Simultaneous Voltage and Calcium Mapping of Genetically Purified Human Induced Pluripotent Stem Cell–Derived Cardiac Myocyte Monolayers

Detailed Methods are provided in the Online Supplement. Here, we use a novel optical mapping system for simultaneous V_m and ratiometric intracellular calcium ([Ca²⁺]_i) imaging of cardiac preparations, including human iPSC-CM monolayers. The optical mapping system presented here (Figure 1) offers several advantages over most existing systems for multiparametric imaging. First is the use of multiple light-emitting-diodes (LEDs) as excitation light sources, which provide a stable, flexible, and economical alternative to established methods. Unlike traditional light sources, LED light intensity can be modulated with exceedingly fast response times, thus making high-speed excitation light switching possible without the need for moving parts. Based on recent advances in LED fabrication technology, powerful LEDs are now available that cover the entire spectrum from deep ultraviolet to infrared, making them an increasingly attractive alternative for multiparametric cardiac imaging.⁵ Second, the imaging method presented here provides a means for quantitative (ratiometric) calcium wave imaging in cardiac tissue of various levels of structural complexity. The use of fluorescence for quantitative measurement is difficult because fluorescence intensity is affected by a range of ill-controlled factors, such as regional differences in dye loading, internalization in subcellular compartments, and photo-bleaching. These limitations are particularly relevant when utilizing single-excitation/single-emission fluorescent dyes. To address this, ratiometric Ca²⁺ dyes are utilized that, on binding Ca²⁺, display spectral shifts with opposite polarity on exposure to different excitation wavelengths.⁶ The ratio of the two emission intensities is independent of fluorescence signal intensity, thereby providing a means of quantifying [Ca²⁺]_i and allowing reliable comparison between regions and samples.⁶ Third, we have utilized a low-affinity ratiometric Ca²⁺ dye, fura-4F. To minimize perturbation of the [Ca²⁺]_i dynamics in cardiac tissue, the choice of Ca²⁺ dye is critical for acquiring accurate measurements of the amplitude and time course of [Ca²⁺]_i transients. For cardiac myocytes and tissue, which show large and rapid changes in [Ca²⁺]_i, a low-affinity and rapidly responding dye is needed.⁷ Other commonly used ratiometric Ca²⁺ dyes, such as fura-2, have a relatively high-affinity for Ca²⁺. This can lead to an artificial prolongation of the Ca²⁺ transient and confound interpretation (ie, the dye acts as a chelator that releases Ca²⁺ with delay). Fourth, the new system uses a single sensor with a multiband transmission filter and no moving parts. The use of a single sensor obviates the technical complexity of multisensor alignment for simultaneous multiparametric imaging.⁸ More complete details of the optical mapping set-up and electronics are available in the Online Supplement.

We have thoroughly validated the performance of this optical mapping system using cardiac tissue of various levels of structural complexity from monolayers of cardiac myocytes to native cardiac tissue slices and the whole heart (Figures 2, 3). Details about each cardiac preparation can be found in the Online Supplement. Figure 2A demonstrates the utility of the system for studying cardiac arrhythmia mechanisms in neonatal rat ventricular myocyte monolayers, a commonly used in vitro model.⁹ Using our mapping system, we were able to monitor simultaneously V_m and [Ca²⁺]_i wave propagation of electric rotors (re-entrant electric activity that can underlie cardiac arrhythmias; Online Movie I). Comparison of the data, also illustrated in the 2 snapshots presented in Figure 2A, reveals that the rotating V_m wavefront precedes the Ca²⁺ wavefront by approximately 10 ms. Next, we demonstrate simultaneous V_m and [Ca²⁺]_i imaging of thin (≤350-μm-thick) ventricular tissue slices (Figure 2B; Online Movie II). Cardiac tissue slices represent an exciting pseudo-two-dimensional model that balances structural complexity (cells within the slice are in an environment that is as close to native as possible in vitro) and ease of data interpretation (observed signals can be related more reliably to their histological sources).^10,11 As illustrated in Figure 2C, we also used the same imaging system to map V_m and [Ca²⁺]_i wave propagation in Langendorff-perfused rat hearts (Online Movie III). Normalized fluorescence intensity maps for V_m and [Ca²⁺] were recorded during sinus rhythm and are shown at two time points 9.72 ms apart. The entire anterior epicardial surface of the ventricles activated almost simultaneously, whereas the peak of [Ca²⁺]_i occurred with a delay of approximately 16 ms.

Next, we generated human cardiac monolayers to enable the simultaneous mapping of V_m and [Ca²⁺]_i wave propagation in a human model system. Human iPSC-CMs were obtained from Cellular Dynamics International (Madison, WI). The iCell cardiomyocytes are highly purified human cardiac myocytes (>98% pure cardiomyocytes) derived from iPS cells using cardiac-directed differentiation and purification protocols.¹² Importantly, the high purity of these cardiomyocyte cultures remains unchanged over the course of at least 2 weeks in culture. The iCell cardiomyocytes are differentiated from human iPS cells reprogrammed from a nonembryonic terminally differentiated cell source, thus avoiding the ethical controversy surrounding embryonic stem cell use. Cryopreserved vials (liquid nitrogen) of iCell human cardiac myocytes were thawed and subsequently plated on bovine fibronectin-coated (20 μg/mL; Invitrogen) transparent silastic membranes at a density of 125,000 cells per monolayer in differentiation media¹³ (embryoid body differentiation media, commonly referred to as embryoid body-20, comprising 80% Dulbecco Modified Eagle Medium [DMEM/F12], 0.1 mmol/L⁻¹ nonessential amino acids, 1 mmol/L⁻¹ L-glutamine, 0.1 mmol/L⁻¹ β-mercaptoethanol, and 20% fetal bovine serum; Gibco) supplemented with 10 μmol/L⁻¹ blebbistatin. After 24 hours in embryoid body-20,¹³ the medium was switched to iCellmaintenance medium, supplemented with 10 μmol/L⁻¹ blebbistatin, and cells were cultured for an additional 96 hours at 37°C, in 5% CO₂, with the medium changed once daily. Monolayers (n=4) were subsequently processed for electrophysiological analysis by optical mapping as described for neonatal rat cardiomyocyte monolayers or for immunocytochemistry analysis (n=7). The cellular structure and composition of typical monolayers generated in this study are presented in Figures 3 and 4.

Cardiomyocytes were plated on fibronectin-coated transparent silastic membranes as described. Immunostaining was performed essentially as described recently.¹² For immunostaining, cells were washed with phosphate-buffered saline, fixed with 4% paraformaldehyde for 10 minutes, and rinsed twice before blocking with 5% donkey serum in phosphate-buffered saline plus 0.1% Triton X-100 (Sigma) for 1.5 hours at room temperature. Primary antibodies including mouse monoclonal anti-MLC-2a (IgG2b, 1:200 dilution; Synaptic Systems), rabbit polyclonal anti-MLC-2v (IgG, 1:100 dilution; ProteinTech Group), mouse monoclonal α-actinin (1:500; Sigma), and rabbit polyclonal connexin 43 (1:100; Millipore) were added to 5% donkey serum in phosphate-buffered saline plus 0.1% Triton X-100 and incubated overnight at 4°C with constant agitation. Subsequently, samples were washed three times in phosphate-buffered saline plus 0.1% Triton X-100. The secondary antibodies donkey antirabbit DyLight 488 and donkey antimouse DyLight 594 (1:500 dilution; Jackson ImmunoResearch) were diluted in the same solution as primary antibodies and incubated at room temperature in the dark for 1.5 hours. Negative control experiments using only the secondary antibodies were performed to ensure specificity of immunostaining and fluorescence detection. Samples were then washed three times with phosphate-buffered saline plus 0.1% Triton X-100 and once with phosphate-buffered saline only. Finally, nuclei were stained with DAPI (1:1000 dilution; Invitrogen) for 10 minutes at room temperature in the dark. Cover slips were mounted on microscope slides for confocal imaging.

Immunofluorescence imaging was performed using a 20× or 60× objective on a Nikon A1R laser scanning confocal microscope system (Nikon Instruments, Melville, NY). Figure 3 shows the cellular and subcellular structure of representative cardiac monolayers that were used in this study. Figure 3A shows a cardiomyocyte monolayer immunostained for α-actinin (red) and nuclei (DAPI, blue). Consistent with a recent report using iCell cardiomyocytes,¹² nuclei were evenly distributed and α-actinin positive staining was present throughout the field of view. Higher magnification (60×) image in Figure 3B shows the sarcomere structure of the Z-line protein α-actinin (red) and also shows that the gap junction protein connexin 43 (green) is localized at the points of myocyte–myocyte contact. This suggests that impulse propagation in these human monolayers may be mediated through cell-to-cell communication via connexin 43. Figures 3C to 3E show the result of coimmunolabeling for α-actinin (sarcomeric Z-line) and MLC-2v (sarcomeric A-band). Figure 3C demonstrates that not all myofilaments stained positive for the ventricular-specific myosin light chain (MLC-2v). For MLC-2v–positive myofilaments, alternating Z-line and A-band staining was observed (Figure 3D). A fluorescence intensity plot over the distance denoted by the white arrow in Figure 3D is shown in Figure 3E. Proper spatial organization of the sarcomeric Z-line (red) and A-band (green, MLC-2v staining is localized in each half of the sarcomeric A-band) suggests a degree of functional maturity of the cardiac sarcomere.¹⁴ The average sarcomere length, measured by multiple single measures of the Z-line to Z-line distance, is 1.98±0.03 μm (n=51 myocytes) for myocytes in these monolayers (Figure 3E).

Next, we used immunofluorescence and flow cytometry to quantify the cardiomyocyte population in iCell cardiomyocyte monolayers. Expression of the two cardiac MLC-2 isoforms of the heart, MLC-2a and MLC-2v, provides information about the diversity of the cardiomyocyte population. The MLC-2a expression is detected in all chambers of developing mouse and human hearts. Postnatally, MLC-2a expression is restricted to the atria in mice, whereas MLC-2a expression is found in both the atria and, to a lesser extent in the ventricles in humans.^15,16 However, MLC-2v expression is restricted to the ventricular chambers in humans, and this chamber specificity persists into adulthood.^15,16 In mouse and human embryoid bodies, MLC-2a expression precedes MLC-2v expression, thereby suggesting that MLC-2v expression is also a marker of myocyte maturity.^14,15 Immunofluorescence and laser confocal scanning analysis demonstrate that these monolayers are comprised of at least three phenotypes expressing primarily MLC-2a (red), MLC-2v (green), and both MLC-2a and MLC-2v (yellow in Figure 4A, B). Cells expressing MLC-2v likely represent ventricular-like cells, whereas MLC-2a–only expressing cells can identify a range of potential cardiomyocyte types, including atrial-like cells.

Results of flow cytometry analysis using MLC-2a–specific and MLC-2v–specific antibodies are shown in Figures 4C and 4D. For flow cytometry, cells were washed with phosphate-buffered saline and then incubated with 1 mL/well of 0.25% Trypsin-EDTA plus 2% chicken serum for 5 minutes. Equal volume of embryoid body-20 medium, containing 20% fetal bovine serum, was subsequently used to inhibit trypsinization. Cells were detached from the cover slips and aggregates were disrupted to singularize the cells by pipetting up and down. Cell suspensions were centrifuged for 5 minutes at 1,000 rpm; the supernatant was discarded and cell pellets were resuspended in 1 mL 1% paraformaldehyde and incubated at 37°C for 10 minutes to fix the cells. Samples were centrifuged and pellets were resuspended in 1 mL ice-cold methanol before incubation on ice for 30 minutes to permeabilize the cells. Cells were washed once with FACS buffer (phosphate-buffered saline without Ca²⁺/Mg²⁺, 1% bovine serum albumin) plus 0.1% Triton and centrifuged, and the supernatant was discarded, leaving a volume of 100 μL. Primary antibodies for MLC-2a (Synaptic Systems, mouse monoclonal, dilution 1:200) and MLC-2v (ProteinTech Group, rabbit polyclonal, dilution 1:400) were dissolved in 100 μL/sample FACS buffer plus 0.1% Triton for total sample volume of 200 μL. Samples were incubated with primary antibodies overnight at 4°C. Cells were washed twice in 3 mL FACS buffer plus 0.1% Triton and centrifuged, and supernatant was discarded, leaving approximately 100 μL. Secondary antibodies (donkey antimouse Dylight 488 and donkey antirabbit Dylight 594, dilution 1:500; Jackson ImmunoResearch) were dissolved in 100 μL/sample FACS buffer plus 0.1% Triton for total sample volume of 200 μL and incubated for 45 minutes in the dark at room temperature. Cells were washed twice in FACS buffer plus 0.1% Triton, centrifuged, resuspended in 200 μL FACS buffer plus 0.1% Triton, and stored on ice until FACS analysis. To define the thresholds for positive 488 and 594 fluorescence, negative control samples were incubated with 488 or 594 secondary antibodies only. Black peaks in Online Figure I represent both the background fluorescence of the secondary antibodies and the autofluorescence of the cells. Green and red peaks show, respectively, 488 and 594 fluorescence for positive control samples (Online Figure I). Both negative and positive control samples allow gating precisely for 488 and 594 positive fluorescence thresholds. Data were collected on a FACS system (MoFlo XDP) and analyzed using FlowJo version 9.4.11. Flow cytometry confirms the immunofluorescence staining of MLC-2a and MLC-2v as shown in Figures 4C and 4D. Three distinct populations of CMs were identified: MLC-2a⁺/MLC-2v⁻ (upper left quadrant, 8.4%); MLC-2a⁺/MLC-2v⁺ (upper right quadrant, 13.6%); and MLC2a-/MLC2v⁺ (lower right quadrant, 44.6%). This is similar to the subpopulations of atrial and ventricular myocytes identified using electrophysiology criteria of action potential (AP) morphology of iCell cardiomyocytes.¹²

We generated large (approximately 1-cm diameter) human iPSC-CM monolayers for optical mapping using genetically selected iPSC-CMs (iCell; Cellular Dynamics International). These iPSC-CM monolayers exhibit spontaneous pacemaker activity (1.22±0.09 Hz; n=4; Figure 5A and Online Movie IV), and they also can be electrically stimulated (Figure 5A and Online Movie V). Uniform propagation of V_m and [Ca²⁺]_i through the entire monolayer was observed in each case (n=4), as shown in the representative activation maps of Figure 6 (conduction velocity=21.9±2.47 cm/s⁻¹ and a V_m to [Ca²⁺]_i peak delay of approximately 33 ms). The V_m activation maps of spontaneous pacemaker activity recorded in two separate monolayers are shown in Figures 6A and 6C. Figures 6B and 6D show activation maps of paced beats in where the stimulator was placed approximately at the center of the monolayer, and where the V_m wave spread uniformly through the monolayer in all directions.

Figure 7 shows the uniformity of AP and [Ca]_i characteristics across the monolayer. The action potential duration (APD) at 90% repolarization (APD₉₀) and the [Ca]_i transient duration at 90% return to baseline (CaT₉₀) are shown from four spatially distinct locations during spontaneous and electrically paced propagation (indicated by blue, red, green, and cyan squared regions). The right panel of Figure 7A presents APD₉₀ and CaT₉₀ during spontaneous activity (cycle length of 1,267±12 ms), the left panel of Figure 7B presents APD₉₀ and CaT₉₀ during electrically paced activity (cycle length of 500 ms), and the right panel of Figure 7B presents APD₉₀ and CaT₉₀ during electrically paced activity (cycle length of 750 ms). As can be seen, the transient duration characteristics for each parameter are uniform across the plate and increase with cycle length. Three consecutive transients, after equilibration, were taken for each data point. The same analysis for another representative monolayer can be found in Online Figure II.