Abstract
Human embryonic stem cells (hESCs) and induced pluripotent stem cells (hiPSCs) can differentiate to cardiomyocytes in vitro, offering unique opportunities to investigate cardiac development and disease as well as providing a platform to perform drug and toxicity tests. Initial cardiac differentiation methods were based on either inductive co-culture or aggregation as embryoid bodies, often in the presence of fetal calf serum. More recently, monolayer differentiation protocols have evolved as feasible alternatives and are often performed in completely defined culture medium and substrates. Thus, our ability to efficiently and reproducibly generate cardiomyocytes from multiple different hESC and hiPSC lines has improved significantly.
We have developed a directed differentiation monolayer protocol that can be used to generate cultures comprising ~50 % cardiomyocytes, in which both the culture of the undifferentiated human pluripotent stem cells (hPSCs) and the differentiation procedure itself are defined and serumfree. The differentiation method is also effective for hPSCs maintained in other culture systems. In this chapter, we outline the differentiation protocol and describe methods to assess cardiac differentiation efficiency as well as to identify and quantify the yield of cardiomyocytes.
Keywords: Human embryonic stem cells, Human induced pluripotent stem cells, Cardiac differentiation, Cardiomyocyte characterization
1.Introduction
The differentiation of human embryonic stem cells (hESCs) and induced pluripotent stem cells (hiPSCs) into cardiomyocytes has provided exciting opportunities to examine these cells which otherwise can only be obtained through invasive biopsies of patients. These cardiomyocytes can be used not only to study heart development and diseases but also to perform drug and toxicity testing . This type of safety pharmacology is important because the heart is remarkably sensitive to the side effects of drugs and other compounds. The generation of hiPSCs from patients with genetic cardiac diseases also allows for the production of cardiomyocytes that capture the genome of the individual, offering the prospect to model these diseases in vitro. These patient specific cells can be used to study the mechanisms underlying the disease pathogenesis and as platforms to explore new therapies.
One of the first protocols to generate cardiomyocytes from hESCs was the co-culture of these cells on mouse visceral endoderm like cells (END-2). Although this method is effective for many human pluripotent stem cell lines (hPSCs), the yield of cardiomyocytes is low and is only effective with mechanically passaged hPSC lines on mouse embryonic fibroblasts (MEFs). The adaptation of hESCs to single cell enzymatic passaging has provided alternative approaches to derive cardiomyocytes. One of these involves forming small aggregates of enzymatically adapted hPSCs in suspension by the centrifugation of these undifferentiated cells in low attachment 96-well plates.
This method, commonly referred to as either “forced aggregation” or “spin embryoid bodies,” has significantly increased the yield of cardiomyocytes in culture as the initial number of hPSCs in each aggregate is controlled and identical across all wells. These protocols necessitate that the culture medium is serum free and is usually a chemically defined medium that contains albumin, polyvinyl alcohol, and essential lipids. An animal component free version of this basal medium containing only recombinant human proteins (APEL medium) is also commercially available. Alternatively, a micro-textured surface, also known as micro or Aggrewells, has provided a scalable method for the initial formation of embryoid bodies of uniform size.
Early protocols to generate cardiomyocytes using this technology required the use of serum, but the hPSCs can now be differentiated in serum free medium with the addition of growth factors. Since 2012 significant progress has also been made with monolayer based differentiation protocols leading to further improvements in efficiency and homogeneity of the cardiomyocytes generated, and this is rapidly becoming the preferred cardiac differentiation approach.
Common among all of these serum free methods is the sequential mimicking in culture of key embryonic developmental signals to induce cardiac differentiation. Signals induced by nodal (using Activin A as a substitute), bone morphogenic proteins (BMPs), Wnts, and fibroblast growth factors (FGFs) are first required to generate cardiac mesoderm like cells.
Subsequent inhibition of the Wnt/β-catenin pathway, initially by Dickkopf-1 (Dkk-1), is then required for cardiac specification . Recently it has been shown that small molecules that lead to the activation and inhibition the Wnt/β-catenin pathway can replace some or all of the cytokines previously used. However, the success of these small molecule based methods is both highly concentration and time dependent, can vary between different cell lines, and is also sensitive to the method by which the undifferentiated cells are maintained.
In this chapter, we describe a monolayer based protocol for the generation of cardiomyocytes from hPSCs that have been maintained undifferentiated using a range of different approaches: either as single cell passaged cultures on MEFs or in completely defined Essential 8 (E8) medium; or as mechanically passaged aggregates in mTeSR1 medium or on MEFs. We use BMP4 and Activin A together with Laduviglusib CHIR-99021 GSK-3 inhibitor axon, a glycogen synthase kinase (GSK-3) small molecule inhibitor that activates Wnt/β-catenin signaling, to induce cardiac mesoderm. This is followed by the temporal treatment with the tankyrase inhibitor XAV939, which inhibits Wnt/β-catenin signaling and leads to the efficient generation of cardiac progenitors.
Approximately 1 week after initiating the differentiation, we observe spontaneously contracting cardiomyocytes. Combining both cytokines and small molecules in our differentiation protocol has proven to be very robust in our laboratory, producing high yields of cardiomyocytes from multiple hPSC lines. These cardiomyocytes can be used in many different assays including electrophysiology, optical mapping and force of contraction measurements for functional analysis, cell imaging and immunohistochemistry for structural and phenotypic analysis, as well as next generation sequencing and mass spectrometry for global transcriptome and proteome analysis. We also describe here three established protocols to identify and quantify cardiomyocytes within the differentiated culture.
2.Materials
For culture recipes and steps, use sterile tissue culture grade water. For other protocol steps, use deionized water or equivalent. All cells are maintained and differentiated in a 37 ◦C humidified 5 % CO2 incubator.All differentiations are performed using 12 or 6 well tissue culture plates.
2.1Cell Culture Medium and Reagents for Differentiation and Dissociation
Phosphate buffered saline (PBS; Gibco) without CaCl₂ and MgCl₂ was used in combination with 0.5 mM EDTA (Invitrogen) prepared in PBS. Dimethyl sulfoxide (DMSO; Sigma Aldrich) served as the solvent for small molecules. Human FGF-2 (100 ng/mL; Miltenyi Biotec) was reconstituted from lyophilized powder following the manufacturer’s instructions, kept cold at all times, and stored as aliquots at −80 °C for long-term storage of up to one year, while thawed aliquots were stored at 4 °C for up to two weeks.
Recombinant human Activin A (25 ng/μL; R&D) and recombinant human BMP4 (25 ng/μL; R&D) were also reconstituted according to the manufacturer’s protocols, kept cold, and stored as aliquots at −80 °C for up to one year, with thawed aliquots stable at 4 °C for up to two weeks. CHIR99021 (4 mM; Axon Medchem) was prepared by dissolving powder in DMSO, and aliquots were stored at −20 °C for up to one year, with thawed aliquots stored at 4 °C for up to three weeks. Similarly, XAV939 (5 mM; Tocris) was dissolved in DMSO, stored as aliquots at −20 °C for long term storage, and kept at 4 °C for up to three weeks after thawing.
Human embryonic stem cell (hESC) medium consisted of DMEM/F12 (Gibco) supplemented with 20% KnockOut™ Serum Replacement (Gibco), 10 mM non-essential amino acids (Gibco), 2 mM L-glutamine (Gibco), 50 mM 2-mercaptoethanol (Gibco), 0.5% penicillin/streptomycin (25 units penicillin G and 25 μg streptomycin sulfate; Gibco), and 10 ng/mL FGF-2. The medium was filter sterilized and stored at 4 °C for up to two weeks. Additional media included mTeSR™1 (STEMCELL Technologies) and Essential 8™ medium (Gibco). Dispase (1 mg/mL; Gibco) was prepared by first dissolving powder in DMEM/F12 to obtain a 5 mg/mL solution, which was aliquoted and stored at −20 °C for up to one year. Thawed aliquots were diluted to 1 mg/mL in DMEM/F12 and stored at 4 °C for up to two weeks. TrypLE™ Select (1× or 10×; Gibco) was used as required, with the 10× stock diluted to 2× or 5× with PBS containing 1 mM EDTA.
Bovine serum albumin (BSA) stock solution (10% w/v; Bovostar, Bovogen Biologicals) was prepared by dissolving 4.5 g of BSA in 35 mL of IMDM at 37 °C in a 50 mL conical tube with vortexing, followed by adjusting the volume to 45 mL with IMDM. The solution was filter sterilized and stored at 4 °C for up to three months. α-Monothioglycerol (α-MTG; Sigma Aldrich) was prepared as a 150 mM solution by adding 13 μL of α-MTG to 1 mL of IMDM, and stored at 4 °C for up to four weeks. The differentiation medium consisted of a 1:1 mixture of Iscove’s Modified Dulbecco’s Medium (IMDM; Gibco; L-glutamine, 25 mM HEPES, no phenol red) and Ham’s F12 Nutrient Mixture with GlutaMAX (Gibco).
This basal medium was supplemented with 5% (v/v) Protein Free Hybridoma Medium (PFHM-II; Gibco), 0.25% (w/v) BSA stock solution, 1× Chemically Defined Lipid Concentrate (100× stock; Gibco), 0.1× Insulin-Transferrin-Selenium-X Supplement (100× stock; Gibco), 450 μM α-MTG, 0.05 mg/mL L-ascorbic acid 2-phosphate (Sigma Aldrich), 2 mM GlutaMAX supplement (100× stock; Gibco), and 0.5% penicillin/streptomycin (25 units penicillin G and 25 μg streptomycin sulfate). The differentiation medium was filter sterilized and stored at 4 °C for up to two weeks or at −20 °C for up to six months, and fresh medium less than one week old was used for initiating differentiation.
Matrigel™ Matrix Growth Factor Reduced (BD) was prepared by diluting 0.5 mg of Matrigel in 6 mL of cold DMEM/F12. Using a chilled pipette, the solution was dispensed into 6 or 12 well plates (1 or 0.5 mL per well, respectively) and left to polymerize at room temperature for at least 45 minutes. Plates were either used immediately or stored at 4 °C for up to two weeks, while Matrigel stock solutions were stored as 0.5 mg aliquots at −20 °C. Finally, dilution medium was prepared using DMEM/F12 supplemented with 10% (v/v) fetal bovine serum (FBS; Gibco).
2.2Flow Cytometry
The wash buffer consisted of 2% (v/v) fetal bovine serum (FBS) in PBS. The blocking solution was prepared with PBS containing 2% (v/v) FBS, 2% normal rabbit serum (Sigma Aldrich), and 1% normal goat serum (DAKO). This solution could be stored at 4 °C for up to three months or at −20 °C indefinitely. Fixation and permeabilization were performed using the FIX & PERM Cell Permeabilization Kit (Invitrogen). For nuclear staining, 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen) was prepared at 0.1 mg/mL in wash buffer, while propidium iodide (PI; MP Biomedicals) was prepared at 1 mg/mL in deionized water. Antibodies specific for cardiac Troponin T (TNNT2) detection are summarized in Table 1 (see Notes 2 and 3), and antibodies for SIRPA and VCAM1 detection are listed in Table 2.
2.3Immunofluorescence
The fixation solution consisted of 2% paraformaldehyde in 0.2 M phosphate buffer (pH 7.4). Permeabilization was carried out using 0.1% Triton X-100 (Sigma Aldrich) in PBS. For blocking, a freshly prepared solution of 4% (v/v) swine serum (DAKO) in PBS was used. Washing steps were performed with 0.05% (v/v) Tween-20 (Merck) in PBS. Mounting was carried out using Mowiol 4-88 (Calbiochem), prepared according to the manufacturer’s instructions. DAPI was used as a nuclear stain from a 5 mg/mL stock solution in deionized water. The antibodies employed for immunofluorescence are listed in Table 3.
3.Methods
3.1Seeding hPSCs for Differentiation
From Mechanically Passaged hPSC Cultures Maintained on MEFs.Human pluripotent stem cells (hPSCs) were maintained as colonies on irradiated mouse embryonic fibroblasts (MEFs) in hESC medium as previously described. One to two days before initiating differentiation, each colony was cut into approximately 15 pieces using a 26-gauge needle and carefully dislodged from the dish. The resulting pieces were transferred in hESC medium onto a Matrigel coated culture dish at a density of about 10 pieces per 4 cm² (Fig. 1a). The hESC medium was refreshed daily. Once the culture reached approximately 70% confluence, differentiation was initiated following the protocol described in Section 3.2.
3.1.2From Enzymatically Passaged hPSC Cultures Maintained on MEFs
Human pluripotent stem cells (hPSCs) were maintained on irradiated mouse embryonic fibroblasts (MEFs) in hESC medium as previously described. The day before differentiation (day 0 minus 1), the hPSC culture was adjusted to approximately 85% confluence. To harvest the cells, the hESC medium was aspirated, the culture was rinsed once with PBS, and enough 1× TrypLE Select was added to cover the surface.
The culture was incubated for 5 minutes at 37 °C, after which five volumes of hESC medium were added. The cells were gently dissociated into single cells by pipetting up and down 3–5 times with a 5 mL Pasteur pipette. The cell suspension was then centrifuged for 3 minutes at 250 × g to pellet the cells. The supernatant was aspirated, and the cells were resuspended in hESC medium. Cells were plated onto Matrigel coated plates at a density designed to reach approximately 40% confluence the following day (Fig. 1b). Differentiation was then initiated according to the protocol described in Section 3.2.
3.1.3From hPSC Cultures Maintained in E8 Medium
Human pluripotent stem cells (hPSCs) were maintained in E8 medium on tissue culture dishes
precoated with vitronectin as previously described. Three to four days before differentiation, the hPSC culture was adjusted to approximately 85% confluence. To harvest the cells, the E8 medium was aspirated, the culture was rinsed once with PBS, and 0.5 mM EDTA was added to cover the surface area. The culture was incubated for 5 minutes at 37 °C, and the EDTA was aspirated once the colonies rounded up and small gaps began to appear. Sufficient E8 medium was then added to the dish, and the hPSCs were dissociated into small clusters by gently pipetting up and down 3–5 times with a 5 mL Pasteur pipette. The cells were seeded in E8 medium onto Matrigel coated culture dishes at a density chosen to reach 70–90% confluence within 72–96 hours after passaging (Fig. 1c).Refresh the E8 medium 48 h after passaging and repeat daily. Proceed to the differentiation protocol.
Fig. 1 Bright field images of hPSC morphology when seeded on Matrigel coated dishes from (a) mechanically passaged hPSC cultures maintained on MEFs, (b) enzymatically passaged hPSC cultures maintained on MEFs,(c) hPSC cultures maintained in Essential 8 medium, (d) hPSC cultures maintained in mTeSR1. Scale bar: 500 μm
3.1.4From hPSC Cultures Maintained in mTeSR1
Human pluripotent stem cells (hPSCs) were maintained in mTeSR1 on Matrigel coated tissue culture dishes following the manufacturer’s protocol. Three to four days before differentiation, the hPSC culture was adjusted to approximately 80% confluence. Each colony was cut into four pieces using a 26-gauge needle. To harvest the cells, the mTeSR1 medium was aspirated, and dispase was added to cover the surface. The culture was incubated for 5 minutes at 37 °C, and the dispase was aspirated once the colonies had rounded up. The well was rinsed twice with twice the volume of DMEM/F12 medium and aspirated. Sufficient DMEM/F12 was then added to dislodge the colony pieces from the dish, and the pieces were transferred to a Matrigel coated culture dish at a density of approximately 25–30 pieces per 4 cm² in mTeSR1 medium, ensuring the culture would reach about 80% confluence within 72–96 hours after passaging (Fig. 1d). The medium was refreshed daily with mTeSR1 until the start of differentiation, after which the protocol described in Section 3.2 was followed.
3.2Differentiating hPSCs to Cardiomyocytes
On day 0 of differentiation, the hPSC maintenance medium was aspirated and replaced with differentiation medium supplemented with 20 ng/mL Activin A, 20 ng/mL BMP4, and 1.5 μM CHIR99021 (Fig. 2). For a 12-well culture plate, 1.5 mL of differentiation medium was added per well, while for a 6 well plate, 3.5 mL was added per well. By day 3 of differentiation, the cells formed a monolayer covering the entire surface of the well (Fig. 2). The medium was aspirated and replaced with an equal volume of differentiation medium supplemented with 5 μM XAV939. On day 6, the medium was again aspirated and replaced with fresh, unsupplemented differentiation medium. This medium was refreshed every 3–4 days as described, and between days 7 and 9 of differentiation, spontaneously contracting areas typically appeared. Wells containing these contracting areas could be maintained for at least three months. These regions could be dissociated (see Sect. 3.3) to obtain cardiomyocytes for downstream assays such as flow cytometry and immunohistochemistry.
Fig. 2 Schematic representation of the protocol to differentiate hPSCs to cardiomyocytes. Bright field (BF) and green fluorescence (eGFP) images show the typical morphology of the differentiating cells at key stages during the differentiation. By day 7 of differentiation, web like colonies begin to appear that subsequently start to contract by day 10 of differentiation (upper panel of images). If using an NKX2-5–eGFP reporter hESC or hiPSC line (8) (van den Berg, Davis, unpublished), these contracting cells will be GFP+ (lower panel of images).
Scale bar: 500 μm
3.3Dissociating Cultures Containing hPSC-Derived Cardiomyocytes
The medium was aspirated, and cells were washed with 1 mL PBS per well for a 12 well plate or 3 mL PBS per well for a 6 well plate, followed by aspiration of the PBS. TrypLE Select was then added at 0.5 mL per well for a 12 well plate or 1 mL per well for a 6 well plate, and the culture was incubated for 5–15 minutes at 37 °C.
To detach the cells, the plate was gently tapped with the palm; if gaps appeared in the cell sheet or detachment occurred, a 1 mL pipette was used to gently pipette the suspension up and down 3–5 times. If cells had not begun to detach, the plate was returned to 37 °C for an additional 5 minutes and reassessed, repeating this process as needed until detachment occurred. The resulting single cell suspension was transferred to a 15 mL conical tube containing 5 mL of dilution medium, followed by centrifugation at 450 × g for 3 minutes at 4 °C. The supernatant was aspirated, and the cells were resuspended in cold wash buffer for flow cytometric evaluation (see Sect. 3.4 or 3.5) or in differentiation medium for immunohistochemical analysis (see Sect. 3.6).
3.4 Flow Cytometric Analysis for Cardiac TNNT2 (Intracellular Cardiac Marker)
The medium was aspirated, and the cells were washed with 1 mL PBS per well of a 12 well plate or 3 mL PBS per well of a 6 well plate, followed by aspiration of the PBS solution. TrypLE Select was added at 0.5 mL per well for a 12 well plate or 1 mL per well for a 6 well plate, and the culture was incubated for 5–15 minutes at 37 °C. To detach the cells, the plate was gently tapped with the palm; if gaps appeared in the cell sheet or detachment occurred, a 1 mL pipette was used to carefully pipette the suspension up and down 3–5 times. If the cells had not begun to detach, the plate was returned to 37 °C for an additional 5 minutes and reassessed, repeating this process until detachment was complete. The single cell suspension was transferred to a 15 mL conical tube containing 5 mL of dilution medium and centrifuged for 3 minutes at 450 × g at 4 °C.
The supernatant was aspirated, and the cells were resuspended in cold wash buffer for flow cytometric evaluation or in differentiation medium for immuno histochemical analysis. The cell suspension was then filtered using a 5 mL tube with a 35 μm cell strainer cap, and the cells were pelleted by centrifugation for 3 minutes at 450 × g at 4 °C. After aspirating the wash buffer, the cell pellet was resuspended in 100 μL Fixation Medium and incubated for 15 minutes at room temperature according to the manufacturer’s protocol. Subsequently, 3 mL of cold wash buffer was added, and the suspension was centrifuged for 3 minutes at 450 × g at 4 °C before aspirating the supernatant. The cell pellet was resuspended in 300 μL Permeabilization Medium by vortexing, and the suspension was evenly divided across three tubes, with 100 μL per tube.
3.5Flow Cytometric Analysis for SIRPA and VCAM1 (Cell Surface Markers)
The primary antibodies were diluted 1:10 in Permeabilization Medium, and 1 μL of the prediluted antibodies was added to each tube as indicated in Table 4, with tubes 1 and 2 serving as control samples. The tubes were briefly vortexed and incubated for 30 minutes at room temperature. Cells were washed by adding 3 mL of cold wash buffer to each tube, followed by centrifugation for 3 minutes at 450 × g at 4 °C, and aspiration of the supernatant. This washing step was repeated once.
Tube 1 was resuspended in 300 μL of wash buffer and kept cold until flow cytometry measurement. For tubes 2 and 3, 2 μL of secondary antibody was diluted in 200 μL Permeabilization Medium, and 100 μL of this solution was added to each tube. The tubes were briefly vortexed to resuspend the cell pellet and incubated for 20 minutes in the dark at room temperature. Cells were washed twice by adding 3 mL of cold wash buffer to each tube, centrifuging for 3 minutes at 450 × g at 4 °C, and aspirating the supernatant. The cell pellet was then resuspended in 300 μL of wash buffer, and the tubes were kept cold. The flow cytometer was set up using tubes 1 and 2 as instrument and gating controls, followed by flow cytometric evaluation of TNNT2 expression.
The cell suspension was filtered using a 5 mL tube with a 35 μm cell strainer cap, and the cells were pelleted by centrifugation for 3 minutes at 450 × g at 4 °C before aspirating the wash buffer. The cell pellet was resuspended in 500 μL of blocking buffer by vortexing, and the suspension was evenly divided across five tubes (100 μL per tube) labeled according to Table 5, with tubes 1–4 serving as control samples for flow cytometer setup.
The appropriate dilution of each antibody (Table 2) was added to the tubes, followed by brief vortexing and a 20-minute incubation in the dark on ice. Cells were washed twice by adding 3 mL of cold wash buffer to each tube, centrifuging for 3 minutes at 450 × g at 4 °C, and aspirating the supernatant each time. Tubes 1–3 were resuspended in 300 μL of wash buffer and kept cold until flow cytometry measurement. For tubes 4 and 5, the cell pellet was resuspended in 100 μL blocking buffer, and 5 μL of the appropriate antibody was added according to Table 2. Tubes were briefly vortexed and incubated for 20 minutes in the dark on ice, then washed twice with cold wash buffer under the same conditions. The cell pellets of tubes 4 and 5 were resuspended in 300 μL wash buffer and kept on ice. Flow cytometry measurement was performed, with tubes 1–4 used as instrument, gating, and compensation controls as illustrated in Figure 3b.
Fig. 3 Characterization and quantification of hPSC-derived cardiomyocytes by flow cytometry.
(a) Representative histograms of the proportion of TNNT2+ cells at day 21 of differentiation. The graph on the left shows the isotype control and was used to establish the gating strategy. The percentage of TNNT2+ cells is indicated in the graph on the right. (b) Time course of SIRPA and VCAM1 expression between days 0 and 14 of differentiation for the whole cell population (upper panel) or within the GFP+ (NKX2.5+) population (lower panel) of an NKX2-5–eGFP reporter hPSC line.
3.6Immunohistochemical Analysis of Sarcomeres in hPSC-Derived Cardiomyocytes
A sterile glass coverslip (15 mm) was placed in each well of a 12-well plate and coated with Matrigel. Dissociated cells were counted using a hemocytometer, and a suspension of 1×1061×106 cells/mL was prepared. The Matrigel solution was removed, and 5–7.5×1055–7.5×10 5 cells (50–75 μL) were plated per well in a total of 1.5 mL of differentiation medium (see Note 7).
Five days after seeding, the medium was replaced with 1.5 mL of fresh differentiation medium. Approximately 10–15 days after seeding, cells were fixed. Differentiation medium was aspirated, and cells were washed once with 1 mL PBS. PBS was aspirated, and 1 mL fixation solution was added per well in a fume hood. Cells were incubated for 30 minutes at room temperature. Fixation solution was removed, and cells were washed three times with 1 mL PBS. Fixed cells could either be immunoassayed immediately for sarcomere markers or stored at 4 °C for 2–3 weeks in 1.5 mL PBS per well.
Cells were briefly rinsed with PBS and permeabilized with 1 mL permeabilization solution per well for 8 minutes at room temperature. The permeabilization solution was aspirated, and coverslips were washed three times with PBS. Coverslips were removed and placed on parafilm strips in a sealed humidified container. Cells were preincubated with 100 μL blocking solution per coverslip for 1 hour at room temperature.
A solution of 100 μL blocking buffer per coverslip containing both primary antibodies (Table 3) was prepared. The blocking solution was removed from each coverslip, and 100 μL of the primary antibody solution was applied to each coverslip. Coverslips were incubated overnight at 4 °C. Cells were washed with 200 μL washing solution per coverslip for 10 minutes; washing was repeated twice. A solution of 100 μL blocking buffer per coverslip containing both secondary antibodies (Table 3) was prepared. 100 μL of the diluted secondary antibody solution was added to each coverslip and incubated at room temperature for 1 hour. Coverslips were washed with 200 μL washing solution per coverslip for 20 minutes; washing was repeated twice.
Cells were incubated for 5 minutes at room temperature with 100 μL DAPI diluted 1:1,000 in blocking solution to stain nuclei. Coverslips were briefly rinsed in H₂O and sealed to glass microscope slides using ~10 μL Mowiol. Slides were left to dry overnight in the dark. Sarcomeric organization of hPSC-derived cardiomyocytes was examined using an epifluorescence or confocal microscope (Fig. 4).
Fig. 4 Identification of hPSC-derived cardiomyocytes by immunohistochemistry. Confocal immunofluorescence images of Troponin I (TNNI3; green) and α-actinin (ACTN2; red) to visualize the sarcomeres in the cardiomyocytes. All nuclei were stained with DAPI (blue). The images in the bottom panel are magnifications of the area framed in the upper images. Scale bar: 25 μm
4.Notes
The quality of BSA can vary between different batches and companies, so several sources should be tested initially. To reduce cytotoxicity, a deionization step using Resin Beads (Bio-Rad, AG 501-X8(D) Resin #142-6425, 20–50 dry mesh size) can be performed if necessary. It is preferable to use primary antibodies that are directly conjugated to fluorescent labels; however, primary antibodies can also be detected using labeled secondary antibodies. When expression of an epitope is low, this approach can improve the signal to background intensity.
Two protocols are described for flow cytometric analysis of either intracellular or cell surface proteins expressed in hPSC-derived cardiomyocytes. TNNT2 expression is frequently examined by intracellular staining to assess cardiomyocyte yield, but this cannot be used to purify cardiomyocytes for further culturing. While TNNT2 and NKX2-5 (mirrored by GFP) are usually co-expressed in hPSC-derived cardiomyocytes, differentiations older than 21 days have shown a population of TNNT2⁺ cells that are NKX2-5⁻ (GFP⁻). In such cases, a second cardiomyocyte marker, such as α-actinin, could also be considered. Alternatively, combined labeling for the cell surface proteins SIRPA and VCAM1 enriches for viable cardiomyocytes that can be isolated by flow cytometry and subsequently cultured further. Dissociated single cardiomyocytes can also be stained for two sarcomeric markers; Troponin I combined with α-actinin displays the characteristic overlap of sarcomeres.
Cultures of undifferentiated hPSCs cannot be maintained in hESC medium without MEFs for more than 48 hours. If cultures do not reach 70% confluence within this timeframe, either the initial cell seeding density should be increased or hPSCs should be seeded in mTeSR1 or E8 medium. Cells can be maintained in these defined media for up to five days before starting differentiation.
The density of cells at day 0 of differentiation is a key factor in determining differentiation efficiency and obtaining a high yield of cardiomyocytes. Cultures should be approximately 50–80% confluent at the start of differentiation, but this should be independently assessed for each hPSC line. From enzymatically passaged hPSCs maintained on MEFs, seeding is typically at 2.5 × 10⁴ hPSCs/cm², while hPSCs maintained in E8 medium are passaged between 1:6 and 1:10. Because growth rates vary between hPSC lines, the optimal seeding density must be determined for each line individually, and it is recommended to test three different densities initially.
Inhibiting Rho associated protein kinase (ROCK) with either 10 μM Fasudil (HA1077) or 10 μM Y-27632 during seeding of hPSCs or hPSC derived cardiomyocytes can reduce dissociation induced apoptosis. Medium should be refreshed 24 hours after seeding to remove the inhibitor. For some hPSC lines, significant cell death and detachment occur once differentiation begins, particularly with hPSCs maintained in E8 medium. In such cases, treating seeded cells with E8 medium containing 1–2% DMSO for 24–30 hours before starting differentiation can improve cell survival and cardiac differentiation efficiency.
The bioactivity of cytokines can vary between batches and suppliers; optimal concentrations of BMP4 and Activin A for cardiomyocyte generation should be determined by cross titration. If differentiated hPSCs form a contracting sheet across the entire well, the culture can be overlaid with Matrigel to reduce detachment. This additional layer can be applied at any time after day 3 of differentiation, including when cells are treated with XAV939. To do so, add 1 mg thawed Matrigel to 12 mL cold differentiation medium, aspirate existing medium, and replace it with an equal volume of the Matrigel containing medium.
Genetically engineered reporter hESC and hiPSC lines expressing fluorescent proteins, such as eGFP or mCherry, under the control of cardiac specific promoters are valuable tools for evaluating differentiation progress and efficiency in real time. These reporters also facilitate isolation of relatively pure cardiomyocyte populations for further culturing. The concentration of TrypLE Select and incubation time required to dissociate differentiated hPSCs depend on the age of the culture. Typically, 1× TrypLE Select (≈5 min) is used for cultures under eight days; 2× TrypLE Select (≈10 min) for days 9–12; 5× TrypLE Select (≈10 min) for days 13–18; and 10× TrypLE Select (minimum 10 min) for cultures older than 19 days. When performing flow cytometric analysis on live cells, a viability marker such as DAPI (0.1 μg/mL) or PI (1 μg/mL) should be used to exclude dead cells.
When seeded on coverslips, differentiated hPSCs should not exceed ~70% confluence due to proliferation of non-cardiomyocytes. Cells should be fixed when cardiomyocytes are still present as individual cells or small clusters to ensure proper immunostaining.
