Abstract
Congenital heart defects constitute the most common human birth defect,
however understanding of how these disorders originate is limited by
our ability to model the human heart accurately in vitro. Here we
report a method to generate developmentally relevant human heart
organoids by self-assembly using human pluripotent stem cells. Our
procedure is fully defined, efficient, reproducible, and compatible
with high-content approaches. Organoids are generated through a
three-step Wnt signaling modulation strategy using chemical inhibitors
and growth factors. Heart organoids are comparable to age-matched human
fetal cardiac tissues at the transcriptomic, structural, and cellular
level. They develop sophisticated internal chambers with well-organized
multi-lineage cardiac cell types, recapitulate heart field
formation and atrioventricular specification, develop a complex
vasculature, and exhibit robust functional activity. We also show that
our organoid platform can recreate complex metabolic disorders
associated with congenital heart defects, as demonstrated by an in
vitro model of pregestational diabetes-induced congenital heart
defects.
Subject terms: Cardiovascular models, Differentiation, Stem-cell
differentiation, Congenital heart defects, Heart development
__________________________________________________________________
There is a pressing need to develop representative organ-like platforms
recapitulating complex in vivo phenotypes to study human development
and disease in vitro. Here the authors present a method to generate
human heart organoids by self-assembly using pluripotent stem cells,
compare these to age-matched fetal cardiac tissues and recreate a model
of pregestational diabetes.
Introduction
Cardiovascular disorders, including cardiovascular disease and
congenital heart defects (CHD), are the leading cause of death in the
developed world, and the most common type of congenital defect in
humans, respectively. Despite the importance of understanding human
cardiovascular disorders for treatment and prevention, progress on the
creation of human heart organoid models for cardiovascular disease
studies has been limited and lags significantly behind other organs
(e.g., kidney, colon, intestine, brain)^[50]1–[51]4. Human pluripotent
stem cells (hPSCs) enable us to recapitulate important developmental
steps in vitro to produce specific cardiac cell types with relative
ease, high purity, and in large amounts^[52]5–[53]7. However, current
cell models are still far away from the structural and cellular
complexity of the tissues and organs they intend to represent (e.g.,
lack of 3D matrix, disorganized cells, and absence of multicell-type
interactions). These models frequently study isolated cell types and
minimize or ignore other heart cells (e.g., epicardial cells,
endocardial cells) or the contribution of cell-cell communication to a
disease phenotype. There is a strong demand to bridge this
technological and knowledge gap, as producing more faithful in vitro
models of the human heart will allow us to better study healthy and
diseased states for research and translational applications.
Significant attempts have been made over the last decade to address the
lack of relevant heart-on-a-chip or heart organoid models, particularly
using tissue engineering techniques^[54]8–[55]15. While these
approaches allow for high control of the end construct, they tend to be
expensive, work-intensive, and not readily scalable. Furthermore, they
frequently do not faithfully represent the original cell composition
(e.g., use of dermal fibroblasts or HUVECs)^[56]8,[57]9 and
organization (e.g., cardiospheres^[58]10,[59]11) of the heart. These
approaches yield functional tissues but fall short in terms of
physiological and structural relevance, as well as cell and ECM
complexity. In more recent times, self-assembling organoid technologies
have become available for the heart. These approaches are exploring the
differentiation of PSC embryo-like aggregates in an attempt to
recapitulate early cardiogenesis in vitro. Recently, mouse embryonic
stem cells (ESCs) were used to generate precardiac organoids showing
distinct heart field specifications^[60]16, cardiac crescent-like
structures juxtaposed with primitive gut tube^[61]17, and atrial and
ventricular cardiomyocyte lineages^[62]18. These studies provided us
with a great deal of understanding and information of early heart
development in vitro, but are faced with limitations associated with
mouse models. Hybrid cardiac-foregut organoids from human PSCs have
also been reported very recently, with external heart layers and an
internal endodermal core^[63]19 and cardioids (an alternative
nomenclature for heart or cardiac organoids) have been reported with a
large internal chamber, relevant cardiac cell lineages and single cell
transcriptomic analysis^[64]20. While demonstrating a large leap in
self-assembling capabilities, these cardioids still relied heavily on
complex growth factor mixtures, co-culture approaches to include
epicardial clusters, and independent differentiation protocols to
achieve their final results.
Here, we report a small molecule-based methodology to create highly
complex and physiologically relevant self-assembling human heart
organoids (hHOs) using hPSCs by manipulating cardiac developmental
programs. Our protocol relies mainly on three sequential Wnt modulation
steps (activation/inhibition/activation) at specific time points on
suspension embryoid bodies (EBs), and produces significant heart-like
structures in terms of structure, organization, functionality, cardiac
cell type complexity, ECM composition, and vascularization. Our method
is low-cost when compared to growth factor-based approaches and
involves less steps and manipulation than currently existing methods.
It is also automatable, scalable, and amenable to
high-content/high-throughput pharmacological screenings. As a
proof-of-concept of the value of this system to model human cardiac
disease, we utilized our organoid system to model the effects of
pregestational diabetes (PGD) — clinically defined as diabetes before
pregnancy and present during at least the 1^st trimester of fetal
development — on the developing embryonic heart.
Results
Self-assembling human heart organoids generated by Wnt signaling modulation
Our method is designed to meet four initial milestones: (1) high
organoid quality and reproducibility; (2) high-throughput/high-content
format; (3) relative simplicity (no need for special equipment outside
of traditional cell culture instrumentation); (4) defined chemical
conditions for maximum control and versatility for downstream
applications. We started by assembling hPSCs into embryoid bodies (EBs)
by centrifugation in ultra-low attachment 96-well plates followed by a
48-h incubation at 37 °C and 5% CO[2] prior to induction (Fig. [65]1a).
After induction, two-thirds of the spent medium was removed and
replaced with fresh medium at each medium change, resulting in gradual
transitions in exposure to the different signals employed and
minimizing agitation of the organoids at the bottom of the well.
Induction of mesoderm and cardiogenic mesoderm was achieved by
sequential exposure to CHIR99021, a canonical Wnt pathway activator
(via specific GSK3 inhibition), and Wnt-C59, a Wnt pathway inhibitor
(via PORCN inhibition) (Fig. [66]1a), in a modification of previously
described protocols^[67]14,[68]21,[69]22. Brightfield and
immunofluorescence imaging of hHOs showed a significant increase in
size throughout the differentiation protocol (Fig. [70]1b). Confocal
microscopy for the cardiomyocyte-specific marker TNNT2 showed that
organoids developed sarcomeres as early as day 7 (Fig. [71]1b), with
much clearer sarcomere banding and fiber assembly by day 15
(Fig. [72]1c). Beating hHOs appeared as early as day 6 of the
differentiation protocol, with robust and regular beating by day 10 in
all organoids and lasting for at least 8 weeks in culture
(Supplementary Movies [73]1–[74]3, corresponding to the longest time
tested). We found out that optimal conditions for the first Wnt
activation were critical for successful heart organoid formation and
differed from those reported for cardiac lineage monolayer
differentiation. We exposed EBs to different concentrations of
CHIR99021 (1 µM, 2 µM, 4 µM, 6.6 µM, and 8 µM) on day 0 for 24 h and
then evaluated hHOs for cardiac lineage formation by confocal
microscopy at day 15 of differentiation (Supplementary Fig. [75]1a).
Optimal cardiogenic mesoderm induction for all human embryonic stem
cell (hESC) and induced pluripotent stem cell (hiPSC) lines was found
at 1–4 μM CHIR99021 concentrations, rather than the typical 10–12 µM
range reported for monolayer methods^[76]14,[77]21–[78]27. 4 µM
CHIR99021 exposure resulted in the highest cardiomyocyte content with
64.9 ± 5.3% TNNT2 ^+ cells at day 15 (Fig. [79]1d, Supplementary
Fig. [80]1a). We believe enhanced differentiation at lower CHIR
concentrations can be attributed to endogenous morphogen production due
to the self-assembling conditions provided^[81]28–[82]32. Time-course
RNA sequencing analysis between days 0 and 19 of differentiation
supports this hypothesis by revealing the stepwise production of
fundamental cardiac development morphogens and growth factors and their
respective receptors (Suppl. Fig. [83]1b). hHOs treated with 4 µM
CHIR99021 also displayed the best functional properties out of all
tested concentrations (Suppl. Fig. [84]1c, d). Our hHO differentiation
protocol was reproducible across multiple hPSC lines (iPSC-L1,
AICS-37-TNNI1-mEGFP, iPSCORE_16_3, H9), all of which exhibited similar
differentiation efficiencies, beat metrics, growth rates, and sizes
(Fig. [85]1e, f).
Fig. 1. Three-step WNT signaling modulation triggers heart organoid formation
in 3D culture.
[86]Fig. 1
[87]Open in a new tab
a A schematic diagram depicting the protocol used to differentiate
TNNT2 ^+ cardiomyocytes in embryoid bodies. CHIR99021 concentration is
variable at day 0 and day 7. (Blue cells represent PSCs, red cells
represent cardiomyocytes, all other colors represent non-myocyte
cells). b Brightfield images of developing organoid over 15 days of
differentiation (top), confocal immunofluorescence images for DAPI
(blue), and TNNT2 (red), of representative organoids by day from day 0
to 15; scale bar: 500 µm, inset: 50 µm (n = 20). c Confocal
immunofluorescence images for DAPI (blue) and TNNT2 (red), in day 15
organoids differentiated using 4 µM CHIR99021 showing sarcomere bands;
Scale bar: 25 µm. d Cell quantification of cardiomyocytes within
organoids taken at multiple z-planes as a percentage of TNNT2 ^+ cells
to total cells of each organoid for the five CHIR99021 concentrations
(n = 5 organoids). Value = mean ± s.d., 1 way ANOVA multiple comparison
test; *p = 0.02, ****p < 0.0001, ns: no significance p = 0.08. e
Percentage of TNNT2 ^+ cells normalized to total cells in confocal
images of hHOs (n = 20 organoids), and f beating frequency (n = 7
organoids), in 3 iPSC lines and 1 ESC line. Value = mean ± s.d. g
Confocal immunofluorescence images of hHOs at differentiation day 15
for DAPI (blue), WT1 (green), TNNT2 (red), and TJP1 (white) after day 7
epicardial induction with 2 µM CHIR99021 (n = 12 organoids). h Cell
quantification of cardiomyocyte (TNNT2 ^+) and epicardial cells (WT1 ^+
and TJP1 ^+) within organoids taken at multiple z-planes as a
percentage of total cells of each day 15 organoid (n = 5 organoids).
Value = mean ± s.d., 1 way ANOVA multiple comparison test; *p = 0.04,
**p = 0.0023, ****p < 0.0001, otherwise ns: no significance p = 0.9.
E8: Essential 8 media, RPMI + B27 – ins: RPMI with B27 supplements
without insulin. i Sketch of hHOs showing surface (top) and
cross-section (bottom) features of the hHOs. Source data are provided
as a Source Data file.
To increase organoid complexity and produce more developmentally
relevant structures, we developed a method to induce proepicardial
organ specification based on a second Wnt activation step^[88]22 on
differentiation days 7–9. To determine if this second activation would
prime our hHOs to increase complexity and better recapitulate heart
development, we tested the effects of a second CHIR99021 exposure on
day 7 and continued culturing the hHOs to day 15 for fixation and
imaging (Fig. [89]1a). Conditions for the second Wnt activation with
CHIR99021 were determined by testing developing hHOs at varying
concentrations (2, 4, 6, and 8 µM), and exposure durations (1, 2, 12,
24, and 48 h). The efficiency of epicardial cell and cardiomyocyte
formation was evaluated using confocal imaging and quantification for
well-established epicardial (WT1, TJP1) and cardiomyocyte (TNNT2)
markers at day 15 (Fig. [90]1g, h; Supplementary Fig. [91]2a–c). We
found that the treatment robustly promoted the formation of
proepicardium and epicardial cells (Fig. [92]1g; Supplementary
Fig. [93]2a, b) on the organoid’s surface, however, high concentrations
or long exposure times inhibited cardiac cell type formation and
provoked an undesired extensive epicardial expansion (Supplementary
Fig. [94]2a, c, d). A single 2 µM CHIR99021 exposure for 1 h on
differentiation day 7 produced the most physiologically relevant
epicardial to myocardial ratio (60–65% cardiomyocytes, 10–20%
epicardial cells) (Fig. [95]1g, h; Supplementary Fig. [96]2).
Structurally, a significant part of the epicardial tissue was found on
external layers of the organoid and adjacent to well-defined myocardial
tissue (TNNT2 ^+ ) (Fig. [97]1g, Supplementary Fig. [98]2a, b), thus
recapitulating the structural organization found in the heart. The
robust expression of TJP1 on epicardial cell membranes also confirmed
the epithelial phenotype of these cells (Supplementary Fig. [99]2b).
Overall, the resulting hHOs contained significant myocardial tissue
with epicardial tissue clustered near the outer surface of the
organoids, mimicking the anatomical structure of the developing
embryonic heart (Fig. [100]1i).
Transcriptomic analysis reveals hHOs closely model human fetal cardiac
development and produce all main cardiac cell lineages
To characterize the developmental steps and molecular identity of the
cellular populations present in hHOs, we performed transcriptomic
analysis throughout hHO formation. hHOs were collected for
RNA-sequencing at different time points (day 0 through day 19) of
differentiation (Fig. [101]2, Supplementary Fig. [102]3). Unsupervised
K-means clustering analysis revealed organoids progressed through three
main developmental stages: day 0–day 1, associated with pluripotency
and early mesoderm commitment; day 3–day 7, associated with early
cardiac development; and day 9–day 19, associated with fetal heart
maturation (Fig. [103]2a, Supplementary Fig. [104]3). Gene ontology
biological process analysis identified important genetic circuitry
driving cardiovascular development and heart formation (Fig. [105]2a
and Supplementary Dataset [106]1; raw data deposited in GEO under
“[107]GSE153185”). To compare cardiac development in hHOs to that of
previously existing methods, we performed RNA-seq on monolayer
iPSC-derived cardiac differentiating cells using previously established
protocols^[108]3. We also compared our RNA-seq results to publicly
available datasets from previously reported monolayer cardiac
differentiation protocols and human fetal heart tissue (gestational age
days 57–67)^[109]33 (“[110]GSE106690”). In all instances, hHO cardiac
development transcription factor expression regulating first and second
heart field specification (FHF, SHF, respectively) was similar to that
observed in monolayer PSC-derived cardiac differentiation and
corresponded well to that observed in fetal heart tissue (Fig. [111]2b,
Supplementary Fig. [112]3a). Gene expression profiles showed hHOs had
higher cardiac cell lineage complexity than cells that underwent
monolayer differentiation, especially in the epicardial, endothelial,
endocardial, and cardiac fibroblast populations (Fig. [113]2c,
Supplementary Fig. [114]3b, c). These data suggest a significant
enrichment in the structural and cellular complexity of our hHOs, thus
bringing them in line with fetal hearts and further away from
monolayer-based differentiation. This was confirmed by extending our
gene expression analysis to look at several widespread critical gene
clusters involved in classic cardiac function, including conductance,
contractile function, calcium handling, and cardiac metabolism, among
others (Fig. [115]2d). Of special interest, expression of
heart-specific extracellular matrix genes was high in hHOs and fetal
hearts but completely absent in monolayer differentiation protocols
(Fig. [116]2d, Supplementary Fig. [117]2d). Markers for pluripotency
were not found in hHOs beyond day 1 (Supplementary Fig. [118]3e).
Principal component analysis showed a clear progression in development
in the hHOs from day 0 to 19 (Supplementary Fig. 3f). Taken together,
these data suggest hHO expression profiles are similar to those of
fetal hearts, and their global transcriptomes are closer to those of
fetal hearts than monolayer protocols, as determined by hierarchical
clustering (Fig. [119]2e).
Fig. 2. Heart organoids contain multi-cell type complexity and possess
developmental and maturation characteristics similar to embryonic fetal
hearts.
[120]Fig. 2
[121]Open in a new tab
a K-means cluster analysis of heart organoid transcriptomes by RNA-seq.
Clusters strongly associated with fetal heart development (e.g. 2, 10,
and 14) appear from day 9 onwards. Pathway enrichment analysis is
provided below for representative cardiac-specific clusters. b Gene
expression analysis (log[2] fold-change vs. day 0) of first and second
heart field markers over heart organoid differentiation process (FHF,
SHF respectively). c Gene expression analysis (log[2] fold-change vs.
day 0) for cardiac-specific cell type populations in heart organoids,
including epicardial cells, fibroblasts, endocardial cells, and
endothelium. d Normalized comparison of key genes involved in cardiac
function across heart organoids, monolayer differentiation methods, and
fetal hearts at gestational day 57–67^[122]33. e Hierarchical
clustering analysis of heart organoids, monolayer differentiation, and
fetal hearts. FG57/67: fetal gestation day 57/67, FHF: first heart
field, MONO_D19: monolayer cardiomyocytes at day 19 of differentiation,
ORG_D19: human heart organoids at day 19 of differentiation, OXPHOS:
oxidative phosphorylation, SHF: second heart field. Heatmap colors are
relative intensity representing gene expression.
Heart organoids produce multiple cardiac-specific cell lineages
Results from the transcriptomic analysis (Fig. [123]2) suggested that
the second CHIR99021 exposure led to the formation of other mesenchymal
lineages and higher complexity in hHOs due to induction of
proepicardial organ formation. To evaluate this finding, we performed
immunofluorescence analysis for secondary cardiac cell lineages.
Confocal imaging confirmed the presence of cardiac fibroblasts positive
for THY1 and VIM (Fig. [124]3a), similar to the composition of the
fetal heart^[125]34. Immunofluorescence analysis for the endocardial
marker NFATC1 (an endocardial specific cell marker^[126]19) revealed
the formation of endocardial layers, similar to the endocardial lining
of heart chambers (Fig. [127]3b). Further imaging revealed a robust
interconnected network of endothelial cells (PECAM1 ^+ ), and
vessel-like tube formation throughout the organoid assembling between
day 11 and 13 (Fig. [128]3c, d, Supplementary Fig. [129]4a). Higher
magnification images uncovered a complex web of endothelial cells
adjacent to or embedded in myocardial tissue (Fig. [130]3e,
Supplementary Movies [131]4, [132]5). These results strongly indicate
that during hHO development, endothelial vascular structures emerge,
adding a vascular network to the organoids. Figure [133]3f shows a
quantification of the contribution of the different cardiac
cell populations to the organoids, with a composition of 12.49 ± 1.01%
cardiac fibroblasts, 13.82 ± 1.54% endocardial cells, and 1.63 ± 0.21%
endothelial cells. It should be noted that these non-myocyte cardiac
cells were often intermixed within TNNT2 ^+ myocyte regions
(Fig. [134]3a–e) as seen in vivo^[135]35. Lastly, extracellular matrix
proteins common in cardiac tissue, such as collagen type 1, collagen
type 4, and fibronectin, were all observed in our hHO model
(Supplementary Fig. [136]4b–d). Together, these observations depict a
complex and sophisticated human heart organoid model with endocardial
lined chambers within myocardial tissue, interspersed with cardiac
fibroblasts and a network of endothelial cells, and complete with
external epicardial tissue, features that are highly recapitulative of
the developing human heart (Fig. [137]3g).
Fig. 3. hHO cardiac cell lineage composition.
[138]Fig. 3
[139]Open in a new tab
a–e Immunofluorescence images of various cell lineages composing the
hHOs. a, Cardiac fibroblast markers THY1 (green) and VIMENTIN (white)
present throughout the hHOs, TNNT2 (red), DAPI (blue); scale bar:
500 µm, inset: 50 µm. b Endocardial marker NFATC1 (green) highly
expressed within chambers of TNNT2 ^+ tissue (red); scale bar: 500 µm,
inset: 50 µm. c–e Endothelial marker PECAM1 (green) showing a defined
network of vessels throughout the organoid and adjacent to TNNT2 ^+
tissue (red), DAPI (blue); showing a single confocal plane (c), a
maximum intensity projection to visualize the vascular network
throughout the organoid (d), and a high magnification 3D reconstruction
showing tubular endothelial structures (e); red dotted circle in (d)
indicates area of high vascular branching. Scale bar: c, d: 500 µm, e
50 µm. f Pie chart of average cell composition in hHOs, calculated as
the percentage of cells with respective cell marker over all cells by
nuclei counting using ImageJ. g Sketch of hHO surface (top) and
cross-section (bottom) showing the organization of cell types and
features of the hHOs. Source data are provided as a Source Data file.
Human heart organoids recapitulate heart field development and
atrioventricular specification
The first and second heart fields (FHF, SHF) are two cell populations
found in the developing heart. Cells from the FHF contribute to the
linear heart tube formation, followed by migrating cells belonging to
the SHF that contribute to further expansion and chamber
formation^[140]36. We found evidence of cells representing both heart
fields in our organoids. HAND1 (FHF) and HAND2 (SHF) are members of the
Twist family of basic helix-loop-helix (bHLH) transcription factors
that play key roles in the developing heart^[141]37. Immunofluorescence
of day 7 hHO cryosections showed well-differentiated regions of HAND1
and HAND2 (Fig. [142]4a) cells in the same organoid, suggesting that
both FHF and SHF progenitors are present and segregated into their
respective heart fields. FHF markers were observed as early as day 3
(prior to TNNT2 detection), confined to specific regions on the
organoid, and reducing in expression around day 9 (Fig. [143]4b,
Supplementary Fig. [144]5a). SHF markers appeared later in the process,
only becoming prominent around day 7, and expressing throughout the
organoid up to day 9 (Fig. [145]4b, Supplementary Fig. [146]5b). In
human hearts, the left ventricle ultimately forms from FHF progenitors,
and the atria form from SHF progenitors^[147]38. Therefore, we sought
to determine if our hHOs contain cardiomyocytes committed to either the
atrial or ventricular lineages. Immunofluorescence for MYL2 (encodes
myosin light chain-2, ventricular subtype) and MYL7 (encodes myosin
light-chain 7, atrial subtype) in day 15 hHOs showed cardiomyocytes
positive for both subtypes. The two different populations localized to
different regions of the organoid and were in close proximity, which
mirrors the expression pattern seen in human hearts (Fig. [148]4c, d).
Atrial cardiomyocytes made up most of the cell population (~48%) while
ventricular cardiomyocytes made up about one fifth (~18%) of the total
cells in the organoids at day 15 (Fig. [149]4e). The expression of
HAND1, HAND2, and MYL7 transcripts was also observed throughout the
differentiation protocol by RNA-seq and was similar to that observed in
human fetal hearts (Supplementary Fig. [150]3a, c). We added a contrast
dye (India ink) to organoid medium to reveal structural detail and
record beating organoids under a light microscope. This revealed the
presence of large central chamber-like structures surrounded by beating
tissue, as suggested before by confocal imaging (Supplementary
Movie [151]6). Taken together, these data suggest that the
differentiation of our hHOs involves heart field formation,
atrioventricular specification and chamber formation, all of which
further emphasizes their recapitulation of human cardiac development.
Fig. 4. Heart field development and cardiomyocyte specification in human
heart organoids.
[152]Fig. 4
[153]Open in a new tab
a Confocal immunofluorescent images of two representative hHO
cryosections on day 8 of differentiation showing robust HAND1 (FHF) and
HAND2 (SHF) transcription factor expression (green) in different
sections of the same organoid; TNNT2 (red), DAPI (blue); scale bar:
500 µm, (n = 12 organoids). b Day 3 to day 9 hHOs showing formation of
FHF (HAND1, left) and SHF (HAND2, right); scale bar: 500 µm. c Confocal
immunofluorescence images of three representative day 15 hHOs
containing well-differentiated ventricular (MYL2, green) and atrial
(MYL7, red) regions, DAPI (blue); scale bar: 500 µm (n = 10). d Inset
images of organoid 1 from (c) scale bar: 50 µm. e Pie chart showing the
percentage of atrial cardiomyocytes (MLC2a + DAPI), ventricular
cardiomyocytes (MLC2v + DAPI), and non-myocyte cells (DAPI only); value
= mean ± SD, (n = 6 organoids). FHF: first heart field, Org: organoid,
SHF: second heart field. Source data are provided as a Source Data
file.
BMP4 and Activin A improve heart organoid chamber formation and
vascularization
The growth factors bone morphogenetic protein 4 (BMP4) and Activin A
have frequently been used as alternatives to small molecule-mediated
Wnt signaling manipulation, since they are the endogenous morphogens
that pattern the early embryonic cardiogenic mesoderm and determine
heart field specification in vivo^[154]16,[155]39–[156]41. We suspected
that BMP4 and Activin A, in combination with our small molecule CHIR
activation/inhibition protocol, could synergistically improve the
ability of hHOs to recapitulate cardiac development in vitro. We tested
the effect of BMP4 and Activin A in the context of our optimized
protocol by adding the two morphogens at 1.25 ng/ml and 1 ng/ml,
respectively^[157]16, at differentiation day 0 for 24 h in conjunction
with 4 µM CHIR99021. No significant differences were found in the
formation of myocardial (TNNT2^+) or epicardial (WT1^+/TJP1^+) tissue
between control and treated hHOs (Fig. [158]5a). However, significant
differences in organoid size were observed as hHOs treated with growth
factors were about 15% larger in diameter (Fig. [159]5b, c). This
difference may correspond with increased chamber connectivity, as
BMP4/Activin A-treated hHOs had internal chamber-like cavities that
were ~50% more interconnected with other chambers compared to control
hHOs (Fig. [160]5d, e). Notably, immunofluorescence analysis of
organoids treated with BMP4 and Activin A showed a 160% increase in
PECAM1^+ cells too, indicating a significant effect on organoid
vascularization (Fig. [161]5f, g). These results suggest improved
structural organization in the developing organoids under BMP4/ActA, in
agreement with previously reported differentiation methods using BMP4
and Activin A alone^[162]16.
Fig. 5. BMP4 and Activin A improve heart organoid differentiation and
development.
[163]Fig. 5
[164]Open in a new tab
a–g All panels compare hHOs differentiated with CHIR99021 alone
(control) and with CHIR + BMP4 + Activin A (treated). a Percent of
cardiomyocyte and epicardial positive cells as a percentage of total
DAPI + nuclei (n = 7 organoids per condition, value = mean ± s.d.,
2-way ANOVA Sidak’s multiple comparisons test, ns: p = 0.76 for TNNT2
and p = 0.97 for WT1 + TJP1) and b organoid diameter, (n = 13 organoids
per condition, value = mean ± s.d., two-tailed, unpaired t-test,
*p = 0.012). c Dashed lines showing the diameter of a control (left)
and treated (right) organoid averaged to determine the diameter. d
Interconnectivity of chambers measured by their separation by thin
TNNT2 ^+ filaments or by thin channels showing clear connection (n = 10
organoids per condition, value = mean ± s.d., two-tailed, unpaired
t-test, *p = 0.041). e immunofluorescence images of hHOs showing
interconnected chambers (yellow arrows), TNNT2 ^+ filaments (white
arrows), and channels connecting chambers (green arrows), DAPI (blue),
TNNT2 (red), scale bar: 500 µm, inset: 100 µm. f PECAM1 ^+ cells as a
percentage of total DAPI + nuclei, (n = 7 organoids per condition,
value = mean ± s.d., two-tailed, unpaired t-test, **p = 0.0023).
g Immunofluorescence images of hHOs showing DAPI (blue), PECAM1 tissue
(green), and TNNT2 tissue (red), scale bar: 500 µm, inset: 50 µm
(n = 12).
Heart organoids exhibit functional and structural features of the developing
human heart
Traditional imaging methods, such as confocal imaging, are poorly
suited for the study of the complex 3D structures of the size present
in hHOs. Thus, we employed optical coherence tomography (OCT) to
characterize chamber properties using minimally invasive means, thereby
preserving chamber physical and morphological properties. OCT showed
clear chamber spaces within day 15 hHOs (Fig. [165]6a, Supplementary
Fig. [166]6a–c). 3D reconstruction of the internal hHO topology
revealed a high degree of interconnectivity between these chambers
(Supplementary Movies [167]7–[168]10), revealing 4–6 chambers near the
center of the organoid ranging from 5.5e^−4 mm^3 to 1.3e^−2 mm^3
(Supplementary Fig. [169]6d, Supplementary Movie [170]10). Given the
relatively large size of our heart organoids (up to 1 mm in diameter,
~0.45 mm^3), we decided to verify whether these chambers could be
attributed to internal cell death. We created a transgenic hiPSC line
expressing FlipGFP, a non-fluorescent engineered GFP variant that turns
fluorescent upon effector caspase activation and is thus a reporter for
apoptosis^[171]42. FlipGFP organoids in control conditions exhibited no
fluorescence indicating that there is no significant programmed cell
death (Supplementary Fig. [172]6e). Doxorubicin-treated hHOs were used
as a positive control for apoptosis (Supplementary Fig. [173]6e), with
evident signs of cell death.
Fig. 6. Heart organoids recapitulate functional and structural features of
the developing heart.
[174]Fig. 6
[175]Open in a new tab
a Optical coherence tomography images showing cross-sections through an
organoid, revealing chambers; scale bar: 500 µm. b TEM images of hHOs
showing endoplasmic reticulum (ER), gap junctions (Gj), glycogen
granules (Gy), lipid droplets (Ld), mitochondria (Mi), nucleus (N), and
sarcomeres (S); scale bars: 2 µm (top), 1 µm (bottom). c
Immunofluorescence images of myocardial tissue in hHOs showing WGA
staining of T-tubule-like structures (green); white arrowheads indicate
representative T-tubule-like structures between cardiomyocytes; scale
bar: 50 µm, inset: 20 µm. d Electrophysiology recordings of 4 organoids
on microelectrode array spanning 15 s and a representative action
potential wave (inset). e Ca^2 + transients in 4 representative hHOs
after two weeks of differentiation.
Ultrastructural analysis of hHOs showed similar features to those found
in early human fetal hearts, with well-defined sarcomeres surrounded by
mitochondria, gap junctions and the presence of tubular structures
reminiscent of T-tubules (Fig. [176]6b), also confirmed by
immunofluorescence staining with WGA (Fig. [177]6c, Supplementary
Fig. [178]6f). We also measured electrophysiological activity to
determine the hHO functionality. Utilizing a multi-electrode array
(MEA) (Supplementary Fig. [179]7), we could detect robust beating and
normal electrophysiological activity with well-defined action potential
waves reminiscent of QRS complexes, T and P waves, and regular action
potentials across multiple organoids (Fig. [180]6d). We also performed
live calcium imaging in whole organoids to determine calcium activity.
We generated hHOs from an iPSC line expressing the rapid calcium
indicator GCaMP6f^[181]43,[182]44, and imaged fluorescence variation
over time as a result of calcium entry and exit from the cells. hHOs
presented strong regular calcium waves typical of cardiac muscle and in
agreement with our electrophysiology data (Fig. [183]6e, Supplementary
Movie [184]11).
Modeling pregestational diabetes induced CHD
As proof-of-concept on the utility of our system, we used our hHO model
to study the effects of pregestational diabetes (PGD) on cardiac
development. Diabetes affects a large sector of the female population
in reproductive age and comes associated with significant
epidemiological evidence linking it to CHD during the first trimester
of pregnancy (up to 12-fold risk increase, 12% vs. 1% risk for healthy
females), but little understanding of the underlying mechanisms exists,
especially in humans. To model this condition, we modified hHO culture
conditions to reflect reported physiological levels of glucose and
insulin in normal mothers (3.5 mM glucose, 170 pM insulin,
normoglycemic hHOs or NHOs)^[185]45, and reported diabetic conditions
for females with type I and type II pregestational diabetes (11.1 mM
glucose and 1.14 nM insulin, pregestational diabetes hHOs or
PGDHOs)^[186]45,[187]46. NHOs and PGDHOs showed significant
morphological differences as early as day 4 of differentiation. NHOs
were slower to grow and exhibited patterning and elongation between
days 4 and 8, while PGDHOs remained spherical throughout the two-week
period (Fig. [188]7a; Supplementary Fig. [189]8a). PGDHOs were also
significantly larger after 1 week of differentiation (Fig. [190]7b),
suggesting macrosomia, a common outcome of newborns born to diabetic
mothers^[191]47. Electrophysiological analysis showed irregular
frequency of action potentials in PGDHOs suggesting arrhythmic events
(Fig. [192]7c and Supplementary Fig. [193]8b, c). Metabolic assays for
glycolysis and oxygen consumption revealed decreased oxygen consumption
rate in PGDHOs and increased glycolysis when compared to cells
dissociated from NHOs (Fig. [194]7d, e, Supplementary Fig. [195]8d).
TEM imaging revealed PGDHOs had a reduced number of mitochondria
surrounding sarcomeres and a significantly higher number of lipid
droplets, suggesting dysfunctional lipid metabolism and a more
glycolytic profile (Fig. [196]7f, Supplementary Fig. [197]8e). None of
these phenotypes were found in NHOs. Compared with normal glycemia
conditions, diabetic hHOs showed decreased MYL2 ^+ ventricular
cardiomyocytes and enlarged MYL7 ^+ atrial cardiomyocyte regions,
indicative of structural defects such as those observed in CHD
(Fig. [198]7g, Supplementary Fig. [199]8f). Immunofluorescence for
myocardial and epicardial markers revealed a drastic difference in the
morphological organization as PGDHOs contained epicardial tissue
surrounded by myocardial tissue, whereas NHOs contained epicardial
tissue on top of or beside myocardial tissue as physiologically
expected (Fig. [200]7h). These impairments in structural/developmental
organization and lipid metabolism in PGDHOs are consistent with
expected phenotypes found in PGD-induced CHD. Taken together, our data
suggest significant molecular and metabolic perturbations between NHOs
and PGDHOs consistent with previous studies on PGD suggesting increased
oxidative stress, cardiomyopathy, and altered lipid
profiles^[201]48–[202]50, constituting a significant step forward
towards modeling metabolic disorders in human organoids.
Fig. 7. Human heart organoids faithfully recapitulate hallmarks of
pregestational diabetes-induced congenital heart disease.
[203]Fig. 7
[204]Open in a new tab
a Brightfield images following the development of 10 hHOs under normal
glycemic conditions (NHOs, left) and under diabetic conditions (PGDHOs,
right) over two weeks of differentiation. b Area of hHOs under a light
microscope over the first two weeks of differentiation (value =
mean ± s.d.; n = 12; 2-way ANOVA Sidak’s multiple comparisons test,
exact p-values: day 0: p > 0.99, day 2: p = 0.017, day 4: p = 0.94, day
6: p = 0.99, day 8: p = 0.0008, day 10: p = 0.0003, days 12 and 14:
p < 0.0001). c Electrophysiology was performed on NHOs and PGDHOs at 15
days. Arrows indicate arrhythmic events. d Seahorse analysis for oxygen
consumption rate (OCR), and e extracellular acidification rate (ECAR)
of normal and diabetic hHOs (Value = mean ± s.d; n = 3 organoids per
condition). f Ultrastructural analysis by TEM of NHOs and PGDHOs
showing endoplasmic reticulum (ER), gap junctions (Gj), glycogen
granules (Gy), lipid droplets (Ld), mitochondria (Mi), nucleus (N), and
sarcomeres (S); scale bars: 2 µm. g Confocal immunofluorescence images
at differentiation day 15 for cardiac (TNNT2, red) and epicardial (WT1,
green) formation, scale bar: 500 µm. h Confocal imaging for ventricular
(MYL2, green) and atrial (MYL7, red) chamber formation under normal and
diabetic-like conditions, scale bar: 500 µm. 2-DG: 2-deoxy-d-glucose,
ECAR: extracellular acidification rate, NHOs: normal heart organoids,
OCR: oxygen consumption rate, PGDHOs: pregestational diabetes heart
organoids, Rot/AA: Rotenone and Antimycin A. Source data are provided
as a Source Data file.
Discussion
In recent years, hPSC-derived cardiomyocytes have become critically
useful tools to model aspects of heart
development^[205]12,[206]26,[207]51, human genetic cardiac
disease^[208]52–[209]55, therapeutic screening^[210]14,[211]56, and
cardiotoxicity testing^[212]57–[213]60. Nonetheless, the complex
structural morphology and multitude of tissue types present in the
human heart impose severe limitations to current in vitro models.
Previous attempts at generating 3D human cardiac tissues typically
included cardiomyocytes and only one or two other cardiac cell
lineages^[214]61–[215]63. Here, we sought to create a highly
reproducible, scalable, and cost-effective differentiation protocol
that yields physiologically relevant human heart organoids with high
structural and multicell type complexity using hPSCs. We created and
optimized multistep manipulation conditions for canonical WNT signaling
using GSK3 and PORCN inhibitors in multiple PSC lines. These conditions
lead to the formation of most cardiac lineages in a self-assembling
heart organoid with similar properties to the fetal heart. This method
consistently yields cardiac organoids comprised of approximately 59%
cardiomyocytes, 16% epicardial cells, 14% endocardial cells, 12%
cardiac fibroblasts, and 1.6% endothelial cells and shows robust
beating throughout the entire structure within a week from
differentiation initiation and up to at least 8 weeks in culture
(longer culture times were not attempted). The organization and
specification of these cell types may be related to HAND transcription
factor expression, as HAND1 and HAND2 lineage-derived cells contribute
to the developing myocardium, epicardium, endocardium, and
vasculature^[216]37,[217]64–[218]66. The fact that both FHF and SHF
HAND markers are present suggests that they could play a role in the
development of the structural and cell type complexity seen in our
hHOs. Notably, hHOs were successfully derived from four independent
iPSC lines and one ESC line, demonstrating reproducibility. When
compared with existing cardiomyocyte monolayer differentiation methods,
hHOs showed higher expression of genes associated with conduction,
contractile function, calcium handling, and various cardiac cell
populations, which better resemble gene expression data retrieved from
human fetal hearts. The depiction of a complex transcriptome highly
recapitulative of human fetal heart tissue further strengthens the
potential use of hHOs as models of human heart development.
The epicardium, an epithelial layer that encapsulates the human heart,
is involved in many important heart processes, including heart
development, metabolism, lipid homeostasis, and myocardial injury
responses^[219]67,[220]68. Epicardial signaling cascades are essential
for cardiac lineage specification^[221]67. During embryonic
development, cells from the proepicardial organ (PEO), an extra-cardiac
cluster of embryonic cells^[222]68, migrate to the surface of the heart
to form the epicardium. Some of these cells can undergo
epithelial-mesenchymal transition (EMT) to generate other cardiac
lineages including cardiac fibroblasts^[223]36,[224]67–[225]69. Due to
its capacity to communicate with the myocardium and its ability to
mobilize stem cell populations, the epicardium has become a key focus
of research in cardiac regeneration and repair^[226]22,[227]67,[228]68.
The epicardium also plays a fundamental but underexplored role in
multiple types of cardiovascular and metabolic disease, including
diabetic cardiomyopathy, coronary artery disease, and metabolic
syndrome. In this last condition, epicardial-derived fat experiences a
significant expansion and correlates strongly with morbidity,
highlighting the potential relevance of the epicardium to human
disease. To increase the complexity of our system, and inspired by a
previous epicardial monolayer differentiation method^[229]22, we
created and optimized conditions for producing heart organoids with
well-defined regions of epicardial tissue adjacent to myocardial
tissue. These epicardial-myocardial interactions are important in
mammalian heart development and function, as epicardial cells increase
cardiomyocyte growth in 3D engineered heart tissues, and
co-transplantation of both cell types into rat hearts increases
endothelial cell production^[230]63. Our hHO protocol will facilitate
the study and modeling of epicardial-myocardial interactions in vitro.
Together with the use of small-molecule inhibitors that manipulate
canonical Wnt signaling pathways, successful cardiomyocyte
differentiation has been achieved in the past using morphogens such as
BMP4 and Activin A^[231]21,[232]26. These growth factors lead to the
induction of cardiac mesoderm in the embryo^[233]70 and established
differentiation protocols using them show effective differentiation to
various cardiac mesoderm progenitors^[234]23,[235]70,[236]71. Recently,
gradient exposures to specific concentrations of BMP4 and Activin A
have been studied in the specification of first and second heart field
formation^[237]16. The addition of these growth factors to the initial
CHIR exposure in our hHO differentiation protocol led to improved
morphological features, such as increased chamber interconnectivity and
vascularization.
The important role that cardiac fibroblasts play on cardiac development
and cardiac matrix production/organization is often overlooked in in
vitro models. Most cardiac fibroblasts in embryonic development arise
from the PEO^[238]36,[239]72,[240]73, highlighting the necessity of
epicardial induction in developmental heart models. These fibroblasts
facilitate cardiomyocyte functionality in hPSC-derived 3D cardiac
microtissues, and as such, their inclusion in any in vitro human heart
model is paramount^[241]61. Immunofluorescence analysis of our hHOs
revealed the presence of cardiac fibroblast markers including the
membrane glycoprotein THY1, which is involved in cell-cell and
cell-matrix adhesion^[242]73,[243]74, and the intermediate filament
protein Vimentin, typically seen in cells of mesenchymal
lineage^[244]74. Other cardiac fibroblast markers were found in the
hHOs via RNA-sequencing analysis, including DDR2 which plays an
important role in EMT^[245]73, and the FHF marker PDGFRα, which is also
crucial for vascularization during development^[246]74. These data
provide a strong indication of the increased complexity of our hHO
system and its close resemblance to fetal heart tissue. It should be
noted that while our immunofluorescence and bulk RNA sequencing data
suggest the presence of the cardiac cell types described above, these
techniques may not fully capture the heterogeneity of our organoids. In
recent years, significant advances have been made in single-cell and
single-organoid “omics” technologies^[247]75–[248]77 as well as the
ability to image 3D engineered tissues with unprecedented quality and
resolution^[249]9,[250]78–[251]80. Further studies employing these
techniques will add significantly more detail and information to the
cell type compositions at different time points of differentiation.
An acute limitation of many organoid systems is a lack of a functional
vascular network to facilitate the exchange of nutrients and removal of
waste material, as they instead rely solely on
diffusion^[252]4,[253]81,[254]82. Several vascularized organoids have
been described in the literature modeling the brain^[255]4,
kidney^[256]83, and blood vessel^[257]84, however, none have been
described modeling the heart. In these studies, various techniques are
used to induce vascularization including implantations in mice^[258]4,
culturing the organoids under flow^[259]83, and embedding endothelial
cells in a Matrigel/collagen matrix, and inducing their
migration^[260]84 to create a vascular network. Remarkably, we observed
the formation of vascular structures in our final protocol for hHOs
without any additional steps. We also observed a point of high vascular
branching that merits future exploration into the developmental point
of origin in heart vascularization. Further studies into the
functionality of this vascular tissue will be necessary, particularly
to determine the maturity of the vessels, their levels of connectivity,
and if they closely resemble coronary vasculature. This latter feature
would open the door to modeling coronary vasculature pathologies that
arise due to CVD and metabolic disorders.
In addition to endothelial structures, we also observed spontaneous hHO
reorganization into interconnected chambers, a powerful 3D feature that
recapitulates fetal-like organogenesis. Previous studies of
microchamber formation in vitro utilized micropatterning of hPSCs into
a confined area to generate 3D cardiac microchambers with cell-free
regions, a myofibroblast perimeter, and nascent trabeculae^[261]15.
Other reports have produced 3D bio-printed hPSC-laden scaffolds and
differentiated them to beating cardiac microtissues with two
chambers^[262]85. While the structures generated in these studies
showed some fetal-like formation of cardiac microchambers, they lacked
endocardial tissue^[263]25, a crucial player in heart maturation and
morphogenesis^[264]86. The hHOs reported here form multiple chambers
lined with NFATC1^+ endocardial cells which are interconnected as seen
in the OCT cross-sectional imaging (Supplementary
Movies [265]7–[266]10). Expression of specific ECM genes in the hHOs
resembling the fetal heart matrix, such as COL1A1, COL4A1, COL5A2,
FBN1, EMILIN1, HSPG2, and LAMA2 (Supplementary Fig. [267]3d) might be
an important factor in chamber organization, as ECM components have
been shown to mediate the formation of chambered mouse cardiac
organoids^[268]18. Therefore, the expression of these genes in our hHOs
deserves further examination in the future. The chambers may also
specify further into atrial-like and ventricular-like regions, as
cardiomyocytes from both lineages are seen in separate regions in our
hHOs.
In the past few years, 3D human cardiac tissues have been used to model
genetic and non-genetic conditions (myocardial infarction, drug
cardiotoxicity)^[269]9,[270]87. We provide evidence that heart
organoids can be valuable models to study CHD in pregestational
diabetes-like conditions. Maternal diabetes is one of the most common
causes of newborn CHD (up to 12% of newborns from diabetic mothers have
some form of CHD^[271]88). Using healthy and diabetic levels of glucose
and insulin in the differentiation media, we demonstrate the effects of
diabetic conditions on the developmental process of hHOs. Organoids
developing in healthy conditions displayed active structural changes
including patterning, while hHOs in diabetic conditions developed
largersizes reminiscent of macrosomia. The larger size of diabetic hHOs
also suggests potential signs of cardiac hypertrophy, a hallmark of
maternal PGD^[272]89. We also observed a reduction in mitochondria,
dysfunctional lipid metabolism, and impaired structural organization.
These data suggest hHOs might be useful tools for the study of
PGD-induced CHD, and might pave the way for the identification of
pharmacological agents aimed at treating or preventing this condition.
Although our technology offers exciting opportunities to model human
congenital heart disease in vitro, significant limitations still exist.
First, organoids tend to deviate from their normal developmental
pathway as a function of time, becoming less relevant the longer they
are cultured. Second, their ability to recapitulate heart development
is still limited when compared to other existing models, such as mice,
even though they have the significant advantage of being human in
origin rather than a surrogate animal model. There is large room for
improvement in the technology, particularly in trying to better
recapitulate morphological and anatomical features and inducing the
formation of effective vascular networks that can provide nutrients.
In summary, we describe here a highly reproducible and high-throughput
human heart organoid generation method relying on self-assembly
triggered by developmental cues and provide proof-of-concept for the
modeling of a congenital heart disorder. Heart organoids present
multicell type and morphological complexity reminiscent of the
developing human fetal heart, including chamber formation,
atrioventricular specification, electrophysiological activity and
vascularization. Heart organoids can be used to model features of
pregestational diabetes-induced congenital heart disease and might thus
constitute useful models for the study of the molecular pathology of
congenital heart disease in humans in the future.
Methods
Stem cell culture
The following human iPSC lines were used in this study: iPSC-L1,
AICS-0037-172 (Coriell Institute for Medical Research, alias AICS),
iPSCORE_16_3 (WiCell, alias iPSC-16; UCSD013i-16-3), iPSC
GCaMP6f^[273]43,[274]44, and human ESC line H9 (WiCell, WA09). All PSC
lines were validated for pluripotency and genomic stability. hPSCs were
cultured in Essential 8 Flex medium containing 1%
penicillin/streptomycin (Gibco) on 6-well plates coated with growth
factor-reduced Matrigel (Corning) in an incubator at 37 °C, 5% CO[2]
until 60–80% confluency was reached, at which point cells were split
into new wells using ReLeSR passaging reagent (Stem Cell Technologies).
hPSC monolayer cardiac differentiation
Differentiation was performed using the small molecule Wnt modulation
strategy adapted from a previous protocol^[275]21 (referred to as
monolayer 1 in the text), with small modifications. Briefly,
differentiating cells were maintained in RPMI with B27 minus insulin
from day 0–7 of differentiation and RPMI with B27 supplement (Thermo)
from day 7–15 of differentiation. Cells were treated with 10 µM
CHIR99021 (Selleck) for 24 h on day 0 of differentiation and with 2 µM
Wnt-C59 (Selleck) for 48 h from day 3–5 of differentiation.
Self-assembling human heart organoid differentiation
A step-by-step protocol describing the fabrication and differentiation
of the human heart organoids can be found at Protocol Exchange^[276]90.
Accutase (Innovative Cell Technologies) was used to dissociate PSCs for
spheroid formation. After dissociation, cells were centrifuged at 300 g
for 5 min and resuspended in Essential 8 Flex medium containing 2 µM
ROCK inhibitor Thiazovivin (Millipore Sigma). hPSCs were then counted
using a Moxi Cell Counter (Orflo Technologies) and seeded at 10,000
cells/well in round bottom ultra-low attachment 96-well plates (Costar)
on day −2 at a volume of 100 µl per well. The plate was then
centrifuged at 100 g for 3 min and placed in an incubator at 37 °C, 5%
CO[2]. After 24 h (day −1), 50 µl of media was carefully removed from
each well, and 200 µl of fresh Essential 8 Flex medium was added for a
final volume of 250 µl/well. The plate was returned to the incubator
for an additional 24 h. On day 0, 166 µl (~2/3 of total well volume) of
media was removed from each well and 166 µl of RPMI 1640/B-27, minus
insulin (Gibco) containing CHIR99021 (Selleck) was added at a final
concentration of 4 µM/well along with BMP4 at 0.36 pM (1.25 ng/ml) and
Activin A at 0.08 pM (1 ng/ml) for 24 h. On day 1, 166 µl of media was
removed and replaced with fresh RPMI1640/B-27, minus insulin. On day 2,
RPMI/B-27, minus insulin, containing Wnt-C59 (Selleck) was added for a
final concentration of 2 µM Wnt-C59 and the samples were incubated for
48 h. The media was changed again on day 4 and day 6, but insulin was
not added to the RPMI1640/B-27 (Gibco) mixture until day 6, because it
has been shown to decrease cardiomyocyte yield before this time
point^[277]21. On day 7, a second 2 µM CHIR99021 exposure was conducted
for 1 h in RPMI1640/B-27. Subsequently, media was changed every 48 h
until organoids were ready for analysis. Diabetic conditions were
simulated by using basal RPMI media with 11.1 mM glucose and 1.14 nM
insulin and compared with control media containing 3.5 mM glucose and
170 pM insulin. 40.5 µM Oleate-BSA (Sigma), 22.5 µM
Linoleate-BSA(Sigma), and 120 µM l-Carnitine (Sigma) were added at day
7 to increase fatty acid metabolism, concentrations previously
described^[278]34.
Lentiviral transduction
For lentiviral production HEK293T (Horizon Inspired Cell Solutions,
HCLXXXX) cells were transfected with the Flip-GFP plasmid
(VectorBuilder) and the packaging plasmids (pMD2; psPAX2) using
lipofectamine with Plus reagent (Thermo). Lentivirus was added to
iPSC-L1 cells with 8 μg/ml polybrene (Fisher Scientific) and incubated
overnight. Puromycin selection was carried out for ~3–5 days. Surviving
clones were collected, replated, and expanded to give rise to the
FlipGFP line.
Immunofluorescence
hHOs were transferred to microcentrifuge tubes (Eppendorf) using a cut
1000 μL pipette tip to avoid disruption to the organoids and fixed in
4% paraformaldehyde solution. Fixation was followed by washes in
PBS-Glycine (20 mM) and incubation in blocking/permeabilization
solution containing 10% Donkey Normal Serum, 0.5% Triton X-100, 0.5%
bovine serum albumin (BSA) in PBS on a thermal mixer (Thermo
Scientific) at minimum speed at 4 °C overnight. hHOs were then washed 3
times in PBS and incubated with primary antibodies (Suppl.
Table [279]1) in Antibody Solution (1% Donkey Normal Serum, 0.5% Triton
X-100, 0.5% BSA in PBS) on a thermal mixer at minimum speed at 4 °C for
24 h. Primary antibody exposure was followed by 3 washes in PBS and
incubation with secondary antibodies (Supplementary Table [280]1) in
Antibody Solution on a thermal mixer at minimum speed at 4 °C for 24 h
in the dark. T-tubules staining was conducted using Wheat Germ
Agglutinin (WGA) lectins conjugated with FITC (Millipore Sigma). The
stained hHOs were washed 3 times in PBS before being mounted on glass
microscope slides (Fisher Scientific) using Vectashield Vibrance
Antifade Mounting Medium (Vector Laboratories). 90 µm Polybead
Microspheres (Polyscience, Inc.) were placed between the slide and the
coverslip (No. 1.5) to preserve the 3D structure of the organoids.
Confocal microscopy and image analysis
Samples were imaged using confocal laser scanning microscopy (Nikon
Instruments A1 Confocal Laser Microscope; Zeiss LSM 880 NLO Confocal
Microscope System). Images were analyzed using Fiji
([281]https://imagej.net/Fiji). For cell quantification in the
organoids, DAPI positive cells were counted and used for normalization
against the target cell marker of interest across at least three
z-planes throughout each organoid for each target cell marker.
RNA sequencing and transcriptomic analysis
RNA was extracted at 11 different time points throughout the hHO
fabrication and differentiation protocol shown in Fig. [282]1a. The
time points are as follows: days 0, 1, 3, 5, 7, 9, 11, 13, 15, 17, and
19. At each time point, eight organoids were removed and stored in
RNAlater (Qiagen) at −20 °C until all samples were collected. RNA was
extracted using the Qiagen RNEasy Mini Kit according to manufacturer
instructions (Qiagen), and the amount of RNA was measured using a Qubit
Fluorometer (Thermo). RNA samples were sent to the MSU Genomics Core,
where the quality of the samples was tested using an Agilent 2100
Bioanalyzer followed by RNA sequencing using an Illumina HiSeq 4000
system. For RNA-seq sample processing, a pipeline was created in
Galaxy. Briefly, sample run quality was assessed with FASTQC, and
alignment to hg38 was carried out using HISAT2. Counts were obtained
using featureCounts and differential expression analysis was performed
with EdgeR. Further downstream bioinformatic analysis was performed in
Phantasus 1.11.0 (artyomovlab.wustl.edu/phantasus) and ToppGene Suite
([283]http://toppgene.cchmc.org).
Optical coherence tomography analysis
A customized spectral-domain OCT (SD-OCT) system was developed to
acquire 3D images of the cardiac organoids. As shown in Suppl.
Fig. [284]9, a superluminescent diode (SLD 1325, Thorlabs) was used as
the light source to provide broadband illumination with a central
wavelength of 1320 nm and a spectral range of 110 nm. The output of the
SLD was split 50/50 with a fiber coupler and transmitted to the sample
and reference arms, respectively. A galvanometer (GVSM002-EC/M,
Thorlabs) was used to scan the optical beam in transverse directions on
the sample. The SD-OCT setup used a custom-designed spectrometer
consisting of a 1024-pixel line scan camera (SU1024-LDH2, Sensors
Unlimited), an 1145-line pairs per mm diffraction grating (HD 1145-line
pairs per mm at 1310 nm, Wasatch Photonics), and an f = 100 mm F-theta
lens (FTH100-1064, Thorlabs). The sensitivity of the OCT system was
measured as ~104 dB when operating at a 20 kHz A-scan rate. The axial
resolution of the SD-OCT system was measured to be ~7 mm in tissue. A
5X objective lens (5X Plan Apo NIR, Mitutoyo) was used to achieve a
transverse image resolution of ~7 mm, and the scanning range used for
the cardiac organoids imaging was ~2 mm × 2 mm. Fixed hHOs were placed
into a 96-well plate with PBS and imaged at a 20-kHz A-scan rate.
Obtained OCT datasets of the cardiac organoids were first processed to
generate OCT images with corrected scales. Then OCT images were further
de-noised using a speckle-modulation generative adversarial
network^[285]91 to reduce the speckle noise. 3D renderings of OCT
images were performed using Amira software (Thermo Fisher Scientific).
TEM sample preparation and imaging
Organoids were fixed in 4% PFA for 30 min followed by 3 washes in
water, 10 min each. Post fixation was performed in 1% osmium tetroxide
in cacodylate buffer (pH 7.3) for 60 min at room temperature. Organoids
were embedded in 2% agarose in water, solidified using ice, for
manipulation. Then, a serial dilution of acetone was used for
dehydration (25%, 50%, 75%, 90%, and 3 times in 100%) for 10 min each.
Organoids were infiltrated with Spurr resin (Electron Microscopy
Sciences) by immersion in 1:3, 2:2, and 3:1 solutions of resin in
acetone, 3 h each under agitation, following embedding in 100% resin
for 24 h, and polymerization at 60 °C overnight. Ultra-thin sections
(50–70 nm) were cut using RMC PTXL Leica Ultramicrotome and collected
in carbon-coated copper grids 200 mesh. Before observation, all samples
were positively stained in 2% uranyl acetate and 1% lead citrate for 6
and 3 min, respectively. The grids were examined at 100 keV using a
JEOL 1400 Flash transmission electron microscope.
Electrophysiology
An in-house microelectrode array (MEA) system described
previously^[286]92 was used to record the electrical activity of
individual organoids (Supplementary Fig. [287]7). The Microelectrode
array (MEA) was fabricated with the following cleanroom procedures.
First, 10 μm Parylene C was deposited on a cleaned 3-inch silicon wafer
(PDS 2010, Specialty Coating System, Inc). Then, 500 nm Au was
evaporated on the substrate. Next, a photoresist (PR) layer was spun on
Au and photolithographically patterned the areas of 32-channel
microelectrodes, interconnection wires, and contact pads. Finally, 2 μm
Parylene C was deposited on the substrate as an insulating layer, and
then Parylene C on the contact pads and microelectrodes were removed
completely using oxygen plasma dry etching (RIE-1701 plasma system,
Nordson March, Inc). Live organoids were placed on the MEA inside a
PDMS well in culture media supplemented with 15 mM HEPES. The MEA was
placed within a Faraday cage inside an incubator at 37 °C at low
humidity to avoid damage to the MEA system. Each organoid was recorded
for a period of 30 min, and the PDMS well was washed with PBS between
organoids. The recorded signals were amplified and digitalized using a
commercial Intan RHD2132 system (Intan Technologies) and then recorded
with Intan RHD2000 interface and analyzed using the Matlab Chronux
toolbox to extract the electrocardiogram (ECG) from the recordings.
Calcium transient analysis
Calcium transients were observed in heart organoids developed from
iPSCs expressing the fast calcium indicator GCaMP6f^[288]43,[289]44.
Dynamic fluorescence changes in the heart organoid were recorded at 10
frames per second on an inverted fluorescence microscope (IX71,
Olympus). Data analysis of fluorescence recordings was performed in
MATLAB. 10 by 10 pixel binning was applied to the fluorescence
recordings to minimize impact of contraction of heart organoids.
Baseline F[0] of the fluorescence intensities F was calculated using
asymmetric least squares smoothing^[290]93. Fluorescence change ∆F/F[0]
was calculated by:
[MATH:
△FF0=(<
/mo>F−F0<
/mn>)/F0 :MATH]
1
Seahorse metabolic analysis
A Seahorse analyzer (Agilent) was used to conduct glycolysis rate assay
as per manufacturer instructions. Organoids were carefully dissociated
using STEMDiff cardiomyocyte dissociation kit and checked for viability
using a hemocytometer before analysis. Only samples with over 90%
viability were used in the assay. Identical numbers of cells were
plated for the assay. Measurements were performed immediately after
dissociation.
Statistics and reproducibility
All analyses were performed using GraphPad software and all raw data
was collected in Microsoft Excel. All data presented a normal
distribution. Statistical significance was evaluated with a standard
unpaired Student t-test (2-tailed; P < 0.05) when appropriate. For
multiple-comparison analysis, 1-way ANOVA with the Tukey’s or Dunnett’s
post-test correction was applied when appropriate (P < 0.05). All data
are presented as mean ± s.d. and represent a minimum of 3 independent
experiments with at least 3 technical replicates unless otherwise
stated. All micrograph images are representative of at least 6
independent experiments per condition/marker and calcium transient
graphs are representative of 6 independent experiments.
Reporting summary
Further information on research design is available in the [291]Nature
Research Reporting Summary linked to this article.
Supplementary information
[292]Supplementary information^ (5MB, pdf)
[293]Peer Review File^ (3.2MB, pdf)
[294]41467_2021_25329_MOESM3_ESM.docx^ (13.9KB, docx)
Description of Additional Supplementary Files
[295]Supplementary video 1^ (28.4MB, avi)
[296]Supplementary video 2^ (28.1MB, avi)
[297]Supplementary video 3^ (59.8MB, avi)
[298]Supplementary video 4^ (28.4MB, avi)
[299]Supplementary video 5^ (3.4MB, avi)
[300]Supplementary video 6^ (22.7MB, avi)
[301]Supplementary video 7^ (13.5MB, avi)
[302]Supplementary video 8^ (13.4MB, avi)
[303]Supplementary video 9^ (8.8MB, avi)
[304]Supplementary video 10^ (57.1MB, avi)
[305]Supplementary video 11^ (8.3MB, avi)
[306]Supplementary Dataset 1^ (374.6KB, xlsx)
[307]Reporting Summary^ (384KB, pdf)
Acknowledgements