Significance This study opens avenues to improve the ability of adult skin cells to form a fully functional skin, with clinical applications. Our investigation elucidates a relay of molecular events and biophysical processes at the core of the self-organization process during tissue morphogenesis. Molecules key to the multistage morphological transition are identified and can be added or inhibited to restore the stalled process in adult cells. The principles uncovered here are likely to function in other organ systems and will inspire us to view organoid morphogenesis, embryogenesis, and regeneration differently. The application of these findings will enable rescue of robust hair formation in adult skin cells, thus eventually helping patients in the context of regenerative medicine.

Abstract Organoids made from dissociated progenitor cells undergo tissue-like organization. This in vitro self-organization process is not identical to embryonic organ formation, but it achieves a similar phenotype in vivo. This implies genetic codes do not specify morphology directly; instead, complex tissue architectures may be achieved through several intermediate layers of cross talk between genetic information and biophysical processes. Here we use newborn and adult skin organoids for analyses. Dissociated cells from newborn mouse skin form hair primordia-bearing organoids that grow hairs robustly in vivo after transplantation to nude mice. Detailed time-lapse imaging of 3D cultures revealed unexpected morphological transitions between six distinct phases: dissociated cells, cell aggregates, polarized cysts, cyst coalescence, planar skin, and hair-bearing skin. Transcriptome profiling reveals the sequential expression of adhesion molecules, growth factors, Wnts, and matrix metalloproteinases (MMPs). Functional perturbations at different times discern their roles in regulating the switch from one phase to another. In contrast, adult cells form small aggregates, but then development stalls in vitro. Comparative transcriptome analyses suggest suppressing epidermal differentiation in adult cells is critical. These results inspire a strategy that can restore morphological transitions and rescue the hair-forming ability of adult organoids: (i) continuous PKC inhibition and (ii) timely supply of growth factors (IGF, VEGF), Wnts, and MMPs. This comprehensive study demonstrates that alternating molecular events and physical processes are in action during organoid morphogenesis and that the self-organizing processes can be restored via environmental reprogramming. This tissue-level phase transition could drive self-organization behavior in organoid morphogenies beyond the skin.

Recent studies have made substantial progress in 3D organoid cultures. Multiple epithelial organoids have been generated that resemble their counterparts in vivo, such as the mammary gland (1, 2), salivary gland (3), stomach, colon, pancreas ducts, and liver bile ducts (4). Using stem cell biology approaches, scientists have also generated the cerebral cortex (5) and optical cup (6). Common features of these organoids are that they are generated by 3D culture of isolated tissue progenitors or pluripotent stem cells and that a proper environmental context is provided to guide cells to differentiate into multiple cell types with proper tissue organization. These cultures start from dissociated cells that have lost external cues; amazingly, they can still reform organized tissues similar to those produced during embryonic development in vivo, albeit with different degrees of tissue organization compared with the normal organ morphology.

Organoid cultures have been used as a disease model and can provide organized tissues for regenerative medicine (7). However, organoid formation also provides a unique opportunity, which is not fully developed, to decipher fundamental principles of the self-organization processes (8). Self-organization is the spontaneous formation of ordered structures from a group of progenitor cells that have no ordered prepattern. This can be viewed as a developmental biology question: how do embryonic cells organize in different ways to generate diverse organs and body forms (9)? It is clear that we do not fully understand how 1D DNA codes generate 3D organized topologies (10). The genetic codes do not encode morphology directly; instead complex tissue architectures are achieved through several intermediate layers of interactions involving physical and genetic mechanisms (11, 12). How such genetic information and physical processes cross talk and intertwine with one another to achieve morphological phenotypes remains to be elucidated. However, this knowledge has significant implications for improving our ability to deliver more complex organoids.

The skin organ is a consummate model for studying self-organization processes because of its accessibility to experimentation, its relatively flat configuration, and the availability of genetic tools in mice (13⇓–15). Previous studies have shown that a mixture of dissociated newborn mouse epidermal and dermal cells can reconstitute and form de novo hair follicles in vivo (16⇓⇓–19). These grafts formed a reconstituted organized skin with orientated hair follicles that undergo cyclic renewal and can respond to injury and regenerate (19). However, the principles underlying self-organizational behavior by stem cell collectives remain elusive. Moreover, although the skin cells derived from newborn mice and adult mice share the same genome, adult cells lose this regenerative ability (20). Thus, it has been difficult to generate organized tissues derived from adult cells. Skin reconstituted from human cells does form hairs, but not as robustly as that reconstituted from cells derived from newborn mice (21⇓–23). Thus, there is a great need to learn more about the fundamental conditions required by skin cells to regenerate a functional skin and to identify key environmental factors which will facilitate the hair-forming ability in more easily obtained adult mouse cells and eventually in human cells.

In the present study we developed a two-step method to produce hair-bearing skin: We elucidated in vitro culture conditions that enable skin organoids to form from dissociated cells and transplant techniques that allow organoid explants to form skin with hairs that show normal hair architecture (24). The in vitro step provides two major advantages. First, it allows time-lapse analysis of cell behaviors occurring within the 3D droplet, i.e., 4D analyses of the dynamic temporal changes in tissue morphospace. To our surprise, cells undergo a series of unexpected, complex morphological transitional processes to go from dissociated cells into a planar layer of presumptive skin. These findings led us to develop the idea that a series of phase transition-like processes takes place at the level of the cell population and that these phase transitions could constitute one of the major physical processes used in tissue self-organization. The second advantage of in vitro culture is that it allows experimental manipulation of the molecular mechanisms involved. Transcriptome profiling reveals four stages of molecular expression and allows us to identify molecules that enhance or suppress morphological transitions between each stage. To validate our hypotheses for tissue self-organization, we used dissociated cells generated from adult skin, which normally does not form hairs, and were able to restore the hair-forming ability of adult mouse cells.

Our results offer a promise to improve the ability of human skin cells to form more hair follicles in a fully functional skin, which has clear clinical applications. For basic science, this work demonstrates that a relay of molecular events and physical processes may be core to the self-organization process during tissue morphogenesis. In this case, morphogenetic behavior analyses prompted us to borrow the biophysical concepts of coagulation and self-assembly to explain the morphological phase transitions observed. It seems genes do not encode morphology directly. Instead, complex tissue patterns are achieved through several intermediate layers of interactions involving “physico-genetic” mechanisms (11, 12). This concept can be applied to understand self-organizing processes in organoid morphogenesis beyond the skin and is discussed further.

Discussion The Self-Organizing Process of Planar Skin Formation from Dissociated Cells Is Counterintuitive. We have developed a 3D in vitro organoid model in which dissociated newborn mouse skin cells are cultured at high density. This gives us the unique opportunity to visualize the process leading from individual cells to skin with time-lapse movies. Within a 10-d period, we observed that the dissociated cells progress through a series of morphological phase transitions to achieve a planar layer of presumptive skin with hair primordia (Fig. 7). Grafting of this explant to nude mice with a full-thickness skin wound leads to well-formed reconstituted skin with robust hair growth. These hair follicles have normal architecture and can undergo cyclic regeneration, fulfilling the definition of tissue-engineered hair follicles (19, 24). Fig. 7. Hypothetical morphospace showing the many possible multicellular configurations that take place during morphogenesis of organoid skin cultures. In this space, each axis represents a major change of cell properties and distinct configuration. The self-organization process from dissociated cells to hairy skin can be viewed as a trajectory. Switching of molecular activity is required to move cell collectives from one phase to the next (represented by open arrows). However, close inspection of this process revealed two surprises. First, in skin development, presumptive skin covers the body surface; then periodically arranged dermal condensation and epidermal placode start to emerge around embryonic day 14. In vitro, it is remarkable, because the dissociated cells, having lost all external cues in developing embryos, can reroute and traverse a different morphogenetic path to acquire the same phenotype they have in vivo. This is not what one would expect if morphogenesis were to occur based on a simple molecular blueprint. Instead, it indicates that more fundamental self-organization principles are followed by the dissociated cells to achieve their final morphology. Second, planar skin has a simple configuration consisting of an epidermis and a dermis, with a basement membrane in between. Thus, one may intuitively consider that dissociated epidermal and dermal cells, mixed in suspension, could simply sort themselves out and form sandwich-like cellular layers. Instead, the cells in our assay take a tortuous route from dissociated cells → aggregate → polarized cysts → coalescing cysts → planar hair-bearing skin. We postulate that direct formation of the final, layered skin may well violate the physical constraints imposed by the nature of the active material that the mixture of cells constitutes. Instead, the morphological phase-transition–like events we describe here may represent the most efficient, feasible way for cells to self-organize along a path of least resistance. Intuitively, the straight line is the shortest distance between two points. However, in the morphospace of multicellular configurations (Fig. 7), a straight path may not be the shortest path: It may be easier for cells to take a winding route through a landscape of possible tissue architectures. What, then, are the guiding principles that determine this route? Tissue-Level Morphological Phase Transitions. Here we look at the phenomenon of multicellular self-organization from the biophysicists’ perspective, borrowing the concepts of phase transitions at the tissue level. The term “phase transition” is most commonly used to describe the transformation of matter from one phase/state to another as a function of changes in internal variables or the environment. Phase transitions have recently been shown to mediate cytoplasmic organization at the subcellular level. The assembly and disassembly of nucleoli and other nuclear bodies cycle between liquid-phase droplets and solid-phase condensations and can be modulated by rRNA transcription (33). Phase transition of the microtubule-associated zinc finger protein plays an essential role in the assembly of the spindle apparatus and its associated components (34). It is compelling to extend this biophysical concept to multicellular self-organization. We consider cells in our assay as particles with certain surface properties, performing a random walk (approximately, see Fig. 2E) in a crowded 3D environment. There are two major particle categories, epidermal (E) and dermal (D), so the major interparticle interactions are E–E, D–D, and E–D. Initially epidermal cells form aggregates. When these aggregates reach a certain size, apical–basal polarity develops, leading to the formation of a cyst-like structure. This apical–basal polarity means the inner core and the outer shell of the cyst exhibit different affinities to the environment. Outside the cyst, the interaction of basement membrane with dermal fibroblasts and the presence of MMPs destabilize the cyst structure. The merging of cysts, the fusion of smaller lamellar planes, and thus the eventual large-scale planar configuration may simply be a straightforward consequence of interactions between cell aggregates whose physical properties are changing over time. The biophysical analogy we draw here, although the size scale and dynamics are different from those in soft-matter systems, may shed light on how the system self-organizes into different multicellular configurations. Molecular and Physical Events of Organoid Formation from Newborn Skin Inspire a Strategy to Restore Hair Formation in Adult Mice. To profile the molecular events associated with the observed morphological transitions, we analyzed the skin cells’ transcriptome. We found that there are four peaks of molecular expression, each corresponding to a tissue phase transition: growth factors for the aggregate formation stage (days 0–2); ECM including collagens for apical–basal polarity and cyst formation (days 1–4); Wnts and MMPs for coalescence of the aggregates (days 3–6); and NFkb and laminin for tissue remodeling in the planar formation stage (days 6–10) (Figs. 3 and 4). Functional perturbation with inhibitors of the key molecules at each phase-transition stage can suppress or accelerate the phase-transition process (Fig. 5). The components of these organoid cultures are difficult to dissect. Instead of getting into spatial dissection, we decided to do time-point analyses of the whole culture first. This analysis will provide us with the first level of information about which molecular pathways are important for the morphological transition between stages. Cells from adult mouse skin are quiescent and normally fail to form hairs. Within the logic of our biophysical analogy, we reasoned that we should be able to restore the phenotype by supplying the necessary molecules to the adult cells to reestablish phase-transition–like self-organization behaviors. We examined the cellular and molecular properties of adult cells to find ways to restore their morphogenetic ability. By mapping the cell configuration back to the morphospace of Fig. 7, we can appreciate that adult cell cultures are stuck in the aggregation phase (x axis). By RNA-seq, we found that epidermal differentiation genes appear at an early stage. To restore the ability of adult cells to form hair-bearing skin, we designed a protocol based on the knowledge derived from newborn cell cultures. First, we added inhibitors to PKC to keep cells in undifferentiated states longer. Then we sequentially added three categories of molecules: (i) IGF2/Vegf, (ii) Wnt3a and Wnt10b, and (iii) MMP14 recombinant proteins at days 0, 1, and 3, respectively, to facilitate progression through the morphogenetic stages. In this way, keratinocyte differentiation is reduced. Thus, under the induction of these sequentially added proteins, the morphological transitions are reestablished, and adult cells become competent to reconstitute skin that, upon transplantation, forms hairs robustly. We identified multiple positive and negative regulatory modules directing the progression of morphological transitions during skin organoid formation (Fig. 5I). Instead of focusing on a single molecule, we think the successive phase-transition–like events are the key to successful self-organization (Fig. 6D). More generally, dissociated cells can self-assemble to form many possible multicellular configurations (cell aggregates, cysts, tubes, sheets, and other configurations) in a hypothetical morphospace (Fig. 7). Between each phase, activators and inhibitors work as a feedback control to stop the earlier phase and initiate the next phase. Thus, simply getting bigger cell aggregates is not useful if they do not progress into cyst stage. The morphospace in Fig. 7 can also be useful to appreciate the diverse morphogenetic phenotypes by different epithelial organoids. For example, reconstituted primary mammary myoepithelial cells and luminal cells can form glandular cystic aggregates when placed on Matrigel substrates (2, 35). However, when FGF2 is provided, the cysts undergo branching morphogenesis (36) and do not coalesce toward a planar configuration. In the present study, the end point is a planar skin with hairs. These results suggest there are molecular specificities among different cell types that steer the transitions through morphospace to different multicellular configurations. In summary, we propose that the combined use of molecular signals and biophysical processes may be a basic principle used by nature to drive morphological transitions from one phase to the next. It is the progression of these phase switches, not the specific molecules, that is the key to the success of self-organization. By analyzing more examples of organoid morphogenesis in this context, we stand to learn more about how different physical principles are combined with intrinsic cellular properties to achieve self-organization, thus enhancing our ability to apply these principles to advance tissue engineering.

Materials and Methods In Vitro Assay. As shown in Fig. 1A, cells were prepared according to our previously described method (19). Usually, the E:D ratio in a piece of back skin from a newborn mouse is about 1:9 when we dissociate the back skin into single cells. For the preparation of adult cells, the skins (n ≥ 3) from 2-mo-old mice, in which hair follicles are at refractory telogen phase, were peeled off before the hair fibers were plucked through waxing. Then the s.c. fat was removed from the skins by scissors, and the skins were floated on a 0.25% trypsin solution at 4 °C for overnight digestion. The epidermis was scrapped off the dermis by a scalpel. Then epidermis and dermis were dissociated into single cells as in the preparation of newborn cells. The dissociated epidermal cell and dermal cells were mixed at a ratio of 1:9 and were dropped onto to a Transwell culture insert (Fisher Scientific) that was put in a six-well culture plate. The lower part of the culture insert was filled with 1.5 mL DMEM/F12 (1: 1) (Gibco) culture medium containing 10% FBS (Gibco). The cells were cultured in a humidified atmosphere containing 5% CO 2 at 37 °C, with the culture medium being changed every other day. All animal procedures were performed upon approval of the University of Southern California (USC) Institutional Animal Care and Use Committee. Live Imaging and Analysis. As shown in SI Appendix, Fig. S3A, cells were cultured on a Transwell insert placed in a glass culture plate. The plate was covered by a latex membrane to avoid evaporation of the culture medium. The system was kept at 37 °C by placing the plate on a heating platform and by heating the lens, which was immersed in the culture medium. A LSM5 meta confocal microscope was used to film the cellular behaviors. The resulting 4D (3D space plus time) cellular images were then tracked using commercially available Imaris software (Bitplane) at the Broad California Institute for Regenerative Medicine (CIRM) Center at the University of Southern California. Complete methods and any associated references are available in SI Appendix.

Acknowledgments We thank Drs. Qing Liu and Justin Ichida at the CIRM Center of the USC for supporting the small molecule inhibitors; the USC Epigenome Core Facility for conducting Illumina transcriptome sequencing; the USC Norris Medical Library Bioinformatics Service for assisting with sequencing data analysis; and Prof. Philip Maini of the University of Oxford, Dr. Philip Murray of the University of Dundee, members of the devBio discussion group at the Wolfson Centre for Mathematical Biology, Dr. Christoph Weber of the Max Planck Institute for the Physics of Complex Systems, Drs. Tian Yang and Xiaohua Lian of the Third Military Medical University, Dr. Chin-Lin Guo of Academia Sinica, and Dr. Maksim V. Plikus of the University of California, Irvine for helpful discussions. C.-M.C., R.B.W., T.-X.J., and P.W. are supported by NIH Grants AR42177 and AR60306. M.L. is supported by Project 2016M590866 funded by the China Postdoctoral Science Foundation, Fundamental Research Funds for the Central Universities Grant 106112015CDJRC231206, Special Funding for Postdoctoral Research Projects in Chongqing Grant Xm2015093, and Fellowship 2011605042 from the China Scholarship Council. L.Y. is supported by Innovation and Attracting Talents Program for College and University (111 Project) Grant B06023 and National Nature Science Foundation of China Grants 11532004 and 31270990. W.-T.J. is supported by the Academia Sinica Research Project on Nanoscience and Technology and the Ministry of Science and Technology of Taiwan. L.J.S. was funded by UK Engineering and Physical Sciences Research Council Grant EP/F500394/1 through a studentship at the Life Sciences Interface Programme of the University of Oxford’s Doctoral Training Centre.