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, 30 (13), 2507-15

Homogeneous and Organized Differentiation Within Embryoid Bodies Induced by Microsphere-Mediated Delivery of Small Molecules

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Homogeneous and Organized Differentiation Within Embryoid Bodies Induced by Microsphere-Mediated Delivery of Small Molecules

Richard L Carpenedo et al. Biomaterials.

Abstract

Cell specification and tissue formation during embryonic development are precisely controlled by the local concentration and temporal presentation of morphogenic factors. Similarly, pluripotent embryonic stem cells can be induced to differentiate in vitro into specific phenotypes in response to morphogen treatment. Embryonic stem cells (ESCs) are commonly differentiated as 3D spheroids referred to as embryoid bodies (EBs); however, differentiation of cells within EBs is typically heterogeneous and disordered. In this study, we demonstrate that in contrast to soluble morphogen treatment, delivery of morphogenic factors directly within EB microenvironments in a spatiotemporally controlled manner using polymer microspheres yields homogeneous, synchronous and organized ESC differentiation. Degradable PLGA microspheres releasing retinoic acid were incorporated directly within EBs and induced the formation of cystic spheroids uniquely resembling the phenotype and structure of early streak mouse embryos (E6.75), with an exterior of FOXA2+ visceral endoderm enveloping an epiblast-like layer of OCT4+ cells. These results demonstrate that controlled morphogen presentation to stem cells using degradable microspheres more efficiently directs cell differentiation and tissue formation than simple soluble delivery methods and presents a unique route to study the spatiotemporal effects of morphogenic factors on embryonic developmental processes in vitro.

Figures

Fig. 1
Fig. 1
EB structure and microsphere incorporation. (A) Day 10 EBs exhibited a smooth exterior in SEM micrographs, while bisected EBs displayed a dense outer layer (B, C). (D–F), Microspheres labeled with CellTracker Red were observed in multiple focal planes (30 μm between images), while soluble treatment of CellTracker Green labeled the outermost 1–2 cell layers. (G–I) EBs formed at 40 rpm with 1:2 (G), 2:1 (H) and 5:1 (I) ratios of CellTracker Red microspheres-to-cells displayed increasing levels of microsphere incorporation. (J) Quantification of microsphere incorporation in populations of lysed EBs demonstrated that lower rotary speed and higher microsphere to cell ratios increased microsphere incorporation (n = 3, mean ± standard deviation).
Fig. 2
Fig. 2
Small molecule release within EBs. EBs formed at 40 rpm with a 2:1 microsphere to cell ratio were imaged after 3 (A–C) and 10 days (D–F). (A–C) After 3 days, microspheres were visible within EBs as individual, punctuate spheres. (D–F) EBs imaged after 10 days contained diffuse fluorescence throughout and fewer individual spheres were apparent. (G) Fluorescent intensity plots across day 3 and day 10 EBs (dashed lines in C and F, respectively) revealed the presence of individual microspheres in day 3 EBs and uniform fluorescence throughout day 10 EBs. (H) CellTracker was released from microspheres over the course of 14 days, and a burst phase (up to one day), sustained release phase (1–7 days), and plateau phase (7–14 days) were observed (n = 3, mean ± standard deviation). (A, B, D, E bar = 200 μm; C, F bar = 100 μm).
Fig. 3
Fig. 3
Delivery of retinoic acid to EBs. Untreated EBs as well as EBs treated with soluble RA, unloaded microspheres, or RA-loaded microspheres were formalin-fixed, sectioned and stained with H&E after 10 days of differentiation. (A) Untreated EBs and (C) soluble RA treated EBs formed solid spheroids while (B) EBs containing unloaded microspheres contained small void spaces. (D, E) EBs containing RA microspheres frequently were completely cystic. (F) Cystic EB formation was significantly enhanced in RA MS EBs compared to all other treatments (n = 3, mean ± standard deviation). (G, H) Cystic RA MS EBs contained bi-epithelial morphology, with a columnar, pseudo-stratified inner cell layer (black arrows) and an adjacent, flattened outermost cell layer (red arrows). *Denotes p < 0.05 vs. untreated, soluble RA; **denotes p = 6 × 10−6 vs. all treatments.
Fig. 4
Fig. 4
Gene expression analysis. Transcriptional analysis of EBs was performed using GeneChip microarrays (A) and quantitative PCR (B). (A) 410 genes were identified as up-regulated greater than threefold in RA microsphere EBs compared to unloaded microsphere EBs after 10 days of differentiation, while 647 genes were observed to decrease in RA MS EBs. Many genes with large fold changes in RA MS EBs were associated with early embryonic structures, including visceral endoderm, epiblast and primitive streak. (B) Time course quantitative PCR analysis of i) Oct4, ii) Fgf5, and iii) Brachyury T gene expression. The epiblast markers Oct4 and Fgf5 were significantly enhanced in RA MS EBs after 7 days, and Fgf5 was enhanced after 10 days as well. Expression of Brachyury T, a marker for primitive streak, in RA MS EBs was delayed until day 10, when a small increase was observed (n = 3, mean ± standard deviation). *Denotes p = 0.02 vs. untreated; **denotes p < 0.05 vs. all treatments; ***denotes p = 5.2 × 10−6 vs. all treatments.
Fig. 5
Fig. 5
Immunostaining of EBs. (A–C) OCT4 staining was performed on day 10 untreated (A), unloaded MS (B) and RA MS EBs (C). Untreated and unloaded MS EBs contained clusters of OCT4+ cells, while OCT4+ cells in RA MS EBs were localized to the columnar cell layer. (D–E) FOXA2, a marker of visceral endoderm, was also expressed in clusters of untreated (D) and unloaded MS EBs (E), but was localized to the outermost layer of cell in RA MS EBs (F). A–C, bar = 50 μm; D–F, bar = 100 μm.
Fig. 6
Fig. 6
RA MS EB ultrastructure. (A, B) SEM micrographs of day 10 RA MS EBs reveal the structure of the distinct squamous endoderm and pseudo-stratified epiblast layers. (C) A dense coat of microvilli were observed on the surface of the endoderm cells, similar to that observed on early streak stage mouse embryos.

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