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© 2014. Published by The Company of Biologists Ltd | Development (2014) 141, 752-760 doi:10.1242/dev.097386
PRIMER
PRIMER SERIES
How to make an intestine
James M. Wells1,2,* and Jason R. Spence3,4,5
KEY WORDS: Directed differentiation, Embryonic stem cells,
Gastrointestinal disease, Gut tube, Intestinal morphogenesis,
Pluripotent stem cells
Introduction
The intestinal tract is one of the most architecturally and
functionally complex organs of the body. The human intestine is 8 m
in length (Hounnou et al., 2002) and is subdivided into five
functional domains along the proximal-to-distal axis: the duodenum,
jejunum and ileum are segments of the small intestine; and the
cecum and colon make up the large intestine. The intestine
comprises specialized cell and tissue types from all three germ
layers. Intestinal tissues include endoderm-derived epithelium,
which houses specialized intestinal stem cells (ISCs) (Sato and
Clevers, 2013), mesoderm-derived smooth muscle, vasculature,
lymphatics and immune cells, and the ectoderm-derived enteric
nervous system (ENS). The contributions from all of the germ layers
are required to help coordinate a myriad of complex intestinal
functions.
The intestine is best known for its digestive function to orchestrate
the breakdown of macronutrients, regulate absorption and secrete
waste. Less appreciated is the central role of the intestinal endocrine
system in coordinating systemic nutrient levels and feeding behavior
with other organ systems via endocrine hormones. In addition, the
epithelium of the intestine acts as a selective barrier by restricting
microbes to the gut lumen (Turner, 2009). These epithelial functions
are largely carried out by four cell types: absorptive enterocytes and
the three secretory lineages. Secretory cells involved in barrier
function include goblet and Paneth cells, which secrete mucin and
antimicrobial factors. The enteroendocrine cells are a rare but
diverse population of cell types that secrete hormones that regulate
satiety, motility, absorption, β-cell proliferation, secretion of other
1
Division of Developmental Biology, Cincinnati Children’s Hospital Medical Center,
3333 Burnet Avenue, Cincinnati, OH 45229-3039, USA. 2Division of
Endocrinology, Cincinnati Children’s Hospital Medical Center, 3333 Burnet
Avenue, Cincinnati, OH 45229-3039, USA. 3Department of Internal Medicine,
University of Michigan Medical School, Ann Arbor, MI 48109, USA. 4Department
of Cell and Developmental Biology, University of Michigan Medical School, Ann
Arbor, MI 48109, USA. 5Center for Organogenesis, University of Michigan Medical
School, Ann Arbor, MI 48109, USA.
*Author for correspondence ([email protected])
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hormones and digestive enzymes, among other things (Engelstoft et
al., 2013). The epithelium of the intestine turns over every 5 days in
mice, and this regenerative capacity is driven by a resident
population of ISCs (Creamer et al., 1961; Sato et al., 2009). The
mechanical functions of the intestine are controlled by complex
interactions between the epithelium, smooth muscle and ENS, which
regulate peristalsis to ensure unidirectional movement of luminal
contents.
Given the cellular and functional complexity of the intestine, it is
no wonder that there are a staggering number of people impacted by
intestinal dysfunction. Common intestinal disorders include
infections, irritable bowel syndrome (IBS), malabsorption and fecal
incontinence. Other debilitating diseases include inflammatory
bowel disease (IBD), colon cancer and diseases that, in some cases,
have a genetic basis, such as Hirschsprung’s Disease. Additionally,
since most oral drugs are absorbed in the intestine, the most
common drug side effects are intestinal. Given the plethora of
functions carried out by the intestine, there is significant interest in
preventing or reversing intestinal disease by manipulation of
intestinal cell biology; for example, by stimulating ISC growth as a
means to protect or re-establish epithelial integrity and barrier
function following injury (Zhou et al., 2013). However,
gastrointestinal (GI) disease research has largely relied on in vivo
animal studies, which are intrinsically low throughput and
sometimes do not adequately mimic human physiology. Therefore,
in vitro-derived human intestinal tissues represent a powerful tool
for functional modeling of congenital defects in human intestinal
development, preclinical screening for drug efficacy and toxicity
testing, and for establishing models to study the mechanistic basis
of diseases including IBD and cancer.
In this Primer, we discuss the current understanding of intestine
development and how this information has been used to direct the
differentiation of human pluripotent stem cells (PSCs) into intestinal
tissue in vitro. This approach requires the temporal manipulation of
signaling pathways in a step-wise manner that recapitulates early
intestinal development (Fig. 1). These main developmental steps
include the formation of definitive endoderm, posterior endoderm
patterning, gut tube formation, and intestinal growth and
morphogenesis (Fig. 2). Success in this area is largely due to a shift
from two- to three-dimensional growth conditions and the presence
of mesenchymal cell types that result in a level of tissue complexity
that more closely resembles the developing intestine in vivo. We will
also evaluate how these tissues can be used to model human
intestinal development and disease.
The first step: endoderm formation
Generating intestinal cells from PSCs requires a step to mimic the
process of gastrulation and formation of definitive endoderm (DE)
(Fig. 1). Studies of gastrulation using frog, fish, chick and mouse
embryos have been essential in identifying a conserved molecular
pathway that directs gastrulating cells into the endoderm and
mesoderm lineages (reviewed in Zorn and Wells, 2009; Zorn and
Wells, 2007). Central to these processes is the Nodal family of
Development
ABSTRACT
With the high prevalence of gastrointestinal disorders, there is great
interest in establishing in vitro models of human intestinal disease
and in developing drug-screening platforms that more accurately
represent the complex physiology of the intestine. We will review how
recent advances in developmental and stem cell biology have made
it possible to generate complex, three-dimensional, human intestinal
tissues in vitro through directed differentiation of human pluripotent
stem cells. These are currently being used to study human
development, genetic forms of disease, intestinal pathogens,
metabolic disease and cancer.
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Development (2014) doi:10.1242/dev.097386
A
Human
pluripotent
stem cells
Human definitive
endoderm
B
Gut tube
morphogenesis
CDX2/DAPI
FOXA2
SOX17/FOXA2
Acitvin A
WNT3A
FGF4
3 days
4 days
Human intestinal
organoid (HIO)
3D spheroids
Transfer
to 3D
cultures
RSPO1
EGF
NOG
1 month
TGFβ signaling proteins, which in frog embryos act both to initiate
gastrulation and to guide the specification of the mesoderm and
endoderm lineages in a dose-dependent manner (Green and Smith,
1990). Nodal acts through its cognate Type I (Alk4/7) and type II
(ActRIIA or B) receptors, and also requires the Crypto co-receptor
for robust activation of signaling in vivo (Gritsman et al., 1999). The
serine-threonine kinase activity of the Nodal-receptor complex
mediates phosphorylation and nuclear localization of a Smad2/3/4
complex. Once in the nucleus, Smads interact with transcriptional
co-factors such as FoxH1/FAST1 or Mix-like homeodomain
proteins to activate a highly conserved endoderm gene regulatory
network (Hoodless et al., 2001; Tremblay et al., 2000). The core of
this transcriptional network in vertebrates contains the transcription
factors Sox17, Foxa2, Mix, Gata4/6 and Eomes. Although the exact
roles of these factors might vary slightly between vertebrates, they
act to coordinate the induction the endoderm lineage (Sinner et al.,
2006).
Establishing posterior identity from the DE
At the end of gastrulation there are four major signaling pathways
that promote the posterior patterning of the vertebrate embryo:
Wnt, Fgf, retinoic acid (RA) and Bmp. In Xenopus embryos, high
levels of β-catenin dependent Wnt (Wnt/β-catenin) signaling
promotes posterior gene expression in endoderm while at the same
time inhibiting anterior gene expression in the posterior (McLin et
al., 2007; Zorn and Wells, 2009). Conversely, repression of Wnt
by the Sfrp family of secreted Wnt antagonists is required to help
establish the stomach-duodenum boundary (Kim et al., 2005).
Bmp ligands are expressed in the posterior mesoderm and act in a
paracrine manner to promote posterior fate of endoderm in frog,
fish and chick embryos (Kumar et al., 2003; Rankin et al., 2011;
Roberts et al., 1995; Tiso et al., 2002). Similarly, Fgf ligands that
are expressed in posterior mesoderm act on adjacent ectoderm and
endoderm to promote a posterior fate (Dessimoz et al., 2006; Wells
and Melton, 2000). Finally, RA is also known to promote posterior
patterning of endoderm (Bayha et al., 2009; Huang et al., 1998;
Niederreither et al., 2003; Stafford and Prince, 2002; Wang et al.,
2006b).
One of the primary means by which these pathways promote a
posterior fate is through direct regulation of Caudal homeobox (Cdx)
genes, first identified as master regulators of posterior fate in
Drosophila (Macdonald and Struhl, 1986; Mlodzik et al., 1985).
One family member, Cdx2, is expressed in a temporally and
spatially dynamic manner, first being expressed in all three germ
layers in early gestation, then becoming restricted to the intestinal
epithelium by midgestation. Initial insights into the critical role for
Cdx2 during gut/intestine development came through tetraploid
complementation assays (Chawengsaksophak et al., 2004). These
experiments showed that Cdx2-null mesoderm and neurectoderm
were properly specified, whereas the hindgut failed to express
endodermal Shh and had delayed formation of the hindgut.
Subsequent studies showed that conditional deletion of Cdx2 in
endoderm using Foxa3-Cre resulted in mutant mice that formed a
gut tube that was truncated at the cecum, completely lacked the
colon and terminated in a blind-ended sac. Mutant intestines failed
to undergo villus morphogenesis and expressed foregut-specific
genes and an esophageal program, with the epithelium
morphologically resembling the squamous epithelium of the
esophagus. Collectively, these data suggested that in the absence of
Cdx2, a foregut program is ectopically activated in the region of the
intestine (Gao et al., 2009).
Studies in vitro and in vivo suggest that Wnt signaling mediates
posterior fate by regulating Cdx2 expression. For example,
electroporation of a constitutively active form of β-catenin in the
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Development
Fig. 1. Intestinal differentiation and morphogenesis in a dish. (A) Directed differentiation of human PSCs into human intestinal organoids (HIOs).
Pluripotent stem cells (PSCs) are first differentiated into definitive endoderm (DE) (yellow) and, during differentiation, a small population of cells differentiate
into mesoderm (red). Upon the activation of WNT and FGF signaling, endoderm begins to express gut-specific transcription factors (CDX2, red nucleus), which
persists in the epithelium throughout intestinal development. In addition, the mesenchymal cells proliferate and coalesce with the endoderm to form threedimensional (3D) ‘spheroids’, consisting of a mesenchymal layer and a polarized epithelial layer with a lumen. Spheroids are then grown under 3D conditions
in vitro and form HIOs. HIOs contain most epithelial cell types of the developing intestine, including goblet cells, Paneth cells, enteroendocrine cells and
enterocytes. Other cell types have not been explicitly identified. The mesenchyme (light red) also differentiates into smooth muscle and fibroblastic cell types.
(B) Directed differentiation of human PSCs (far left image shows one colony) is induced by the nodal mimetic activin A, resulting in the formation of
SOX17+/FOXA2+ DE. Human DE is then differentiated into CDX2+ gut/intestinal tissue by inducing high levels of FGF and WNT signaling (for example, by
using recombinant FGF4+WNT3A). During this differentiation process, endoderm and mesoderm undergo morphogenesis to form CDX2+ tube-like structures
(middle panel) and 3D spheres that delaminate from the tissue culture dish (fourth image from the left shows a delaminated spheroid). Three-dimensional
spheroids contain CDX2+ endoderm and mesoderm, and are grown in a 3D matrix in the presence of EGF, noggin (NOG) and R-spondin (RSPO1). Over the
course of 1 month, spheroids expand and differentiate into HIOs, which contain multiple differentiated cell types. HIOs can be repeatedly passaged every 1014 days for more than 1 year.
PRIMER
Development (2014) doi:10.1242/dev.097386
E8.5
Hindgut
E9.5
Primitive gut tube
Mes
Mes
En
En
Mes
Ec
En
Squamous epithelium
Cuboidal epithelium
Pseudo-stratified epithelium
mouse foregut endoderm, genetic activation of β-catenin or
stabilization of β-catenin using a Gsk3b inhibitor at E8.25 resulted
in ectopic induction of Cdx2 and repression of Sox2 in the foregut
(Sherwood et al., 2011). Fgf, RA and BMP signaling pathways
similarly regulate posterior endoderm specification by regulating
expression of the Cdx genes (Bayha et al., 2009; Dale et al., 1992;
Dessimoz et al., 2006; Keenan et al., 2006; Kinkel et al., 2008;
Kumar et al., 2003; Lickert and Kemler, 2002; Northrop and
Kimelman, 1994). In many cases, these pathways directly regulate
Cdx expression through Wnt, RA and Fgf responsive elements
(Haremaki et al., 2003; Rankin et al., 2011; Tiso et al., 2002).
Finally, Cdx factors feed back on these posteriorizing pathways by
regulating expression of key signaling components such as Wnt3,
Fgf8 and the RA synthesizing enzyme Cyp26a1. These data suggest
that there is a posterior regulatory network involving the synergistic
activities of Fgf, Wnt and RA, which act to regulate expression of
the Cdx family of transcription factors.
Using embryonic patterning and inductive cues to generate
intestinal tissue in vitro
Activation of Nodal signaling has been the primary approach to
generate DE cells from mouse and human embryonic stem cells
(ESCs) (D’Amour et al., 2005; Kubo et al., 2004). This requires
culturing ESCs in high levels of activin A (100 ng/ml) that, over the
course of 3 or 4 days, result in monolayer cultures of DE cells
expressing endoderm markers such as Sox17 and Foxa2. The reason
most protocols use the nodal mimetic activin is because it does not
require the crypto co-receptor and is many-fold more active than
nodal in promoting endoderm formation from human PSCs or in
Xenopus animal caps (Tada et al., 2005). Although the DE derived
from Nodal verses activin treatment is remarkably similar, recent
studies indicate that there are some differences in gene expression
and the in vivo differentiation potential of activin versus nodalgenerated DE (Chen et al., 2013). In part, this could be due to the
significant differences in the activity of activin versus nodal as the
levels of nodal signaling can impact the anterior-posterior (A-P)
nature of the endoderm. For example, high levels of nodal/activin
754
Fig. 2. Gut tube formation in the
developing mouse embryo. Three primary
germ layers, mesoderm (Mes), ectoderm (Ec)
and endoderm (En), are specified during
gastrulation. The gut tube is specified from
the endoderm, which has a flat and
squamous-like morphology (left, E7.5). The
developing gut tube epithelium forms along
the anterior-posterior axis (middle, E8.5),
stained here by whole-mount in situ
hybridization for NM_029639 (Moore-Scott et
al., 2007). At this point in time, the anterior
endoderm has formed the foregut tube, the
posterior endoderm has formed hindgut tube
and the middle region is forming the midgut.
Stained sections of the mid/hindgut region
show that the epithelium has acquired a
cuboidal-like morphology. Gut tube formation
is completed by E9.5 in the mouse embryo
(right), with the pseudo-stratified epithelium
fully encased in mesoderm-derived
mesenchyme. Staining for desmoplakin
highlights the pseudo-stratified morphology of
the gut tube. The boxed area and lines
indicate the regions or sections enlarged in
the panels underneath.
signaling in the anterior primitive streak of mice promote anterior
endoderm fate (Chen et al., 2013; Spence et al., 2011b). Consistent
with this, longer activin treatment directs ESCs into an anterior
definitive endoderm fate. Further differences between nodal and
activin-generated DE could be due their differential ability to
synergize with Wnt signaling to promote anterior endoderm fate, as
has been shown in fish and frogs (Ho et al., 1999; Zorn et al., 1999),
and in ESCs (Sumi et al., 2008).
Manipulation of the posterior regulatory network has been the
focus of efforts to direct the intestinal differentiation of mouse and
human PSC-derived DE (Ameri et al., 2009; Cao et al., 2010;
McCracken et al., 2011; Ogaki et al., 2013; Sherwood et al., 2011;
Spence et al., 2011b; Ueda et al., 2010). Activation of the Wnt
pathway in DE cultures derived from mouse ESCs directs a
posterior fate, as demonstrated by Cdx2 expression. However, Cao
and colleagues reported that Cdx2 expression required the presence
of fibroblast-conditioned medium (Cao et al., 2010), suggesting that
other signaling factors secreted by fibroblasts were acting with Wnt
to promote posterior specification. Indeed, WNT and FGF pathways
were shown to act synergistically to induce a posterior fate in human
PSC cultures, as marked by broad CDX2 expression (Fig. 1B)
(McCracken et al., 2011; Spence et al., 2011b). Moreover, these
studies demonstrated there was a temporal signaling requirement
such that prolonged activation of the FGF and WNT signaling
pathways together was required to irreversibly specify a posterior
fate. For example, extended treatment of human DE cultures with
FGF4 and WNT3A proteins for 4 days was required for stable and
irreversible CDX2 expression, and was crucial for intestinal
specification. By contrast, shorter treatment for 2 days resulted in
transient, reversible induction of CDX2 and was not sufficient for
intestinal specification (Spence et al., 2011b). As discussed above,
the WNT and FGF pathways may have multiple nodes of
intersection with RA and BMP signaling during establishment of the
posterior axis in embryos. Thus, it seems reasonable to expect a
combinatorial role for these pathways in activating the posterior
regulatory network in PSC cultures; however, this has not yet been
demonstrated.
Development
E7.5
Late gastrula
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Development (2014) doi:10.1242/dev.097386
The intestines are assembled from progenitors arising from the
three germ layers. Although most studies focus on the endodermderived epithelium, the mesoderm and ectoderm-derived tissues
play central roles in intestinal development and function. The
mesoderm that gives rise to the intestinal mesenchyme is generated
during gastrulation and is distributed laterally along the trunk of
the vertebrate embryo. This lateral plate mesoderm separates into
somatic and splanchnic mesoderm, the latter of which is a distinct
population of cells adjacent to the developing intestinal epithelium
prior to gut tube closure in the avian embryo (Thomason et al.,
2012). As the gut tube closes and the epithelium matures, the
splanchnic mesoderm forms a mesenchyme that encases the
primitive gut tube. This mesenchyme forms multiple layers with
distinct cell types and functions, including subepithelial
myofibroblasts and fibrobasts, as well as circular and longitudinal
smooth muscle. In addition, the mesoderm-derived endothelial
cells and pericytes form a complex vascular plexus that requires a
close proximity to epithelial cells for nutrient and hormone
transport (Powell et al., 2011; Zorn and Wells, 2009; Zorn and
Wells, 2007).
As well as giving rise to multiple cell types, the mesenchyme is
also a crucial player in villus formation (Mathan et al., 1976;
Karlsson et al., 2000) (see Fig. 3). Aggregates of mesenchyme
adjacent to the intestinal epithelium are first visible at E14.5, and
express platelet-derived growth factor A (Pdgfra) (Karlsson et al.,
2000). At later stages, as the epithelium remodels to give rise to the
villi projecting into the lumen, the mesenchymal clusters remain
associated with the tip of the villus epithelium and express several
signaling ligands, including Bmp2 and Bmp4. In addition, both the
epithelium adjacent to the mesenchymal clusters and the cluster
itself appear to be non-proliferative, suggesting that clusters may act
as a signaling center to coordinate villus emergence and withdraw
from the cell cycle (Karlsson et al., 2000). Consistent with this,
inhibition of Bmp signaling leads to excessive proliferation and
ectopic formation of crypt-like structures on the villus epithelium,
similar to what is seen in human juvenile polyposis (Haramis, 2004;
Batts et al., 2006).
Pseudostratified
epithelium
Villus
morphogenesis
More recent work has demonstrated that mesenchymal clusters
respond to, and are dependent on, epithelial hedgehog (Hh)
signaling (Walton et al., 2012). Hh signaling has been previously
shown to be crucial for villus morphogenesis (Madison et al., 2005;
Mao et al., 2010). The study by Walton and colleagues described a
mechanism by which Hh signaling controls the size and formation
of Pdgfra+ mesenchymal clusters, thus allowing villus
morphogenesis to proceed. In the absence of Hh signaling, both the
mesenchymal clusters and the villi fail to form (Walton et al., 2012).
In addition to cluster formation, new work has also revealed the
importance of the mesenchyme-derived intestinal smooth muscle
layers, which generate compressive forces necessary for intestinal
folding seen in a variety of vertebrates, and which leads to an
increased intestinal surface area (Shyer et al., 2013).
Ectoderm-derived tissue is also crucial for intestine development
and function. Within the intestinal mesoderm are two neural
plexuses that control intestinal motility. These neurons, collectively
called the enteric nervous system (ENS), coordinate the muscular
contractions of peristalsis, which control the mixing of food with
digestive enzymes and the movement of luminal contents through
the GI tract. The peripheral nervous system is derived from a subset
of trunk neural crest cells (NCCs) referred to as vagal neural crest
cells, which originate just caudal to the hindbrain in the region
between somites 1 and 7 (Burns et al., 2002; Durbec et al., 1996).
Vagal NCCs migrate ventrally and invade the mesenchyme
surrounding the primitive foregut tube in early somite stage embryos
(reviewed by Kuo and Erickson, 2010). NCCs migrate caudally to
colonize the developing small and large intestine, and migration is
regulated by several pathways including netrin, Bmp and endothelin
signaling (Goldstein et al., 2005; Jiang et al., 2003; Nataf et al.,
1996). Last, NCCs differentiate into neuronal and glial lineages, and
this involves the repression of several signaling pathways, including
Bmp and Notch (Chalazonitis et al., 2004; Goldstein et al., 2005).
Harnessing embryonic morphogenesis and tissue-tissue
interactions to generate more functional intestinal tissues
from PSCs
Multilineage contributions and tissue-tissue interactions are critical
for the development a functional intestine. There is an emerging
Villus morphogenesis
Columnar epithelium
Maturation/crypt
emergence
Bmp
Villus
Hh
Pdgfa
Mouse: E13.5
HIO: 14 days
Mouse: E14.5
HIO: 14-28 days
Crypt
Mouse: E16.5
HIO: 28 days
Mouse: P14
HIO: 45-60 days
Fig. 3. Milestones in murine and HIO intestinal development. The intestinal epithelium is present as a pseudostratified epithelium at ~E12.5 through E13.5
in the mouse, and within 14 days of HIO specification. After this point, the appearance of mesenchymal clusters (orange), which respond to hedgehog (Hh) and
platelet-derived growth factor A (Pdgfa) secreted from the epithelium, is coincident with the onset of villus morphogenesis. In the mouse, villus morphogenesis
proceeds between E14.5 and E16.5, whereas in HIOs this stage takes considerably longer. As the epithelium remodels, the columnar epithelium forms with
stereotypical villus (green) and proliferative intervillus (yellow) domains. Mesenchymal clusters remain associated with the villus tip and act as a postmitotic
signaling center, signaling to the epithelium via secreted growth factors (including Bmp ligands, among others), which prevent epithelial proliferation. The fetal
intervillus domain gives rise to the adult crypt by postnatal day 14 in the mouse, and after approximately 8 weeks in HIOs. The mature adult crypt houses the
proliferative intestinal stem cells (yellow) and secretory Paneth cells (red), as well as several additional cell types, the most prevalent of which are enterocytes
(green), goblet cells (blue) and enteroendocrine cells (purple).
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Development
Contribution of other germ layers to the developing
intestine
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Development (2014) doi:10.1242/dev.097386
concept that one reason for the lack of functionality of many PSCderived cells and tissues is due the absence of organ morphogenesis
in PSC monolayer cultures. For example, monolayer culture-derived
pancreatic endocrine cells lack the ability to secrete insulin in
response to glucose (D’Amour et al., 2006) and liver hepatocytes
have an enzyme expression profile that is fetal in nature
(DeLaForest et al., 2011). These non-physiological, twodimensional (2D) monolayer cultures do not recapitulate the many
morphogenetic processes that occur during endoderm organ
development. Efforts to use more three-dimensional (3D)
approaches for differentiation of PSCs have largely used
spontaneous aggregation into embryoid bodies. However the
stochastic nature of this approach makes it hard to efficiently direct
differentiation into specific cell and tissue types. Moreover, there is
little evidence that normal morphogenesis occurs in embryoid
bodies. Here, we will discuss morphogenetic processes that are
important for intestinal development and how they have been use to
direct the morphogenesis of PSCs into complex, 3D human
intestinal ‘organoids’ (HIOs; see Box 1).
Transition from 2D to 3D: formation of the primitive gut tube in the
embryo
Shortly after gastrulation, all three germ layers undergo
morphogenesis that will result in the formation of complex 3D
structures such as the neural tube, somites and the gut tube. Gut tube
morphogenesis in birds and mammals initiates at the anterior and
posterior ends of the embryo with the formation of the foregut and
hindgut, and is largely completed within 36 hours. During this
process, the 2D DE undergoes dramatic cell shape changes, starting
as flat squamous cells, but then transitioning into a cuboidal
epithelium during the morphogenesis of the foregut and hindgut.
The entire process of gut tube formation is completed within 2 days
after the end of gastrulation in mice, at which time the gut tube
epithelium has a pseudostratified morphology (Fig. 2). In addition,
cell lineage tracing experiments and embryo imaging experiments
demonstrate that endoderm and mesoderm cells are actively
migrating laterally and along the anterior-posterior axis between the
gastrula and gut tube stages of development (Kimura et al., 2006;
Lawson et al., 1991; Lawson et al., 1986; Lawson and Pedersen,
1987).
The signaling pathways that control posterior fate, such as FGF
and WNT, also regulate endoderm and mesoderm cell behaviors that
are involved in morphogenesis of the gut tube. Non-canonical Wnt
signaling is involved in hindgut morphogenesis and elongation.
Mutations in genes encoding either Wnt5a or the downstream
mediators of non-canonical Wnt signaling, such as the disheveledinteracting protein Dapper1 (Dact1), perturb both hindgut formation
and subsequent intestinal elongation (Cervantes et al., 2009; Marlow
et al., 2004; Rauch et al., 1997; Tai et al., 2009; Yamaguchi et al.,
1999). Fgf signaling may also play a role in morphogenesis by
regulating migration of presumptive hindgut mesoderm. In chick,
Fgf8 and Fgf4 act as a chemoattractant and repellant, respectively,
to regulate mesoderm migration laterally and posteriorly as the
embryo and hindgut extend caudally (Yang et al., 2002).
Triggering gut tube morphogenesis in PSC cultures
Although the Wnt and Fgf pathways play distinct roles during
embryonic hindgut morphogenesis, studies using human PSC
Box 1. PSC-derived versus primary intestinal epithelial cultures: what is the difference?
Pluripotent
stem cells
Blastocyst
Gut tube
spheroid
Differentiation and
morphogenesis
Growth and
morphogenesis
Induced
pluripotency
Growth
morphogenesis
Intestinal
crypt
Surgery
biopsy
I
Intestinal
i l
organoid
Intestinal
enteroid
Congential
dysfunction
Adult epithelial
function
Adult intestinal
stem cell biology
Cancer
Drug screening
Organ development
Tissue interactions
Tissue engineering
Host/pathogen
interaction
One of the major advances in gastrointestinal biology includes long-term growth and expansion of individual adult ISCs in three-dimensional ‘enteroid’
cultures (Ootani et al., 2009; Sato et al., 2009; Sato and Clevers, 2013; Sato et al., 2011). Enteroids are also commonly referred to as ‘mini-guts’, as well
as ‘organoids’ (Sato and Clevers, 2013). Human intestinal organoids (HIOs), however, are derived from pluripotent stem cells and are significantly different
from enteroids, as recently discussed at length (Finkbeiner and Spence, 2013). Briefly, enteroids are derived from fully committed adult intestinal crypts
(or single ISCs) that are isolated from human or mouse intestine (Gracz et al., 2013; Wang et al., 2013). Enteroids consist of epithelium only and are biased
towards an undifferentiated stem-cell fate due to the culture conditions (Miyoshi et al., 2012). Enteroids have proven to be an unparalleled model in which
to study molecular mechanisms by which adult intestinal stem cells are regulated and maintained, and are well suited to adult disease studies (de Lau et
al., 2011; Dekkers et al., 2013; Zhou et al., 2013). HIOs, however, are generated by recapitulation of a developmental program and are therefore an
unparalleled model in which to study human gastrointestinal development (Du et al., 2012). Both HIOs and adult enteroids have great potential as new in
vitro models for drug screening, infectious disease and cancer.
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cultures suggest that FGF and WNT synergistically act both to
promote expression of CDX2 and to initiate a morphogenetic
process that is similar to gut tube morphogenesis in vivo (Figs 1, 2)
(Spence et al., 2011b). For example, in vivo, embryonic DE forms
as a flat sheet of cells with a squamous-like morphology. However,
as gut tube morphogenesis commences, DE cells transition to the
cuboidal type of epithelium that lines the developing mid- and
hindgut (Fig. 2). Similar transitions occur with PSC-derived DE,
which starts as a flat sheet of cells, much like a squamous
epithelium. However, following 4 days of treatment with both FGF4
and WNT3a, DE cells condense into the cuboidal epithelium and
form gut tube-like structures.
In addition to the epithelial morphogenesis observed in PSC
cultures, the mesoderm undergoes transitions that are similar to
those observed during gut tube morphogenesis in vivo: FGF ligands
mediate mesoderm specification, proliferation and migration (Yang
et al., 2002). In PSC cultures, FGF4 mediated an expansion of
mesoderm that formed the mesenchyme surrounding the epithelium
of these gut tube-like structures. It is likely that these gut tube
structures were fully committed to the intestinal lineage, as growth
in 3D pro-intestinal culture conditions (Sato et al., 2009) resulted in
the formation intestinal tissue, and not other gut tube derivatives.
In addition to intestinal epithelium, PSC-derived HIOs have a
significant level of mesenchymal complexity. The mesenchyme in
HIOs comes from mesoderm, which populates the early DE cultures
and expands in response to FGF4 to form a layer of primitive
mesenchyme surrounding the developing organoid (Spence et al.,
2011b) (see Fig. 1). This primitive mesenchyme differentiates in
parallel with the epithelium and forms several differentiated cell
types, such as smooth muscle, subepithelial myofibroblasts and
fibroblasts, consistent with in vivo development (Spence et al.,
2011b).
(McCracken et al., 2011; Spence et al., 2011b). The 3D cuboidal
epithelium of the early intestinal organoids gives rise to a
pseudostratified epithelium, which undergoes a process mimicking
villus morphogenesis, with a few exceptions. As organoids continue
to grow in culture, the pseudostratified epithelium transitions to
columnar epithelium; however, true villi do not form. Instead, villuslike structures form, which are made up of epithelial protrusions
extending into the lumen. These structures are not true villi due their
lack of the lamina propria, which is the underlying mesenchymal
tissue made up of nerve, vascular, immune and support cells normally
found in the core of the villus. Although true villi do not seem to
develop in HIOs, the villus-like structures still behave similarly to
their in vivo counterpart and proliferation is restricted to the base of
the villus-like structure, equivalent to the intervillus progenitor
domain, and preferentially expresses molecular markers of the
intervillus domain, including Sox9. Furthermore, HIOs generate cells
that express markers for the major intestinal cell types: enterocytes,
goblet, enterendocrine and Paneth cells. The co-expression of
Gata4/Gata6 and presence of Paneth cells in the HIOs suggest that
they are most similar to the small intestine. In addition to the major
cell types mentioned above, the small intestine in vivo possesses other
cell types, including tuft, cup and M-cells; however, it is unclear at
this point whether HIOs possess these cell types. HIOs also appear to
undergo a process similar to crypt emergence. In HIOs, the intestinal
stem cell marker Lgr5 is not expressed robustly during early stages of
organoid ‘development’, but is expressed in discreet epithelial
domains after prolonged culture. It is important to point out that to
date, functional validation of all cell types in organoids has not been
carried out, and many of the conclusions we have drawn are based on
marker analysis. Therefore, as this system becomes more widely used,
it will be interesting to see additional levels of functional validation
for individual cell types.
Milestones during midgestational intestine development are mirrored
in PSC-derived intestine
Future directions
Building a better intestine in vitro
In the mouse embryo, immediately after gut tube formation (around
E9.0), the hindgut is a simple cuboidal epithelium that undergoes a
series of changes over the next 6-7 days that culminates in the
emergence of the stereotypical villus structures of the postnatal
intestine (Fig. 3) (Spence et al., 2011a). Between E9.5 and E12.5,
the intestinal epithelium proliferates and expands in cell number,
girth and luminal size, with the intestinal epithelium and lumen
shaped like an oval (Sbarbati, 1982). Recent studies using genetic
lineage tracing and 3D confocal microscopy demonstrate that the
epithelium becomes pseudostratified around E12.5 (Grosse et al.,
2011). Villus morphogenesis begins at E14.5, and gives rise to a
columnar epithelium and the stereotypical villus structures that
project into the lumen. The intestinal mesenchyme is also involved
and acts as a signaling center to drive villus morphogenesis
(Karlsson et al., 2000; Madison et al., 2005; Walton et al., 2012).
Shortly after the villi have formed, around E16.5 in mouse,
proliferative intervillus progenitor cells give rise to newly
differentiated cells on the villi, which include enterocytes, goblet
and enteroendocrine cells. Of note, Paneth cells do not differentiate
until around postnatal day 14 (P14) in mice and their differentiation
occurs simultaneously with tissue remodeling events that are
responsible for crypt emergence (Calvert and Pothier, 1990; Kim et
al., 2012). In humans, crypts and Paneth cells are present by 22
weeks of gestation (Trier and Moxey, 1979).
Many of the epithelial milestones that occur at these stages of
intestinal development also take place during PSC-derived intestine
‘development’ in vitro, i.e. during the formation of HIOs (Fig. 3)
PSC-derived HIOs exhibit several basic functional properties; for
example, they secrete mucus into the lumen and they are competent
to absorb dipeptides, indicating a functional dipeptide transport
system. Despite these observations, it is still unclear whether HIOs
represent a ‘mature’ or ‘immature’ tissue. Recent studies have
suggested that human PSC-derived intestinal tissues are more
similar to fetal human intestine than to adult human intestine
(Fordham et al., 2013). Importantly, however, different methods
used to generate HIOs and maintain their growth in vitro confound
comparisons from one study to another. In general, the generation
of functional, mature cells or tissues from human PSC cultures has
proven challenging across many tissue types (Dolnikov et al., 2005;
Si-Tayeb et al., 2010; D’Amour et al., 2006; Kroon et al., 2008).
Recently, the transcription factor Blimp1 (also know an Prdm1) was
identified as a key regulator of the transition from fetal to mature
intestinal epithelium, including the formation of crypts (Harper et
al., 2011; Muncan et al., 2011). Therefore, identification of signaling
pathways that repress Blimp1 may be one approach to promote
maturation of HIOs.
One potential reason for lack of maturation and/or functionality
is because in vitro-derived tissues lack important cell types that are
crucial for organ function. For example, even though HIOs contain
important mesoderm-derived cell types that are crucial for
intestinal function, such as smooth muscle myocytes, fibroblasts
and subepithelial myofibroblasts (Powell et al., 2011), they are
missing a vascular plexus and the enteric nervous system (ENS)
that coordinates the muscular contractions of peristalsis. The lack
757
Development
PRIMER
PRIMER
Development and disease research
Although generating transplantable segments of intestine in vitro is
still years (or decades) away, in vitro-derived human intestinal tissue
allows for unprecedented studies of human development,
homeostasis and disease. Agonists and antagonists can be used to
dissect the role of signaling pathways during intestinal
morphogenesis, patterning and cell fate specification. Using standard
genetic gain- and loss-of-function approaches, it is straightforward
to study gene function and identify gene regulatory networks
involved in lineage development and morphogenesis. Alternatively,
iPSC technology can be used to identify the molecular basis of
human congenital disorders affecting the intestines such as cystic
fibrosis and genetic forms of malabsorption (Wang et al., 2006a).
Because of the ease of manipulation and imaging, human organoidbased approaches are a powerful tool to study wide variety of other
disease processes. Introduction of viral and bacterial pathogens into
the lumen of organoids can allow for new studies of infectious
disease and facilitate identification of pathways mediating hostpathogen interactions (Finkbeiner et al., 2012). Organoids derived
from PSCs or adult tissues may also be useful to study the
environmental and genetic causes of cancer initiation and
progression (Ootani et al., 2009). Last, many commonly prescribed
drugs have intestinal side effects, making it a possibility that HIOs
could be used as a rapid screening tool for drugs with good
absorption and fewer intestinal complications.
As for using in vitro-derived intestinal tissues in transplantationbased therapies, current efforts have focused on seeding damaged
intestinal epithelia with adult ISCs (Yui et al., 2012). This approach
has resulted in engraftment and expansion of transplanted cells;
however, it is technically challenging and might be a difficult
approach to use broadly, particularly in more proximal regions of
the intestinal tract. Moreover, this approach is not helpful for
individuals suffering from short gut syndrome. Although in vitro
derivation of transplantable intestinal segments is not likely to occur
in the immediate future, recent advances in tissue engineering have
made it possible to generate an artificial trachea for transplantation
(Jungebluth et al., 2011). Similar approaches using endogenous or
biosynthetic scaffolds should be possible for intestine.
Human organoids are fueling a renaissance of in vitro studies of
human intestinal development and disease. Organoid-based research
is especially powerful when combined with new technologies such
as gene targeting, real-time high content imaging, and fluidics-based
platforms to grow multiple tissue-type organoids on chips to mimic
systemic organ interactions.
Acknowledgements
We thank our colleagues for many helpful discussions and apologize for not
including additional references due to space limitations.
758
Competing interests
The authors declare no competing financial interests.
Funding
J.R.S. is supported by the University of Michigan Center for Organogenesis, the
University of Michigan Biological Sciences Scholar Program, The National Institute
of Diabetes and Digestive and Kidney Diseases and a March of Dimes Basil
O’Connor new investigator award. J.M.W. is supported by the National Institutes of
Health. Deposited in PMC for release after 12 months.
References
Ameri, J., Ståhlberg, A., Pedersen, J., Johansson, J. K., Johannesson, M. M.,
Artner, I. and Semb, H. (2009). FGF2 specifies hESC-derived definitive endoderm
into foregut/midgut cell lineages in a concentration-dependent manner. Stem Cells
28, 45-56.
Bajpai, R., Chen, D. A., Rada-Iglesias, A., Zhang, J., Xiong, Y., Helms, J., Chang,
C. P., Zhao, Y., Swigut, T. and Wysocka, J. (2010). CHD7 cooperates with PBAF to
control multipotent neural crest formation. Nature 463, 958-962.
Batts, L. E., Polk, D. B., Dubois, R. N. and Kulessa, H. (2006). Bmp signaling is
required for intestinal growth and morphogenesis. Dev. Dyn. 235, 1563-1570.
Bayha, E., Jørgensen, M. C., Serup, P. and Grapin-Botton, A. (2009). Retinoic acid
signaling organizes endodermal organ specification along the entire antero-posterior
axis. PLoS ONE 4, e5845.
Burns, A. J., Delalande, J. M. and Le Douarin, N. M. (2002). In ovo transplantation of
enteric nervous system precursors from vagal to sacral neural crest results in
extensive hindgut colonisation. Development 129, 2785-2796.
Calvert, R. and Pothier, P. (1990). Migration of fetal intestinal intervillous cells in
neonatal mice. Anat. Rec. 227, 199-206.
Cao, L., Gibson, J. D., Miyamoto, S., Sail, V., Verma, R., Rosenberg, D. W., Nelson,
C. E. and Giardina, C. (2010). Intestinal lineage commitment of embryonic stem
cells. Differentiation 81, 1-10.
Cervantes, S., Yamaguchi, T. P. and Hebrok, M. (2009). Wnt5a is essential for
intestinal elongation in mice. Dev. Biol. 326, 285-294.
Chalazonitis, A., D’Autréaux, F., Guha, U., Pham, T. D., Faure, C., Chen, J. J.,
Roman, D., Kan, L., Rothman, T. P., Kessler, J. A. et al. (2004). Bone
morphogenetic protein-2 and -4 limit the number of enteric neurons but promote
development of a TrkC-expressing neurotrophin-3-dependent subset. J. Neurosci.
24, 4266-4282.
Chawengsaksophak, K., de Graaff, W., Rossant, J., Deschamps, J. and Beck, F.
(2004). Cdx2 is essential for axial elongation in mouse development. Proc. Natl.
Acad. Sci. USA 101, 7641-7645.
Chen, A. E., Borowiak, M., Sherwood, R. I., Kweudjeu, A. and Melton, D. A. (2013).
Functional evaluation of ES cell-derived endodermal populations reveals differences
between Nodal and Activin A-guided differentiation. Development 140, 675-686.
Creamer, B., Shorter, R. G. and Bamforth, J. (1961). The turnover and shedding of
epithelial cells. I. The turnover in the gastro-intestinal tract. Gut 2, 110-116.
D’Amour, K. A., Agulnick, A. D., Eliazer, S., Kelly, O. G., Kroon, E. and Baetge, E.
E. (2005). Efficient differentiation of human embryonic stem cells to definitive
endoderm. Nat. Biotechnol. 23, 1534-1541.
D’Amour, K. A., Bang, A. G., Eliazer, S., Kelly, O. G., Agulnick, A. D., Smart, N. G.,
Moorman, M. A., Kroon, E., Carpenter, M. K. and Baetge, E. E. (2006). Production
of pancreatic hormone-expressing endocrine cells from human embryonic stem
cells. Nat. Biotechnol. 24, 1392-1401.
Dale, L., Howes, G., Price, B. M. and Smith, J. C. (1992). Bone morphogenetic
protein 4: a ventralizing factor in early Xenopus development. Development 115,
573-585.
de Lau, W., Barker, N., Low, T. Y., Koo, B.-K., Li, V. S. W., Teunissen, H., Kujala, P.,
Haegebarth, A., Peters, P. J., van de Wetering, M. et al. (2011). Lgr5 homologues
associate with Wnt receptors and mediate R-spondin signalling. Nature 476, 293297.
Dekkers, J. F., Wiegerinck, C. L., de Jonge, H. R., Bronsveld, I., Janssens, H. M.,
de Winter-de Groot, K. M., Brandsma, A. M., de Jong, N. W. M., Bijvelds, M. J.
C., Scholte, B. J. et al. (2013). A functional CFTR assay using primary cystic
fibrosis intestinal organoids. Nat. Med. 19, 939-945.
DeLaForest, A., Nagaoka, M., Si-Tayeb, K., Noto, F. K., Konopka, G., Battle, M. A.
and Duncan, S. A. (2011). HNF4A is essential for specification of hepatic
progenitors from human pluripotent stem cells. Development 138, 4143-4153.
Dessimoz, J., Opoka, R., Kordich, J. J., Grapin-Botton, A. and Wells, J. M. (2006).
FGF signaling is necessary for establishing gut tube domains along the anteriorposterior axis in vivo. Mech. Dev. 123, 42-55.
Dolnikov, K., Shilkrut, M., Zeevi-Levin, N., Danon, A., Gerecht-Nir, S., ItskovitzEldor, J. and Binah, O. (2005). Functional properties of human embryonic stem
cell-derived cardiomyocytes. Ann. N. Y. Acad. Sci. 1047, 66-75.
Du, A., McCracken, K. W., Walp, E. R., Terry, N. A., Klein, T. J., Han, A., Wells, J. M.
and May, C. L. (2012). Arx is required for normal enteroendocrine cell development
in mice and humans. Dev. Biol. 365, 175-188.
Durbec, P. L., Larsson-Blomberg, L. B., Schuchardt, A., Costantini, F. and
Pachnis, V. (1996). Common origin and developmental dependence on c-ret of
subsets of enteric and sympathetic neuroblasts. Development 122, 349-358.
Engelstoft, M. S., Egerod, K. L., Lund, M. L. and Schwartz, T. W. (2013).
Enteroendocrine cell types revisited. Curr. Opin. Pharmacol. 13, 912-921.
Finkbeiner, S. R. and Spence, J. R. (2013). A gutsy task: generating intestinal tissue
from human pluripotent stem cells. Dig. Dis. Sci. 58, 1176-1184.
Development
of the vascular and neural plexus limits the physiological relevance
of drug absorption studies and makes it impossible to study
intestinal motility disorders. Given that protocols exist for
generating vascular endothelial cells and NCCs from human PSCs
(Bajpai et al., 2010; Lee et al., 2007; Levenberg et al., 2002), it
should be possible to incorporate vascular and ENS progenitors
into developing HIOs in an attempt to generate vascular and neural
networks. An example of this approach was recently demonstrated
whereby vascular progenitors were added during the differentiation
of PSCs into hepatocyte progenitors (Takebe et al., 2013). The
addition of vascular cells resulted in the formation of 3D liver buds
with a well-formed vascular plexus and greatly improved
functionality in vivo.
Development (2014) doi:10.1242/dev.097386
Finkbeiner, S. R., Zeng, X.-L., Utama, B., Atmar, R. L., Shroyer, N. F. and Estes, M.
K. (2012). Stem cell-derived human intestinal organoids as an infection model for
rotaviruses. MBio 3, e00159-12.
Fordham, R. P., Yui, S., Hannan, N. R. F., Soendergaard, C., Madgwick, A.,
Schweiger, P. J., Nielsen, O. H., Vallier, L., Pedersen, R. A., Nakamura, T. et al.
(2013). Transplantation of expanded fetal intestinal progenitors contributes to colon
regeneration after injury. Cell Stem Cell 13, 734-744.
Gao, N., White, P. and Kaestner, K. H. (2009). Establishment of intestinal identity and
epithelial-mesenchymal signaling by Cdx2. Dev. Cell 16, 588-599.
Goldstein, A. M., Brewer, K. C., Doyle, A. M., Nagy, N. and Roberts, D. J. (2005).
BMP signaling is necessary for neural crest cell migration and ganglion formation in
the enteric nervous system. Mech. Dev. 122, 821-833.
Gracz, A.D., Fuller, M.K., Wang, F., Li, L., Stelzner, M., Dunn, J.C.Y., Martín, M.G.
and Magness, S.T. (2013). CD24 and CD44 mark human intestinal epithelial cell
populations with characteristics of active and facultative stem cells. Stem Cells 31,
2024-2030.
Green, J. B. and Smith, J. C. (1990). Graded changes in dose of a Xenopus activin A
homologue elicit stepwise transitions in embryonic cell fate. Nature 347, 391-394.
Gritsman, K., Zhang, J., Cheng, S., Heckscher, E., Talbot, W. S. and Schier, A. F.
(1999). The EGF-CFC protein one-eyed pinhead is essential for nodal signaling. Cell
97, 121-132.
Grosse, A. S., Pressprich, M. F., Curley, L. B., Hamilton, K. L., Margolis, B.,
Hildebrand, J. D. and Gumucio, D. L. (2011). Cell dynamics in fetal intestinal
epithelium: implications for intestinal growth and morphogenesis. Development 138,
4423-4432.
Haramis, A. P. G. (2004). De novo crypt formation and juvenile polyposis on BMP
inhibition in mouse intestine. Science 303, 1684-1686.
Haremaki, T., Tanaka, Y., Hongo, I., Yuge, M. and Okamoto, H. (2003). Integration of
multiple signal transducing pathways on Fgf response elements of the Xenopus
caudal homologue Xcad3. Development 130, 4907-4917.
Harper, J., Mould, A., Andrews, R. M., Bikoff, E. K. and Robertson, E. J. (2011).
The transcriptional repressor Blimp1/Prdm1 regulates postnatal reprogramming of
intestinal enterocytes. Proc. Natl. Acad. Sci. USA 108, 10585-10590.
Ho, C. Y., Houart, C., Wilson, S. W. and Stainier, D. Y. (1999). A role for the
extraembryonic yolk syncytial layer in patterning the zebrafish embryo suggested by
properties of the hex gene. Curr. Biol. 9, 1131-S4.
Hoodless, P. A., Pye, M., Chazaud, C., Labbé, E., Attisano, L., Rossant, J. and
Wrana, J. L. (2001). FoxH1 (Fast) functions to specify the anterior primitive streak in
the mouse. Genes Dev. 15, 1257-1271.
Hounnou, G., Destrieux, C., Desmé, J., Bertrand, P. and Velut, S. (2002).
Anatomical study of the length of the human intestine. Surg. Radiol. Anat. 24, 290294.
Huang, D., Chen, S. W., Langston, A. W. and Gudas, L. J. (1998). A conserved
retinoic acid responsive element in the murine Hoxb-1 gene is required for
expression in the developing gut. Development 125, 3235-3246.
Jiang, Y., Liu, M. T. and Gershon, M. D. (2003). Netrins and DCC in the guidance of
migrating neural crest-derived cells in the developing bowel and pancreas. Dev. Biol.
258, 364-384.
Jungebluth, P., Alici, E., Baiguera, S., Le Blanc, K., Blomberg, P., Bozóky, B.,
Crowley, C., Einarsson, O., Grinnemo, K.-H., Gudbjartsson, T. et al. (2011).
Tracheobronchial
transplantation
with
a
stem-cell-seeded
bioartificial
nanocomposite: a proof-of-concept study. Lancet 378, 1997-2004.
Karlsson, L., Lindahl, P., Heath, J. K. and Betsholtz, C. (2000). Abnormal
gastrointestinal development in PDGF-A and PDGFR-(alpha) deficient mice
implicates a novel mesenchymal structure with putative instructive properties in villus
morphogenesis. Development 127, 3457-3466.
Keenan, I. D., Sharrard, R. M. and Isaacs, H. V. (2006). FGF signal transduction and
the regulation of Cdx gene expression. Dev. Biol. 299, 478-488.
Kim, B. M., Buchner, G., Miletich, I., Sharpe, P. T. and Shivdasani, R. A. (2005).
The stomach mesenchymal transcription factor Barx1 specifies gastric epithelial
identity through inhibition of transient Wnt signaling. Dev. Cell 8, 611-622.
Kim, T.-H., Escudero, S. and Shivdasani, R. A. (2012). Intact function of Lgr5
receptor-expressing intestinal stem cells in the absence of Paneth cells. Proc. Natl.
Acad. Sci. USA [Epub ahead of print] doi:10.1073/pnas.1113890109.
Kimura, W., Yasugi, S., Stern, C. D. and Fukuda, K. (2006). Fate and plasticity of the
endoderm in the early chick embryo. Dev. Biol. 289, 283-295.
Kinkel, M. D., Eames, S. C., Alonzo, M. R. and Prince, V. E. (2008). Cdx4 is required
in the endoderm to localize the pancreas and limit beta-cell number. Development
135, 919-929.
Kroon, E., Martinson, L. A., Kadoya, K., Bang, A. G., Kelly, O. G., Eliazer, S.,
Young, H., Richardson, M., Smart, N. G., Cunningham, J. et al. (2008).
Pancreatic endoderm derived from human embryonic stem cells generates glucoseresponsive insulin-secreting cells in vivo. Nat. Biotechnol. 26, 443-452.
Kubo, A., Shinozaki, K., Shannon, J. M., Kouskoff, V., Kennedy, M., Woo, S.,
Fehling, H. J. and Keller, G. (2004). Development of definitive endoderm from
embryonic stem cells in culture. Development 131, 1651-1662.
Kumar, M., Jordan, N., Melton, D. and Grapin-Botton, A. (2003). Signals from lateral
plate mesoderm instruct endoderm toward a pancreatic fate. Dev. Biol. 259, 109122.
Kuo, B. R. and Erickson, C. A. (2010). Regional differences in neural crest
morphogenesis. Cell Adh. Migr. 4, 567-585.
Lawson, K. A. and Pedersen, R. A. (1987). Cell fate, morphogenetic movement and
population kinetics of embryonic endoderm at the time of germ layer formation in the
mouse. Development 101, 627-652.
Development (2014) doi:10.1242/dev.097386
Lawson, K. A., Meneses, J. J. and Pedersen, R. A. (1986). Cell fate and cell lineage
in the endoderm of the presomite mouse embryo, studied with an intracellular tracer.
Dev. Biol. 115, 325-339.
Lawson, K. A., Meneses, J. J. and Pedersen, R. A. (1991). Clonal analysis of
epiblast fate during germ layer formation in the mouse embryo. Development 113,
891-911.
Lee, G., Kim, H., Elkabetz, Y., Al Shamy, G., Panagiotakos, G., Barberi, T., Tabar,
V. and Studer, L. (2007). Isolation and directed differentiation of neural crest stem
cells derived from human embryonic stem cells. Nat. Biotechnol. 25, 1468-1475.
Levenberg, S., Golub, J. S., Amit, M., Itskovitz-Eldor, J. and Langer, R. (2002).
Endothelial cells derived from human embryonic stem cells. Proc. Natl. Acad. Sci.
USA 99, 4391-4396.
Lickert, H. and Kemler, R. (2002). Functional analysis of cis-regulatory elements
controlling initiation and maintenance of early Cdx1 gene expression in the mouse.
Dev. Dyn. 225, 216-220.
Macdonald, P. M. and Struhl, G. (1986). A molecular gradient in early Drosophila
embryos and its role in specifying the body pattern. Nature 324, 537-545.
Madison, B. B., Braunstein, K., Kuizon, E., Portman, K., Qiao, X. T. and Gumucio,
D. L. (2005). Epithelial hedgehog signals pattern the intestinal crypt-villus axis.
Development 132, 279-289.
Marlow, F., Gonzalez, E. M., Yin, C., Rojo, C. and Solnica-Krezel, L. (2004). No tail
co-operates with non-canonical Wnt signaling to regulate posterior body
morphogenesis in zebrafish. Development 131, 203-216.
Mao, J., Kim, B. M., Rajurkar, M., Shivdasani, R. A. and McMahon, A. P. (2010).
Hedgehog signaling controls mesenchymal growth in the developing mammalian
digestive tract. Development 137, 1721-1729.
Mathan, M., Moxey, P. C. and Trier, J. S. (1976). Morphogenesis of fetal rat duodenal
villi. Am. J. Anat. 146, 73-92.
McCracken, K. W., Howell, J. C., Wells, J. M. and Spence, J. R. (2011). Generating
human intestinal tissue from pluripotent stem cells in vitro. Nat. Protoc. 6, 19201928.
McLin, V. A., Rankin, S. A. and Zorn, A. M. (2007). Repression of Wnt/beta-catenin
signaling in the anterior endoderm is essential for liver and pancreas development.
Development 134, 2207-2217.
Miyoshi, H., Ajima, R., Luo, C. T., Yamaguchi, T. P. and Stappenbeck, T. S. (2012).
Wnt5a potentiates TGF-β signaling to promote colonic crypt regeneration after tissue
injury. Science 338, 108-113.
Mlodzik, M., Fjose, A. and Gehring, W. J. (1985). Isolation of caudal, a Drosophila
homeo box-containing gene with maternal expression, whose transcripts form a
concentration gradient at the pre-blastoderm stage. EMBO J. 4, 2961-2969.
Moore-Scott, B. A., Opoka, R., Lin, S.-C. J., Kordich, J. J. and Wells, J. M. (2007).
Identification of molecular markers that are expressed in discrete anterior-posterior
domains of the endoderm from the gastrula stage to mid-gestation. Dev. Dyn. 236,
1997-2003.
Muncan, V., Heijmans, J., Krasinski, S. D., Büller, N. V., Wildenberg, M. E.,
Meisner, S., Radonjic, M., Stapleton, K. A., Lamers, W. H., Biemond, I. et al.
(2011). Blimp1 regulates the transition of neonatal to adult intestinal epithelium. Nat.
Commun. 2, 452.
Nataf, V., Lecoin, L., Eichmann, A. and Le Douarin, N. M. (1996). Endothelin-B
receptor is expressed by neural crest cells in the avian embryo. Proc. Natl. Acad.
Sci. USA 93, 9645-9650.
Niederreither, K., Vermot, J., Le Roux, I., Schuhbaur, B., Chambon, P. and Dollé,
P. (2003). The regional pattern of retinoic acid synthesis by RALDH2 is essential for
the development of posterior pharyngeal arches and the enteric nervous system.
Development 130, 2525-2534.
Northrop, J. L. and Kimelman, D. (1994). Dorsal-ventral differences in Xcad-3
expression in response to FGF-mediated induction in Xenopus. Dev. Biol. 161, 490503.
Ogaki, S., Shiraki, N., Kume, K. and Kume, S. (2013). Wnt and Notch signals guide
embryonic stem cell differentiation into the intestinal lineages. Stem Cells 31, 10861096.
Ootani, A., Li, X., Sangiorgi, E., Ho, Q. T., Ueno, H., Toda, S., Sugihara, H.,
Fujimoto, K., Weissman, I. L., Capecchi, M. R. et al. (2009). Sustained in vitro
intestinal epithelial culture within a Wnt-dependent stem cell niche. Nat. Med. 15,
701-706.
Powell, D. W., Pinchuk, I. V., Saada, J. I., Chen, X. and Mifflin, R. C. (2011).
Mesenchymal cells of the intestinal lamina propria. Annu. Rev. Physiol. 73, 213-237.
Rankin, S. A., Kormish, J., Kofron, M., Jegga, A. and Zorn, A. M. (2011). A gene
regulatory network controlling hhex transcription in the anterior endoderm of the
organizer. Dev. Biol. 351, 297-310.
Rauch, G. J., Hammerschmidt, M., Blader, P., Schauerte, H. E., Strähle, U.,
Ingham, P. W., McMahon, A. P. and Haffter, P. (1997). Wnt5 is required for tail
formation in the zebrafish embryo. Cold Spring Harb. Symp. Quant. Biol. 62, 227234.
Roberts, D. J., Johnson, R. L., Burke, A. C., Nelson, C. E., Morgan, B. A. and
Tabin, C. (1995). Sonic hedgehog is an endodermal signal inducing Bmp-4 and Hox
genes during induction and regionalization of the chick hindgut. Development 121,
3163-3174.
Sato, T. and Clevers, H. (2013). Growing self-organizing mini-guts from a single
intestinal stem cell: mechanism and applications. Science 340, 1190-1194.
Sato, T., Vries, R. G., Snippert, H. J., van de Wetering, M., Barker, N., Stange, D.
E., van Es, J. H., Abo, A., Kujala, P., Peters, P. J. et al. (2009). Single Lgr5 stem
cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459,
262-265.
759
Development
PRIMER
PRIMER
Tremblay, K. D., Hoodless, P. A., Bikoff, E. K. and Robertson, E. J. (2000).
Formation of the definitive endoderm in mouse is a Smad2-dependent process.
Development 127, 3079-3090.
Trier, J. S. and Moxey, P. C. (1979). Morphogenesis of the small intestine during fetal
development. Ciba Found. Symp. 1979, 3-29.
Turner, J. R. (2009). Intestinal mucosal barrier function in health and disease. Nat.
Rev. Immunol. 9, 799-809.
Ueda, T., Yamada, T., Hokuto, D., Koyama, F., Kasuda, S., Kanehiro, H. and
Nakajima, Y. (2010). Generation of functional gut-like organ from mouse induced
pluripotent stem cells. Biochem. Biophys. Res. Commun. 391, 38-42.
Walton, K. D., Kolterud, A., Czerwinski, M. J., Bell, M. J., Prakash, A., Kushwaha,
J., Grosse, A. S., Schnell, S. and Gumucio, D. L. (2012). Hedgehog-responsive
mesenchymal clusters direct patterning and emergence of intestinal villi. Proc. Natl.
Acad. Sci. USA 109, 15817-15822.
Wang, J., Cortina, G., Wu, S. V., Tran, R., Cho, J.-H., Tsai, M.-J., Bailey, T. J.,
Jamrich, M., Ament, M. E., Treem, W. R. et al. (2006a). Mutant neurogenin-3 in
congenital malabsorptive diarrhea. N. Engl. J. Med. 355, 270-280.
Wang, Z., Dollé, P., Cardoso, W. V. and Niederreither, K. (2006b). Retinoic acid
regulates morphogenesis and patterning of posterior foregut derivatives. Dev. Biol.
297, 433-445.
Wang, F., Scoville, D., He, X. C., Mahe, M. M., Box, A., Perry, J. M., Smith, N. R.,
Lei, N. Y., Davies, P. S., Fuller, M. K. et al. (2013). Isolation and characterization of
intestinal stem cells based on surface marker combinations and colony-formation
assay. Gastroenterology 145, 383, e1-e21.
Wells, J. M. and Melton, D. A. (2000). Early mouse endoderm is patterned by soluble
factors from adjacent germ layers. Development 127, 1563-1572.
Yamaguchi, T. P., Bradley, A., McMahon, A. P. and Jones, S. (1999). A Wnt5a
pathway underlies outgrowth of multiple structures in the vertebrate embryo.
Development 126, 1211-1223.
Yang, X., Dormann, D., Münsterberg, A. E. and Weijer, C. J. (2002). Cell movement
patterns during gastrulation in the chick are controlled by positive and negative
chemotaxis mediated by FGF4 and FGF8. Dev. Cell 3, 425-437.
Yui, S., Nakamura, T., Sato, T., Nemoto, Y., Mizutani, T., Zheng, X., Ichinose, S.,
Nagaishi, T., Okamoto, R., Tsuchiya, K., Clevers, H. and Watanabe, M. (2012).
Functional engraftment of colon epithelium expanded in vitro from a single adult
Lgr5(+) stem cell. Nat. Med. 18, 618-623.
Zhou, W.-J., Geng, Z. H., Spence, J. R. and Geng, J.-G. (2013). Induction of
intestinal stem cells by R-spondin 1 and Slit2 augments chemoradioprotection.
Nature 501, 107-111.
Zorn, A. M. and Wells, J. M. (2007). Molecular basis of vertebrate endoderm
development. Int. Rev. Cytol. 259, 49-111.
Zorn, A. M. and Wells, J. M. (2009). Vertebrate endoderm development and organ
formation. Annu. Rev. Cell Dev. Biol. 25, 221-251.
Zorn, A. M., Butler, K. and Gurdon, J. B. (1999). Anterior endomesoderm
specification in Xenopus by Wnt/beta-catenin and TGF-beta signalling pathways.
Dev. Biol. 209, 282-297.
Development
Sato, T., Stange, D. E., Ferrante, M., Vries, R. G. J., Van Es, J. H., Van den Brink,
S., Van Houdt, W. J., Pronk, A., Van Gorp, J., Siersema, P. D. et al. (2011). Longterm expansion of epithelial organoids from human colon, adenoma,
adenocarcinoma, and Barrett’s epithelium. Gastroenterology 141, 1762-1772.
Sbarbati, R. (1982). Morphogenesis of the intestinal villi of the mouse embryo: chance
and spatial necessity. J. Anat. 135, 477-499.
Sherwood, R. I., Maehr, R., Mazzoni, E. O. and Melton, D. A. (2011). Wnt signaling
specifies and patterns intestinal endoderm. Mech. Dev. 128, 387-400.
Shyer, A. E., Tallinen, T., Nerurkar, N. L., Wei, Z., Gil, E. S., Kaplan, D. L., Tabin, C.
J. and Mahadevan, L. (2013). Villification: how the gut gets its villi. Science 342,
212-218.
Si-Tayeb, K., Noto, F. K., Nagaoka, M., Li, J., Battle, M. A., Duris, C., North, P. E.,
Dalton, S. and Duncan, S. A. (2010). Highly efficient generation of human
hepatocyte-like cells from induced pluripotent stem cells. Hepatology 51, 297305.
Sinner, D., Kirilenko, P., Rankin, S., Wei, E., Howard, L., Kofron, M., Heasman, J.,
Woodland, H. R. and Zorn, A. M. (2006). Global analysis of the transcriptional
network controlling Xenopus endoderm formation. Development 133, 1955-1966.
Spence, J. R., Lauf, R. and Shroyer, N. F. (2011a). Vertebrate intestinal endoderm
development. Dev. Dyn. 240, 501-520.
Spence, J. R., Mayhew, C. N., Rankin, S. A., Kuhar, M. F., Vallance, J. E., Tolle, K.,
Hoskins, E. E., Kalinichenko, V. V., Wells, S. I., Zorn, A. M. et al. (2011b).
Directed differentiation of human pluripotent stem cells into intestinal tissue in vitro.
Nature 470, 105-109.
Stafford, D. and Prince, V. E. (2002). Retinoic acid signaling is required for a critical
early step in zebrafish pancreatic development. Curr. Biol. 12, 1215-1220.
Sumi, T., Tsuneyoshi, N., Nakatsuji, N. and Suemori, H. (2008). Defining early
lineage specification of human embryonic stem cells by the orchestrated balance of
canonical Wnt/beta-catenin, Activin/Nodal and BMP signaling. Development 135,
2969-2979.
Tada, S., Era, T., Furusawa, C., Sakurai, H., Nishikawa, S., Kinoshita, M., Nakao,
K., Chiba, T. and Nishikawa, S. (2005). Characterization of mesendoderm: a
diverging point of the definitive endoderm and mesoderm in embryonic stem cell
differentiation culture. Development 132, 4363-4374.
Tai, C. C., Sala, F. G., Ford, H. R., Wang, K. S., Li, C., Minoo, P., Grikscheit, T. C.
and Bellusci, S. (2009). Wnt5a knock-out mouse as a new model of anorectal
malformation. J. Surg. Res. 156, 278-282.
Takebe, T., Sekine, K., Enomura, M., Koike, H., Kimura, M., Ogaeri, T., Zhang, R.
R., Ueno, Y., Zheng, Y. W., Koike, N. et al. (2013). Vascularized and functional
human liver from an iPSC-derived organ bud transplant. Nature 499, 481-484.
Thomason, R. T., Bader, D. M. and Winters, N. I. (2012). Comprehensive timeline of
mesodermal development in the quail small intestine. Dev. Dyn. 241, 1678-1694.
Tiso, N., Filippi, A., Pauls, S., Bortolussi, M. and Argenton, F. (2002). BMP
signalling regulates anteroposterior endoderm patterning in zebrafish. Mech. Dev.
118, 29-37.
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