent effect of FCCP. As illustrated in Figure 4D, mitochondria in H9C2 cells depolarized significantly faster (time constant: 40 s vs. 19,291 s) and to a greater extent (93% vs. 16%) when treated with the same concentration of FCCP (1 μmol/L) compared to adult cardiomyocyte mitochondria. The cell type-dependent effect of FCCP is supported by the studies by Johnson-Cadwell et al. showing that FCCP caused rapid and substantial mitochondrial depolarization in cultured cerebellar granule neurons at a concentration as low as 0.25 μmol/L (Johnson-Cadwell et al., 2007). Altogether, our data indicate that MitoView 633 can be used as a reliable ΔΨ_m indicator for measuring stress-induced mitochondrial depolarization.

To explore the compatibility of MitoView with other fluorescent dyes of interest, adult cardiomyocytes were co-stained with Fluo4-AM, MitoSOX Red, and MitoView 633 (Figure 5A), and continuously perfused with Tyrode’s solution. A selected cell was subjected to local laser flash (indicated by the yellow square in Figure 5A) to induce regional oxidative stress, as previously described (Goh et al., 2016), and was subsequently imaged over time. The analysis of the data revealed that ΔΨ_m within the flash zone experienced a rapid and substantial depolarization, while in the distant zone, it remained nearly unchanged (Figure 5B). Accordingly, mitochondrial ROS levels significantly increased in the laser-flashed zone, as evidenced by the MitoSOX Red fluorescence (Figure 5B). Notably, the MitoSOX Red fluorescent intensity exhibited a gradual decrease after the initial burst (Figure 5C), likely attributed to substantial disruption of the electron transport chain. Fluo4-AM imaging depicted an elevation in local Ca²⁺ within the flashed zone (Figure 5D), which could be attributed to mitochondrial depolarization inhibiting mitochondrial Ca²⁺ uptake and buffering. The change in Fluo4-AM fluorescent intensity in the distant zone was less pronounced. This increase in Fluo4-AM fluorescence upon laser flash might also be linked to the elevation of ROS levels. As shown in Figure 5E, Ca²⁺ spark frequency in the flashed zone was significantly higher compared to that in the distant zone, confirming increased Ca²⁺ release. This effect aligns with previous reports on the stimulatory impact of mitochondrial-derived ROS on sarcoplasmic reticulum Ca²⁺ release and spark (Yan et al., 2008; Zhou et al., 2011). The ability of MitoView 633 to be used in conjunction with MitoSOX Red could prove invaluable for laboratories employing conventional filter-based fluorescent microscopes, as the spectral characteristics of TMRM largely overlap with those of MitoSOX Red. It's worth noting that while the local flashes triggered regional mitochondrial depolarization in cardiomyocytes in our study, other reports have indicated that this approach can induce cell-wide (Papanicolaou et al., 2011), inter-cell mitochondrial depolarization (Slodzinski et al., 2008), or even sustained mitochondrial oscillations (Aon et al., 2003; Zhou et al., 2011; Goh et al., 2016), suggesting that this phenomenon is species-specific and cell status-dependent.