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Embryonic origin of adult stem cells required for tissue homeostasis and regeneration

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Cite as: eLife 2017;6:e21052 doi: 10.7554/eLife.21052

Abstract

Planarian neoblasts are pluripotent, adult somatic stem cells and lineage-primed progenitors that are required for the production and maintenance of all differentiated cell types, including the germline. Neoblasts, originally defined as undifferentiated cells residing in the adult parenchyma, are frequently compared to embryonic stem cells yet their developmental origin remains obscure. We investigated the provenance of neoblasts during Schmidtea mediterranea embryogenesis, and report that neoblasts arise from an anarchic, cycling piwi-1+ population wholly responsible for production of all temporary and definitive organs during embryogenesis. Early embryonic piwi-1+ cells are molecularly and functionally distinct from neoblasts: they express unique cohorts of early embryo enriched transcripts and behave differently than neoblasts in cell transplantation assays. Neoblast lineages arise as organogenesis begins and are required for construction of all major organ systems during embryogenesis. These subpopulations are continuously generated during adulthood, where they act as agents of tissue homeostasis and regeneration.

https://doi.org/10.7554/eLife.21052.001

eLife digest

Flatworms are masters of regeneration. If virtually any piece of a flatworm is cut off, a new fully functional individual will grow from it within two weeks. This is no simple task since flatworms contain a wide variety of organ systems, including a brain, nervous system, eyes, kidneys, gut, muscle and skin.

Flatworms owe their regenerative abilities to adult stem cells called neoblasts. Like embryonic stem cells, neoblasts can replicate themselves and they can develop into any type of cell found in an adult worm. In contrast, adult stem cells in fruit flies, zebrafish, mice and humans can only produce the type of cells found in the organ or tissue they live in.

Now, Davies et al. have tracked how and when neoblasts develop in embryos of the flatworm species Schmidtea mediterranea by documenting the distinct gene expression signatures in flatworm embryos at various stages of development. An atlas of the genes that are expressed in various embryonic tissues and in major organs as they begin to develop was also created. These tools, and the results of cell transplantation experiments, revealed that neoblasts emerge from embryonic stem cells as the major organs start to form. As the emerging neoblasts start to express the same combination of genes as adult neoblasts, they also begin to behave just like these cells. The populations of neoblasts remain present throughout the life of the flatworm, helping to maintain, repair and regenerate tissues.

In the future, work that builds on the results presented here by Davies et al. will help researchers to understand more about how stem cells are maintained and regulated. By learning more about the genetic differences between neoblasts and human adult stem cells scientists may be able to explain why humans and other mammals have a limited ability to regenerate. This information could potentially help to develop treatments that stimulate regeneration in patients with degenerative diseases or traumatic injuries.

https://doi.org/10.7554/eLife.21052.002

Introduction

Neoblasts are planarian adult somatic stem cells that exhibit levels of plasticity and pluripotency comparable to embryonic and induced pluripotent stem cells (Elliott and Sánchez Alvarado, 2013; Rink, 2013; Wagner et al., 2011). In flies, fish, mice and humans, adult somatic stem cells are fate-restricted, sustaining production of cell lineage(s) in resident tissues (Fuchs and Segre, 2000; Wagers and Weissman, 2004). Although embryonic stem cells cultured ex vivo remain capable of producing a diversity of tissue types from different germ layers, such plasticity is typically lost from most somatic cells as development proceeds. In contrast, the planarian neoblast population is wholly responsible for the production of all differentiated cell types in these bilaterally symmetric, triploblastic animals (Baguñà and Auladell, 1989). In fact, transplantation of a single neoblast into a stem cell deficient host was sufficient for rescue and long-term reconstitution (Wagner et al., 2011), confirming the pluripotency of planarian somatic stem cells. Despite longstanding discussion of the similarities between neoblasts and embryonic stem cells, a comparison explicitly stated in the original definition of the term (Randolph, 1892), the provenance of neoblasts during embryogenesis was unknown.

Neoblasts are abundant and widely distributed across the anteroposterior axis, occupying the parenchymal space surrounding the gut (Reddien et al., 2005). All neoblasts contain chromatoid bodies (Auladell et al., 1993; Hay and Coward, 1975; Hori, 1982; Morita et al., 1969) and express nuage genes, including piwi-1, and factors implicated in germ cell identity, genome surveillance and post-transcriptional regulation of gene expression (Guo et al., 2006; Palakodeti et al., 2008; Reddien et al., 2005; Rouhana et al., 2010, 2012; Salvetti et al., 2005; Shibata et al., 1999; Solana et al., 2009; Wagner et al., 2012; Yoshida-Kashikawa et al., 2007). Neoblasts are the only cycling somatic cells in adults (Baguñà, 1976; Newmark and Sánchez Alvarado, 2000; Orii et al., 2005; Salvetti et al., 2000); quiescent neoblasts were not observed in BrdU pulse chase experiments (Newmark and Sánchez Alvarado, 2000). Mounting evidence suggests that the neoblast population contains pluripotent stem cells as well as cycling, lineage-primed progenitors (Reddien, 2013). Heterogeneous expression of developmental transcription factors (TFs) in neoblasts has been reported and likely reflects the diversity of lineage-primed progenitors within the compartment (Adler et al., 2014; Cowles et al., 2013; Currie and Pearson, 2013; Lapan and Reddien, 2011, 2012; März et al., 2013; Pearson and Sánchez Alvarado, 2010; Scimone et al., 2014, 2011; van Wolfswinkel et al., 2014; Wenemoser et al., 2012).

Schmidtea mediterranea (Smed) freshwater flatworms are stable diploids that exist as two biotypes: asexual animals that reproduce by fission, and obligate cross-fertilizing hermaphrodites that reproduce sexually (Newmark and Sánchez Alvarado, 2002; Newmark et al., 2008). Both biotypes mount robust regeneration responses following amputation, and similarly rely on neoblasts for homeostatic maintenance and regeneration of all tissues. The asexual clonal line CIW4 (C4) has received the most scrutiny in studies examining the molecular mechanisms underlying regeneration, neoblast maintenance, pluripotency and lineage commitment (Newmark and Sánchez Alvarado, 2002). However, neoblasts are ever-present in C4 animals, precluding investigation of their developmental origin. Neither a normal table for Smed embryonic development nor functional studies have been reported.

Our work establishes Smed as a developmental model system and leverages the novel, unexploited context of embryogenesis to hone the molecular and operational definition of the planarian neoblast. We generated a molecular staging resource for Smed embryogenesis that associates unique gene expression signatures with chronological age, embryo morphology, representative images and written summaries of key developmental events to holistically describe and define prototypes for each stage. We also provide an atlas of molecular markers describing temporary embryonic tissue types and definitive organ system development. These data, found in the supplementary material, are also searchable online at https://planosphere.stowers.org.

We investigated the developmental origin of neoblasts during Smed embryogenesis and show that early embryonic cells are molecularly and functionally distinct from the adult neoblast population. Pluripotent neoblasts and lineage-dedicated progenitors arise as organogenesis begins. Our results suggest that the framework for understanding cell fate specification and organ formation during Smed embryogenesis diverges radically from existing developmental paradigms. Here, in a bilaterally symmetric, triploblastic animal not thought to undergo gastrulation (Cardona et al., 2005; Le Moigne, 1963; Sánchez Alvarado, 2003; Stevens, 1904), heterogeneous expression of key developmental regulators within a pluripotent, cycling blastomere population generates the panoply of lineage-dedicated progenitors required for organogenesis. Moreover, neoblasts perpetuate embryonic developmental programs during adulthood, where they are required for continued maintenance and rebuilding of tissues during homeostasis and regeneration.

Results

A molecular staging series for Smed embryogenesis informed by single embryo RNA-Seq

Smed flatworms are direct developers: newborn hatchlings grow and mature into adult worms without an intervening larval stage (Sánchez Alvarado, 2003). At hatching, juveniles are sexually immature but otherwise possess a body plan grossly similar to that of adult hermaphrodites (Sánchez Alvarado, 2003; Wang et al., 2007). Smed embryos undergo an evolutionarily divergent mode of development that bears little resemblance to the ancestral Spiralian cleavage programs utilized by many Lophotrochozoans. Smed embryos are ectolecithal: yolk is not contained within oocytes, but rather is produced by somatic vitellaria (yolk glands) arrayed ventrolaterally beneath the testes (Chong et al., 2011; Steiner et al., 2016; Stevens, 1904). Oocytes are fertilized internally by sperm from a partner. Zygote(s) are packaged, along with yolk cells, into an egg capsule in the genital atrium (Figure 1A) (Chong et al., 2011; Hyman, 1951; Newmark et al., 2008; Stevens, 1904).

Figure 1 with 19 supplements see all
A molecular staging series for Smed embryogenesis informed by single embryo RNA-Seq.

(A) Top: Cartoon depicting the reproductive system of a sexually mature Smed hermaphrodite. Ventral view. G, gonopore; O, ovary; OD, oviduct; P, penis papilla; SD, sperm duct . Oocytes are fertilized internally and zygote(s) are packaged with yolk produced by vitellogenic gland cells into a developing egg capsule in the genital atrium (purple). Capsules are laid through the gonopore. Bottom: Brightfield image of a live Smed hermaphrodite. Anterior: left. Dorsal view. White asterisk: pharynx. (B) Developmental timeline and staging designations for Smed embryogenesis at 20°C. Timeline: days (d) post egg capsule deposition. Gray bars and letters C–I indicate time windows, and corresponding panels (C–I), for RNA-Seq samples. Double-headed arrows: time windows for stages (S) S1–S8. (CI) Brightfield images of live embryos harvested for RNA-Seq (top), hematoxylin- and eosin-stained sections (middle), and heat maps for enriched transcripts (bottom). Scale bars: 100 µm. Yellow arrowheads: temporary embryonic pharynx. Black arrowheads: definitive pharynx. Heat maps depict cohorts of enriched transcripts at indicated stages. (J) Principal component analysis demonstrates clustering of replicates and separation of developmental time points in expression space. (K) Correlation matrix for single embryo sequencing replicates. Total transcripts with a row sum >1 CPM: 31,248. (CK) Y,yolk. 2, S2. 3, S3. 4, S4. 5, S5. 6, S6. 7, S7. 8, S8.

https://doi.org/10.7554/eLife.21052.003

In contrast to the synchronous, oriented blastomere cleavage patterns of Spiralian embryos (Lambert, 2010), blastomeres in freshwater planarian embryos undergo dispersed cleavage among yolk cells: they divide asynchronously and are not in direct contact with one another (Bardeen, 1902; Cardona et al., 2005; Hallez, 1887; Ijima, 1884; Le Moigne, 1963; Metschnikoff, 1883; Vara et al., 2008). During sphere formation, some blastomeres differentiate into temporary embryonic cell types that provide form and function to the embryo, including the primitive ectoderm, temporary embryonic pharynx and primitive gut (Cardona et al., 2005; Hallez, 1887; Le Moigne, 1963; Sánchez Alvarado, 2003). Temporary embryonic tissues are not thought to contribute to the juvenile body plan; they are thought to degenerate as the definitive organs form and morphogenesis proceeds (Cardona et al., 2005; Le Moigne, 1963; Vara et al., 2008). In contrast, undifferentiated blastomeres remaining after sphere formation are thought to give rise to all definitive tissues found in juvenile worms (Hallez, 1887; Hyman, 1951; Le Moigne, 1963; Sánchez Alvarado, 2003; Stevens, 1904).

Several attributes of ectolecithal development challenge efforts to accurately stage live embryos including internal fertilization, dispersed cleavage, lack of morphological landmarks, and inherent variability in embryo size and developmental timing. Extant staging series for Polycelis nigra (Le Moigne, 1963), Schmidtea polychroa (Cardona et al., 2005; Martín-Durán et al., 2010) and Girardia tigrina (Vara et al., 2008) rely upon gross morphological criteria gleaned from live animals and fixed, histologically or antibody-stained specimens. As this is the first systematic characterization of Smed embryogenesis, we established an accurate and objective staging method based on unique gene expression signatures, cohorts of enriched transcripts identified through single embryo RNA-Seq , associated with chronological age and embryo morphology (Figure 1B–I, Figure 1—source data 1). When appropriate, efforts were made to integrate and adapt extant staging schema from other planarian species.

Smed embryos gestate for approximately two weeks at 20°C prior to hatching. We generated total RNA replicates from single Smed embryos for seven chronologically and/or morphologically distinct stages (S), S2-–S8 (Figure 1B–I); S1 samples (zygotes, Figure 1—figure supplement 1) were not queried by RNA-SSeq. Yolk (Y) replicates were prepared from egg capsules lacking developing embryos at 8 days post capsule deposition. In addition, single animal replicates were prepared from C4 and virgin sexually mature adults (SX). RNA-Seq libraries were analyzed for four biological replicates (i.e., four individuals) per stage (Materials and methods). Identification of appropriate reference(s) and normalization methods was challenging due to vast differences in sample composition and complexity among different stages. However, clustering of replicates and discrete separation of stages was seen in a multidimensional scaling plot (Figure 1J). Replicates for a given stage generally showed strong correlation among themselves despite not having controlled for differences in genetic background or embryo size (Figure 1K). Notably the S2 and S3 replicates, generated from embryos undergoing sphere formation and nascent spheres, respectively, were intermingled in expression space, showed the greatest variability and few significant differences in gene expression (Figure 1J–K, Figure 1—source data 1, Figure 1—figure supplement 2). Discrete stages were retained due to apparent differences in embryo morphology (Figure 1C–D). Similarities among S2 and S3 samples may be due to maternal RNA contribution or to difficulty detecting labile S2 embryos, such that only well-developed S2 embryos were prospected by RNA-Seq.

Two approaches were used to identify differentially expressed transcripts: pairwise comparisons of adjacent stages (Figure 1—source data 1, Figure 1—figure supplement 2)and comparisons of each stage relative to a mixed stage reference generated by averaging the read counts for the remaining replicates (Y, S2–S8) (Materials and methods, Figure 1—source data 1, Figure 1—figure supplement 3). The goal of the pairwise comparisons was to identify transcripts with the starkest changes in expression in either direction, without constraints on transcript behavior at other points during embryogenesis. In contrast, the mixed stage reference comparisons maximized the likelihood of identifying transcripts exhibiting stage-specific expression, and only upregulated transcripts were analyzed. Stringent criteria were applied in both scenarios for flagging differentially expressed transcripts, including thresholds based on the Benjamini-Hochberg adjusted p-value, fold-change, normalized RPKM expression level for time points, and identification of at least one open reading frame in the transcript (Figure 1—source data 1). More upregulated transcripts were identified in the mixed stage reference than in pairwise comparisons, perhaps due to increased sequencing depth of the averaged reference samples, which may enable more reliable detection of lowly expressed transcripts. Furthermore, identification of S2-enriched transcripts suggested that the whole embryo sequencing approach was sensitive enough to detect transcripts expressed in rare cell populations (i.e., blastomeres and differentiated cells making up the embryo proper). Non-redundant lists of upregulated transcripts, resulting from the union of the pairwise and mixed reference comparisons, served as molecular fingerprints for each time point for downstream analyses, including hierarchical clustering and GO analysis (Figure 1—source data 1). The molecular staging resource (Figure 1—source data 1) incorporates representative images (Figure 1C–I, Figure 1—figure supplement 1), gene expression signatures (Figure 1—figure supplements 410, Figure 1—source data 28) and written summaries of key developmental events (Figure 1—source data 1), defining S1–S8. An expression atlas describing temporary embryonic tissue types and development of the definitive organ systems is also provided (Figure 1—source data 9, Figure 1—figure supplements 1119). The molecular staging resource and expression atlas are also available and searchable online (https://planosphere.stowers.org).

Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development

The molecular staging series identified S2- and S3-enriched transcripts (Figure 1C–D, Figure 1—source data 23), including elongation factor 1a-like-1 (EF1a-like-1), expressed in all known embryonic cell populations in nascent spherical embryos (Figure 2A–B). During S2, primitive ectoderm cells are the first to differentiate (Hallez, 1887; Ijima, 1884; Le Moigne, 1963; Metschnikoff, 1883). EF1a-like-1 was expressed in primitive ectoderm cells (Figure 2A–B, Video 1), which flatten, elaborate numerous cytoplasmic processes, and form a single-cell layer bounding the sphere (Figure 1—source data 9, Figure 1—figure supplement 11). EF1a-like-1 expression was also detected in the temporary embryonic pharynx (Figure 2A–B, Video 1) an innervated pump containing neurons, radial muscle and associated epithelial cells (Figure 1—source data 9, Figure 1—figure supplement 12), and in the primitive endoderm (Figure 2A), which consists of an inner gut cavity and phagocytic cells associated with the temporary embryonic pharynx (Figure 1—source data 9, Figure 1—figure supplement 13). A population of undifferentiated blastomeres and yolk cells remain in the embryonic wall, the parenchymal space between the primitive ectoderm and endoderm, in nascent S3 spheres (Figure 1D) (Hyman, 1951; Sánchez Alvarado, 2003). EF1a-like-1 was expressed in undifferentiated blastomeres, but not yolk cells, in S3 embryos (Figure 2A–B, Video 1).

Blastomere anarchy drives Smed embryogenesis.

(AB) Architectural features of S2 and S3 embryos. (A) Expression of the pan embryonic cell marker EF1a-like-1 (blue) in S2 (left) and S3 (right) embryos. Cyan arrowheads: primitive ectoderm cells. Cyan arrows: undifferentiated blastomeres. Red arrowhead: temporary embryonic pharynx. Red arrows: primitive gut cells. (B) S3 embryo stained with EF1a-like-1 riboprobes (red) and sytox green nuclear counterstain (green). Cyan arrowheads: primitive ectoderm cells. Cyan arrows: undifferentiated blastomeres. Yellow arrowhead: temporary embryonic pharynx. (C) Confocal Z-slice of an ovary from a sexually mature Smed hermaphrodite stained with piwi-1 riboprobes (green) and DAPI (blue). Yellow arrows: oocytes. (D) Dispersed cleavage. S2 embryo stained with piwi-1 riboprobes (red, blastomeres) and antibodies raised against the mitotic epitope H3S10p (green). Nuclei stained with DAPI (blue). Yellow arrow: dividing blastomere. (E) piwi-1 is expressed in undifferentiated blastomeres of S3 embryos. S3 embryo costained with riboprobes complementary to piwi-1 (red) and EF1a-like-1 (green). 100% piwi-1+ blastomeres coexpressed EF1a-like-1. n = 159 cells scored, n = 5 S3 embryos. Cyan arrows: undifferentiated blastomeres. Yellow arrowhead: temporary embryonic pharynx. Red arrows: fiduciary beads used for SPIM reconstruction. (F) piwi-1+ cells are located in the embryonic wall. Paraffin-embedded cross-section of a S3 sphere stained with piwi-1 riboprobes (blue) and eosin (pink). Cyan arrows: piwi-1+ cells. GC: yolk-filled gut cavity. Inset: magnified view of a piwi-1+ cell. Scale: 25 µm. (G) Left: Average RPKM per embryo for piwi-1 (S2–S8). Right: WISH developmental time course with piwi-1 riboprobes (blue) (S2–S8). O, oral hemisphere; V, ventral. (AG) Scale: 100 µm. Left: Observed distribution of piwi-1+ cells in S3–S4 embryos (blue bars) relative to the oral-aboral axis (0–3.14 radians). Maximum likelihood analysis best described distribution by the function ((1-exp(-θ/θ’))*sin(θ), blue line). The optimal calculated θ’ was 0.45 ± 0.045 radians, based on simulations with comparably sized data sets, and was several orders of magnitude more likely to explain the observed distribution than the theoretical normal distribution, sin(θ), (θ’ = 0), red line. S3: n = 32 embryos, n = 1,746 piwi-1+ cells scored. S4: n = 8 embryos, 2,665 piwi-1+ cells scored. Right: observed piwi-1+ cell distributions for individual S3 (top) and S4 (bottom) embryos. (CG) piwi-1+ cells are detected throughout embryogenesis. (H) piwi-1+ cell positions are not stereotyped in S3–S4 embryos.

https://doi.org/10.7554/eLife.21052.032
Video 1
S3 embryo architecture.

SPIM reconstructed S3 embryo costained with EF1a-like-1 (red) and sytox green nuclear counterstain. EF1a-like-1 is a pan-embryonic cell marker that stains primitive ectoderm cells, the temporary embryonic pharynx and undifferentiated blastomeres in the embryonic wall. EF1a-like-1 staining is absent from yolk cells in the embryonic wall and gut cavity.

https://doi.org/10.7554/eLife.21052.033

piwi-1+ cells were present throughout embryogenesis. piwi-1 expression was detected in oocytes (Figure 2C), suggesting that zygotes contain piwi-1 mRNA. Zygote-derived blastomeres undergoing dispersed cleavage among yolk cells during S2 also expressed piwi-1 (Figure 2D). Costaining with piwi-1 riboprobes and antibodies raised against the G2-M phase mitotic marker H3S10p showed that piwi-1+ blastomeres divide asynchronously during S2 (Figure 2D).

As spheres form during S2, some blastomeres downregulate piwi-1 expression and differentiate into temporary embryonic cell types. piwi-1 expression was restricted to, and expressed throughout, the undifferentiated blastomere population in S3 embryos, as demonstrated by double fluorescent WISH with EF1a-like-1 and piwi-1 riboprobes (Figure 2E, Video 2). piwi-1+ blastomeres were always located in the embryonic wall (Figure 2F). piwi-1 expression was never detected in the primitive ectoderm, temporary embryonic pharynx or primitive gut (Figure 2E,G). During S3–S5, piwi-1+ cell number clearly increased, effectively blanketing the sphere (Figure 2G). As definitive gut development proceeded during S6–S8, piwi-1+ cells occupied the parenchyma between the developing gut branches (Figure 2G). Notably, piwi-1+ cells were not detected in the definitive pharynx, and the compartment receded from the anterior margin as head structures developed (Figure 2G). By S8, the spatial distribution of piwi-1+ cells was indistinguishable from that of the adult neoblast compartment (Figure 2G).

Video 2
piwi-1 is expressed in all undifferentiated blastomeres of S3 embryos.

SPIM reconstructed S3 embryo costained with piwi-1 (red) and EF1a-like-1 (green). piwi-1 is expressed in all undifferentiated blastomeres in the embryonic wall (piwi-1+, EF1a-like-1+ cells). piwi-1 is not expressed in differentiated tissues marked by EF1a-like-1 alone, including the primitive ectoderm and temporary embryonic pharynx (green). Several fluorescent beads used for three-dimensional reconstruction are visible (red).

https://doi.org/10.7554/eLife.21052.034

Analysis of piwi-1+ cell positions in S3–S4 embryos revealed that undifferentiated blastomeres were spatially disordered, or ‘anarchic,’ as described for other freshwater flatworms (Bardeen, 1902; Cardona et al., 2005; Hallez, 1887; Ijima, 1884; Le Moigne, 1963; Metschnikoff, 1883; Vara et al., 2008). The observed distribution of piwi-1+ cells in S3–S4 embryos with respect to the temporary embryonic pharynx (Figure 2H, blue line) was nearly identical to that of a random distribution (Figure 2H, red line). Fewer piwi-1+ cells were located adjacent to the oral pole than was predicted for a theoretical normal distribution, perhaps due to spatial constraints imposed by the temporary embryonic pharynx and associated primitive gut cells (Figure 2H). We used a simple dampening term and maximum likelihood estimation to account for the deviation between the observed and theoretical normal distributions (Figure 2H). However, the distribution of piwi-1+ cells in the embryonic wall was not stereotyped; it varied greatly across individuals (Figure 2H).

Strikingly, cell cycle activity was restricted to piwi-1+ cells at all developmental stages assayed, and all piwi-1+ cells in early embryos were cycling. Expression of the cell cycle regulators PCNA and RRM2-2 closely mimicked that of piwi-1 during embryogenesis, both with respect to the spatial distribution of positive cells and trends observed in the RNA-Seq data (Figures 2G and 3A–B). Double fluorescent WISH on S3–S5 embryos revealed that PCNA and RRM2-2 were expressed exclusively in piwi-1+ blastomeres, all of which were cycling (Figure 3C–D, Videos 34). S3–S5 embryos costained with piwi-1 and H3S10p antibodies confirmed that mitotic activity was restricted to piwi-1+ blastomeres (Figure 3E, Video 5). Consistent with observations made in S2 embryos (Figure 2D), cell divisions were asynchronous during S3–S5. The mitotic index for S3–S5 piwi-1+ blastomeres was stable, with no statistically significant difference in the calculated division rate (Figure 3F). Analysis of mitotic (piwi-1+, H3S10p+) cell positions along the oral-aboral axis in S3–S4 embryos did not reveal regional biases in mitotic activity across samples (Figure 3G).

Cell cycle activity is restricted to the piwi-1+ compartment.

(AB) Left: Colorimetric WISH depicting expression of PCNA (A) or RRM2-2 (B) during stages S2–S8. Right: Average RPKM values per embryo for PCNA (A) or RRM2-2 (B) in Y (yolk) and S2–S8. V, ventral. Scale: 100 µm. (CD) S3 (top), S4 (middle) and S5 (bottom) embryos costained with piwi-1 (red) and PCNA (green [C]) or RRM2-2 (green [D]) riboprobes. The percentage of piwi-1+ cells coexpressing the indicated cell cycle marker (red) and the percentage of PCNA+ or RRM2-2+ cells coexpressing piwi-1 (green) appear in the lower left corner of merged images. Scale bars: 100 µm. (C) S3: n = 273 cells, n = 6 embryos. S4: n = 1,267 cells, n = 4 embryos. S5: n = 734 cells, n = 3 embryos. (D) S3: n = 130 cells, n = 4 embryos. S4: n = 1,295 cells, n = 5 embryos. S5: n = 350 cells, n = 3 embryos. (E) Mitotic activity is restricted to the piwi-1+ cell compartment in S3–S5 embryos. Left: S4 embryo costained with piwi-1 and the embryonic pharynx marker LYAG-like (both in green) and antibodies against the mitotic epitope H3S10p (red). White arrows: dividing blastomeres. White arrowhead: temporary embryonic pharynx. Scale bar: 100 µm. Right: Bar graph depicting the percentage of mitotic cells scored that expressed piwi-1 in S3–S5 embryos. (F) The mitotic index for the piwi-1+ cell compartment did not vary significantly during S3–S5. Average percentage of piwi-1+ cells in mitosis during S3–S5. Error bars represent the standard deviation of the mean. Observed distribution of mitotic (piwi-1+, H3S10p+) cells in S3-–S4 embryos (blue bars) along the oral-aboral axis (0–3.14 radians). Using the function derived with maximum likelihood estimation for the piwi-1+ cell distribution, (1-exp(-θ/θ’))*sin(θ) (blue line), and simulations using equivalent sample sizes, the optimal θ’ was calculated to be 0.58 ± 0.33, and was 50-fold more likely to explain the observed trend than a simple normal distribution, sin(θ), where θ’=0 (red line). S3: n = 82 mitotic cells, n = 18 embryos. S4: n = 110 mitotic cells, n = 8 embryos. (G) Mitotic cell positions are not stereotyped in early embryos.

https://doi.org/10.7554/eLife.21052.035
Video 3
Cell cycle activity is restricted to piwi-1 blastomeres, and all blastomeres are cycling.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and PCNA (green). PCNA expression is restricted to piwi-1+ blastomeres, and all piwi-1+ cells co-express PCNA.

https://doi.org/10.7554/eLife.21052.036
Video 4
Cell cycle activity is restricted to piwi-1 blastomeres, and all blastomeres are cycling.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and RRM2-2 (green). RRM2-2 expression is restricted to piwi-1+ blastomeres, and all piwi-1+ cells co-express RRM2-2. Several fluorescent beads used for three-dimensional reconstruction are visible (red).

https://doi.org/10.7554/eLife.21052.037
Video 5
Mitotic activity is restricted to piwi-1+ blastomeres, which cycle asynchronously.

SPIM reconstructed S4 embryo costained with piwi-1 and LYAG-like (both in green) and H3S10p antibodies (red). LYAG-like marks the temporary embryonic pharynx and is not expressed in piwi-1+ blastomeres. Several examples of piwi-1+, H3S10p+ cells are evident.

https://doi.org/10.7554/eLife.21052.038

Several striking parallels may be drawn regarding cell cycle behavior of the piwi-1+ population during embryogenesis and adulthood. First, cell cycle activity is largely restricted to this compartment in both contexts, with exception of the male and female germline in sexually mature adults (Baguñà, 1976; Newmark and Sánchez Alvarado, 2000; Reddien et al., 2005; Wagner et al., 2011; Wang et al., 2007). Second, cycling cells and mitotic figures do not display obvious positional biases within the parenchyma in either S3–S5 embryos or adults during homeostasis (Newmark and Sánchez Alvarado, 2000). Little substantiating evidence exists in support of a quiescent piwi-1+ cell population during embryogenesis or adulthood (Newmark and Sánchez Alvarado, 2000), and differences in cell cycle length were not observed between neoblast subclasses in adult asexual animals (van Wolfswinkel et al., 2014). Finally, the staggering net increase in piwi-1+ blastomeres during S3–S5 suggests a capacity for self-renewal, a property possessed by neoblasts.

Many adult asexual stem cell genes are expressed throughout embryonic development

Numerous neoblast-enriched transcripts, identified through whole asexual animal irradiation studies and cell sorting (Eisenhoffer et al., 2008; Labbé et al., 2012; Rossi et al., 2007; Solana et al., 2012; Wagner et al., 2012; Wurtzel et al., 2015), have been vetted for co-expression with piwi-1+ and ascribed function(s) in neoblast proliferation, maintenance or cell fate commitment. Some neoblast-enriched transcripts, including those encoding nuage components and cell cycle regulators, are expressed in all neoblasts, whereas others are predominantly expressed in subpopulation(s) of cells that may be primed to adopt differentiated fates (van Wolfswinkel et al., 2014). To address similarities in the gene expression profiles of the piwi-1+ population during embryogenesis and adulthood, the expression trends of 242 adult asexual neoblast-enriched transcripts were examined using the molecular staging series data. Neoblast-enriched transcript membership was determined by sequences downregulated in whole animals following lethal irradiation across three independent experiments (Duncan et al., 2015; Wagner et al., 2012) (Chen and Sánchez Alvarado, personal communication). Strikingly, most adult asexual neoblast-enriched transcripts were expressed throughout embryogenesis (Figure 4A, Figure 4—source data 1). 74% (n = 180) of the neoblast-enriched transcripts had average RPKM values per embryo ≥1.0 in S2 embryos and 52% (n = 128) transcripts had five-fold or greater expression levels in S2 embryos versus yolk, raising the possibility that other adult stem cell genes were expressed in blastomeres. Consistent with this idea, 41% of the adult asexual neoblast-enriched transcripts (n = 99) were present in the molecular expression signature(s) for S2–S5 embryos (Figure 1—source data 25, Figure 4—source data 1). Expression of neoblast-enriched transcripts usually peaked during S4 or S5, prior to construction of the definitive organ systems, and diminished thereafter (Figure 4A, Figure 4—source data 1). The apparent decrease in expression after S5 was likely attributable to drastic changes in the complexity of the single embryo RNA samples during organogenesis, and was similarly observed for piwi-1 and the cell cycle regulators PCNA and RRM2-2 (Figures 2G and 3A–B, Figure 4—source data 1).

Figure 4 with 1 supplement see all
Many adult neoblast markers are similarly expressed throughout the piwi-1+ compartment during embryogenesis.

(A) Many transcripts with adult asexual neoblast-enriched expression are expressed throughout embryogenesis. Hierarchical clustering of 242 adult asexual neoblast-enriched transcripts during embryonic development using normalized mixed stage reference comparison data. Left: Heat map. Colored bars (left) denote clusters. Right: Normalized average RPKM values per embryo, plotted as a function of developmental time, for Clusters 1–8. Y, yolk; S2–S8, Stages 2–8. (BE) Colorimetric WISH depicting expression of piwi-2 (B), piwi-3 (C), tud-1 (D) and bruli-1 (E) during embryogenesis (blue) (S2–S8). V, ventral. Black arrowheads: temporary embryonic pharynx. Red arrowheads: definitive pharynx. Scale bars: 100 µm. (FI) Many markers of the adult asexual neoblast compartment are also expressed in piwi-1+ blastomeres. Fluorescent WISH on S4 embryos with riboprobes against piwi-1 (red) and piwi-2 (F), piwi-3 (G), tud-1 (H) or bruli-1 (I) (green). Percentage of piwi-1+ cells coexpressing the indicated marker (red) and the percentage of the indicated adult asexual neoblast marker coexpressing piwi-1 (green) appears in the lower left corner of merged images. Scale bars: 100 µm. (F) n = 435 cells, n = 9 S3–S4 embryos. (G) n = 535 cells, n = 5 S3–S4 embryos. (H) n = 1,867 cells, n = 8 S3–S5 embryos. (I) n = 1,353 cells, n = 3 S4 embryos.

https://doi.org/10.7554/eLife.21052.039

Hierarchical clustering of the 242 adult asexual neoblast-enriched transcripts revealed correlated expression of genes associated with DNA replication (e.g., the replication licensing factors MCM2 and MCM5), DNA repair (e.g., fancd2-like, msh2 and msh6) and cell cycle progression (e.g., cyclin D-like, cyclin-B1 and cyclin-B2) during embryogenesis, and the expression trends for these genes mimicked those of PCNA and piwi-1 (Figure 4A [Cluster 1], Figure 4—source data 1). Notably, transcripts encoding the nuage-associated factors piwi-2 and piwi-3, the RNA-binding protein bruli-1, and the transcription factors SoxP-1 and junL1-1, all genes previously implicated in neoblast maintenance or function (Guo et al., 2006; Palakodeti et al., 2008; Reddien et al., 2005; Wagner et al., 2012), were also coregulated with piwi-1 during embryogenesis (Figure 4A [Cluster 3], Figure 4—source data 1). Consistent with the expression trends detected by RNA-Seq and the previously reported expression pattern for Schmidtea polychroa (Spol) tud-1 (Solana et al., 2009), piwi-2, piwi-3, tud-1, bruli-1, Sox-P1 and Sox-P2 were expressed in cells with similar morphology and distribution to the piwi-1+ population during embryogenesis (Figure 4B–E, Figure 4—figure supplement 1A–C). Double fluorescent WISH revealed coincident expression of piwi-1 and the adult stem cell markers piwi-2, piwi-3, tud-1 and bruli-1 in S3–S5 embryos (Figure 4F–I, Figure 4—figure supplement 1D–F, Videos 69). Y12 antibodies, which label chromatoid bodies in adult neoblasts (Rouhana et al., 2012), stained piwi-1+ blastomeres during S4-–S5 (Figure 4—figure supplement 1G, Video 10). Taken together, these findings suggest that many adult asexual neoblast markers, including genes implicated in DNA replication and repair, cell cycle control, chromatin remodeling and/or modification, genome surveillance and pluripotency, are also likely to be expressed throughout the piwi-1+ population during embryogenesis. Moreover, shared elements of the blastomere and neoblast expression signatures, including DNA replication and repair pathway, cell cycle, nuage and RNA processing genes, are prominent features of an evolutionarily conserved gene expression signature for metazoan primordial stem cells (Alié et al., 2015).

Video 6
piwi-1+ blastomeres co-express the adult asexual neoblast-enriched gene piwi-2.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and piwi-2 (green). piwi-1+ blastomeres co-express the nuage factor piwi-2, and virtually all piwi-2+ cells co-express piwi-1. Several fluorescent beads used for three-dimensional reconstruction are visible (green).

https://doi.org/10.7554/eLife.21052.042
Video 7
piwi-1+ blastomeres co-express the adult asexual neoblast-enriched gene piwi-3.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and piwi-3 (green). piwi-1+ blastomeres co-express the nuage factor piwi-3, and virtually all piwi-3+ cells co-express piwi-1.

https://doi.org/10.7554/eLife.21052.043
Video 8
piwi-1+ blastomeres co-express the adult asexual neoblast-enriched gene tud-1.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and tud-1 (green). piwi-1+ blastomeres co-express the nuage factor tud-1, and virtually all tud-1+ cells co-express piwi-1.

https://doi.org/10.7554/eLife.21052.044
Video 9
piwi-1+ blastomeres co-express the adult asexual neoblast-enriched gene bruli-1.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and bruli-1 (green). piwi-1+ blastomeres co-express the stem cell maintenance gene bruli-1, and virtually all bruli-1+ cells co-express piwi-1. Several fluorescent beads used for three-dimensional reconstruction are visible (red).

https://doi.org/10.7554/eLife.21052.045
Video 10
piwi-1+ blastomeres possess chromatoid bodies.

SPIM reconstructed S4 embryo costained with piwi-1 (red) and Y12 antibodies (green). Y12 antibody staining was restricted to, and present throughout, the piwi-1+ blastomere population.

https://doi.org/10.7554/eLife.21052.046

Early-embryo-enriched transcripts expressed throughout piwi-1+ blastomeres are downregulated as organogenesis begins

Hierarchical clustering of S2-–S4-enriched transcripts using scaled RPKM values identified 1,048 sequences in Clusters 5, 6 and 8 that were downregulated by S5 and remained lowly expressed through S8; these sequences are referred to as early-embryo-enriched (EEE) transcripts (Figure 5A, Figure 5—source data 1). EEE transcripts were likely expressed in blastomeres and/or temporary embryonic tissues, as 98% of the sequences had average expression values at least five-fold greater in S2 embryos than in Y (Figure 5—source data 1). Most EEE transcripts were expressed at low levels in intact adults regardless of biotype: average RPKM values less than 1.0 were recorded for 65% and 59% of the EEE transcripts in C4 or SX, respectively (Figure 5—source data 1).

Figure 5 with 1 supplement see all
Early-embryo-enriched transcripts are downregulated as organogenesis begins.

(A) Hierarchical clustering of S2–S4-enriched transcripts (n = 1,756) using scaled RPKM data. Left: Heat map. Y, yolk. Colored bars (left) denote Clusters 5, 6 and 8 containing early-embryo-enriched (EEE) transcripts. Cluster f5 sequences (blue, n = 413) were expressed at roughly equivalent levels during S2 and S3, with 66% (n = 275) transcripts showing five-fold or greater declines in average expression values between S3 and S5. Cluster 6 sequences (red, n = 523) exhibited maximal expression during S2, and average expression levels declined more than five-fold between S2 and S4 for 81% (n = 426) of these transcripts. Cluster 8 sequences (green, n = 112) showed peak expression during S4, with 52% (n = 60) of the transcripts showing five-fold or greater declines in average expression values by S5. Right: Normalized expression trends for EEE transcripts in Clusters 5 (blue), 6 (red) and 8 (green) plotted as a function of developmental time. Median 50% of transcripts based on expression maxima are plotted. (BE) Colorimetric WISH depicting expression of the EEE transcripts tct-like (B), BTF3-like (C), DDX5-like (D) and eIF4a-like (E) (blue) in S2–S8 embryos and C4 asexual adults. Black arrowheads: temporary embryonic pharynx. Red arrowheads: definitive pharynx. O, oral; V, ventral. Scale bars: 100 µm. (FI) EEE transcripts were expressed throughout the piwi-1+ compartment in S3–S4 embryos. Fluorescent double WISH with riboprobes against piwi-1 (red) and the EEE transcripts tct-like (F), BTF3-like (G), DDX5-like (H) and eIF4a-like (I) (green) in S4 embryos. Percentage piwi-1+ cells coexpressing the indicated EEE marker (red) and percentage EEE+ cells coexpressing piwi-1 (green) appear in the lower left corner of merged images. (F) n = 895 piwi-1+ cells, n = 905 tct-like+ cells, n = 7 S3–S4 embryos. (G) n = 692 piwi-1+ cells, n = 728 BTF3+ cells, n = 6 S3–S4 embryos. (H) n = 676 piwi-1+ cells, n = 681 DDX5-like+ cells, n = 5 S3–S4 embryos. (I) n = 312 piwi1+ cells, n = 332 eIF4a+ cells, n = 4 S3–S4 embryos.

https://doi.org/10.7554/eLife.21052.047

EEE transcript expression trends during embryonic development and adulthood were validated using the Nanostring nCounter platform (Geiss et al., 2008). Total RNA replicates from single S2–S8 embryos, Y, C4 and SX adults were queried for expression of 108 EEE transcripts using a custom probe set (Materials and methods, Figure 5—source data 2). Curiously, S2 replicates had the lowest summed read counts across the experiment (Figure 5—figure supplement 1A), and expression of piwi-1, the cell cycle marker H2B, and EEE transcripts was not detected during S2 (Figure 5Figure 1—figure supplement 1E–F, Figure 5—source data 2), perhaps due to lack of template amplification in the nCounter assays. Apart from the S2 samples, expression trends for piwi-1, H2B, the pro-differentiation factor prog-1 and more than 90% of the EEE transcripts mirrored those observed in the RNA-Seq time course (Figure 5—figure supplement 1E–F). Most EEE transcripts queried on the nCounter platform showed peak expression during S3 and/or S4, downregulation during S5, and low to undetectable expression in late stage embryos, C4 or SX adults (Figure 5—figure supplement 1F). Hierarchical clustering using normalized Nanostring count data identified three cohorts of EEE transcripts that hadcomposition and decay kinetics similar to those observed in the RNA-Seq time course; 65% of the EEE transcripts assayed co-clustered in both the RNA-Seqand nCounter analyses (Figure 5—figure supplement 1F).

To determine which cell types express EEE transcripts, and to examine spatiotemporal changes in EEE transcript expression, colorimetric WISH was performed on S2–S8 embryos and intact C4 adults (Figure 5—source data 3). Some EEE transcripts were expressed exclusively in differentiated temporary embryonic tissues. VAL-like, MPEG1-like-1, MPEG1-like-2 and netrin-like were solely expressed in the temporary embryonic pharynx until S6, whereas gelsolin-like and 4XLIM-like were expressed in both the primitive ectoderm and temporary embryonic pharynx during S3–S4 (Figure 5—source data 3, Figure 1—figure supplements 12C–G and 11D–E). Expression of these EEE transcripts is likely under zygotic control, occurring during or after the cell fate decisions to downregulate piwi-1, exit the cell cycle and differentiate.

Most EEE transcripts queried by WISH (n = 15, 75% assayed) were expressed in both undifferentiated blastomeres and temporary embryonic tissue(s) during S3–S4 (Figure 5B–E, Figure 5—source data 3). Some of these transcripts may be maternally deposited, albeit we cannot ascertain the relative contribution(s) of maternal and zygotic expression from our RNA-Seq data. Expression of these EEE transcripts diminished greatly by S5, with EEE transcripts that had been expressed at moderate or low levels becoming undetectable; specific expression of robustly expressed transcripts sometimes persisted until S6 (Figure 5B–E, Figure 5—source data 3). Consistent with the RNA-Seq and Nanostring nCounter results, EEE transcript expression was not detected by colorimetric WISH in S7 or S8 embryos or in C4 adults (Figure 5B–E, Figure 5—source data 3). Fluorescent double WISH performed with riboprobes complementary to piwi-1 and the EEE transcripts tct-like, BTF3-like, DDX5-like and eIF4a-like revealed coincident expression throughout the S4 blastomere compartment (Figure 5F–I). Intriguingly, EEE transcript expression often decayed quicker in differentiated cells than in undifferentiated blastomeres, raising the possibility that regulation of EEE transcription and/or transcript stability may vary by cell type. Robust expression of many EEE transcripts was detected in blastomeres during S3–S4, EEE transcript expression in temporary embryonic tissues was present during S3 and drastically diminished by S4 (Figure 5B–E, Figure 5—source data 3).

EEE transcripts expressed throughout the undifferentiated piwi-1+ blastomere population in S3–S4 embryos are downregulated as definitive organogenesis begins during S5. These transcripts likely represent a key temporal shift in the expression profile of piwi-1+ cells during embryogenesis. Moreover, EEE transcript expression provides a molecular metric to distinguish piwi-1+ blastomeres from adult neoblasts.

Adult lineage progenitors arise within piwi-1+ blastomeres as organogenesis begins

Many developmental regulators implicated in lineage commitment and differentiation were expressed at low levels in S2–S4 embryos, and were upregulated dramatically as definitive organogenesis began during S5. Key regulators of cell fate specification for many tissues, including the epidermis (p53 and zfp-1) (Pearson and Sánchez Alvarado, 2010; van Wolfswinkel et al., 2014; Wagner et al., 2012), nervous system (coe, sim, pax3/7 like, lhx1/5–1 and pitx) (Cowles et al., 2013; Currie and Pearson, 2013; März et al., 2013; Scimone et al., 2014), excretory system (pou2/3, six1/2–2, eya, sal1 and osr) (Scimone et al., 2011), photoreceptor neurons (eya, six-1/2, otxA and soxB), pigment cup cells (eya, six-1/2, sp6-9 and dlx) (Lapan and Reddien, 2011, 2012) and primordial germ cells (nos) (Wang et al., 2007) were among the S5- and/or S6-enriched transcripts (Figure 1—source data 56). Additional validated or putative drivers of cell fate determination in muscle (myoD) (Cowles et al., 2013; Scimone et al., 2014), the gastrovascular system (prox-1 and foxA1) (Adler et al., 2014; Scimone et al., 2014; van Wolfswinkel et al., 2014), nervous system (lhx2/9, six3-1, nkx6-like, otxB-like, otxA, pax6a and pax6b) (Pineda et al., 2002; Scimone et al., 2014) and eyes (ovo) (Lapan and Reddien, 2012) showed statistically significant upregulation during S5 and/or S6, albeit with adjusted p-values and fold-changes above the stringent thresholds set for inclusion in the S5–S6-enriched transcript lists. GO analysis on S5-enriched transcripts showed statistically significant enrichment of terms associated with patterning and cell fate specification, transcriptional regulation, and development of organ systems including the epidermis, central and peripheral nervous system, muscle, digestive and excretory systems (Figure 1—source data 5). Taken together, these observations suggest that formation of many definitive organ systems begins during S5, a supposition bolstered by WISH developmental time course data depicting expression patterns for numerous progenitor and cell type-specific markers during embryogenesis (Figure 1—source data 9, Figure 1—figure supplements 1319).

Adult asexual knockdown phenotypes for many developmental TFs upregulated during S5 and S6 suggest that these genes are required for lineage specification, tissue maintenance, and production of new tissue during regeneration, with correspondence between affected tissues and site(s) of expression. Heterogeneous expression of TFs in neoblasts and post-mitotic progenitors informed the hypothesis that the neoblast population contains pluripotent stem cells as well as cycling, lineage-primed progenitors (Reddien, 2013). Single cell sequencing (SCS) studies suggest that the zeta (ζ) and gamma (γ) neoblast subclasses are epidermal and gut progenitors, respectively (van Wolfswinkel et al., 2014; Wurtzel et al., 2015), while the sigma (σ) neoblast subclass likely contains both pluripotent stem cells and progenitors for other lineages, including neural subtypes, protonephridia and primordial germ cells (van Wolfswinkel et al., 2014). In practice, coexpression of pan-neoblast markers (e.g., piwi-1) and developmental TFs is used for neoblast subclass identification.

Smed embryos are wholly reliant on cycling piwi-1+ cells for creation of new tissues, and heterogeneous expression of key developmental TFs within piwi-1+ blastomeres is predicted to generate the diverse array of progenitors required for organogenesis. While only a small fraction of the piwi-1+ compartment is predicted to be double positive for any given lineage marker, the entire population of lineage-positive cells is predicted to be positive for piwi-1 at its inception. As development proceeds, the fraction of lineage-positive cells coexpressing piwi-1 will decrease as cells downregulate expression of piwi-1 and differentiate further.

If parallels with neoblasts hold true, then piwi-1+ blastomeres should self-renew and give rise to differentiating progeny during S5. To assay for cells exiting the piwi-1+ compartment during S5, embryos were costained with piwi-1 riboprobes and PIWI-1 antibodies. In adults, piwi-1 mRNA is restricted to the neoblast population, whereas PIWI-1 protein perdures in early post-mitotic progeny (Guo et al., 2006; Scimone et al., 2011; Wagner et al., 2011). Indeed, recent work suggests that mechanisms exist to sequester piwi-1 mRNA and chromatoid bodies within one daughter cell during neoblast division, producing one cell that maintains neoblast identity and one cell that differentiates (Lei et al., 2016). Virtually all S5 piwi-1+ blastomeres also contained PIWI-1 protein (Figure 6—figure supplement 1). Rare cells positive for PIWI-1 protein but in which piwi-1 mRNA was undetectable were observed in S5 embryos, suggesting that some of the division progeny exited the piwi-1+ blastomere population (Figure 6—figure supplement 1).

To address whether lineages required for organogenesis arise within piwi-1+ blastomeres during S5, expression of four evolutionarily conserved TFs implicated in tissue differentiation across three germ layers was examined singly and in combination with piwi-1. p53 and pax6a, regulators of epidermal and neural fates, respectively, were proxies for ectodermal derivatives. Populations expressing myoD, a master regulator of muscle fate, were considered mesodermal derivatives, and populations expressing gata456a, a regulator of gut development, represented endodermal derivatives. While expression of these TFs was occasionally detected in a small number of cells during S4, robust expression of p53 (Figure 1—figure supplement 15B), gata456a (Figure 1—figure supplement 13F), myoD (Figure 1—figure supplement 17A) and pax6a (Figure 1—figure supplement 16B) manifest in scattered parenchymal cells during S5. Indeed, fluorescent double WISH with piwi-1 and p53, gata456a, myoD or pax6a identified prospective epidermal, gut, muscle and neural progenitor populations that coexpressed piwi-1 and the developmental TFs in S5 embryos, as well as single positive populations for all of the markers assayed (Figure 6A–D, Videos 1114).

Figure 6 with 1 supplement see all
Adult lineages arise within the piwi-1+ blastomere population as organogenesis begins.

(AD) Developmental transcription factors implicated in tissue specific differentiation programs are expressed in subpopulations of piwi-1+ cells during S5. Fluorescent WISH with piwi-1 (red) and p53 (A), gata456a (B), myoD (C) and pax6a (D) (green) riboprobes on S5 embryos. Embryos in (B-D) were costained with VAL-like, a temporary embryonic pharynx specific marker (also in red). Right: Venn diagrams depict percentages of cells that were single or double positive for piwi-1 and the indicated TFs. Scale bars: 100 µm. (EG) Hierarchical clustering of zeta (ζ, E), gamma (γ, F) and sigma (σ, G) neoblast subclass-enriched transcripts during embryogenesis (Y and S2–S8), and in asexual (C4) and virgin sexual (SX) adults.

https://doi.org/10.7554/eLife.21052.052
Video 11
Definitive epidermal progenitors arise in the piwi-1+ blastomere population during S5.

SPIM -reconstructed S5 embryo costained with piwi-1 (red) and p53 (green). Definitive epidermal progenitors, coexpressing piwi-1 and p53, are dispersed in the embryonic wall. As definitive epidermal progenitors differentiate, they are predicted to downregulate piwi-1 and to retain expression of p53.

https://doi.org/10.7554/eLife.21052.055
Video 12
Definitive gut progenitors arise in the piwi-1+ blastomere population during S5.

SPIM reconstructed S5 embryo costained with piwi-1 and VAL-like (both in red) and gata456a (green). Definitive gut progenitors, coexpressing piwi-1 and gata456a, are dispersed in the embryonic wall. As definitive gut progenitors differentiate, they are predicted to downregulate piwi-1 and to retain expression of gata456a. VAL-like is expressed the temporary embryonic pharynx and is not detected in piwi-1+ blastomeres.

https://doi.org/10.7554/eLife.21052.056
Video 13
Muscle progenitors arise in the piwi-1+ blastomere population during S5.

SPIM reconstructed S5 embryo costained with piwi-1 and VAL-like (both in red) and myoD (green). Muscle progenitors, coexpressing piwi-1 and myoD, are dispersed in the embryonic wall. As muscle progenitors differentiate, they are predicted to downregulate piwi-1 and to retain expression of myoD. VAL-like is expressed the temporary embryonic pharynx and is not detected in piwi-1+ blastomeres.

https://doi.org/10.7554/eLife.21052.057
Video 14
Neural progenitors arise in the piwi-1+ blastomere population during S5.

SPIM reconstructed S5 embryo costained with piwi-1 and VAL-like (both in red) and pax6a (green). Neural progenitors, coexpressing piwi-1 and pax6a, are dispersed in the embryonic wall. As neural progenitors differentiate, they are predicted to downregulate piwi-1 and to retain expression of pax6a. VAL-like is expressed the temporary embryonic pharynx and is not detected in piwi-1+ blastomeres.

https://doi.org/10.7554/eLife.21052.058

The advent of developmental TF expression within piwi-1+ blastomeres during S5, coupled with downregulation of EEE transcripts, may signal the emergence of molecularly distinct subpopulations akin to the neoblast subclasses. Whole embryo expression trends for EEE, σ, γ and ζ class transcripts suggest that large scale, cell-intrinsic shifts in gene expression occur within blastomeres between S4 and S5. Moreover, the developmental output of piwi-1+ blastomeres, as described by molecular fate mapping, diversifies greatly during S5 and S6.

The expression signatures of the ζ and γ neoblast subclasses emerge during S5. ζ neoblasts require p53 and zfp-1 activities for production of post-mitotic mesenchymal progenitors, which simultaneously differentiate, migrate and ultimately integrate into the epidermis during homeostasis (van Wolfswinkel et al., 2014). p53, zfp-1, tcf15, sox-P3, fgfr1, egr-1, six6*, gata123* (van Wolfswinkel et al., 2014) and 15 additional transcripts used as classifiers of ζ subgroup identity in SCS experiments (Wurtzel et al., 2015) displayed statistically significant upregulation in whole embryos during S5 or later developmental stages (Figure 6E, Figure 6—source data 1, Figure 1—figure supplement 15A, Figure 1—source data 58). Furthermore, the ζ neoblast transcripts soxP3, egr-1, fgfr1 and prog-1 clustered together in the analysis of adult asexual neoblast-enriched transcripts, indicating that they displayed similar expression profiles during embryogenesis (Figure 4A, Figure 4—source data 1). Transcripts specifically expressed in post-mitotic epidermal progenitors downstream of ζ neoblasts, including prog-1, AGAT-1 and zpuf-6, were enriched and first detected by WISH during S5 (Figure 1—figure supplement 15C,E–G, Figure 1—source data 5). γ neoblasts are identified by enriched expression of gata456a, hnf4, prox-1 and nkx2.2* (van Wolfswinkel et al., 2014; Wagner et al., 2011). prox-1 and nkx2.2 were expressed at low levels in S2–S4 embryos and showed statistically significant upregulation during S5, and 15 additional γ neoblast-enriched transcripts identified in SCS experiments were enriched during S5 or later in development (Figure 6F, Figure 6—source data 1, Figure 1—figure supplement 13E, Figure 1—source data 58). The molecular staging series detected expression of gata456a and hnf4 in yolk and early embryos (S2–S4) (Figure 6F). However, gata456a and hnf4 expression were solely detected in the embryo proper by WISH, first in the developing temporary embryonic pharynx during S2 and later in parenchymal cells during S5 (Figure 1—figure supplement 13F–G).

Charting the emergence of the σ neoblast subclass is hampered by limitations of the subclass designation: σ neoblasts are presumed to be an amalgamation of several progenitor populations and pluripotent stem cells. Several σ -class genes, notably SoxP-1 and SoxP-2, were expressed in parenchymal cells throughout embryogenesis, similar to piwi-1 (Figure 4A, Figure 4—source data 1, Figure 4—figure supplement 1A–C, Figure 6—source data 1). SoxP-1 is required for neoblast maintenance (Wagner et al., 2012), suggesting that σ-class transcripts with expression profiles to SoxP-1 and SoxP-2 during embryogenesis may also be expressed in pluripotent neoblasts. In contrast, lineage-primed progenitor factions within the σ subclass probably arise during S5. Genes with functions in tissue-specific differentiation programs, including soxB1 (Lapan and Reddien, 2012; Monjo and Romero, 2015), pou2-3 (Scimone et al., 2011), nos (Wang et al., 2007), pitx (Currie and Pearson, 2013; März et al., 2013), sim (Cowles et al., 2013), and smad6/7–1 (González-Sastre et al., 2012) were expressed at low levels in S2–S4 embryos and were upregulated during S5 (Figure 6G, Figure 6—source data 1, Figure 1—source data 5).

The gene expression signature of the adult neoblast compartment is an emergent property of the piwi-1+ population during embryogenesis. First, EEE transcripts are uniquely associated with the expression signature(s) of undifferentiated piwi-1+ blastomeres in early embryos. Second, adult neoblast subclasses arise as lineages are born within piwi-1+ blastomeres during S5. Molecular heterogeneity within the neoblast compartment is largely attributed to the diverse array of lineage-dedicated progenitors within the population, a hypothesis supported by our finding that subclass marker expression was dramatically upregulated as organogenesis began. Progenitor subpopulations required for organ formation during embryogenesis persist into adulthood, where steady-state output from different lineages maintains tissue homeostasis. At present, we cannot distinguish whether lineages perpetually re-emerge due to asymmetric division of pluripotent stem cells, or whether progenitor populations established during embryogenesis are maintained by self-renewal. These observations beg the question: do piwi-1+ cells behave similarly to neoblasts throughout embryogenesis? Or alternatively, is neoblast activity an acquired feature that emerges in tandem with the adult molecular expression signature?

Embryos undergoing organogenesis contain cells with cNeoblast activity

Neoblasts are completely and irreversibly eliminated following treatment with 6,000 Rads (Reddien et al., 2005; Wagner et al., 2011), causing irradiation sickness and ultimately death. Transplantation of wildtype adult tissue grafts or cell suspensions into lethally irradiated adult hosts that are devoid of stem cells results in engraftment and expansion of donor-derived piwi-1+ cells, production of differentiated progeny, reconstitution of the neoblast compartment and rescue from lethality (Baguñà J and Auladell, 1989; Guedelhoefer and Sánchez Alvarado, 2012; van Wolfswinkel et al., 2014; Wagner et al., 2011). piwi-1+ cells that form pluripotent, expanding colonies following sublethal irradiation or transplantation into a lethally irradiated host are called clonogenic neoblasts (cNeoblasts) (Wagner et al., 2011). cNeoblasts are predicted to have a widespread distribution in the parenchyma, and this population contains within it the most primitive stem cells. At present, the operational definition for cNeoblast exists apart from the gene expression signatures for the neoblast subclasses; cNeoblasts are likely contained within, but may not be exclusive to, the σ class.

To assess whether Smed embryos harbor piwi-1+ cells with cNeoblast activity, heterochronic, heterotopic transplantations were performed. S4, S5, S6, S7 and S8 embryonic cell suspensions were injected into the tail parenchyma of lethally irradiated sexual adult hosts at 3 days post-irradiation (dpi) (Figure 7A, Materials and Methods). To determine whether comparable numbers of piwi-1+ cells were introduced per host for the developmental stages assayed, transplanted hosts were fixed at 1 hr post-transplant (hpt) and stained with piwi-1 riboprobes. More than 85% of S4–S8 transplants fixed at 1 hpt contained piwi-1+ cell(s) in the tail parenchyma, suggesting that the cell injection technique was robust and reliable (Figure 7B). S5, S6, S7 and S8 transplants contained comparable numbers of piwi-1+ cells per host at 1 hpt, while significantly fewer piwi-1+ cells were introduced per S4 embryonic cell transplant (Figure 7C,F).

Embryonic cells acquire the ability to engraft, persist and proliferate in an adult microenvironment as organogenesis proceeds.

(A) Schematic depicting the workflow for heterochronic transplantation experiments. S4, S5, S6, S7 or S8 embryonic cell suspensions were injected into the tails of lethally irradiated sexual adult hosts at 3 days post-irradiation (dpi). Cohorts of transplanted animals were fixed at 1 hr and 5 days post-transplantation (1 hpt and 5 dpt, respectively) for staining with piwi-1 riboprobes and H3S10p antibodies. Lethally irradiated, uninjected host controls were fixed and stained at 5 dpt. (B) Percentage of transplanted animals fixed at 1 hpt (blue bars) or 5 dpt (red bars) containing one or more donor-derived piwi-1+ cell(s). X-axis: stage (S) of donor cells. (C) Number of donor-derived piwi-1+ cell(s) per transplant at 1 hpt and 5 dpt. Each point represents one transplanted animal. Mean ± standard deviation (black bars) are shown. Statistical tests were performed using a generalized linear model, assuming that the counts followed a Poisson distribution. S4 transplants contained significantly fewer piwi-1+ cells at 1 hpt than S5, S6, S7 or S8 transplants (Tukey post-hoc comparisons, S4 vs S5, S4 vs S6 and S4 vs S7, S4 vs S8: p<0.001). Group differences in the number of piwi-1+ cells at 1 hpt for S5 and S6 transplants were not statistically significant (p=0.21). Significantly fewer S4 and S5 donor-derived piwi-1+ cells persisted at 5 dpt than were observed for later stages (Tukey post-hoc comparisons: S4 vs S5, S4 vs S6, S4 vs S7 and S4 vs S8: p<0.001. S5 vs S6, S5 vs S7, S5 vs S8: p<0.001). (D) Percentage of transplants with mitotic piwi-1+ cell(s) at 5 dpt (green bars). X-axis: Donor cell stage. (E) Mitotic index for donor-derived piwi-1+ cells at 5 dpt. Stage-specific differences were not observed for S4–S8 embryonic cell populations using a generalized linear model, assuming counts followed a Poisson distribution and the number of piwi-1+ cells as a covariate. (BE) Numbers of transplants scored: S4: n = 36 (1 hpt), n = 43 (5 dpt), four independent experiments. S5: n = 15 (1 hpt), n = 16 (5 dpt), two independent experiments. S6: n = 31 (1 hpt), n = 29 (5 dpt), four independent experiments. S7: n = 31 (1 hpt), n = 30 (5 dpt), four independent experiments. S8: n = 19 (1 hpt), n = 20 (5 dpt), three independent experiments. (F) Confocal maximal projections of S4, S5, S6, S7 and S8 embryonic cell transplants fixed at 1 hpt and 5 dpt. Animals were stained with piwi-1 riboprobes (green), antibodies against the mitotic marker H3S10p (red, 5 dpt only) and DAPI nuclear counterstain (blue). S6, S7 and S8 insets: mitotic piwi-1+ cells. Red arrows indicate mitotic cells magnified in insets. Yellow arrows: mitotic piwi-1+ cells. Scale bar (inset): 20 µm. Scale bar (panel): 100 µm. (BC) S4–S8 embryonic piwi-1+ cells were reliably introduced into hosts. S6–S8 embryonic piwi-1+ cells persisted in an adult microenvironment. (D-E) S6–S8 embryonic piwi-1+ cells proliferated in an adult microenvironment.

https://doi.org/10.7554/eLife.21052.059

To assay whether embryonic piwi-1+ cells persisted and proliferated in an adult microenvironment, cohorts of transplanted animals and lethally irradiated uninjected hosts were fixed at 5 days post-transplantation (dpt) and stained with piwi-1 riboprobes and H3S10p antibodies (Figure 7B–F). Persistent piwi-1+ cells from S6, S7 and S8 embryos were observed in the vast majority of transplants scored at 5 dpt (Figure 7B), and no statistically significant difference among stages was detected in the mean number of piwi-1+ cells present per host (Figure 7C). Moreover, most S6, S7 and S8 embryonic donor cell transplants contained dividing piwi-1+ cell(s) at 5 dpt (Figure 7D–F), and no statistically significant difference among stages was detected in the mitotic index of donor-derived piwi-1+ cells per host at 5 dpt (Figure 7E). In contrast, S4 and S5 embryonic piwi-1+ cells were significantly less likely to persist in adult hosts at 5 dpt than were S6, S7 or S8 embryonic piwi-1+ cells (Figure 7B–C). Persistent S4 derived piwi-1+ cells were rarely observed at 5 dpt, and cell division was not observed (Figure 7D–E). As expected, piwi-1+ cells were never observed in uninjected lethally irradiated hosts (n = 82 hosts scored at 3 dpi).

Fewer S5 embryonic cell transplants contained persistent piwi-1+ cells at 5 dpt (Figure 7B), and the number of S5 derived piwi-1+ cells per host at 5 dpt was significantly lower than for later stages (Figure 7C). Similarly, the fraction of S5 embryonic cell transplants containing mitotic piwi-1+ cell(s) was reduced relative to S6 and later stages (Figure 7D). The reduced persistence of S4 and S5 derived piwi-1+ cells in an adult microenvironment was probably not attributable to technical variability or to the absolute number of embryonic piwi-1+ cells introduced per host, since comparable numbers of S5, S6, S7 and S8 piwi-1+ cells were introduced per transplant (Figure 7C). Stage-specific, cell autonomous factors likely underlie the starkly different responses of S5 and S6 embryonic cells following transplantation into the adult parenchyma. These results suggest that S4 and S5 piwi-1+ blastomeres are functionally distinct from piwi-1+ cells present at S6 and later stages. piwi-1+ blastomeres acquire competency to engraft and respond appropriately to adult environmental cues as development proceeds during S5.

To assess whether embryos undergoing organogenesis harbor cells capable of rescuing and reconstituting lethally irradiated adult hosts, S5, S6, S7 and S8 embryonic cell suspensions were injected into lethally irradiated sexual adult hosts at 1 dpi (Figure 8A, Materials and methods). Consistent with previous results, persistent piwi-1+ cells from S6, S7 and S8 embryos were observed in the vast majority of samples scored at 5 dpt (Figure 8B), and no statistically significant difference was detected in either the mean number or the mitotic index of piwi-1+ donor-derived cells per host at 5 dpt (Figure 8C–D). Likewise, donor-derived piwi-1+ cells from S5 embryos were far less likely to persist and divide in adult hosts (Figure 8B–D). As expected, piwi-1+ cells were never observed in uninjected lethally irradiated hosts (n = 81 hosts scored at 6 dpi).

Figure 8 with 1 supplement see all
Embryos undergoing organogenesis contain cNeoblasts.

(A) Schematic for heterochronic transplantation experiments. S5, S6, S7 or S8 embryonic cell suspensions were injected into the tail parenchyma of lethally irradiated sexual adult hosts at 1 day post irradiation (dpi). Cohorts of transplanted animals and uninjected host controls were fixed at 5 days post-transplantation (dpt) for staining with piwi-1 riboprobes and H3S10p antibodies. The remaining animals were monitored for 70 dpt for survival and rescue. (B) Percentage of transplants with persistent, donor-derived piwi-1+ cell(s) (blue) or donor-derived mitotic (piwi-1+, H3S10p+) cell(s) (red) at 5 dpt. X-axis: Embryonic donor cell stage. (C) Number of piwi-1+ cells per transplanted host at 5 dpt for S5–S8 embryonic cell transplants. Each point represents one transplanted animal. Means ± standard deviation (SD) are shown (black bars). Statistically significant differences in the number of persistent piwi-1+ cells per transplant at 5 dpt were observed using a generalized linear model, assuming that count data followed a Poisson distribution. S5 transplants contained fewer persistent piwi-1+ cells than S6 or S7 transplants (Tukey post-hoc comparisons, S5 vs S6: p<0.0001, S5 vs S7: p<0.0001, S5 vs S8: p<0.0001). (D) Mitotic index for donor-derived piwi-1+ cells at 5 dpt for S5–S8 embryonic cell transplants. Each point represents one transplanted animal. Means ± standard deviation (SD) are shown (black bars). Statistically significant differences in the piwi-1+ cell mitotic index were observed using a generalized linear model with piwi-1+ cell counts as a covariate, assuming that count data followed a Poisson distribution. S5 transplants contained significantly fewer cycling cells than S6, S7 or S8 transplants (Tukey post-hoc comparisons, S5 vs S6: p<0.01, S5 vs S7: p<0.01, S5 vs S8: p<0.001). (E) Confocal maximal projections for S5, S6, S7 and S8 embryonic cell transplants fixed at 5 dpt and stained with piwi-1 riboprobes (green), H3S10p antibodies (red) and DAPI (blue). S6, S7 and S8 insets show mitotic piwi-1+ cells. Red arrows indicate mitotic cells magnified in the insets. Yellow arrows: mitotic piwi-1+ cells. Scale bar (inset): 20 µm. Scale bar (panel): 100 µm. (BE) Numbers of transplants scored in four independent experiments: S5 n = 22; S6 n = 24; S7 n = 21; S8 n = 27 in (C), n = 21 in (D). (F) Survival curves for S5, S6, S7 and S8 embryonic cell transplants and uninjected 6,000-Rad-irradiated host controls as a function of time (days) post-transplant. (G) Live images of regenerating S6, S7 and S8 rescue animals. Left: Tail fragment after self-amputation of head and trunk tissue. Middle: Tail fragment with unpigmented anterior blastema (yellow arrowheads). Right: Animal with new head tissue and developing eyes (yellow arrows) and a regenerated pharynx (yellow asterisk). Animals from different experiments are shown in the S7 panels; the same animals are shown in the S6 and S8 panels. Dorsal view. Anterior: top. Scale: 100 µm. (FG) Numbers of transplants scored in four independent experiments: host controls n = 89; S5 n = 105; S6 n = 90; S7 n = 92; S8 n = 85. Rescue animals were obtained in two experiments for S6 and S7 transplants, and four experiments for S8 transplants. (BE) S6, S7 and S8 embryonic donor cells persist and divide in the adult parenchyma. (FG) S6, S7 and S8 embryonic cells can rescue lethally irradiated adult hosts.

https://doi.org/10.7554/eLife.21052.060

Survival of S5–S8 transplants and host controls was monitored for 70 dpt (i.e., 71 dpi). Worms were scored for irradiation-induced phenotypes, including head and tail regression, lesion formation, ventral curling and death. Rescue, anterior blastema formation and subsequent regeneration of an entire individual from injected tail fragment, was also scored (Guedelhoefer and Sánchez Alvarado, 2012; Wagner et al., 2011) (Figure 8F–G, Figure 8—figure supplement 1). Irradiation-induced phenotypes manifest in control and transplanted individuals alike between 7–14 dpt (i.e., 8–15 dpi) (Figure 8—figure supplement 1A–E), and none of the controls survived long-term (Figure 8F, Figure 8—figure supplement 1E). Remarkably S6, S7 and S8 embryonic donor cells were capable of rescuing lethally irradiated adult hosts (Figure 8F–G). Rescued animals underwent complete head regression, self-amputation posterior to the pharynx, and tail fragments that formed anterior blastemas which promoted regeneration of individuals containing two visible eyes, a central pharynx and triclad gut (Figure 8F–G, Figure 8—figure supplement 1B–D). None of the S5 embryonic cell recipients mounted a rescue response (Figure 8F, Figure 8—figure supplement 1A). The rescue assay results suggest that S6, S7 and S8 embryos harbor cNeoblasts, cells that are capable of self-renewing and producing the diverse array of cell types required for whole animal regeneration.

Taken together, stage-dependent molecular and functional distinctions exist among piwi-1+ blastomeres before and after organogenesis begins. S4 piwi-1+ cells, which express EEE and adult asexual neoblast enriched transcripts, are largely incapable of persisting and dividing in an adult microenvironment (Figure 7B–C). During S5, as dramatic shifts in gene expression occur, piwi-1+ cells become competent to persist and proliferate in the adult parenchyma (Figures 7B–C and 8B–D). We propose that cNeoblasts arise during S5. Heterochronic transplantation experiments revealed that S6, S7 and S8 embryos possess cells that behaved similarly to adult cNeoblasts: they consistently engrafted into adult hosts, proliferated and were ultimately capable of mounting a rescue response (Figure 8B–G). Acquisition of cNeoblast activity during embryogenesis correlates temporally with large-scale changes in gene expression observed at the outset of organogenesis, suggesting that pluripotent stem cells and lineage-primed progenitors first emerge during S5 (Figure 9). The ontogeny of the adult neoblast compartment can therefore be traced back to the piwi-1+ zygote, which gives rise to anarchic, cycling piwi-1+ blastomeres, some of which persist in the S3–S4 embryonic wall and establish pluripotent neoblasts and progenitor subpopulations during S5. The remarkable developmental plasticity of adult planarians is likely due to the singular ability of neoblasts to perpetuate and redeploy embryonic developmental programs.

Ontogeny of the adult neoblast compartment.

Asynchronously cycling piwi-1+ cells fuel embryogenesis, giving rise to all temporary and definitive tissues. During S2, some piwi-1+ blastomeres (purple cells) exit the cell cycle and differentiate into temporary embryonic tissues (primitive ectoderm, temporary embryonic pharynx and primitive endoderm). The remaining piwi-1+ blastomeres, located in the embryonic wall (purple cells, S3-S4), continue to divide and express both EEE transcripts (turquoise arrow) and adult asexual neoblast enriched transcripts (red arrow). As organogenesis begins during S5, EEE transcripts are downregulated throughout the compartment (purple cells transition into red). Concomitantly, progenitor subpopulations required for definitive organ formation are specified via the heterogeneous expression of developmental transcription factors within the piwi-1+ population (colored cells denote different progenitor subpopulations). Adult pluripotent neoblasts, themselves a lineage, are established during S5 (red cells). Embryonic donor cells harvested during or after S6 function similarly to adult neoblasts (cNeoblast activity, gray arrow). Pluripotent and lineage-primed neoblasts established during embryogenesis are maintained throughout the lifetime of the animal. Neoblasts are required for tissue maintenance during homeostasis and the formation of new tissue during regeneration.

https://doi.org/10.7554/eLife.21052.062

Discussion

Regeneration remains one of the most poorly understood processes in developmental biology. The origin and regulation of cells that make regeneration possible are largely obscure. How similar or distinct developmental processes are during embryogenesis and regeneration remains to be determined. In fact, few experimental systems are available to study these issues systematically. Here, we lay the foundation for the formal comparison of molecular processes and gene functions during embryogenesis and regeneration in Smed, an organism uniquely suited to address the relationship between developmental plasticity and regeneration competency.

S. mediterranea: a developmental system for comparative studies of embryogenesis and regeneration

To identify similarities and key differences between embryogenesis and regeneration, careful consideration must be given to distinctions in context, chronology, scope and type of regeneration (homeostatic or facultative), and to the ontogeny of effector cell type(s). Which aspects of embryonic development are recapitulated during regeneration, and which are context-specific? What factors influence the competency and extent of regenerative responses, and do these factors have embryonic origins? The workhorses of modern developmental biology, C. elegans, D. melanogaster, D. rerio, and M. musculus have limited, if any, regenerative potential during adulthood, precluding or severely limiting comparative inquiries. In contrast, Smed adults exhibit robust homeostatic and facultative regenerative potential, both neoblast-dependent phenomena that have largely been studied using a clonal, asexually reproducing strain. Neither descriptive nor functional studies of Smed embryogenesis have been reported.

We generated two foundational resources: a molecular staging series for Smed embryogenesis and an expression atlas describing temporary and definitive organ development. To investigate the embryonic origins of regeneration, we provide an ontogeny for the adult neoblast compartment. The molecular staging series facilitated the identification of embryonic predecessors of adult neoblast lineages and the developmental transition when neoblast specification occurs. Adult neoblast lineages, including pluripotent stem cells and lineage-primed progenitors, are established as definitive organogenesis begins. Neoblast lineages, required for organ formation during embryogenesis, persist into adulthood, where they are redeployed for homeostatic maintenance of all differentiated tissues, including the germline, and formation of missing tissues during regeneration. Moving forward, investigation of neoblast dynamics, particularly the regulation of self-renewal and commitment to differentiation, will be a central, unifying feature of comparative studies on organ formation, maintenance, repair and replacement.

Deciphering the origin and identity of neoblasts, agents of development, tissue homeostasis and regeneration

Unraveling the molecular mechanism of neoblast specification during embryogenesis will provide insight into the identity of adult pluripotent stem cells indispensible for homeostatic and facultative regeneration in planarians. Intensive efforts to define the term neoblast, and to characterize the presumed plurality of cell types within the asexual adult neoblast population, have generated numerous, unreconciled molecular and functional criteria. At present, gene expression signatures for neoblast subpopulations cannot be correlated with functional distinctions (should they exist) in self-renewal, pluripotency or cell fate restriction. It is not known whether hierarchical relationships exist between pluripotent cells and progenitors, whether cycling progenitors can dedifferentiate and/or interconvert, how stable subpopulations are over time, or when and how progenitors make an irreversible commitment to differentiation. Furthermore, preferred usage of Smed C4 asexual animals to classify and examine relationships among subpopulations precludes comprehensive examination of the lineage repertoire.

The expression of pluripotency factors is probably necessary, but not sufficient, for the assumption of neoblast fate. We report that many genes known to regulate adult asexual neoblast self-renewal and maintenance were expressed throughout embryogenesis, and showed that several of these genes were expressed throughout S3–S5 piwi-1+ blastomeres. Functional differences between blastomeres and neoblasts were observed in heterochronic, heterotopic transplantation experiments. S4 and S5 embryonic donor cells were far less likely to persist and proliferate in stem cell deficient adult hosts than were embryonic donor cells harvested during S6 or later. Moreover, S5 embryonic donor cells did not rescue lethally irradiated hosts, whereas S6–S8 embryonic donor cells functioned like adult neoblasts and were capable of rescue. These results suggest that neoblast specification occurs during S5, and highlight the importance of cell-intrinsic changes in the blastomere to neoblast transition.

Differences in transplanted cell behavior correlated with large-scale changes in gene expression as definitive organogenesis began. EEE transcripts, uniquely expressed in blastomeres, are attractive candidates for repressors of neoblast fate, and/or effectors of molecular changes in blastomeres that are necessary for acquisition of neoblast fate. It is not known whether EEE transcript downregulation in blastomeres is necessary for neoblast specification, or whether expression of EEE transcripts and developmental TFs required for lineage specification are mutually exclusive. Further studies will determine whether and how maternal factors influence the early stages of Smed embryogenesis. Which EEE transcripts are maternally deposited, and which mechanisms effect maternal transcript degradation? When do wave(s) of zygotic genome activation occur, and how does zygotic genome activation in blastomeres relate to cell differentiation and neoblast specification? Additional factors, such as changes in chromatin state, may also correlate with or play a causative role in the blastomere to neoblast transition.

The paradox of pattern formation: robustness of an anarchic, self-organizing system and its implications for understanding regeneration

Ectolecithal development and dispersed cleavage pose unique developmental challenges, making Smed embryogenesis a novel paradigm for regulative development. Herein, the collective pluripotency of an anarchic, cycling piwi-1+ population generates the diversity of cell types required for the development of all temporary and definitive organ systems in these bilaterally symmetric, triploblastic animals. We showed that piwi-1+ blastomeres are spatially disordered within the embryonic wall during S3–S4, and that dispersed epidermal, gut, muscle and neural progenitors arise within the blastomere population during S5. Further investigation is needed to understand how signals from differentiated tissues impact cell fate decisions within the piwi-1+ blastomere compartment and effect progenitor cell migration, communication and interactions necessary to form organ rudiments.

Smed embryogenesis provides a unique vantage point from which to investigate the origin, anatomical composition and signaling logic underlying the neoblast niche. During sphere formation, differentiating blastomeres must interact and self-assemble temporary embryonic tissues, providing structure to the embryo and establishing a microenvironment that promotes maintenance and expansion of piwi-1+ blastomeres. In turn, blastomeres are likely to effect changes that are conducive to the establishment and/or maintenance of the neoblast population. Neoblast lineages are predicted to actively maintain their niche during the lifetime of the animal. Although the adult gastrovascular system has long been suspected of providing trophic support signals to neoblasts (Forsthoefel et al., 2012), the molecular mechanisms underlying this phenomenon remain elusive. Expansion of piwi-1+ blastomeres correlates with that of an ill-defined embryonic gut population (Figure 1—figure supplement 13A–D). Investigating gut development and gut communication with the piwi-1+ population during embryogenesis may uncover key regulators of neoblast specification or regulators of neoblast dynamics that may similarly impact stem cell behavior during adulthood.

We report that many transcripts implicated in lineage commitment and classification of neoblast subclasses wereexpressed at low levels in early embryos and were dramatically upregulated as organogenesis began. This observation is consistent with the hypothesis that neoblast heterogeneity is due to the presence of different subpopulations of cycling, lineage-primed progenitors within the compartment (Reddien, 2013). It also suggests that organ formation during embryogenesis probably utilizes many of the same genetic regulatory networks and transition states elucidated during adult homeostasis and regeneration. Development of techniques to interrogate gene function during embryogenesis will enable us to identify master regulators of organogenesis for different tissues, and to address similarities and differences in their modes of action during embryogenesis and adulthood. Embryogenesis may also provide a vantage point for the identification of upstream activators for these developmental TFs, helping to address how diverse, dispersed patterns of gene activation arise in the blastomere and neoblast compartments.

The key distinction between embryogenesis and regeneration is the de novo formation of tissues in the former, and the presence of preexisting structures and signaling environments in the latter. Studies can now be performed to assess how tissues that are hypothesized to be instructive for regeneration are initially established during embryogenesis, and how the formation of these tissues relates to the acquisition of regenerative potential during development. For example, planarian body wall muscle is hypothesized to be required for the re-specification of axial identities during regeneration (Witchley et al., 2013). At present, technical limitations preclude tissue-specific knockdown experiments that would address requirements for polarity genes in muscle during regeneration. However, we can now address when and how the definitive axes are established during Smed embryogenesis, including the identification of tissues and signals that initially polarize embryos prior to the development of body wall muscle. Examining axis formation during embryogenesis, and comparing the process across different chronological and developmental contexts, may uncover roles for additional tissues and/or novel polarity regulators. Furthermore, we can address which developmental milestones and gene products are required to establish a state of regeneration competency in the embryo.

Sustained effort and continued investment in the adaptation and development of new technologies for the molecular interrogation of Smed embryogenesis will facilitate discoveries that may challenge long-held assumptions about developmental processes, including cell fate specification, pattern formation and adult stem cell regulation. Moreover, utilizing Smed for comparative studies of embryogenesis and regeneration presents an unprecedented opportunity for formal examination of the embryonic origins of regenerative potential.

Materials and methods

Planaria culture and husbandry

Sexually reproducing S. mediterranea (Smed) stocks were descendants of animals collected in Sardinia by Dr. Maria Pala in 1999. Animals from the clonally derived sexual strain S2F1L3F2 (Wagner et al., 2011) and from the asexual clonal strain CIW-4 (C4) (Newmark and Sánchez Alvarado, 2002) were propagated via successive rounds of amputation and regeneration. Animals were maintained in 1x Montjuic water at 20°C in the dark and fed homogenized beef liver as previously described in Cebrià and Newmark (2005). Cultures subjected to intensive cutting and/or feeding regimens were supplemented with 100 µg/mL gentamicin sulfate (Gemini Bioproducts, #400–100P).

Egg capsules were collected daily from outbred cohorts of sexually mature adults cultured at low density (6–8 animals per 400 mL culture), and were stored in dated Petri dishes at 20°C in constant darkness until use. The collection date was considered 1 day post-egg capsule deposition. To maintain optimal fertility levels, sexually mature animals used for egg capsule collections were replaced every 3–4 months with either juveniles (6–8 weeks post hatching) or adult regenerates (6–8 weeks post cut).

Single embryo RNA-Seq

Live embryos were dissected out of egg capsules in 1x Holfreter’s buffer (3.5 g/L NaCl; 0.2 g/L NaHCO3; 0.05 g/L KCl; 0.2 g/L MgSO4; 0.1 g/L CaCl2; 1.0 g/L dextrose, pH 7.0–7.5) for S2–S7 egg capsules, or 1x Montjuic water for S8 hatchlings. Yolk (Y) samples were obtained from 8 d egg capsules that contained neither spherical nor elongating embryos. Single embryos were imaged on a Leica M205 FA dissecting microscope, transferred into microfuge tubes containing 200 µl TRIzol reagent (Thermo Fisher, item #15596–018), homogenized by pipetting, and stored at −80°C. Single animal samples of intact C4 adults and virgin, sexually mature adults were homogenized in 1.0 mL TRIzol using an IKA Ultra Turrax T 25 Basic tissue disruptor prior to storage at −80°C. Total RNA extraction was performed in 1.0 mL TRIzol per sample according to the manufacturer’s protocol, following the recommendations for working with small amounts of tissue. Pellets were resuspended in 25 µl nuclease free water, and 5 µl aliquots were reserved for quality control testing. Total RNA concentration and integrity were determined using Agilent Bioanalyzer 2100 Expert Total RNA Nano or Pico chips (Agilent Technologies, items # 5067–1511 and 5067–1513). Total RNA samples were prepared for ten biological replicates per time point, and total RNA quality and yield were considered along with embryo size and morphology when selecting samples for library construction.

PolyA-selected, single-stranded RNA-Seq libraries were prepared for four biological replicates per stage using the Illumina TruSeq RNA Sample V2 kit (item # RS-122–2001 and RS-122–2002), starting with 500 ng total RNA per sample (C4, virgin sexual adult [SX], Y, S4, S5, S6, S7, S8), or 100 ng total RNA per sample (S2, S3). Library concentrations and insert sizes were determined using Agilent Bioanalyzer DNA 1000 chips (Agilent Technologies, item # 5067–1504), and diluted, pooled samples were reanalyzed with Agilent Bioanalyzer 2100 DNA High Sensitivity chips (Agilent Technologies, item # 5067–4626). Nine barcoded samples, one replicate per time point (S2–S8, Y, C4) were pooled and sequenced per flow cell lane. Single end, 50 bp reads were acquired on an Illumina Hi-Seq 2000 sequencer. Illumina Primary Analysis version RTA 1.13.48.0 and Secondary Analysis version CASAVA-1.8.2 were run to demultiplex reads and generate FASTQ files. Barcoded SX replicates were pooled and run on a single flow cell lane of a HiSeq 2500, and Illumina Primary Analysis version RTA 1.17.21.3 and Secondary Analysis version CASAVA-1.8.2 were used. The RNA-Seq data have been deposited in the GEO database under the accession number GSE82280.

Mapping sequencing reads to the smed_20140614 transcriptome

Sequencing reads were mapped to the smed_20140614 reference transcriptome (n = 36,035 transcripts) (Tu et al., 2015), which contains sequencing data from de novo Trinity assemblies from the C4 and sexual biotypes, staged embryo collections, sorted cycling (X1) cells, and previously published sources (Adler et al. [2014], [Böser et al. [2013]; the Dresden transcript collection at PlanMine [http://planmine.mpi-cbg.de]). Transcripts were consolidated and reduced to a unique set using the CD-HIT program (Fu et al., 2012). smed_20140614 sequences may be downloaded from http://smedgd.stowers.org. Reads were mapped using the Bowtie algorithm, Version 1.0.0 (Langmead et al., 2009), allowing for two mismatches and up to five multi-matches (--best --strata -v 2 m 5). Read counts for transcripts were tabulated from SAM files using a custom script. Of 36,035 transcripts, 32,000 accumulated ≥1 CPM across all 40 samples. Samples were each sequenced to an average depth of 19 million reads, and exhibited an average map rate of 89% to the transcriptome.

RPKM normalization

RPKM (Reads Per Kilobase per Million) values were scaled using TMM normalization (Robinson et al., 2010) in edgeR to account for read depth across samples. In addition, 16s ribosomal RNA transcripts (SMED30032887), which soak up a significant but variable fraction of reads per sample, were removed prior to calculating RPKM values.

Identification of differentially expressed transcripts

Differential gene expression was evaluated using the edgeR library (Robinson et al., 2010), and adjusted p-values were calculated as described in Hochberg (1995). Pairwise comparisons were performed between adjacent time points using edgeR: Y vs S2, S2 vs S3, S3 vs S4, S4 vs S5, S5 vs S6, S6 vs S7 and S7 vs S8. Mapped data were filtered to remove transcripts with less than a sum of 1 CPM across all 32 samples, resulting in 30,766 transcripts. The maximum read sum across samples for omitted transcripts was 14. In addition, transcripts for the 28S (SMED30027845), 18S (SMED30032663) and 16S (SMED30032887) ribosomal subunits were removed. Differentially expressed genes were identified in mixed stage reference comparisons using the GLM approach in edgeR to contrast each treatment group (i.e., developmental stage) to the average of the remaining groups (Y, S2–S8). Non-redundant lists of enriched transcripts from the pairwise and mixed stage reference comparisons, for S2 through S8, were subject to Euclidean distance clustering using scaled RPKM data in edgeR (Figure 1C–I, Figure 1—figure supplements 410, Figure 1—source data 28).

GO analysis

Gene Ontology (GO) terms (Gene Ontology Consortium, 2015) were assigned to smed_20140614 transcripts on the basis of homologous PFAM domains (Finn et al., 2014) and significant Swiss-Prot hits (E-value ≤ 0.001), (UniProt Consortium, 2015). GO term enrichment queries were performed using the R software package topGO, version 2.20.0 (Alexa and Rahnenfuhrer, 2010). GO analysis was performed on the non-redundant lists of enriched transcripts for S2–S8 (Figure 1—source data 1). Categories containing similar and/or related Biological Process (BP) GO ids enriched at one or more time point(s) were generated manually (Figure 1—source data 1). Enriched BP GO ids selected for categorization had Benjamini-Hochberg corrected p-values ≤1e-10 (Benjamini and Hochberg, 1995), and must have been associated with ≥1% of the enriched transcripts for the developmental stage(s) in question. BP GO ids were only assigned to one category. BP GO ids that did not describe a cell and/or tissue type present in Smed (e.g., heart, lung, neural crest) were omitted. Using these categories as a guide, lists of enriched BP GO ids and non-redundant lists of associated transcripts were generated for S2–S8 (Figure 1—source data 28). Transcripts may appear in more than one BP GO id category, just as transcripts may be associated with more than one GO term.

Neoblast enriched transcript analysis

The neoblast enriched transcript list (n = 242) emerged from the downregulated sequences in whole animals at 24 and/or 48 hr post-lethal irradiation in three independent experiments (Duncan et al., 2015; Wagner et al., 2012) (Chen and Sánchez Alvarado, personal communication) (Figure 4A, Figure 4—source data 1). Euclidean distance clustering was performed using the mixed stage reference comparison data.

Cloning

Constructs for riboprobe synthesis were constructed using the pPR-T4P (J. Rink) cloning strategy described in Adler et al. (2014), with the exception that PCR inserts were amplified using mixed stage embryo cDNA (S2–S8) as a template. Primers used for cloning EEE transcripts and insert sequences appear in Figure 5—source data 3.

Whole mount in situ hybridization (WISH) and immunostaining

Colorimetric and fluorescent WISH was performed as described by King and Newmark (2013) and Pearson et al. (2009), with the following modifications:

  1. 1) Egg capsules for S2–S7 embryos (2–10 days post-egg capsule deposition) were punctured with an insect pin and fixed in 4% formaldehyde in 1x PBS-Triton X (PBSTx) 0.5% for 4–6 hr at room temperature. Fixed embryos were dissected out from the egg capsules, washed in 1x PBSTx (0.5%) for 10 min, and subjected to incremental dehydration in methanol (10 min in 50% methanol, 2 × 10 min in 100% methanol). Fixed embryos were stored at −20°C. S2–S7 embryos were not bleached. Proteinase K treatment was increased to 20 min.

  2. S8 embryos (newborn hatchlings, 14–16 days post-egg capsule deposition) were removed from egg capsules prior to fixation. S8 embryos and C4 intact adults (2–4 mm) were incubated in 4% formaldehyde in PBSTx (0.5%) for one hour at room temperature.

  3. 3) S2–S7 embryos were not bleached. S8 embryos and C4 adults were bleached in formamide bleaching solution for 30 min to 1 hr under bright light.

  4. Transplanted S2F1L3F2 hosts and irradiated intact controls (~5–6 mm in length) were fixed for 90 min. Worms were bleached for 1.5–2 hr in formamide bleaching solution under bright light.

Immunostaining was performed after fluorescent WISH development with rabbit polyclonal antibodies against H3S10p (1:1000; Millipore # 06–570), mouse monoclonal antibodies against Smith Antigen (Y12) (1:200, ThermoFisher Scientific, PIMA190490), or mouse monoclonal antibodies against Smed PIWI-1 (1:1000, a generous gift from J. Rink). H3S10p antibodies were detected using preabsorbed Alexa-conjugated secondary antibodies (1:1000, Abcam, ab150086, ab150069, ab150071), while anti-Y12 and anti-PIWI staining was visualized with tyramide development using Goat anti-mouse IgG F(ab')2 HRP (1:1000, Jackson Immunoresearch #115-036-072). Nuclear staining was performed with DAPI (1:5000, 1 mg/mL stock solution, ThermoFisher Scientific, D1306) or with Sytox Green (1:5000, 5 mM stock solution, ThermoFisher Scientific, S7020).

S2-–S8 colorimetric and fluorescent WISH samples that were to be imaged using light sheet microscopy were mounted as described in the Microscopy section, whereas others were mounted in 80% glycerol supplemented with 2.5% DABCO. C4 and sexual adult samples were mounted in Scale A2 mounting media (Hama et al., 2011).

Histology

Embryos (S2–S8) were fixed for 4 hr at room temperature in 4% formaldehyde in 1x PBS, followed by 3 × 10 min washes in 1x PBS and gradual dehydration in 30%, 50%, 70%, 80%, 95% and 100% ethanol. The samples were soaked for 30 min in 5% glycerol diluted in 100% ethanol, cleared in xylene for 10 min, then soaked in two changes of Clear-rite 3 (Richard-Allan Scientific) for a total of 25 min. Paraffin infiltration proceeded with 2 × 45 min incubations, and embedded embryos underwent serial sectioning (5 µm thickness). Paraffin was removed prior to staining by heating slides at 60°C for 20 min, then performing 3 × 2 min washes in xylene, 3 × 1 min washes in 100% ethanol, 3 × 1 min washes in 80% ethanol before rinsing in tap water. Hematoxylin and eosin staining was performed using the ST Infinity H and E Staining System (Leica Biosystems) in a Leica Autostainer. Slides were incubated for 30 s in Hemalast, then for 2 min in hematoxylin, and were rinsed for 2 min in tap water. Next, slides were incubated for 45 s in differentiator and for 1 min in bluing agent, with each step followed by a 1 min tap water rinse and a 1 min incubation in 80% ethanol. Slides were stained with eosin for 30 s, dehydrated 3 × 1 min in 100% ethanol and cleared in 3 × 1 min incubations in xylene.

Microscopy

A Leica M205 FA stereomicroscope was used to capture images of live animals and colorimetric WISH samples. A Leica DM600B upright microscope was used to capture images of histological sections. A Zeiss LSM-510-VIS confocal and a customized light sheet microscope were used to capture Z-stacks for fluorescent WISH samples.

Fixed, stained Smed embryos were mounted in 1% low melt agarose in 1x PBS along with fluorescent conjugated beads required for image registration and reconstruction (FluoSpheres Polystyrene Microspheres, 1.0 µm, red fluorescent [580/605], Invitrogen/Molecular Probes, F13083; FluoSpheres Carboxylate Modified Microspheres, 0.1 µm, yellow-green fluorescent [505/515], Invitrogen/Molecular Probes, F8803). 1 µM fluorescent bead stock solutions were diluted 1:10,000–1: 360,000, depending on the size of the embryo and the magnification of the detection objective used. Samples were placed in an imaging chamber within a Single Plane Illumination Microscopy (SPIM) system described in Nakajima et al. (2013). S3–S5 embryos were imaged using either a 10x Plan Apochromat or a 5x Plan NeoFluar objective. Z-stacks were taken every 45˚ around the surface of the samples using a rotating stage, producing eight stacks of images per embryo. Multiview data sets were reconstructed using Fiji SPIM plugins for data registration and fusion (Preibisch et al., 2010). Reconstructed data sets were viewed in the Imaris software package, where they were cropped and masked to remove beads. Cell positions and the embryonic pharynx were marked manually using the 3D Spot Finder function, and the three-dimensional coordinates for marked cells were exported into excel for analysis of cell positions. Colocalization was determined manually on S3–S4 whole embryos, or on crop3D sections (100 µm X 200 µm X 100 µm) of S5 embryos.

Cell position analysis on reconstructed SPIM images

Three-dimensional coordinates for piwi-1+ cells or mitotic cells (piwi-1+, H3S10p+), the embryonic pharynx, and the embryo center were exported from SPIM reconstructions of S3 and S4 embryos. Spot positions collected in IMARIS were shifted and rotated in MATLAB to a coordinate system where the embryonic center was at the origin and the embryonic pharynx on the z-axis. Relative theta distribution likelihoods of the form (1-exp(-θ/θ’))*sin(θ) were calculated by assuming a θ’ and calculating the likelihood that such a distribution would produce the observed data by multiplying together the individual probabilities. The uncertainty in the best fit θ’’ was found by simulating several datasets, each with a number of cells equal to those observed in the actual data and having a dampening term of θ’’. For each dataset, the most likely θ’ was found, the standard deviation of which was taken to be the error in θ’’.

Nanostring nCounter digital expression analysis

Total RNA samples from single embryos and adults (S2–S8, Y, C4 and SX; four biological replicates per sample) were prepared as described for single-animal RNA-Seq. 100 ng total RNA per sample was assayed on the Nanostring nCounter platform (Geiss et al., 2008) using a custom-made probe set. Reporter and capture probe sequences can be found in Figure 5—source data 2. Housekeeping control genes were reported in Wenemoser et al. (2012). Nanostring data was normalized using the NanoStringNorm library from Bioconductor (Waggott et al., 2012), using the sum of the positive controls and the sum of the housekeeping genes as independent normalization factors, with the mean of the negative controls used to estimate background. Raw and normalized nanostring data can be found in Figure 5—source data 2.

Bulk cell transplantation

Host animals (5–7 mm in length, ≥7 day starved) were selected from the clonally derived Smed sexual strain S2F1L3F2 (Wagner et al., 2011) and were cultured in 1x Montjuic water supplemented with 100 µg/mL gentamicin sulfate. The host neoblast population was ablated by exposure to 6,000 Rads on a GammaCell 40 Exactor irradiator; cohorts of unirradiated animals were reserved to verify complete elimination of the neoblast population by WISH with riboprobes against piwi-1.

Whole-embryo cell suspensions were created for S4, S5, S6, S7 or S8 by mechanically disrupting embryos in chilled, freshly made 1x Holfreter’s solution +5% heat-inactivated fetal bovine serum (Sigma Aldrich, F4135) via repeated pipetting. Ten embryos were disrupted per cell suspension for S5, S6, S7 and S8, whereas 20–35 embryos were disrupted per S4 cell suspension. S4, S5 and S6 embryos were typically ‘eviscerated’ prior to pipetting by poking with an insect pin and gently squeezing out ingested yolk from the gut. Cell suspension volumes were adjusted to 1.0 mL before samples were filtered through 20 µm (all stages) and 10 µm (S5–S8) cell strainers (Partec CellTrics, Sysmex, 04-0042-2315 [20 µm], 04-0042-2314 [10 µm]) into low-retention microfuge tubes. Cells were pelleted by centrifugation at 310 rcf for 5 min, were resuspended in a final volume of ~10 µL (S5–S8) or ~5 µl (S4), and were kept on ice during transplantation.

Embryo cell suspensions were loaded by mouth pipetting into borosilicate glass needles (Sutter Instrument Co., #B100-75-15) pulled using a flaming/brown micropipette puller (Sutter Instrument Co., Model P-97) and were injected using an Eppendorf FemtoJet at 1.0–1.5 psi, as described by Wagner et al. (2011). Hosts were immobilized on a cold peltier plate, ventral side up, and cells were injected into the tail stripe (i.e., the medial, post-pharyngeal parenchymal space between the two posterior branches of the intestine). Hosts were injected at 1 day post-irradiation (dpi) for rescue experiments and at 3 dpi for short-term experiments. Transplanted animals and uninjected, 6,000-Rad-irradiated hosts for rescue experiments were maintained individually in 3 cm petri dishes at 20°C in the dark, with water exchanges and visual inspection of animals performed every 2–3 days. Transplanted animals slated for fixation were reared in 10 cm petri dishes, with 10 or fewer animals per dish, with water exchanges every 2–3 days.

Online data repository

Original data underlying this manuscript can be accessed from the Stowers Original Data Repository at http://www.stowers.org/research/publications/libpb-1086

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Decision letter

  1. Utpal Banerjee
    Reviewing Editor; University of California, Los Angeles, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Embryonic origin of adult stem cells required for tissue homeostasis and regeneration" for consideration by eLife. Your article has been reviewed by four peer reviewers, and the evaluation has been overseen by Utpal Banerjee as Reviewing Editor and Marianne Bronner as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Diana Laird (Reviewer #2); Peter W Reddien (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. All reviewers agree on the importance of the work and the changes asked for can be addressed by making changes to the manuscript.

Summary:

In this manuscript, Davies and colleagues investigate the developmental origins of adult pluripotent stem cells (neoblasts) in the planarian Schmidtea mediterranea. The authors provide a comprehensive characterization of S. mediterranea embryogenesis by meticulously cataloguing (with corresponding gene expression data and expression mapping) the different stages of embryonic development. This groundwork allowed the authors to ask when neoblasts originate during embryogenesis. The authors demonstrate that neoblasts arise from anarchic, cycling piwi-1+ cells, which generate all temporary and definitive tissues/organs during embryonic development. Interestingly, early embryonic piwi-1+ cells have a distinct molecular signature from neoblasts. Unlike neoblasts, which express transcription factors (TFs) important for development, early embryonic piwi-1+ cells express "early embryo enriched" (EEE) transcripts. Later in embryogenesis when organogenesis initiates, EEE transcripts are downregulated while various developmental TFs are upregulated. Strikingly, in an elegant set of transplantation experiments, the authors demonstrate that piwi-1+ cells are functionally different at different embryonic stages. Similar to transplanted adult neoblasts, transplanted piwi-1+ cells from embryos undergoing organogenesis can rescue lethally irradiated planarians that are devoid of neoblasts, whereas, early EEEhi/TFlo piwi-1+ cells cannot.

Essential revisions:

Usually for minor revisions, we make a summary of the reviewers' comments, but in this case, the majority of questions raised are regarding lack of clarity in the manuscript and the complexity of data presentation. So we are including the detailed comments below. All of these can be addressed by rewriting the manuscript making it more accessible to readers. Also, below I paste an exchange between the reviewers during the post review session:

" I did want to address one point raised by two other reviewers, which I think reflects the way in which the data presentation could be improved. Both raised concerns about the transplant experiments that were easy to miss because some of the key results were buried in the supplement.

The "no transplant" control is shown in Figure 7F. As expected (and shown previously) no planarians survive irradiation. Figure 7—figure supplement 2 shows piwi-1+ cell counts. In panel B the authors show the percentage of transplants with piwi-1+ cells at 1 hpt (directly after transplantation), which indicates that "the cell injection technique was robust and reliable". Panel C shows the number of piwi-1+ cells per transplant at 1 hpt. For example, in Figure 7—figure supplement 2, one can see that comparable numbers of piwi-1+ cells were transplanted from S5 and S6 embryos (1 hpt) but at 5 dpt, very few to zero piwi-1+ cells remain from S5 donors compared to S6 (panel E). Furthermore, irradiated hosts transplanted with S5 piwi-1+ donor cells are not rescued (Figure 7F). The authors mention these data in the text (subsection “Embryos undergoing organogenesis contain cells with cNeoblast activity”), but it might be helpful to the reader if these data were in the main figure and not buried in the supplement. Furthermore, some of the panels can be merged (for example the 1 hpt and 5 dpt piwi-1+ cell transplants) so that it is easier for the reader to compare the data. "

As you can see, the reviewers, as much as they admired the science, were confused by the presentation. So, please make the paper a bit more lucid for the readership of eLife.

Reviewer #1

This impressive body of work will be a valuable resource to planarian biologists. To make the manuscript accessible to a broader audience and more manageable for readers in general, here are some editing suggestions that should be addressed before publication:

1) To appeal to the broader eLife audience, the authors should provide more description/context in the main text to help non-experts understand the results presented. This includes data the authors deem not essential for the narrative of the paper and is found in Supplementary Online Material instead of the main text. For example, sphere formation (subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”) is not defined in the main text, but is described in the SOM as beginning during Stage 2. Furthermore, the distinction between temporary and definitive embryonic organs may not be entirely clear to non-expert readers. The authors do a better job describing this uncommon developmental phenomenon in the SOM (e.g., "The embryonic pharynx degenerates during S6, as the definitive pharynx primordium develops beneath it. Embryonic pharynx markers are no longer detectable by S7").

2) Throughout the manuscript, the authors are urged to present figures in the order they are referred to in the text. Looking up figures (and multiple Supplements) that are "out of order" is cumbersome, especially when considering the sheer volume of data the reader must go through. For example, Figure 1—figure supplement 12 is referred to twice in the text (Results section) before any of the other Figure 1 supplements. Figure 1—figure supplement 12 should be designated as Figure 1—figure supplement 1 (and all other Figure 1 supplements renumbered accordingly thereafter). Also, having two sets of differently named supplemental data (Figure X—figure supplement Y, which is per the journal's style, and Supplementary Online Figure Z, which appears not to be) makes navigating supplementary material unnecessarily difficult for the reader.

3) Introduction section: Remove the extensive discussion of results from the Introduction. This will also help shorten the overall length of the paper.

Reviewer #2

Davies et all provide the first comprehensive description of Smed embryos at a molecular and anatomic level. This is not trivial given that S. med broods its embryos and the authors should be congratulated for this Herculean task. They specifically ask whether neoblasts that have been well studied in the adult planarian exist during embryogenesis and conclude that a cycling piwi1+ precursor population gives rise to organs as well as neoblasts, but is distinct on a molecular level as well as functionally in transplantation assays. Although important for the field, the following issues should be addressed:

Comments:

1) Overall, the accessibility and readability for a generalist journal as this one should be increased. This can be accomplished by presuming less prior knowledge about planaria in the Results section and structuring the writing with summary statements at the end of each section in the Results. It might suggest that authors provide a succinct table with morphological criteria for embryo staging as well as key genes expression changes, accompanied with cartoons.

2) Would it be possible to resolve S2 and S3 by growing the animals at a lower temperature?

3) ISH studies need sense controls. Specifically, in later stages (S6-8), it is unclear whether some of the signal for piwi1 in Figure 1 could result from probe trapping or non-specific staining in the testes and ovary ducts.

4) Subsection “Progenitors for definitive lineages arise within piwi-1+ blastomeres as organogenesis commences” refers to data presumably not shown demonstrating that ~4% of cells expressing PIWI1 by immunostaining at S5 do not harbor piwi1 transcript, and suggest that the protein perdures after transcript degradation upon differentiation. Can the authors please clarify whether all piwi-1+ cells expressed PIWI1 protein in the embryos?

5) My main concern with the authors conclusions lies in the transplant studies. Can the authors please estimate how many cells were transplanted, or how many embryo equivalents. Figure 7B should show a no transplant control (which presumably will not contribute) as well as cNeoblast control from adult (which will presumably contribute more robustly than the embryos). The authors need to demonstrate that piwi+ cells and structures rescued following irradiation are donor-derived, as this will dismiss the possibility that the blastomere graft rejuvenates host cells; this can be accomplished genetically by the phenotype of regenerated structures as well as by using a marker such as PKH26 of the donor cells.

Reviewer #3

Davies et al. present a molecular, cellular, and morphological study of planarian (S. mediterranea) embryogenesis. They report a molecular resource of stage-enriched genes useful for staging embryos and for studying gene function in embryogenesis, including the discovery of embryo-enriched planarian gene expression. They show that piwi+ cells are present throughout embryogenesis, but that lineages for adult tissues only develop within this piwi+ population at a particular stage (S5). Interestingly, in transplant experiments, clonogenic (colony-forming) neoblast activity also develops at a particular stage, approximately when lineages appear. The manuscript is an impressive body of work and well written. This paper will be an important landmark in the study of embryogenesis and its comparison to regeneration in planarians.

Abstract:

"Neoblast subpopulations established during embryogenesis persist into adulthood"

Subpopulations may not persist as subpopulations, rather continuously generated. Since the renewal capacity of subpopulations is unknown, different wording is suggested.

Results section:

"seven morphologically distinct stages (S), S2-S8" were selected for sequencing. Panels D-F contain embryos that look superficially similar; thus it would be helpful for the authors to further describe at this point in the text the criteria for stage separation in the sequencing experiment (I assume time of development was also a factor?).

"pairwise comparisons of adjacent time points"

These aren't really "time points", since stages overlap in timing. Different wording is suggested.

Assessment of most statements of data in the text is difficult, because each one requires a goose chase through excel tables, main figures, figure supplements, and online-only figures. Some items called out as figure supplements are not figures but tables with titles different than their figure supplement title. Take subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”, for example; this single sentence refers to 4 panels in 2 main figures, 4 Figure supplements, and 3 supplementary online figures. I tried to find Figure 5—figure supplement 1 to look at the markers for all known and restricted embryonic cell populations and eventually gave up…

Paragraph four in subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”. The paragraph could use a concluding sentence to clarify message.

Similarly, the point of "…sample sizes, the optimal θ' was calculated to be 0.58 +/- 0.33,and was 50-fold more likely to explain the observed trend than a simple normal distribution." is not clear.

Last paragraph of the subsection: restriction of cell cycle activity to piwi+ cells in adults should reference the piwi-1 2005 paper and could also reference Wagner 2011, which had further analysis of this marker for all dividing cells.

Figure 4—figure supplement 2 Y12 data is interesting, but only S4 staining is apparent. If it is not present in piwi+ cells at earlier stages this merits display. The chromatoid body presence could also be considered for a main figure.

Subsection “Early embryo enriched transcripts expressed throughout piwi-1+ blastomeres are downregulated as organogenesis begins”. Naming clusters "6" "5" can be deciphered from examination from the figure, but is not immediately clear what these are when reading the text. i.e., individual of the multiple clusters of S2-S4-enriched txts that showed downregulation at S5 and thereafter.

The third paragraph in this section presents analysis of expression of the interesting EEE genes. However, the main messages/synthesis is hard to decipher because of the distracting links out to disparate files/figure locations. The narrative/text feels beholden to a set of files and figures as opposed to the other way around.

More discussion of the EEE genes that are expressed broadly in the embryo would be of interest. What types of genes were discovered?

Reviewer #4

This manuscript contains a large amount of information, much of it contained in supplementary materials. It is an important piece of work, because (1) it addresses the expression of genes on a genome wide scale during the development of a planarian flatworm; (2) it addresses the origin and development of neoblasts.

That being said, in its current form, the manuscript is very difficult to digest. The writing is long (section 5 of the Results alone has 6.5 pages!) and "windy: (see example given for section 2 below), and there are some fundamental questions that are either not addressed, or the answers are "hidden" in the long narrative.

Specific comments:

The Introduction is long and detailed on many aspects of neoblast biology, but at the same time unclear and lacking in other aspects. Generally, I am often not sure whether the authors talk about facts known from previous studies, or are already highlighting the findings to be reported in this paper.

Suggestion: shorten the intro by at least 50%. After (in a briefer format than now) summarizing known functions and types of neoblasts in adults/regeneration, summarize literature (as far as it exists) regarding

-embryogenesis in planarians (see below; the statements about embryogenesis that are now found in section 2 of the Results should be in the Introduction)

-specifically, what is said in the literature where/when do neoblasts first appear in embryos (there will probably not be a lot of published information; one example is Ramachandra et al., 1999, on embryonic BrdU -positive presumed neoblasts in an Acoel).

-proliferation, migration, differentiation of neoblasts during embryonic period.

Results

Section 1: surveys molecular staging; stages themselves described in supplementary table and figures. This is a useful and important work of reference for the planarian developmental field.

Please highlight in the text:

-what in terms of morphogenesis and organogenesis is novel compared to the previous embryological study of Schmidtea polychroa (Cardona et al)

-how do stages utilized here (1-8) differ from the 8 previously introduced ones

-what does molecular profiling add to defining stages (clearer temporal distinction?..)

Section 2 – “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”

The section describes the expression pattern of piwi, and concludes that the gene is restricted to undifferentiated cells. The text is very difficult to read. Different types of information are intermingled. For example, paragraph two summarizes a result known from the literature. Talks about general expression patterns of non-specified transcripts, documented only in the supplementary material, in differentiating cells. later in the paragraph you speak to the point: piwi is expressed in undifferentiated cells in the bodywall at the S3 stage. But how does the pan-nuclear marker EF1a-like 1 show that, since this marker appears to be expressed in all cells (including the piwi-positive ones)? The final sentences again talk about findings from the previous literature.

I suggest to remove all references to the previous literature from this (Results) section; put them in the Introduction, and in the Results focus on where exactly piwi is expressed. Specifically:

Paragraphs 1-2

-DAPI shows large number of labeled dots: are these all nuclei of blastomeres (Figure 2A, C)?

-if so, only a small subset of blastomeres expresses piwi (2A): how would one characterize distribution? Clustered, around embryonic pharynx?

-Paragraph 2: sounds like piwi is downregulated in many cells. What is shown (Figure 2A) is that from beginning, piwi is expressed in only few blastomeres (if DAPI really reflects all nuclei). Is there an earlier stage with many more piwi+ cells?

-the pattern and dynamics of piwi in undifferentiated cells ("blastomeres") remains a bit vague. Is the pattern the following: stage 1, before embryonic transient cells differentiate: all cells express piwistage 2/3: as embryonic cells differentiate, they lose piwi; ALL other cells retain it later stages: as adult cells become postmitotic, they lose piwi; ALL other cells retain it

I don't think this is what happens. But the authors do not make it clear in text and figures what exactly happens. I assume that only a small subset of undifferentiated cells express piwi. I understand that these are distributed. This information needs to be settled; it is one of the crucial pieces of data required in subsequent sections (see comments below).

Section 3 reports that genes sets known to be associated with adult neoblasts are also expressed in the embryo. This is an important finding. The main question is:

-are these genes, in the embryo (e.g., S4, S5, S6), restricted to piwi+ cells?

-are they restricted to ALL undifferentiated cells (assuming that not all undifferentiated cells are piwi+; see my questions/comments above)

-are they also expressed in cells that stop dividing and differentiate?

I was not able to find a clear answer to these questions in the text.

Section 4 Early embryonic genes (EEE)

I don't understand the message of this section. One reason for confusion arises right at the beginning:

- in the opening lines the authors talk about "S2-S4 enriched transcripts expressed throughout the piwi blastomere population".

- then the authors introduce the 1048 EEE genes: are these the same as above (i.e., they are expressed in the piwi blastomere population)?

- because, if they are: why do piwi (undifferentiated) blastomeres and differentiated (transient embryonic cells) express the same set of genes? I thought from the previous sections (and on general grounds) that as cells become postmitotic and differentiate, they switch on different genetic programs.

Section 5

It is shown that gene sets associated with specific differentiated cell types are upregulated around stage 5, which is important, but expected, since it confirms the observation that cells differentiate (structurally) at this stage.

The main claim of the heading of this section (that progenitors of specific cell types arise WIHIN the piwi-positive population) needs to be demonstrated, and/or described more clearly (see questions to section 2 above)

Section 6

Cell suspensions derived from embryos at different stages (S5-S8) were injected into irradiated adults, and were able to rescue them. This is also an important experiment. A puzzling finding was that cells from S5 embryos were unable to rescue. Is it possible that S5 embryos did not have a high enough density of piwi+ cells to do the job? Is it possible to get a value for the number of piwi+ cells present in an aliquot of the cell suspension from different stage embryos?

https://doi.org/10.7554/eLife.21052.063

Author response

Reviewer #1

This impressive body of work will be a valuable resource to planarian biologists. To make the manuscript accessible to a broader audience and more manageable for readers in general, here are some editing suggestions that should be addressed before publication:

1) To appeal to the broader eLife audience, the authors should provide more description/context in the main text to help non-experts understand the results presented. This includes data the authors deem not essential for the narrative of the paper and is found in Supplementary Online Material instead of the main text. For example, sphere formation (subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”) is not defined in the main text, but is described in the SOM as beginning during Stage 2. Furthermore, the distinction between temporary and definitive embryonic organs may not be entirely clear to non-expert readers. The authors do a better job describing this uncommon developmental phenomenon in the SOM (e.g., "The embryonic pharynx degenerates during S6, as the definitive pharynx primordium develops beneath it. Embryonic pharynx markers are no longer detectable by S7").

The manuscript was revised extensively, with the goal of making our story accessible to a broad readership. Care has been taken to eliminate unnecessary specialist vocabulary and to clearly define terms as they are introduced. We shortened the Introduction, eliminating the extensive summary of our results. In addition, we emphasized the foundational resources, key results and conceptual breakthroughs stemming from our work. We added two new paragraphs to the start of the Results section, intended to give readers a crash course in freshwater flatworm embryology. We define necessary terminology and introduce key concepts, like direct development, ectolecithal embryos, dispersed cleavage, temporary embryonic tissues and definitive tissues. We reorganized Figures 2 and 7 (now Figures 2 and 79, respectively) and rewrote the associated text and Figure Legends; we also rewrote much of the text accompanying Figure 5. To aid readers, we reorganized all of the supplemental data so that items are noted chronologically in the text. Supplementary files were broken down into supplemental figures, source data and rich media files. We hope this encourages readers to delve into the wealth of data provided with our manuscript. In addition, we generated an online searchable resource (https://planosphere.stowers.org) that houses the Molecular Staging Series and Molecular Fate Mapping Atlas. This resource, hosted by the Stowers Institute, will be freely available upon publication and will reference our published manuscript.

2) Throughout the manuscript, the authors are urged to present figures in the order they are referred to in the text. Looking up figures (and multiple Supplements) that are "out of order" is cumbersome, especially when considering the sheer volume of data the reader must go through. For example, Figure 1—figure supplement 12 is referred to twice in the text (Results section) before any of the other Figure 1 supplements. Figure 1—figure supplement 12 should be designated as Figure 1—figure supplement 1 (and all other Figure 1 supplements renumbered accordingly thereafter). Also, having two sets of differently named supplemental data (Figure X —figure supplement Y, which is per the journal's style, and Supplementary Online Figure Z, which appears not to be) makes navigating supplementary material unnecessarily difficult for the reader.

To make it easier for readers to navigate through the extensive amount of supplemental data associated with our manuscript, we reorganized and renamed the supplemental data files as suggested by reviewer #1. Figure supplement, source data and rich media files were renamed to reflect the order they are referred to in the text. Large excel tables are now listed as “source data” files; SPIM movies are listed as “rich media” files. In addition, the Molecular Fate Mapping Atlas, formerly the Supplementary Online Material and Supplementary Online Figures 19, are now labeled as Figure 1—source data 9 and Figure 1—figure supplements 1119. This was done to consolidate the supplementary data, and to eliminate the confusion created by having “figure supplements” and “Supplementary Online Material.”

3) Introduction section: Remove the extensive discussion of results from the Introduction. This will also help shorten the overall length of the paper.

The Introduction was shortened and reorganized, in accordance with suggestions from reviewers #1 and #4. The lines, which contained a detailed summary of our results, were removed from the Introduction.

Reviewer #2

[…]Comments:

1) Overall, the accessibility and readability for a generalist journal as this one should be increased. This can be accomplished by presuming less prior knowledge about planaria in the Results section and structuring the writing with summary statements at the end of each section in the Results. It might suggest that authors provide a succinct table with morphological criteria for embryo staging as well as key genes expression changes, accompanied with cartoons.

We revised the manuscript to make it more accessible to a general audience, and detailed the changes made in this regard in our response to reviewer #1. To facilitate use of this staging system by others, http://planosphere.stowers.org provides cartoons, time (days post-egg capsule deposition), gene expression data (RNA-Seq and WISH), and written descriptions of embryo morphology for each of the stages queried.

2) Would it be possible to resolve S2 and S3 by growing the animals at a lower temperature?

Early S2 embryos undergoing sphere formation are quite fragile and are difficult to detect in live, unstained samples; it’s doubtful that simply lowering the temperature would make these samples easier to find. We are interested in performing gene expression profiling on oocytes, zygotes and early S2 embryos in the future. Laser capture microdissection, performed on fixed tissue stained with a nuclear dye, may be the best route to obtaining tissue for S2 and earlier stages. It is possible that injection of vital dye into oocytes, and/or EdU incorporation into oocytes or early blastomeres, would enable us to identify tissue for RNA-Seq in live samples, and to do lineage tracing studies. We are interested in identifying maternal RNAs contributed to oocytes, the onset of zygotic transcription – which is likely quite early (i.e., before we started our RNA-Seq time course), and determining when the first asymmetries in gene expression occur among the blastomeres. These experiments and are future directions for us.

3) ISH studies need sense controls. Specifically, in later stages (S6-8), it is unclear whether some of the signal for piwi1 in Figure 1 could result from probe trapping or non-specific staining in the testes and ovary ducts.

Hematoxylin and eosin staining is shown in Figure 1, not in situ hybridization data. piwi-1 expression, shown first in Figure 2, always shows specific expression in parenchymal cells. It is unlikely that the piwi-1+ signal is nonspecifically trapped in the reproductive system during S6-S8, since the testes, ovaries and somatic gonadal tissue are absent from S8 newborn hatchlings and arise weeks later during juvenile development. Incidentally, we detect nanos expression by WISH during S6-S8, in increasing numbers of cells with a noted dorsolateral bias and in the developing eyes (S7-S8). Our suspicion is that the dorsolaterally positioned cells are likely piwi-1+, nanos+ primordial germ cells; we have not followed up on this point yet, and would need to costain with additional PGC markers as well. Consistent with our WISH results, nanos expression was undetectable by RNA-Seq until S5, and was lowly expressed in S5-S8 embryos.

4) Subsection “Progenitors for definitive lineages arise within piwi-1+ blastomeres as organogenesis commences” refers to data presumably not shown demonstrating that ~4% of cells expressing PIWI1 by immunostaining at S5 do not harbor piwi1 transcript, and suggest that the protein perdures after transcript degradation upon differentiation. Can the authors please clarify whether all piwi-1+ cells expressed PIWI1 protein in the embryos?

Yes, piwi-1+ cells also express PIWI-1 protein (99 +/- 0.4% piwi-1+ cells were double positive for PIWI-1 protein, n= 4,152 piwi-1+ cells scored, n= 4 S5 embryos). We amended the text to report this result and created a new figure, Figure 6 —figure supplement 1, containing representative images. The quantification you requested is present in the Figure 6 —figure supplement 1 legend. The revised main manuscript text is included in our comments to reviewer #1.

5) My main concern with the authors conclusions lies in the transplant studies. Can the authors please estimate how many cells were transplanted, or how many embryo equivalents. Figure 7B should show a no transplant control (which presumably will not contribute) as well as cNeoblast control from adult (which will presumably contribute more robustly than the embryos). The authors need to demonstrate that piwi+ cells and structures rescued following irradiation are donor-derived, as this will dismiss the possibility that the blastomere graft rejuvenates host cells; this can be accomplished genetically by the phenotype of regenerated structures as well as by using a marker such as PKH26 of the donor cells.

We reorganized the presentation of the cell transplantation data and revised the associated text extensively to bring out some of the key points formerly house in the supplemental material. The transplantation experiments are now shown in Figures 7 and 8. In Figure 7C, we show quantification for the number of piwi-1+ cells per host at 1 hour and 5 days post-transplant. We show that comparable numbers of S5, S6, S7 and S8 cells are injected per host; an average injection generally delivers between 50-100 embryonic piwi-1+ cells into the tail. We detail how the embryonic cell suspensions were prepared in the Materials and methods. We are able to get far fewer S4 embryonic cells injected per host, which was expected since far fewer piwi-1+ blastomeres are present during S4 versus later stages. The S4 results represent our best efforts to concentrate as many embryo equivalents as possible while minimizing volume of the cell suspension.

We provide scoring data for “no transplant control” data in the text for both sets of transplantation experiments.

We did not run transplantation experiments using adult neoblast enriched fractions as a positive control. We concede that the side by side comparison would be nice to see, though not necessary. There were two reasons for this: 1) We strongly suspected that S8 embryonic cells would rescue since animals at this stage regenerate following amputation (our unpublished observation). The regeneration capacity, coupled with the distribution of piwi-1+ cells in S8 embryos, suggested strongly to us that upon hatching juvenile worms possess neoblasts. Had we not achieved reproducible rescue results with S8 embryonic donor cells, we certainly would have pursued adult neoblast injection experiments. 2) The published cell transplantation papers used C4 asexual adult animals as donors. Differences in transplanted cell behavior using C4 animals and embryos as donors could arguably be attributed to differences in biotype. While sexually mature adult donor tissue has been shown to rescue in plug graft experiments, we presently don’t have a cell sorting protocol for neoblast isolation from sexually mature adults. Our cell sorting protocols are based on DNA content, as well as cell size and morphology; we would expect our neoblast enriched fraction to be contaminated with dividing germline stem cells and transit amplifying germ cells. This would not be insignificant, given that adults possess roughly 100 testes.

Lethally irradiated adults are capable of wound healing, mount an acute transcriptional response to wounding, and transiently reorganize expression domains of key polarity genes along the A-P axis of the fragment. However, these animals never generate a blastema or regenerate new tissue. None of the lethally irradiated, uninjected sexual adult hosts survived long-term, nor did they mount a regenerative response (e.g., head blastema formation) prior to lysing. The most parsimonious explanation for our results is that engrafted, proliferating, donor-derived piwi-1+ cells contributed differentiating progeny that subsequently mounted the observed regenerative response in rescue animals. We have seen production of post-mitotic, definitive epidermal progenitors in S5 donor cell transplants (NB.21.11e+ and AGAT-1+ cells) at 10 dpt (13 dpi); these definitive epidermal progenitors were present in the tail parenchyma in the neighborhood of piwi-1+ cells. Moreover, NB.21.11e+ and AGAT-1+ cells are irreversibly eliminated by lethal irradiation within 5 dpi, suggesting that the cells present in S5 transplanted hosts at 10 dpt (13 dpi) differentiated from donor-derived piwi-1+ cells. Additional transplant experiments would be required to examine production of differentiating progeny for other lineages. SNPs could be assessed in donor and transplanted hosts after long-term reconstitution; however, this would require using genetically defined lines for embryonic donor cells, which we did not do for these experiments. We agree that using vital membrane dyes to track donor cells would be a good way of assessing donor versus host contribution following transplantation, and will pursue this suggestion in the future.

Reviewer #3

[…]Abstract:

"Neoblast subpopulations established during embryogenesis persist into adulthood"

Subpopulations may not persist as subpopulations, rather continuously generated. Since the renewal capacity of subpopulations is unknown, different wording is suggested.

At present we cannot distinguish between the two possibilities mentioned by reviewer #3: self-renewal of committed progenitors within the neoblast population (stable, or persistent subpopulations) or the continual re-emergence of committed progenitors from pluripotent stem cell(s) throughout the lifetime of the animal. The concluding sentences of the Abstract now read:

“Neoblast lineages arise as organogenesis begins and are required for construction of all major organ systems during embryogenesis. These subpopulations are continuously generated during adulthood, where they act as agents of tissue homeostasis and regeneration.”

We also inserted as sentence in to the final paragraph of the Results text associated with Figure 6, which addresses this point:

“At present, we cannot distinguish whether lineages perpetually re-emerge due to asymmetric division of pluripotent stem cells, or whether progenitor populations established during embryogenesis are maintained by self-renewal.”

Results section:

"seven morphologically distinct stages (S), S2-S8" were selected for sequencing. Panels D-F contain embryos that look superficially similar; thus it would be helpful for the authors to further describe at this point in the text the criteria for stage separation in the sequencing experiment (I assume time of development was also a factor?).

Yes, you are correct that a combination of chronology and morphology were used to select samples and define the stages. As stated in the Materials and methods, we rear our animals at 20˚C and collect egg capsules daily. Each collection could theoretically be composed of 0-24 hour old embryos (hours indicating time post egg capsule deposition), but we designate the time of collection as 1 day post-egg capsule deposition and age them out accordingly. The time windows for collection are indicated in Figure 1B, and the RNA-Seq replicates (Figure 1—source data 1) indicate the time point(s) for each of the stages queried.

The main text now reads:

Smed embryos gestate for approximately two weeks at 20˚C prior to hatching. We generated total RNA replicates from single Smed embryos for seven chronologically and/or morphologically distinct stages (S), S2-S8 (Figure 1B-I); S1 samples (zygotes, Figure 1—figure supplement 1) were not queried by RNA-Sequencing.”

"pairwise comparisons of adjacent time points"

These aren't really "time points", since stages overlap in timing. Different wording is suggested.

The wording “time points” was changed to “stages.”

Assessment of most statements of data in the text is difficult, because each one requires a goose chase through excel tables, main figures, figure supplements, and online-only figures. Some items called out as figure supplements are not figures but tables with titles different than their figure supplement title. Take subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”, for example; this single sentence refers to 4 panels in 2 main figures, 4 Figure supplements, and 3 supplementary online figures. I tried to find Figure 5—figure supplement 1 to look at the markers for all known and restricted embryonic cell populations and eventually gave up…

We reorganized and renamed the supplemental data to make it easier for readers to navigate and locate items of interest. Supplemental items are now organized chronologically, as reviewer #1 suggested. All excel data files are now listed as “Figure X—source data X” files. Supplemental figures are listed as “Figure X —figure supplement X.” SPIM movies are now listed as “Rich Media Files.” In addition, the Molecular Fate Mapping Atlas files are now couched under the Figure 1 subheading, along with the RNA-Seq data. This eliminates a second class of supplemental figure names, which other reviewers also found burdensome. Finally, https://planosphere.stowers.org, an online searchable resource, is another venue for examining much of the supplementary data accompanying our manuscript.

Paragraph four in subsection “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”. The paragraph could use a concluding sentence to clarify message.

Similarly, the point of "…sample sizes, the optimal θ' was calculated to be 0.58 +/- 0.33,and was 50-fold more likely to explain the observed trend than a simple normal distribution." is not clear.

Reviewer #1 brought up the same points, and we revised the main text and figure legends to minimize the technical detail and bring forth the main conclusions. The revised text is included earlier in the rebuttal, under the comments to reviewer #1.

Last paragraph of the subsection: restriction of cell cycle activity to piwi+ cells in adults should reference the piwi-1 2005 paper and could also reference Wagner 2011, which had further analysis of this marker for all dividing cells.

We apologize for this oversight and added both references to this passage.

Figure 4—figure supplement 2Y12 data is interesting, but only S4 staining is apparent. If it is not present in piwi+ cells at earlier stages this merits display. The chromatoid body presence could also be considered for a main figure.

Y12 staining was only performed on S4 and S5 embryos. The manuscript was revised as follows:

“Y12 antibodies, which specifically label chromatoid bodies in adult neoblasts {Rouhana, 2012 #482}, stained piwi-1+ blastomeres during S4-S5 (Figure 4—figure supplement 1G, Rich Media File 10).”

Subsection “Early embryo enriched transcripts expressed throughout piwi-1+ blastomeres are downregulated as organogenesis begins”. Naming clusters "6" "5" can be deciphered from examination from the figure, but is not immediately clear what these are when reading the text. i.e., individual of the multiple clusters of S2-S4-enriched txts that showed downregulation at S5 and thereafter.

We altered this paragraph significantly to clarify what we mean by the term “EEE transcripts.” Some of the details regarding expression trends in clusters 5, 6 and 8 were moved the Figure 5A legend.

The main text now reads:

“Hierarchical clustering of S2-S4 enriched transcripts using scaled RPKM values identified 1048 sequences in clusters 5, 6 and 8 that were downregulated by S5 and remained lowly expressed through S8; these sequences are referred to as early embryo enriched (EEE) transcripts (Figure 5A, Figure 5—source data 1). EEE transcripts were likely expressed in blastomeres and/or temporary embryonic tissues, since 98% of the sequences had average expression values at least 5-fold greater in S2 embryos versus Y (Figure 5—source data 1). Most EEE transcripts were expressed at low levels in intact adults regardless of biotype: average RPKM values less than 1.0 were recorded for 65% and 59% of the EEE transcripts in C4 or SX, respectively (Figure 5—source data 1).”

The Figure 5A legend now reads:

Hierarchical clustering of S2-S4 enriched transcripts (n=1756) using scaled RPKM data. Left: Heat map. Y: yolk. Colored bars (left) denote clusters 5, 6 and 8 containing early embryo enriched (EEE) transcripts. Cluster 5 sequences (blue, n=413) were expressed at roughly equivalent levels during S2 and S3, with 66% (n=275) transcripts showing 5-fold or greater declines in average expression values between S3 and S5. Cluster 6 sequences (red, n=523) exhibited maximal expression during S2, and average expression levels declined more than 5-fold between S2 and S4 for 81% (n=426) of these transcripts. Cluster 8 sequences (green, n=112) showed peak expression during S4, with 52% (n=60) of the transcripts showing 5-fold or greater declines in average expression values by S5. Right: Normalized expression trends for EEE transcripts in Clusters 5 (blue), 6 (red) and 8 (green) plotted as a function of developmental time. Median 50% of transcripts based on expression maxima are plotted.”

The third paragraph in this section presents analysis of expression of the interesting EEE genes. However, the main messages/synthesis is hard to decipher because of the distracting links out to disparate files/figure locations. The narrative/text feels beholden to a set of files and figures as opposed to the other way around.

We rewrote the section to more clearly articulate our main points. In addition, moved the summary of the WISH expression screen from Figure 5—source data 1 (formerly Figure 5—figure supplement 1) to a separate file, Figure 5—source data 3. The rewritten text is as follows:

“To determine which cell types express EEE transcripts, and to examine spatiotemporal changes in EEE transcript expression, colorimetric WISH was performed on S2-S8 embryos and intact C4 adults (Figure 5—source data 3). Some EEE transcripts were expressed exclusively in differentiated temporary embryonic tissues. VAL-like, MPEG1-like-1, MPEG1-like-2, and netrin-like were solely expressed in the temporary embryonic pharynx until S6, while gelsolin-like and 4XLIM-like were expressed in both the primitive ectoderm and temporary embryonic pharynx during S3-S4 (Figure 5—source data 3, Figure 1 —figure supplements 11D-E, 12C-G). Expression of these EEE transcripts is likely under zygotic control, occurring during or after the cell fate decisions to downregulate piwi-1, exit the cell cycle and differentiate.

Most EEE transcripts queried by WISH (n=15, 75% assayed) were expressed in both undifferentiated blastomeres and temporary embryonic tissue(s) during S3-S4 (Figure 5B-E, Figure 5— source data 3). Some of these transcripts may be maternally deposited, albeit we cannot ascertain the relative contribution(s) of maternal and zygotic expression from our RNA-Sequencing data. Expression of these EEE transcripts diminished greatly by S5, with moderate and lowly expressed EEE transcripts becoming undetectable; specific expression of robustly expressed transcripts sometimes persisted until S6 (Figure 5B-E, Figure 5—source data 3). Consistent with the RNA-Seq and Nanostring nCounter results, EEE transcript expression was not detected by colorimetric WISH in S7, S8 embryos or C4 adults (Figure 5B-E, Figure 5 – source data 3). Fluorescent double WISH performed with riboprobes complementary to piwi-1 and the EEE transcripts tct-like, BTF3-like, DDX5-like and eIF4a-like revealed coincident expression throughout the S4 blastomere compartment (Figure 5F-I). Intriguingly, EEE transcript expression often decayed quicker in differentiated cells than undifferentiated blastomeres, raising the possibility that regulation of EEE transcription and/or transcript stability may vary by cell type. While robust expression of many EEE transcripts was detected in blastomeres during S3-S4, EEE transcript expression in temporary embryonic tissues was present during S3 and drastically diminished by S4 (Figure 5B-E, Figure 5—source data 3).

EEE transcripts expressed throughout the undifferentiated piwi-1+ blastomere population in S3-S4 embryos are downregulated as definitive organogenesis begins during S5. These transcripts likely represent a key temporal shift in the expression profile of piwi-1+ cells during embryogenesis. Moreover, EEE transcript expression provides a molecular metric to distinguish piwi-1+ blastomeres from adult neoblasts.”

More discussion of the EEE genes that are expressed broadly in the embryo would be of interest. What types of genes were discovered?

We have validated expression of a relatively modest cohort of EEE transcripts by WISH, and will do more extensive work to rigorously identify homologs, validate expression of additional candidates and ultimately will devise RNAi knock-down methodology to query function of these genes during S2-S4. The EEE transcripts that were specifically expressed in temporary embryonic tissues clustered with those that showed pan-embryonic expression during S3, followed by blastomere-specific expression during S4; that may confound attempts to hypothesize much based on the annotations alone. Roughly a third of the EEE transcripts have no BLASTx hit versus the NR or Swiss-Prot databases, and are considered novel; we have done little to pursue these candidates. Of the candidates that have BLASTx-based annotations, many appear to be involved in translational control and proteostasis; these candidates superficially appear similar to the types of maternally deposited transcripts present in other organisms, but of course we cannot confirm or refute that connection at this point. The annotated lists are provided in Figure 5—source data 1.

Reviewer #4

[…]Specific comments:

The Introduction is long and detailed on many aspects of neoblast biology, but at the same time unclear and lacking in other aspects. Generally, I am often not sure whether the authors talk about facts known from previous studies, or are already highlighting the findings to be reported in this paper.

Suggestion: shorten the intro by at least 50%. After (in a briefer format than now) summarizing known functions and types of neoblasts in adults/regeneration, summarize literature (as far as it exists) regarding

-embryogenesis in planarians (see below; the statements about embryogenesis that are now found in section 2 of the Results should be in the Introduction)

-specifically, what is said in the literature where/when do neoblasts first appear in embryos (there will probably not be a lot of published information; one example is Ramachandra et al., 1999, on embryonic BrdU -positive presumed neoblasts in an Acoel).

-proliferation, migration, differentiation of neoblasts during embryonic period.

The Introduction was shortened in accordance with comments from reviewers #1, 2 and 4. The lengthy discussion of our results was removed from the Introduction. Instead, we highlight our main conclusions and conceptual advances at the end of the revised Introduction. Information pertaining to neoblasts – molecular and functional definitions, functions, subclasses, and roles during adulthood – were condensed and reorganized, as suggested by reviewer #4.

In the interest of making the manuscript more accessible to a general audience, we added two new paragraphs to the start of the Results section which introduce some of the essential concepts and terminology for Triclad embryos (e.g., direct developer, ectolecithal embryo, dispersed cleavage, temporary embryonic tissues, definitive organs).

Results

Section 1: surveys molecular staging; stages themselves described in supplementary table and figures. This is a useful and important work of reference for the planarian developmental field.

Please highlight in the text:

-what in terms of morphogenesis and organogenesis is novel compared to the previous embryological study of Schmidtea polychroa (Cardona et al)

-how do stages utilized here (1-8) differ from the 8 previously introduced ones

We are happy to comment on similarities and distinctions between our staging series and the published staging series for S. polychroa (Spol). However, we do not expound on these points in the Results section because they will likely appeal to specialists and not the broader eLife readership. Note that it is difficult to directly compare developmental timing with those of the 8 stages presented in the Cardona et al. 2005 publication because the Spol animals were reared at 17˚C and had a gestation time of around 20 days. To confound matters further, the Romero lab later published another Spol staging series for animals reared at 20˚C (Martin-Duran et al. 2010), in which only 7 stages are articulated.

Smed Stage 1/Spol Stage 1 (Cardona et al. 2005)/Not present in (Martin-Duran et al. 2010): We do not have a molecular signature for Stage1 and we have not looked at the zygote or first cell divisions in detail. Commenting further at this point is difficult. In addition, Spol animals are parthenogenic while Smed reproduce sexually, so gametogenesis, egg activation and zygote formation likely differ between the species.

Smed Stage 2/Spol Stage 2 (Cardona et al. 2005)/Stage 1(Martin-Duran et al. 2010): Both Smed and Spol embryos are ectolecithal developers, have blastomeres that undergo dispersed cleavage and form spheres. We believe asymmetries are present within the blastomere population during S2, but disagree with the “Type 1” and “Type 2” blastomere designations presented in Cardona et al. 2005, and which were eventually disavowed in later Spol publications. We cannot comment on the molecular mechanisms implicated in sphere formation at present.

Smed Stages 3+4/Spol Stage 3(Cardona et al. 2005)/ Spol Stage 2+3 (Martin-Duran et al. 2010): Our molecular profiling data supports our designation for two separate stages. Smed Stage 3 is likely most analogous to Spol Stage 2 (Martin-Duran et al. 2010), while Smed Stage 4 is likely most analogous to Spol Stage 3 (Martin-Duran et al. 2010).

Smed Stage 5/Spol Stage 4-5(Cardona et al. 2005)/Spol Stage 4 (Martin-Duran et al. 2010): Gut development is clearly underway during Stage 5, but since these gastrodermal cells express novel embryonic markers it is difficult to say without further analysis whether this is a temporary or definitive tissue. Mature gut progenitors arise at this stage, but the gastrodermal cells lining the intestinal cavity do not express these markers. While we agree that more blastomeres, and hence more mitotic cells are present during S5 than earlier stages, our analysis of the mitotic index for the blastomere compartment suggests that the S3-S5 Smed blastomeres divide at a constant rate, arguing gainst the “mitotic burst” phenomenon described in Spol references.

Smed Stage 6-8/Spol Stages 6-8 (Cardona et al. 2005): these stages are superficially similar, though distinctions in timing and morphology may be present for different definitive organ systems mentioned.

Smed Stage 6-7/Stage 5 (Martin-Duran et al. 2010): Elongation may occur, on average, much sooner in Spol embryos than Smed embryos.

Smed Stage 7-8/Stage 7 (Martin-Duran et al. 2010): Definitive organ development and maturation may occur, on average, much sooner in Spol embryos than Smed embryos.

It would certainly be possible to conduct similar expression profiling studies in Spol and/or other freshwater planarian embryos, and/or to examine spatiotemporal changes in expression patterns during embryogenesis for homologs of the markers we present in this publication. Comparative analysis of gene function during embryogenesis would also be required for informed comment on many of these issues.

-what does molecular profiling add to defining stages (clearer temporal distinction?..)

The molecular profiling data suggests that the genetic regulatory networks and molecular mechanisms governing Smed embryogenesis are robust, and likely temporally much more precise than was anticipated from previous studies of freshwater flatworm embryogenesis. Our RNA-Sequencing data underscores the profound developmental transition between Stages 4 and 5, one that is not readily apparent by changes in morphology alone. We were also able to identify new classes of embryo enriched transcripts which were not anticipated from previous studies. Moreover, the gene expression signatures for each stage provide a wealth of candidates for novel tissue and/or stage specific markers and putative genetic regulators whose function(s) will be assessed in future studies.

Section 2 – “Anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development”

The section describes the expression pattern of piwi, and concludes that the gene is restricted to undifferentiated cells. The text is very difficult to read. Different types of information are intermingled. For example, paragraph two summarizes a result known from the literature. Talks about general expression patterns of non-specified transcripts, documented only in the supplementary material, in differentiating cells. later in the paragraph you speak to the point: piwi is expressed in undifferentiated cells in the bodywall at the S3 stage. But how does the pan-nuclear marker EF1a-like 1 show that, since this marker appears to be expressed in all cells (including the piwi-positive ones)?

The data presented in Figure 2 was reorganized with EF1a-like-1 expression depicted in Figure 2A-B, and piwi-1 expression depicted thereafter. The text was revised to clarify some of the important points brought up by reviewers #1 and #4. The first paragraph of the section, “An anarchic, cycling piwi-1+ blastomeres fuel Smed embryonic development,” describes the EF1a-like-1 expression in S2 and S3 embryos, and includes detailed descriptions of embryonic cell types and architecture. EF1a-like-1 is a pan-embryonic marker, meaning that it is expressed in differentiated temporary embryonic tissues (the primitive ectoderm, the temporary embryonic pharynx, the primitive gut) and the undifferentiated blastomeres located in the embryonic wall (the parenchymal space between the primitive ectoderm and endoderm) of S3 embryos. EF1a-like-1 is not expressed in yolk cells. Yolk cells are present in the embryonic wall along with the blastomeres; these cells have very small, condensed sytox green+ nuclei. Ingested yolk is also present in the primitive gut cavity. Blastomere nuclei are larger, with diffuse sytox green signal (likely due to the decondensed nature of the chromatin), are easily distinguished from yolk cell nuclei in the embryonic wall. In addition, the blastomere nuclei are readily distinguished from primitive ectodermal nuclei by sytox green staining; latter are stretched and sit at the surface of the sphere. We are confident that the combination of EF1a-like-1 and nuclear staining makes allows for unambiguous identification of undifferentiated blastomeres in the embryonic wall of S3 embryos. Piwi-1 expression was always restricted to, and expressed throughout, the undifferentiated blastomere compartment of S3 embryos in EF1a-like-1, piwi-1 costaining experiments. Quantification is provided in the Figure 2E legend.

The final sentences again talk about findings from the previous literature.

I suggest to remove all references to the previous literature from this (Results) section; put them in the Introduction, and in the Results focus on where exactly piwi is expressed. Specifically:

Paragraphs 1-2

-DAPI shows large number of labeled dots: are these all nuclei of blastomeres (Figure 2A, C)?

-if so, only a small subset of blastomeres expresses piwi (2A): how would one characterize distribution? Clustered, around embryonic pharynx?

No, all of the nuclei are NOT blastomeres in the former Figures 2A and 2C (now 2D and 2B, respectively). Yolk cells are abundant but do not express either EF1a-like-1 or piwi-1. Please refer to the previous paragraph regarding the EF1a-like-1, sytox green costaining results.

The distribution of blastomeres in S3-S4 embryos varies widely, as described in Figure 2H. At present we cannot comment on the distribution of the blastomeres during S2 as the embryos undergo sphere formation; that would require costaining with additional markers specific for the temporary embryonic tissues.

-Paragraph 2: sounds like piwi is downregulated in many cells. What is shown (Figure 2A) is that from beginning, piwi is expressed in only few blastomeres (if DAPI really reflects all nuclei). Is there an earlier stage with many more piwi+ cells?

-the pattern and dynamics of piwi in undifferentiated cells ("blastomeres") remains a bit vague. Is the pattern the following: stage 1, before embryonic transient cells differentiate: all cells express piwistage 2/3: as embryonic cells differentiate, they lose piwi; ALL other cells retain itlater stages: as adult cells become postmitotic, they lose piwi; ALL other cells retain it

I don't think this is what happens. But the authors do not make it clear in text and figures what exactly happens. I assume that only a small subset of undifferentiated cells express piwi. I understand that these are distributed. This information needs to be settled; it is one of the crucial pieces of data required in subsequent sections (see comments below).

We show that piwi-1 mRNA is maternally deposited in oocytes (Figure 2C, revised manuscript), and that piwi-1+ blastomeres undergo dispersed cleavage during S2, prior to sphere formation (Figure 2D, revised manuscript). As noted previously, nuclear dyes mark both blastomere and yolk cell nuclei. Our data suggests that piwi-1 is expressed in the zygote and its descendants, the blastomeres, during S2. During sphere formation, asymmetries in the piwi-1+ blastomere population become evident: some blastomeres differentiate into temporary embryonic tissues, and these cells down-regulate piwi-1 and exit the cell cycle. We do not know when asymmetries within the piwi-1+ blastomeres develop during S2 and would like to explore this in the future. However, we never detect expression of piwi-1 (Figure 2G, revised manuscript) or the cell cycle markers PCNA or RNRM2-2 (Figure 3A-B, revised manuscript) in differentiated tissues of S2 or S3 embryos (compare to the expression pattern for EF1a-like-1 in Figure 2A-B, revised manuscript). During S3 and thereafter, piwi-1 expression is restricted to undifferentiated cells. We agree with reviewer #4 that clear communication of these points is critical, and revised the text associated with Figure 2C-G (revised manuscript) as follows:

“piwi-1+ cells were present throughout embryogenesis. piwi-1 expression was detected in oocytes (Figure 2C), suggesting that zygotes contain piwi-1 mRNA. Zygote-derived blastomeres undergoing dispersed cleavage among yolk cells during S2 also expressed piwi-1 (Figure 2D). Costaining with piwi-1 riboprobes and antibodies raised against the G2-M phase mitotic marker H3S10p showed that piwi-1+ blastomeres divide asynchronously during S2 (Figure 2D).

As spheres form during S2, some blastomeres downregulate piwi-1 expression and differentiate into temporary embryonic cell types. piwi-1 expression was restricted to, and expressed throughout, the undifferentiated blastomere population in S3 embryos, as demonstrated by double fluorescent WISH with EF1a-like-1 and piwi-1 riboprobes(Figure 2E, Rich Media File 2). piwi-1+ blastomeres were always located in the embryonic wall, sandwiched between the primitive ectoderm and endoderm (Figure 2F). piwi-1 expression was never detected in primitive ectodermal cells, the embryonic pharynx or primitive gut cells (Figure 2E, G). During S3-S5, piwi-1+ cell number clearly increased, effectively blanketing the sphere (Figure 2G). As definitive gut development proceeded during S6-S8, piwi-1+ cells occupied the parenchyma between the developing gut branches (Figure 2G). Notably, piwi-1+ cells were not detected in the definitive pharynx, and the compartment receded from the anterior margin as head structures developed (Figure 2G). By S8, the spatial distribution of piwi-1+ cells was indistinguishable from that of the adult neoblast compartment (Figure 2G).”

Section 3 reports that genes sets known to be associated with adult neoblasts are also expressed in the embryo. This is an important finding. The main question is:

-are these genes, in the embryo (e.g., S4, S5, S6), restricted to piwi+ cells?

-are they restricted to ALL undifferentiated cells (assuming that not all undifferentiated cells are piwi+; see my questions/comments above)

-are they also expressed in cells that stop dividing and differentiate?

I was not able to find a clear answer to these questions in the text.

We show colocalization analysis for piwi-1 and several adult asexual neoblast-enriched genes, including piwi-2, piwi-3, bruli-1 and tud-1 in S3-S5 embryos (Figure 4F-I). The staining patterns are coincident at these stages, meaning that all piwi-1+ cells express these markers (and vice versa). These data suggest that all undifferentiated cells express a similar complement of nuage genes in S3-S5 embryos: they are pan-blastomere markers at early stages, just as they are expressed throughout the neoblast population in adulthood. The numbers in the lower right corner of the merged images display the percent colocalization with respect to both populations scored (see Figure 4F-I legend); very few single positive cells were observed. We have not performed colocalization analysis and quantification on S6 and older embryos, but would predict to see virtually all piwi-1+ cells at later stages expressing these markers, as do adult neoblasts.

Some differentiating and/or terminally differentiated cells may retain expression of some of these markers. For example, bruli-1 expression was detected in the four neurons innervating the temporary embryonic pharynx (Figure 4E), and may be expressed in the brain and potentially other neural tissues in late stage embryos, as in adults. Similarly, piwi-2 expression was frequently seen in the S3 temporary embryonic pharynx (Figure 4B), and may also be expressed in neural tissues in late stage embryos, as seen in adults. In the future, RNA-Seq on cycling cells versus differentiated cells or whole embryos (WT or irradiated) may yield a global view of how enriched or exclusive expression of these genes are to the blastomere/neoblast compartment.

Section 4 Early embryonic genes (EEE)

I don't understand the message of this section. One reason for confusion arises right at the beginning:

- in the opening lines the authors talk about "S2-S4 enriched transcripts expressed throughout the piwi blastomere population".

- then the authors introduce the 1048 EEE genes: are these the same as above (i.e., they are expressed in the piwi blastomere population)?

The 1048 EEE transcripts are a subset of the 1756 S2-S4 enriched transcripts that were clustered in Figure 5A. Specifically, these 1048 transcripts comprise clusters 5, 6 and 8. Details regarding the expression profiles and BLASTx-based annotation of the EEE transcripts are found in Figure 5—source data 1 (formerly Figure 5—figure supplement 1).

For clarification, we rewrote the first paragraph of this section and moved some of the detailed description of the cluster 5, 6 and 8 expression trends to the Figure 5A legend. The revised text for both the manuscript and figure legend are detailed in a response to reviewer #3, who also brought up this point.

- because, if they are: why do piwi (undifferentiated) blastomeres and differentiated (transient embryonic cells) express the same set of genes? I thought from the previous sections (and on general grounds) that as cells become postmitotic and differentiate, they switch on different genetic programs.

In response to comments from reviewer #3, we rewrote the section detailing the WISH screen and costaining experiments described in Figure 5B-I; please see the previous section for the revised text. Maternal deposition of some EEE transcripts may explain why undifferentiated blastomeres and differentiated temporary embryonic cell types in S3 embryos co-express some EEE transcripts. However, we note that expression of these EEE transcript diminishes quicker in the temporary embryonic tissues than in the blastomeres. Differential transcript stability, relative contributions of maternal load and zygotic transcription, and different modes of maternal mRNA degradation may all be at play and could be investigated in the future. We also note that some EEE transcripts do appear to be expressed exclusively in differentiated cell types, like the temporary embryonic pharynx and the primitive ectoderm. This is consistent with the idea that cells turn on different genetic programs as they differentiate, and suggests that these EEE transcripts are transcribed zygotically, during or after the cell fate decision to downreguate piwi-1, exit the cell cycle and differentiate.

Section 5

It is shown that gene sets associated with specific differentiated cell types are upregulated around stage 5, which is important, but expected, since it confirms the observation that cells differentiate (structurally) at this stage.

The main claim of the heading of this section (that progenitors of specific cell types arise WIHIN the piwi-positive population) needs to be demonstrated, and/or described more clearly (see questions to section 2 above)

Great care was taken to articulate each point critical to our model of how the diversity of lineages required for organogenesis are born during Smed embryogenesis. Moreover, our results immediately suggest why gastrulation is dispensible for freshwater planarian embryos. While many details remain to be worked out in the future, we are energized by paradigm-challenging ramifications of our work. We made very modest changes to the text for this section, as we contend that reviewer #4’s difficulty understanding the conclusions were rooted in not in our writing, but her/his confusion about ectolecithal development and whether piwi-1 was expressed throughout the undifferented blastomere population (see comments related to Figure 2).

Section 6

Cell suspensions derived from embryos at different stages (S5-S8) were injected into irradiated adults, and were able to rescue them. This is also an important experiment. A puzzling finding was that cells from S5 embryos were unable to rescue. Is it possible that S5 embryos did not have a high enough density of piwi+ cells to do the job? Is it possible to get a value for the number of piwi+ cells present in an aliquot of the cell suspension from different stage embryos?

We show that comparable numbers of piwi-1+ cells were introduced per host for S5 and later stages (see Revised Figure 7C, quantification for the number of piwi-1+ cells per host at 1 hour post-transplant). This data was present in the supplemental material in our initial submission, and was moved to a main figure due to its importance for our conclusion regarding stage-dependent differences in donor cell activity.

https://doi.org/10.7554/eLife.21052.064

Article and author information

Author details

  1. Erin L Davies

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    ELD, Conceptualization, Investigation, Resources, Writing, Original draft, Writing, Review and editing, Visualization, Project administration
    Competing interests
    No competing interests declared.
  2. Kai Lei

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    KL, Investigation
    Competing interests
    No competing interests declared.
  3. Christopher W Seidel

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    CWS, Software, Formal analysis
    Competing interests
    No competing interests declared.
  4. Amanda E Kroesen

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    AEK, Investigation, Visualization
    Competing interests
    No competing interests declared.
  5. Sean A McKinney

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    SAM, Formal analysis, Visualization
    Competing interests
    No competing interests declared.
  6. Longhua Guo

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    LG, Investigation, Resources
    Competing interests
    No competing interests declared.
  7. Sofia MC Robb

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    SMCR, Software
    Competing interests
    No competing interests declared.
  8. Eric J Ross

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    EJR, Software, Formal analysis
    Competing interests
    No competing interests declared.
  9. Kirsten Gotting

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    KG, Software, Formal analysis
    Competing interests
    No competing interests declared.
  10. Alejandro Sánchez Alvarado

    1. Howard Hughes Medical Institute, Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    ASA, Conceptualization, Resources, Writing, Review and editing, Supervision and funding acquisition
    For correspondence
    1. asa@stowers.org
    Competing interests
    ASA: Reviewing editor, eLife
    ORCID icon 0000-0002-1966-6959

Funding

Howard Hughes Medical Institute

  • Kai Lei

National Institute of General Medical Sciences (R37GM057260-17)

  • Alejandro Sánchez Alvarado

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank members of the SIMR Planaria Core Facility MS Merryman, D Baumann, C Abrams, K Evans, J Thaden, J Theis and A Vogelslang for their invaluable assistance with planarian husbandry. N Marsh, N Thomas and S Beckham assisted with histology. K Zapian and J Jenkin assisted with irradiation for transplantation experiments. H Vu, S Elliott, LC Cheng, and K Tu graciously donated constructs; S Jones and C Guerrero assisted with WISH and transplant monitoring, respectively. K Tu and S Nowotarksi provided insightful comments on the manuscript. M Miller assisted with scientific illustration. H Li assisted with statistical analysis of the transplant experiments. Funding from the Stowers Institute for Medical Research, the Howard Hughes Medical Institute, and NIH RO1 R37GM057260-17 to ASA supported this work.

Reviewing Editor

  1. Utpal Banerjee, Reviewing Editor, University of California, Los Angeles, United States

Publication history

  1. Received: August 30, 2016
  2. Accepted: January 4, 2017
  3. Accepted Manuscript published: January 10, 2017 (version 1)
  4. Version of Record published: February 6, 2017 (version 2)

Copyright

© 2017, Davies et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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