Generation of parathyroid glands from pluripotent stem cells

Abstract

The parathyroid glands (PTGs) regulate calcium metabolism by secreting parathyroid hormone (PTH). Patients with hypoparathyroidism require lifelong replacement therapy, which is associated with risks of chronic kidney disease, bone fractures, and a reduced quality of life. Generating PTGs from pluripotent stem cells (PSCs) offers a potential regenerative therapy for this condition. This review first explains PTG organogenesis, followed by an overview of both in vitro and in vivo approaches to PTG generation. In vitro studies have successfully induced PTH-expressing parathyroid cells from human PSCs. However, challenges remain, particularly in achieving sufficient PTH secretion and functional efficacy in vivo. Meanwhile, an in vivo organ generation technique known as blastocyst complementation has successfully produced functional PTGs in rodents. However, whether this technology can be applied using human PSCs and animal embryos remains unclear. Pluripotent stem cell-derived PTGs hold promise for both clinical applications and basic research, but further advancements will be necessary to overcome existing challenges in this field.

Introduction

Parathyroid glands (PTGs) regulate Ca homeostasis by releasing parathyroid hormone (PTH). Disruption of PTG function leads to hypoparathyroidism. The prevalence of hypoparathyroidism is estimated at 22 per 100,000 individuals in Denmark [1], 38.3 per 100,000 individuals in Japan [2], 37 per 100,000 individuals in the United States [1]. The most common cause of hypoparathyroidism is neck surgery, particularly total thyroidectomy for thyroid malignancy, which accounts for about 75% of all cases [1]. A rare cause is genetic hypoparathyroidism, which accounts for less than 10% of all cases [1]. Patients with permanent hypoparathyroidism require lifelong management to prevent hypocalcemia. Conventional treatment for hypoparathyroidism consists of oral calcium and vitamin D analogs [3]. Treatment aims to achieve low to low-normal blood Ca concentration and normal urine Ca concentration. Although conventional therapy can restore blood Ca concentration, it does not restore other PTH actions. Hypoparathyroidism and conventional treatment may increase the risk of hypercalciuria, renal stones, nephrocalcinosis, and chronic kidney disease [4]. Idiopathic hypoparathyroidism patients receiving conventional therapy often have high bone mineral density, but are at a heightened risk of vertebral fractures [5]. The quality of life for hypoparathyroidism patients receiving conventional therapy is lower than that of healthy individuals, even when normal Ca levels are achieved [6]. These results indicate that the deficiency of PTH itself is the problem. Therefore, PTH replacement therapy could offer advantages unachievable with conventional therapy. TransCon PTH (palopegteriparatide) is a long-acting prodrug of PTH 1–34. A phase 3 trial in adults with chronic hypoparathyroidism demonstrated that 95% of participants treated with TransCon PTH achieved independence from conventional therapy by week 52 [7]. It improves renal function [7] and quality of life [8]. However, patients require daily self-injections and the long-term efficacy and safety of the therapy remain unknown.

Pluripotent stem cells (PSCs) are defined by their ability to differentiate into ectodermal, mesodermal, or endodermal tissues or cells. There are two types of PSCs: embryonic stem cells (ESCs), derived from the internal cell mass of a zygote, and induced PSCs (iPSCs), which are obtained by adding Yamanaka factors, POU class 5 homeobox 1 (POU5F1, also known as OCT3/4), SRY-box transcription factor 2 (SOX2), Myelocytomatosis oncogene (c-MYC), and Krüppel-like factor 4 (KLF4), to somatic cells [9]. Generating functional PTGs from patients’ own iPSCs and transplanting them would be clinically significant. The established simple subcutaneous transplantation for PTGs would be a further advantage. This review discusses recent progress in generating PTGs from PSCs using both in vitro and in vivo approaches.

Parathyroid gland organogenesis

During gastrulation (embryonic day [E] 6.5–8.5 in mice), epiblast cells give rise to the three primary germ layers: definitive endoderm (DE), mesoderm, and ectoderm. The definitive endoderm eventually forms a gut tube, from which endodermal organs (i.e., thyroid glands, PTGs, lungs, liver, and small and large intestines) bud off. Forkhead box A2 (Foxa2, also known as HNF3β) [10] and SRY-box 17 (Sox17) [11] are required for DE development. C-X-C motif chemokine receptor 4 (Cxcr4) is expressed in the DE of gastrulating embryos (E7.2–7.8) [12].

The gut tube separates into the foregut, midgut, and hindgut along the anterior-posterior axis (Fig. 1) [13]. The foregut gives rise to the esophagus, trachea, stomach, lungs, thyroid glands, PTGs, liver, biliary system, and pancreas, while the midgut forms the small intestine and the hindgut forms the large intestine. The most rostral foregut endoderm is the anterior foregut endoderm (AFE). The caudal AFE gives rise to the lungs and trachea [13], while the more rostral AFE forms the pharyngeal endoderm (PE). SRY-box 2 (Sox2) is well known for its essential role in maintaining the ICM/epiblast [14] and the pluripotent state in PSCs [15]. In addition, Sox2 plays an important role in AFE development [16].

Fig. 1  Endoderm development

The foregut gives rise to the esophagus, trachea, stomach, lungs, thyroid glands, parathyroid glands, liver, biliary system, and pancreas. The midgut forms the small intestine, while the hindgut forms the large intestine. Parathyroid glands are derived from rostral anterior foregut endoderm.

The pharyngeal arches appear between E8.5–10 of mouse embryogenesis [17], while PTG and thymus development occurs between E10.5–15.5 [18]. The pharyngeal endoderm covers the internal surface of the pharyngeal arches [17]. The internal endodermal pockets between the arches are known as pharyngeal pouches (PP) (Fig. 2). The PTGs and thymus originate from the third and fourth PP [19]. The third PP forms between E9.5–10.5 and generates the inferior PTGs [20]. In most mammals, the fourth PP gives rise to a second pair of PTGs, the superior PTGs. However, in rodents, the fourth PP does not form PTGs, resulting in a single pair of PTGs.

Fig. 2  Developmental genetics of 3rd pharyngeal pouches (PP) and parathyroid glands (PTGs)

In mice, the PTGs develop from the 3rd PP with the thymus. Gata3 and Tbx1 are important for the development of PPs. Hoxa3, Pax9, and Eya1/Six1 are involved in the formation of PTG primordium. Gcm2 is the specific gene for PTG development and maintains the expression of other PTG related genes, including CaSR and Pth.

T-box 1 (Tbx1) is essential for the development of the pharyngeal arches and pouches (Table 1). It is first observed in the egg cylinder at E7.5 [21]. At E9.5, its expression is observed in the mesenchyme of the first, second and third pharyngeal arches, as well as in the epithelium of the first, second, and third PPs [21]. Homozygous Tbx1 deficiency results in a complete loss of the PTGs and thymus, along with aortic arch and cardiac outflow tract defects, similar to 22q11.2 deletion syndrome, also known as DiGeorge syndrome [22].

Table 1 Genes related to parathyroid glands (PTGs) and thymus organogenesis

Gene symbol		Full gene name	Expression site	Expression timing	Phenotypes of mutant mouse	Related human disease
Mouse	Human
Eya1	EYA1	EYA transcriptional coactivator and phosphatase 1	Pharyngeal endoderm (PE)	E9.5–10.5	Loss of PTGs and thymus primordium at E12.5 Loss of PTGs, thymus, ears, and kidneys, and hypoplasia of thyroid glands Reduced sine oculis-related homeobox 1 (Six1) expression in the PE	Branchio-oto-renal (BOR) syndrome
Foxn1	FOXN1	Forkhead box N1	Thymus primordium	E11.25	Hairless and loss of thymus (well-known as nude mouse)	Severe combined immunodeficiency
Gata3	GATA3	GATA binding protein 3	PE PTG primordium	E8.5 E11.5	Small third pharyngeal pouches (PP) at E11.5 Loss of PTGs and thymus primordium at E12.5 Embryonic lethal by E12.5–13.5	HDR syndrome (hypoparathyroidism, sensorineural deafness and renal disease)
Gcm2	GCM2	Glial cells missing2	PE PTG primordium	E9.5 E11.5	Loss of PTGs Arrested expression of CasR and CCL21 at E11.5	Genetic hypoparathyroidism
Hoxa3	HOXA3	Homeobox A3	PE	E10.5	Loss of PTGs and thymus, hypoplasia of thyroid glands, craniofacial abnormalities, malformations of the heart and great vessels Gcm2 was undetectable by E11.5
Pax1	PAX1	Paired box 1	PE	E9.5	Hypoplastic PTGs and thymus Gcm2 expression is reduced by E11.5
Pax9	PAX9	Paired box 9	PE	E9.5	The third and fourth PPs appeared normal at E10, but were retarded from E11.5 Loss of PTGs and thymus primordium at E12.0
Tbx1	TBX1	T-box 1 (mouse) T-box transcription factor 1 (human)	PE	E8.5–9.5	Loss of PTGs and thymus, cardiac outflow tract abnormalities, abnormal facial structures, abnormal vertebrae and cleft palate	22q11.2 deletion syndrome also known as DiGeorge syndrome

GATA binding protein 3 (Gata3) is a dual zinc-finger transcription factor that regulates the development of a number of organs, including PTGs. Homozygous Gata3 mutants showed a very small third pouch at E11.5 and fail to develop PTG-thymus primordia by E12.5 due to the absence of Glial cells missing2 (Gcm2) expression [23]. Moreover, they display severe internal bleeding, growth retardation, and extensive brain and neural tube defects, resulting in embryonic lethality around E12.5–13.5 [24].

Both ectodermal and endodermal epithelial cells of the pharyngeal arches and pouches express several sets of homeobox (Hox) genes. Hox A3 (Hoxa3, also known as Hox-1.5) is expressed in the endodermal epithelial cells of the third and fourth PPs at E10.5 [25]. Hoxa3 homozygous mutants exhibit multiple abnormalities in the pharyngeal tissues derived from the third and fourth PPs, including the absence of PTGs and thymus [25-27]. Expression of both Gcm2 and Tbx1 is reduced at E10.5, and Gcm2 expression becomes undetectable by E11.5 [28].

Paired box 1 (Pax1) and Pax9 show overlapping expression in the endoderm of the PPs. At E9.5, Pax1 is expressed in the first two PPs [29]. By E10.5, its expression is clearly observed in the third PP [29]. Loss of Pax1 function results in hypoplastic PTGs and thymus [30, 31]. Gcm2 expression is reduced in Pax1 mutants by E11.5. The PTGs in Pax1 mutants do not significantly differ from controls until after E13.5 but become severely hypoplastic by the neonatal stage [31]. Loss of Pax9 leads to retardation of the third and fourth PP at E11.5 [32]. From E12.0 onward, PTG and thymus primordia are absent in Pax9 mutants. Both Pax1 and Pax9 expression is downregulated in Hoxa3 mutants, indicating that Hoxa3 plays a master role in initiating PTG development [25, 31].

The eyes absent gene (Eya1) and sine oculis-related homeobox 1 (Six1) play essential roles in the organogenesis of the PPs. At E10.5, Eya1 expression is observed in the epithelium of the second, third, and fourth PPs [33]. Eya1 heterozygous mutants show renal abnormalities and hearing loss, similar to branchio-oto-renal (BOR) syndrome, whereas Eya1 homozygous mutants lack PTGs, thymus, ears, and kidneys [34]. The third and fourth PPs are present, and the expression of Hoxa3, Pax1, and Pax9 remains unaffected [18]. However, Six1 expression is significantly reduced in the endoderm of the second, third, and fourth PPs in Eya1 mutant embryos at E9.5–10.5. As a result, the PTG-thymus primordia fail to form at E12.5.

Gcm2 is an essential and specific gene for PTG development. Gcm2 is first expressed in the third PP as early as E9.5 [35]. By E10.5, Gcm2 expression becomes restricted to the anterior part of the third PP. Forkhead box N1 (Foxn1), a representative thymus marker, is the gene mutated in the classical nude mouse strain, which is hairless and lacks a thymus [36]. By E11.5, Foxn1 is expressed in the ventral-posterior part of the third PP [37], whereas the dorsal-anterior part expresses Gcm2. At E13.5, the thymus migrates caudally, while the PTGs remain lateral to the thyroid glands.

Günther et al. proposed that disruption of Gcm2 leads to a complete loss of PTGs [38]. In Gcm2^–/– mutants, the PTG primordium is present before E12 [39]. However, Gcm2^–/– mutants fail to maintain the expression of other early PTG marker genes, including the Ca sensing receptor (CaSR) and CCL21, and parathyroid hormone (Pth), a late PTG marker, is never detected. Finally, the PTG primordium undergoes programmed cell death between E12–12.5 in Gcm2^–/– mutants [39].

Generation of PTGs from human PSCs in vitro

Various methods have been reported for inducing endocrine cells and tissues from mammalian PSCs, including the hypothalamus, anterior and posterior pituitary, thyroid glands, and adrenal glands. However, only a limited number of methods focus on the induction of PTGs (Table 2).

Table 2 Representative methods to produce parathyroid glands (PTGs) from human pluripotent stem cells (PSCs) in vitro

	Species	Cell types	2-dimensional (2D) or 3-dimensional (3D)	Representative differentiation markers			Parathyroid hormone (PTH) expression at protein level	Hormone regulation
				Definitive endoderm	Pharyngeal endoderm	PTGs
Bingham et al., 2009	Human	Embryonic stem cells (ESCs)	2D	C-X-C motif chemokine receptor 4 (CXCR4), Forkhead box A2 (FOXA2), SRY-box 17 (SOX17) (protein) CXCR4 SOX17	EYA transcriptional coactivator and phosphatase 1 (EYA1)/sine oculis-related homeobox 1 (SIX1), homeobox A3 (HOXA3), paired box 1 (PAX1)	calcium sensing receptor (CASR), glial cells missing transcription factor 2 (GCM2), PTH (protein) CaSR PTH	Yes (determined by ELISA)	Unknown
Green et al., 2011	Human	ESCs	EB formation in 3D culture→2D	FOXA2, SRY-box transcription factor 2 (SOX2), SOX17 (protein) CXCR4, FOXA2 SOX2	PAX1, paired box 9 (PAX9), T-box 1 (TBX1)	GCM2	No	Unknown
Lawton et al., 2020	Human	ESCs and induced PSCs (iPSCs)	2D	FOXA2, SOX17 (protein) SOX17	GATA binding protein 3 (GATA3), HOXA3, PAX9, EYA1/SIX1	CASR, GCM2, PTH	No	Unknown
Nakatsuka et al., 2023	Human	iPSCs	2D	FOXA2, SOX2, SOX17 (protein) CXCR4	HOXA3, TBX1	CASR, GCM2, PTH (protein) CaSR GCM2 PTH	Yes (determined by immunostaining)	Yes
Şenkal-Turhan et al., 2024	Human	iPSCs	EB formation in 3D culture→Matrigel-based 3D organoid	(protein) FOXA2	GATA3	CASR, GCM2, PTH (protein) CaSR GCM2 PTH	Yes (determined by dot blot analysis, immunostaining, western blot)	Yes

In Bingham’s protocol [40, 41], human ESCs were cultured in high-dose activin A (100 ng/mL) and 5% fetal bovine serum (FBS) for 12 days, then in activin A-free medium for an additional two weeks. PTH secretion in the culture medium was confirmed at the protein level using enzyme-linked immunosorbent assay (ELISA). However, one group reported that Bingham’s protocol was not reproducible [42], possibly due to the molecules present in FBS (i.e., growth factors, hormones, and nutrients), which may have affected the experiments.

Green et al. proposed a method for AFE differentiation from human ESCs [43]. Undifferentiated human ESCs were first differentiated into DE using high concentrations of activin A (days 1 to 5), forming embryoid bodies (EBs) on low-adhesion dishes. At day 5, these EBs were dissociated and plated as a monolayer. In the monolayer culture, the DE cells were induced into AFE using noggin (a bone morphogenetic protein (BMP) inhibitor) and SB-431542 (an activin A/nodal and transforming growth factor (TGF)-β inhibitor) from days 5 to 7. The noggin/SB-431542-treated AFE culture expressed SOX2, TBX1, and PAX9 at both RNA and protein levels.

Next, the AFE culture was further differentiated into ventral AFE in the presence of Wnt family member 3a (WNT3a), keratinocyte growth factor (KGF), fibroblast growth factor 10 (FGF10), BMP4, and epidermal growth factor (EGF) (collectively referred to as WKFBE) from days 7 to 10. This culture expressed NK2 homeobox 1 (NKX2-1), a marker for lung and thyroid gland progenitors, as well as PAX1. However, the ventral AFE (noggin/SB-431542 treated AFE culture plus WKFBE) culture did not achieve final differentiation into the thymus, PTGs, thyroid glands or lungs.

Sonic hedgehog (SHH) and FGF8 play important roles in patterning the third pouch [37, 44]. From days 11 to 19, the addition of SHH or FGF8 to ventral AFE cultures (noggin/SB-431542-induced AFE plus WKFBE) led to GCM2 mRNA expression by day 19. However, PTH expression has not been demonstrated at either the RNA or protein level.

In Lawton’s protocol [42], undifferentiated human ESCs and iPSCs were differentiated into DE using high concentrations of activin A, Wnt3a, and the CDK inhibitor PD-0332991 (days 2 to 5), based on previous studies. Wnt3a has been shown to be required for endoderm formation during embryogenesis and in vitro differentiation [45]. The PD-0332991 treatment decreases the pluripotency of human ESCs and promotes endoderm differentiation [46].

Subsequently, AFE induction was performed based on established protocols [43, 47]. The DE culture was treated with noggin/SB-431542 on days 6 and 7, followed by IWP2, an inhibitor of endogenous Wnts, on days 8 and 9. PD-0332991 treatment was continued until the end of AFE induction. Relative expression of NKX2.3, which is essential for PP development [48], increased 19-fold compared to the original human ESCs.

For PE induction, AFE cultures were cultured from days 10 to 23 with a combination of LY-364947 (a TGF-β signaling inhibitor), all-trans retinoic acid (ATRA), FGF10, cyclopamine (a SHH signaling inhibitor), and BMP4. During PTG organogenesis, Bmp4 is expressed in the ventral region of the third PP, which gives rise to the thymus [20]. In contrast, Noggin is expressed in the dorsal region, which forms the PTGs. BMP4 was therefore inhibited with NOGGIN in PE culture after day 24.

Relative expression of PTH increased 23-fold compared to original human ESCs. Furthermore, a culture derived from one iPSC line showed a 196-fold increase in PTH and a 10,900-fold increase in GCM2 using the same protocol. PTH expression at the protein level was not assessed in Lawton’s protocol.

In Nakatsuka’s protocol [49], human iPSCs were induced into DE and AFE using high concentrations of activin A and CHIR-99021 (a Wnt/β-catenin pathway activator) on day 1, followed by the same dose of activin A and LDN-193189 (a BMP receptor inhibitor) on days 2 and 3, based on a previous DE induction method [50]. SOX2 expression appeared to be bimodal, similar to previous in vitro data [51].

For PE induction, AFE cells were cultured with ATRA and IWRl-endo (a Wnt inhibitor) from days 6 to 10. Finally, parathyroid cells were induced by adding SHH and activin A to the PE culture from days 10 to 24. PTH expression was confirmed at both RNA and protein levels. These parathyroid cells expressed CaSR, and the expression of PTH and GCM2 appeared to be downregulated as extracellular Ca increased, indicating that they were functional.

In Şenkal-Turhan’s protocol [52], human iPSCs were differentiated into 3D parathyroid organoids based on Bingham’s protocol [40]. Human iPSCs were cultured as floating cultures in high-dose activin A (100 ng/mL) and FBS for 5 days. On day 5 of differentiation, these spheroids were transferred into a 3D Matrigel dome. From days 10 to 20, 50 ng/mL SHH was added to the culture medium. The parathyroid organoids on day 20 expressed CasR, GCM2, and PTH at both RNA and protein levels. The concentration of PTH in the medium was assessed using Dot Blot analysis. The expression of Erk, p-Erk, and PTH, evaluated by Western blotting, appeared to be regulated in response to extracellular Ca concentration. Parathyroid organoids were transplanted into parathyroidectomized rats. Immunostaining of grafts on day 14 demonstrated expression of PTG-related markers (CasR, CxCr4, Foxn1, Gcm2, and PTH). However, the therapeutic potential of parathyroid organoids has not yet been fully evaluated.

Generation of PTGs in vivo

Blastocyst complementation (BC) is an in vivo organ generation strategy using PSCs. The concept of BC was first proposed in lymphocyte complementation using recombination-activating gene 2 (Rag2)-deficient mouse embryos [53]. Injection of wildtype mouse embryonic stem cells (mESCs) into Rag2-knockout (KO) blastocysts leads to the generation of chimeric mice with mature B and T cells, all of which derive from the injected mESCs. The Rag2-KO BC model was considered useful as a functional test for lymphocytes.

Kobayashi et al. demonstrated that BC can be applied to organ complementation [54]. First, specific organ-deficient animals are produced by knocking out a gene essential for organ development, creating a vacant organ developmental niche during organogenesis. Then, allogenic and xenogeneic PSCs are injected into organ-deficient animal embryos. As a result, the injected donor PSCs differentiate with host-derived cells in a coordinated manner and fill the vacant developmental niche. The resulting chimeric animals develop organs totally derived from donor PSCs.

Our first success was achieved in the pancreas [54]. Pancreatic and duodenal homeobox1 (Pdx1) plays a critical role in pancreatic development [55, 56]. Kobayashi et al. hypothesized that Pdx1^–/– blastocysts could provide a pancreatic developmental niche for BC. When mouse PSCs were injected into Pdx1 KO mouse blastocysts (mouse PSCs→mouse blastocysts), both pancreatic exocrine and endocrine tissues were almost entirely composed of injected mouse PSC-derived cells. The PSC-derived β-cells produced insulin and normalized blood glucose in Pdx1^–/– mice complemented with mPSCs. They further succeeded in interspecific BC between mouse and rat. Injection of rat iPSCs into Pdx1^–/– mouse embryos (rat PSCs→ mouse blastocysts) yielded interspecific chimeric mice with a pancreas derived from rat iPSCs. Pdx1^–/– mice complemented with rat iPSCs demonstrated normal glucose tolerance, indicating that xenogeneic iPSC-derived β-cells are functional.

Yamaguchi et al. performed a reciprocal experiment (mouse PSCs → rat blastocysts) [57]. By injecting mouse PSCs into Pdx1 KO rat blastocysts, we generated a rat-sized pancreas composed of injected mouse PSC-derived cells. Mouse PSC-derived β-cells generated in Pdx1 KO rats normalized blood glucose in streptozotocin-induced diabetic mice through islet transplantation. Of note, the use of immunosuppressants was limited to the first five days following islet transplantation. These research results pave the way for generating organs derived from human PSCs in animal bodies, with the potential for transplantation to be achieved using immunosuppressants for very short periods, or not at all.

Following the success in the pancreas, BC has been reported in other organs, including blood vessels [58], forebrain [59], gametes [60], heart [61], kidneys [62], lungs [63-65], skin [66], and thyroid glands [64] (Fig. 3). All of the studies mentioned above used rodents. Allogeneic BC has also succeeded in pigs through a combination of gene editing and nuclear transfer [67, 68].

Fig. 3  Various organs generated using blastocyst complementation (BC)

In the BC strategy, allogeneic or xenogeneic pluripotent stem cells are injected into specific organ-deficient animals produced by knockout of a gene. The organs generated through this approach, along with their associated target genes, are illustrated. All of these studies were conducted using rodents, specifically mice and rats.

We have recently generated functional PTGs in rodents using BC (Fig. 4) [69]. The experiment consisted of two processes. First, PTG-deficient embryos were generated by CRISPR-Cas9-mediated zygote Gcm2 KO. Mouse ESCs were then injected into PTG-deficient embryos to generate chimeric mice with mESC-derived PTGs (mouse PSCs→ mouse blastocysts).

Fig. 4  Generation of functional parathyroid glands (PTGs) from mouse embryonic stem cells (mESCs) using blastocyst complementation (BC)

(A) Parathyroid gland-deficient embryos were generated by CRISPR-Cas9-mediated zygote Gcm2 knockout (KO). The resulting Gcm2 KO mice die shortly after birth (upper). Blastocyst complementation for generating mESC-derived PTGs in Gcm2 KO aparathyroid mice. The resulting chimeric mice can survive (lower). (B) Mouse-derived PTGs effectively restored plasma Ca levels in mice with hypoparathyroidism following parathyroidectomy.

Mouse Gcm2 has five exons, with exons 2 and 3 encoding the entire DNA-binding domain known as the gcm-motif [70, 71]. The N-terminal regions of the gcm-motif are highly conserved across species. We, therefore, targeted the gcm-motif to delete Gcm2 function entirely using CRISPR-Cas9-mediated KO. The CRISPR-Cas9-mediated zygote Gcm2 KO efficiently produced PTG-deficient embryos. Histological examination and qPCR for Pth sequences confirmed the complete loss of PTGs in Gcm2 KO embryos.

To demonstrate that PTGs generated by BC were derived from donor ES cells, we established mESCs with a knock-in of 2A-linked-tdTomato directly following the coding sequence of Pth in CAG-EGFP transgenic mESCs [57]. The established mESCs were injected into Gcm2 KO embryos. The resulting chimeric mice (Gcm2^–/– mice complemented with PTh-tdTomato mESCs) had tdTomato-positive PTGs and survived into adulthood without the symptoms related to hypoparathyroidism.

Mouse ESC-derived PTGs regulated PTH release in response to external Ca concentration, demonstrating their functionality. Furthermore, mouse ESC-derived PTGs grafted beneath the renal capsule of post-parathyroidectomy mice ameliorated the host’s hypoparathyroidism.

In contrast to the many successes in rodent BC, some hurdles still remain in human organ generation via BC. Although human–animal chimeras (human PSCs→ animal blastocysts) have been reported, the contribution rate of human cells in interspecies chimeras was very low (~0.001–0.01%) [72, 73]. Several reasons have been proposed [74], including differences in the cellular status of PSCs between humans and rodents, as well as the larger evolutionary distance between humans and rodents or humans and pigs, compared to that between rats and mice. Despite the challenges, progress has been made in human-animal chimera generation (Fig. 5). Masaki et al. reported that developmental stage-mismatched cells undergo apoptosis when injected into preimplantation embryos [75]. Overexpression of the anti-apoptotic gene BCL2 enables stage-mismatched cells to contribute to chimeras [76], and this effect is even greater in combination with the cell growth factor MYCN [77]. Therefore, overexpression of these genes in human PSCs would increase the efficiency of chimera formation between human PSCs and porcine embryos. Das et al. successfully generated human endothelium in ETV2-null pig embryos at E17 and E18 [78]. When BCL2-overexpressing human PSCs were injected into ETV2-null pig embryos, which lack the hematopoietic and endothelial lineages, all endothelial cells were of human origin. A subsequent study from the same group demonstrated the generation of human skeletal muscle tissue using TP53-null human iPSCs in MYF5/MYOD/MYF6-null pig embryos at E20 and E27 [79]. They found that expression of TP53, inducing apoptosis, was upregulated in the human embryos compared to the pig embryos. To achieve increased efficiency of interspecies chimerism, they deleted TP53 in human iPSCs. Wang et al. showed that 4CL (4 chemicals + LIF) [80] medium and BCL2/MYCN overexpression (4CL/N/B) improve the interspecies contribution potential of human iPSCs [81]. When 4CL/N/B human PSCs were injected into SIX1/SALL1-null nephric-defective pig embryos, 40–60% of mesonephric cells were of human origin.

Fig. 5  Current attempts in BC between human induced pluripotent stem cells (iPSCs) and pig embryos to generate human organs

(A) Generation of human endothelium in pig embryos deficient in ETV2 (E17 and E18) using human iPSCs that overexpress BCL2.

(B) The same group created human skeletal muscle in MYF5/MYOD/MYF6 KO pig embryos (E20 and E27) using TP53-null human iPSCs.

(C) Humanized mesonephros were produced in SIX1/SALL1 KO nephric-defective pigs using human iPSCs that overexpress BCL2 and MYCN (E25 and E28).

Conclusion

Recent advancements have been made in generating PTGs from PSCs both in vitro and in vivo (Graphical Abstract). PTGs derived from PSCs hold potential not only for regenerative medicine but also as valuable tools for basic endocrine research. For example, attempts have been made to generate pituitary adenomas from human PSCs in vitro using the established induction method [82]. Chang et al. proposed a BC-based approach to generate mouse models for studying forebrain functions [59, 83]. Whichever method is ultimately utilized, future success in generating PTGs from human PSCs could lead to significant advances in the treatment of hypoparathyroidism and enhance our understanding of endocrine research.

Graphical Abstract 

Grants

This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI Grants JP21K16337 and 24K19296; the Japan Neuroendocrine Society; the Japan Medical Women’s Association; the Takeda Science Foundation; the Uehara Memorial Foundation; the Yamaguchi Endocrine Research Foundation; the Yasuda Medical Foundation; and the Yokohama Foundation for Advancement of Medical Science. All these grants were awarded to M.K.

Acknowledgment

BioRender (https://biorender.com) was used in the creation of the figures.

Disclosure

No potential conflict of interest relevant to this article was reported.

References

• 1   Clarke  BL,  Brown  EM,  Collins  MT,  Jüppner  H,  Lakatos  P, et al. (2016) Epidemiology and Diagnosis of Hypoparathyroidism. J Clin Endocrinol Metab 101: 2284–2299.
• 2   Hasegawa  M,  Sakakibara  Y,  Takeuchi  Y,  Sugitani  I,  Ozono  K, et al. (2024) Prevalence and characteristics of postoperative and nonoperative chronic hypoparathyroidism in Japan: a nationwide retrospective analysis. JBMR Plus 8: ziae100.
• 3   Mannstadt  M,  Bilezikian  JP,  Thakker  RV,  Hannan  FM,  Clarke  BL, et al. (2017) Hypoparathyroidism. Nat Rev Dis Primers 3: 17055.
• 4   Mitchell  DM,  Regan  S,  Cooley  MR,  Lauter  KB,  Vrla  MC, et al. (2012) Long-term follow-up of patients with hypoparathyroidism. J Clin Endocrinol Metab 97: 4507–4514.
• 5   Chawla  H,  Saha  S,  Kandasamy  D,  Sharma  R,  Sreenivas  V, et al. (2017) Vertebral fractures and bone mineral density in patients with idiopathic hypoparathyroidism on long-term follow-up. J Clin Endocrinol Metab 102: 251–258.
• 6   Büttner  M,  Musholt  TJ,  Singer  S (2017) Quality of life in patients with hypoparathyroidism receiving standard treatment: a systematic review. Endocrine 58: 14–20.
• 7   Rejnmark  L,  Gosmanova  EO,  Khan  AA,  Makita  N,  Imanishi  Y, et al. (2024) Palopegteriparatide treatment improves renal function in adults with chronic hypoparathyroidism: 1-year results from the phase 3 PaTHway trial. Adv Ther 41: 2500–2518.
• 8   Khan  AA,  Rubin  MR,  Schwarz  P,  Vokes  T,  Shoback  DM, et al. (2023) Efficacy and safety of parathyroid hormone replacement with TransCon PTH in hypoparathyroidism: 26-week results from the phase 3 PaTHway trial. J Bone Miner Res 38: 14–25.
• 9   Takahashi  K,  Yamanaka  S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126: 663–676.
• 10   Ang  SL,  Wierda  A,  Wong  D,  Stevens  KA,  Cascio  S, et al. (1993) The formation and maintenance of the definitive endoderm lineage in the mouse: involvement of HNF3/forkhead proteins. Development 119: 1301–1315.
• 11   Kanai-Azuma  M,  Kanai  Y,  Gad  JM,  Tajima  Y,  Taya  C, et al. (2002) Depletion of definitive gut endoderm in Sox17-null mutant mice. Development 129: 2367–2379.
• 12   McGrath  KE,  Koniski  AD,  Maltby  KM,  McGann  JK,  Palis  J (1999) Embryonic expression and function of the chemokine SDF-1 and its receptor, CXCR4. Dev Biol 213: 442–456.
• 13   Zorn  AM,  Wells  JM (2009) Vertebrate endoderm development and organ formation. Annu Rev Cell Dev Biol 25: 221–251.
• 14   Avilion  AA,  Nicolis  SK,  Pevny  LH,  Perez  L,  Vivian  N, et al. (2003) Multipotent cell lineages in early mouse development depend on SOX2 function. Genes Dev 17: 126–140.
• 15   Boyer  LA,  Lee  TI,  Cole  MF,  Johnstone  SE,  Levine  SS, et al. (2005) Core transcriptional regulatory circuitry in human embryonic stem cells. Cell 122: 947–956.
• 16   Que  J,  Okubo  T,  Goldenring  JR,  Nam  KT,  Kurotani  R, et al. (2007) Multiple dose-dependent roles for Sox2 in the patterning and differentiation of anterior foregut endoderm. Development 134: 2521–2531.
• 17   Cesario  JM,  Malt  AL,  Jeong  J (2015) Developmental genetics of the pharyngeal arch system. Morgan and Claypool, San Rafael, USA.
• 18   Xu  PX,  Zheng  W,  Laclef  C,  Maire  P,  Maas  RL, et al. (2002) Eya1 is required for the morphogenesis of mammalian thymus, parathyroid and thyroid. Development 129: 3033–3044.
• 19   Grevellec  A,  Tucker  AS (2010) The pharyngeal pouches and clefts: development, evolution, structure and derivatives. Semin Cell Dev Biol 21: 325–332.
• 20   Patel  SR,  Gordon  J,  Mahbub  F,  Blackburn  CC,  Manley  NR (2006) Bmp4 and Noggin expression during early thymus and parathyroid organogenesis. Gene Expr Patterns 6: 794–799.
• 21   Chapman  DL,  Garvey  N,  Hancock  S,  Alexiou  M,  Agulnik  SI, et al. (1996) Expression of the T-box family genes, Tbx1-Tbx5, during early mouse development. Dev Dyn 206: 379–390.
• 22   Jerome  LA,  Papaioannou  VE (2001) DiGeorge syndrome phenotype in mice mutant for the T-box gene, Tbx1. Nat Genet 27: 286–291.
• 23   Grigorieva  IV,  Mirczuk  S,  Gaynor  KU,  Nesbit  MA,  Grigorieva  EF, et al. (2010) Gata3-deficient mice develop parathyroid abnormalities due to dysregulation of the parathyroid-specific transcription factor Gcm2. J Clin Invest 120: 2144–2155.
• 24   Pandolfi  PP,  Roth  ME,  Karis  A,  Leonard  MW,  Dzierzak  E, et al. (1995) Targeted disruption of the GATA3 gene causes severe abnormalities in the nervous system and in fetal liver haematopoiesis. Nat Genet 11: 40–44.
• 25   Manley  NR,  Capecchi  MR (1995) The role of Hoxa-3 in mouse thymus and thyroid development. Development 121: 1989–2003.
• 26   Chisaka  O,  Capecchi  MR (1991) Regionally restricted developmental defects resulting from targeted disruption of the mouse homeobox gene hox-1.5. Nature 350: 473–479.
• 27   Chisaka  O,  Musci  TS,  Capecchi  MR (1992) Developmental defects of the ear, cranial nerves and hindbrain resulting from targeted disruption of the mouse homeobox gene Hox-1.6. Nature 355: 516–520.
• 28   Chojnowski  JL,  Masuda  K,  Trau  HA,  Thomas  K,  Capecchi  M, et al. (2014) Multiple roles for HOXA3 in regulating thymus and parathyroid differentiation and morphogenesis in mouse. Development 141: 3697–3708.
• 29   Wallin  J,  Eibel  H,  Neubüser  A,  Wilting  J,  Koseki  H, et al. (1996) Pax1 is expressed during development of the thymus epithelium and is required for normal T-cell maturation. Development 122: 23–30.
• 30   Dietrich  S,  Gruss  P (1995) undulated phenotypes suggest a role of Pax-1 for the development of vertebral and extravertebral structures. Dev Biol 167: 529–548.
• 31   Su  D,  Ellis  S,  Napier  A,  Lee  K,  Manley  NR (2001) Hoxa3 and pax1 regulate epithelial cell death and proliferation during thymus and parathyroid organogenesis. Dev Biol 236: 316–329.
• 32   Peters  H,  Neubüser  A,  Kratochwil  K,  Balling  R (1998) Pax9-deficient mice lack pharyngeal pouch derivatives and teeth and exhibit craniofacial and limb abnormalities. Genes Dev 12: 2735–2747.
• 33   Kalatzis  V,  Sahly  I,  El-Amraoui  A,  Petit  C (1998) Eya1 expression in the developing ear and kidney: towards the understanding of the pathogenesis of Branchio-Oto-Renal (BOR) syndrome. Dev Dyn 213: 486–499.
• 34   Xu  PX,  Adams  J,  Peters  H,  Brown  MC,  Heaney  S, et al. (1999) Eya1-deficient mice lack ears and kidneys and show abnormal apoptosis of organ primordia. Nat Genet 23: 113–117.
• 35   Gordon  J,  Bennett  AR,  Blackburn  CC,  Manley  NR (2001) Gcm2 and Foxn1 mark early parathyroid- and thymus-specific domains in the developing third pharyngeal pouch. Mech Dev 103: 141–143.
• 36   Nehls  M,  Pfeifer  D,  Schorpp  M,  Hedrich  H,  Boehm  T (1994) New member of the winged-helix protein family disrupted in mouse and rat nude mutations. Nature 372: 103–107.
• 37   Gordon  J,  Manley  NR (2011) Mechanisms of thymus organogenesis and morphogenesis. Development 138: 3865–3878.
• 38   Günther  T,  Chen  ZF,  Kim  J,  Priemel  M,  Rueger  JM, et al. (2000) Genetic ablation of parathyroid glands reveals another source of parathyroid hormone. Nature 406: 199–203.
• 39   Liu  Z,  Yu  S,  Manley  NR (2007) Gcm2 is required for the differentiation and survival of parathyroid precursor cells in the parathyroid/thymus primordia. Dev Biol 305: 333–346.
• 40   Bingham  EL,  Cheng  SP,  Woods Ignatoski  KM,  Doherty  GM (2009) Differentiation of human embryonic stem cells to a parathyroid-like phenotype. Stem Cells Dev 18: 1071–1080.
• 41   Woods Ignatoski  KM,  Bingham  EL,  Frome  LK,  Doherty  GM (2010) Differentiation of precursors into parathyroid-like cells for treatment of hypoparathyroidism. Surgery 148: 1186–1189; discussion 1189–1190.
• 42   Lawton  BR,  Martineau  C,  Sosa  JA,  Roman  S,  Gibson  CE, et al. (2020) Differentiation of PTH-expressing cells from human pluripotent stem cells. Endocrinology 161: bqaa141.
• 43   Green  MD,  Chen  A,  Nostro  MC,  d’Souza  SL,  Schaniel  C, et al. (2011) Generation of anterior foregut endoderm from human embryonic and induced pluripotent stem cells. Nat Biotechnol 29: 267–272.
• 44   Moore-Scott  BA,  Manley  NR (2005) Differential expression of Sonic hedgehog along the anterior-posterior axis regulates patterning of pharyngeal pouch endoderm and pharyngeal endoderm-derived organs. Dev Biol 278: 323–335.
• 45   D’Amour  KA,  Bang  AG,  Eliazer  S,  Kelly  OG,  Agulnick  AD, et al. (2006) Production of pancreatic hormone-expressing endocrine cells from human embryonic stem cells. Nat Biotechnol 24: 1392–1401.
• 46   Pauklin  S,  Vallier  L (2013) The cell-cycle state of stem cells determines cell fate propensity. Cell 155: 135–147.
• 47   Huang  SX,  Green  MD,  de Carvalho  AT,  Mumau  M,  Chen  YW, et al. (2015) The in vitro generation of lung and airway progenitor cells from human pluripotent stem cells. Nat Protoc 10: 413–425.
• 48   Li  L,  Ning  G,  Yang  S,  Yan  Y,  Cao  Y, et al. (2019) BMP signaling is required for nkx2.3-positive pharyngeal pouch progenitor specification in zebrafish. PLoS Genet 15: e1007996.
• 49   Nakatsuka  R,  Kato  T,  Zhang  R,  Uemura  Y,  Sasaki  Y, et al. (2023) The induction of parathyroid cell differentiation from human induced pluripotent stem cells promoted via TGF-α/EGFR signaling. Stem Cells Dev 32: 670–680.
• 50   Matsuno  K,  Mae  SI,  Okada  C,  Nakamura  M,  Watanabe  A, et al. (2016) Redefining definitive endoderm subtypes by robust induction of human induced pluripotent stem cells. Differentiation 92: 281–290.
• 51   Brafman  DA,  Moya  N,  Allen-Soltero  S,  Fellner  T,  Robinson  M, et al. (2013) Analysis of SOX2-expressing cell populations derived from human pluripotent stem cells. Stem Cell Reports 1: 464–478.
• 52   Şenkal-Turhan  S,  Bulut-Okumuş  E,  Aydın  M,  Başak Türkmen  N,  Taşlıdere  A, et al. (2024) Induced pluripotent stem cell-derived parathyroid organoids resemble parathyroid morphology and function. Adv Sci (Weinh) 11: e2407567.
• 53   Chen  J,  Lansford  R,  Stewart  V,  Young  F,  Alt  FW (1993) RAG-2-deficient blastocyst complementation: an assay of gene function in lymphocyte development. Proc Natl Acad Sci U S A 90: 4528–4532.
• 54   Kobayashi  T,  Yamaguchi  T,  Hamanaka  S,  Kato-Itoh  M,  Yamazaki  Y, et al. (2010) Generation of rat pancreas in mouse by interspecific blastocyst injection of pluripotent stem cells. Cell 142: 787–799.
• 55   Jonsson  J,  Carlsson  L,  Edlund  T,  Edlund  H (1994) Insulin-promoter-factor 1 is required for pancreas development in mice. Nature 371: 606–609.
• 56   Offield  MF,  Jetton  TL,  Labosky  PA,  Ray  M,  Stein  RW, et al. (1996) PDX-1 is required for pancreatic outgrowth and differentiation of the rostral duodenum. Development 122: 983–995.
• 57   Yamaguchi  T,  Sato  H,  Kato-Itoh  M,  Goto  T,  Hara  H, et al. (2017) Interspecies organogenesis generates autologous functional islets. Nature 542: 191–196.
• 58   Hamanaka  S,  Umino  A,  Sato  H,  Hayama  T,  Yanagida  A, et al. (2018) Generation of vascular endothelial cells and hematopoietic cells by blastocyst complementation. Stem Cell Reports 11: 988–997.
• 59   Chang  AN,  Liang  Z,  Dai  HQ,  Chapdelaine-Williams  AM,  Andrews  N, et al. (2018) Neural blastocyst complementation enables mouse forebrain organogenesis. Nature 563: 126–130.
• 60   Kobayashi  T,  Goto  T,  Oikawa  M,  Sanbo  M,  Yoshida  F, et al. (2021) Blastocyst complementation using Prdm14-deficient rats enables efficient germline transmission and generation of functional mouse spermatids in rats. Nat Commun 12: 1328.
• 61   Coppiello  G,  Barlabé  P,  Moya-Jódar  M,  Abizanda  G,  Pogontke  C, et al. (2023) Generation of heart and vascular system in rodents by blastocyst complementation. Dev Cell 58: 2881–2895.e2887.
• 62   Goto  T,  Hara  H,  Sanbo  M,  Masaki  H,  Sato  H, et al. (2019) Generation of pluripotent stem cell-derived mouse kidneys in Sall1-targeted anephric rats. Nat Commun 10: 451.
• 63   Mori  M,  Furuhashi  K,  Danielsson  JA,  Hirata  Y,  Kakiuchi  M, et al. (2019) Generation of functional lungs via conditional blastocyst complementation using pluripotent stem cells. Nat Med 25: 1691–1698.
• 64   Wen  B,  Li  E,  Ustiyan  V,  Wang  G,  Guo  M, et al. (2021) In vivo generation of lung and thyroid tissues from embryonic stem cells using blastocyst complementation. Am J Respir Crit Care Med 203: 471–483.
• 65   Yuri  S,  Murase  Y,  Isotani  A (2024) Generation of rat-derived lung epithelial cells in Fgfr2b-deficient mice retains species-specific development. Development 151: dev202081.
• 66   Nagano  H,  Mizuno  N,  Sato  H,  Mizutani  E,  Yanagida  A, et al. (2024) Skin graft with dermis and appendages generated in vivo by cell competition. Nat Commun 15: 3366.
• 67   Matsunari  H,  Nagashima  H,  Watanabe  M,  Umeyama  K,  Nakano  K, et al. (2013) Blastocyst complementation generates exogenic pancreas in vivo in apancreatic cloned pigs. Proc Natl Acad Sci U S A 110: 4557–4562.
• 68   Matsunari  H,  Watanabe  M,  Hasegawa  K,  Uchikura  A,  Nakano  K, et al. (2020) Compensation of disabled organogeneses in genetically modified pig fetuses by blastocyst complementation. Stem Cell Reports 14: 21–33.
• 69   Kano  M,  Mizuno  N,  Sato  H,  Kimura  T,  Hirochika  R, et al. (2023) Functional calcium-responsive parathyroid glands generated using single-step blastocyst complementation. Proc Natl Acad Sci U S A 120: e2216564120.
• 70   Akiyama  Y,  Hosoya  T,  Poole  AM,  Hotta  Y (1996) The gcm-motif: a novel DNA-binding motif conserved in Drosophila and mammals. Proc Natl Acad Sci U S A 93: 14912–14916.
• 71   Kim  J,  Jones  BW,  Zock  C,  Chen  Z,  Wang  H, et al. (1998) Isolation and characterization of mammalian homologs of the Drosophila gene glial cells missing. Proc Natl Acad Sci U S A 95: 12364–12369.
• 72   Theunissen  TW,  Friedli  M,  He  Y,  Planet  E,  O’Neil  RC, et al. (2016) Molecular criteria for defining the naive human pluripotent state. Cell Stem Cell 19: 502–515.
• 73   Wu  J,  Platero-Luengo  A,  Sakurai  M,  Sugawara  A,  Gil  MA, et al. (2017) Interspecies chimerism with mammalian pluripotent stem cells. Cell 168: 473–486.e415.
• 74   Yamaguchi  T (2019) Hurdles to generating human islets in animals via blastocyst complementation. Curr Diab Rep 19: 45.
• 75   Masaki  H,  Kato-Itoh  M,  Umino  A,  Sato  H,  Hamanaka  S, et al. (2015) Interspecific in vitro assay for the chimera-forming ability of human pluripotent stem cells. Development 142: 3222–3230.
• 76   Masaki  H,  Kato-Itoh  M,  Takahashi  Y,  Umino  A,  Sato  H, et al. (2016) Inhibition of apoptosis overcomes stage-related compatibility barriers to chimera formation in mouse embryos. Cell Stem Cell 19: 587–592.
• 77   Zhu  Y,  Zhang  Z,  Fan  N,  Huang  K,  Li  H, et al. (2022) Generating functional cells through enhanced interspecies chimerism with human pluripotent stem cells. Stem Cell Reports 17: 1059–1069.
• 78   Das  S,  Koyano-Nakagawa  N,  Gafni  O,  Maeng  G,  Singh  BN, et al. (2020) Generation of human endothelium in pig embryos deficient in ETV2. Nat Biotechnol 38: 297–302.
• 79   Maeng  G,  Das  S,  Greising  SM,  Gong  W,  Singh  BN, et al. (2021) Humanized skeletal muscle in MYF5/MYOD/MYF6-null pig embryos. Nat Biomed Eng 5: 805–814.
• 80   Mazid  MA,  Ward  C,  Luo  Z,  Liu  C,  Li  Y, et al. (2022) Rolling back human pluripotent stem cells to an eight-cell embryo-like stage. Nature 605: 315–324.
• 81   Wang  J,  Xie  W,  Li  N,  Li  W,  Zhang  Z, et al. (2023) Generation of a humanized mesonephros in pigs from induced pluripotent stem cells via embryo complementation. Cell Stem Cell 30: 1235–1245.e1236.
• 82   Matsumoto  R,  Suga  H,  Arima  H,  Yamamoto  T (2022) Disease modeling of pituitary adenoma using human pluripotent stem cells. Cancers (Basel) 14: 3660.
• 83   Dai  HQ,  Liang  Z,  Chang  AN,  Chapdelaine-Williams  AM,  Alvarado  B, et al. (2020) Direct analysis of brain phenotypes via neural blastocyst complementation. Nat Protoc 15: 3154–3181.