1. Developmental Biology
  2. Stem Cells and Regenerative Medicine
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Planarian stem cells sense the identity of the missing pharynx to launch its targeted regeneration

  1. Tisha E Bohr
  2. Divya A Shiroor
  3. Carolyn E Adler  Is a corresponding author
  1. Department of Molecular Medicine, Cornell University College of Veterinary Medicine, United States
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Cite this article as: eLife 2021;10:e68830 doi: 10.7554/eLife.68830

Abstract

In order to regenerate tissues successfully, stem cells must detect injuries and restore missing cell types through largely unknown mechanisms. Planarian flatworms have an extensive stem cell population responsible for regenerating any organ after amputation. Here, we compare planarian stem cell responses to different injuries by either amputation of a single organ, the pharynx, or removal of tissues from other organs by decapitation. We find that planarian stem cells adopt distinct behaviors depending on what tissue is missing to target progenitor and tissue production towards missing tissues. Loss of non-pharyngeal tissues only increases non-pharyngeal progenitors, while pharynx removal selectively triggers division and expansion of pharynx progenitors. By pharmacologically inhibiting either mitosis or activation of the MAP kinase ERK, we identify a narrow window of time during which stem cell division and ERK signaling produces pharynx progenitors necessary for regeneration. These results indicate that planarian stem cells can tailor their output to match the regenerative needs of the animal.

eLife digest

Many animals can repair and regrow body parts through a process called regeneration. Tiny flatworms called planaria have some of the greatest regenerative abilities and can regrow their whole bodies from just a small part. They can do this because around a fifth of their body is made of stem cells, which are cells that continuously produce new cells and turn into other cell types through a process called differentiation.

Measuring the gene activity in stem cells from planaria shows that these cells are not all the same. Different groups of stem cells have specific genes turned on which are needed to regrow certain body parts. It is unclear whether all stem cells respond to injuries in the same way, or whether the stem cells that respond are specific to the type of injury. For example, stem cells needed to repair the gut may respond more specifically to gut injuries than to other damage.

Bohr et al. studied how stem cells in planaria respond to different injuries, by comparing an injury to a specific organ to a more serious injury involving several organs. The specific injury was the loss of the pharynx, the feeding organ of the flatworm, while the more serious injury was the loss of the entire head. Within hours of removing the pharynx, stem cells that were poised to develop into pharyngeal cells became much more active than other stem cell types. When the head was removed, however, a wide range of stem cells became active to make the different cell types required to build a head. This suggests that stem cells monitor all body parts and respond rapidly and specifically to injuries.

These findings add to the understanding of regeneration in animal species, which is of great interest for medicine given humans’ limited ability to heal. Many of the genetic systems that control regeneration in planaria also exist in humans, but are only active before birth. In the long-term, understanding the key genes in these processes and how they are controlled could allow regeneration to be used to treat human injuries.

Introduction

When faced with injury or disease, many animals can repair or even replace damaged tissue. This process of regeneration is observed across animal species, and is often fueled by tissue-resident stem cells (Bely and Nyberg, 2010; Sánchez Alvarado and Tsonis, 2006; Tanaka and Reddien, 2011). In response to injury, stem cells accelerate the production of specific types of differentiated cells to repair damaged tissues. For example, in adult mammals, injuries to the intestine, skin, or lung induce stem cells to increase proliferation rates and alter their differentiation potential (Buczacki et al., 2013; Stabler and Morrisey, 2017; Tetteh et al., 2015; Tumbar et al., 2004). These findings suggest that injury can modify the behavior of stem cells to promote repair, but how these changes contribute to tissue regeneration remains unclear.

The freshwater planarian Schmidtea mediterranea is an ideal model organism to study the interaction between injury and tissue repair due to their virtually endless ability to regenerate (Ivankovic et al., 2019). This ability is driven by an abundant, heterogeneous population of stem cells (Adler and Sánchez Alvarado, 2015; Reddien, 2018; Zhu and Pearson, 2016). Defined by ubiquitous expression of the argonaute transcript piwi-1 (Reddien et al., 2005), the planarian stem cell population consists of pluripotent stem cells capable of reconstituting the entire animal (Wagner et al., 2011) and likely organ-specific progenitors (Figure 1A; Scimone et al., 2014a; van Wolfswinkel et al., 2014; Zeng et al., 2018). These progenitors express organ-specific transcription factors required for the maintenance and regeneration of planarian organs, including a pharynx, primitive eyes, muscle, intestine, an excretory system and a central nervous system (Figure 1A), all enveloped in epithelium (Roberts-Galbraith and Newmark, 2015). Expression of specific progenitor markers in piwi-1+ stem cells (Fincher et al., 2018; Plass et al., 2018; Scimone et al., 2014a; van Wolfswinkel et al., 2014; Zeng et al., 2018) provides an opportunity to link the behavior of organ-specific progenitors with injury by tracking stem cell behavior as organ regeneration initiates.

Figure 1 with 1 supplement see all
Both targeted and non-targeted mechanisms contribute to pharynx regeneration.

(A) Schematic of planarian stem cell lineage. Left, whole-mount in situ hybridization (WISH) for the stem cell marker piwi-1. Right, cartoon depiction of the dashed boxed region showing planarian stem cells consisting of pluripotent stem cells and organ-specific progenitors that produce planarian organs. Markers of organ-specific progenitors are indicated. (B) Live images of planarians before and after pharynx amputation. Arrows = pharynx; scale bars = 500 μm. (C) Models for targeted and non-targeted regeneration after different amputations (indicated by red lines). Progenitors are color coded as in A. (D) Schematic of F-ara-EdU delivery relative to amputation for E and F. (E) Confocal images of F-ara-EdU (yellow) in the pharynx (dashed outline) of intact animals, or 7 days after pharynx or head amputation. Dashed box in D = region imaged; DAPI = DNA (blue); scale bar = 100 μm. (F) Number of F-ara-EdU+ cells in the entire pharynx (dashed outline in E). Cartoon represents incision injuries. Graphs represent mean ± SD; symbols = individual animals; shapes distinguish biological replicates and; *, p≤0.05; **, p≤0.01; ***, p≤0.001; one-way ANOVA with Tukey test. Raw data can be found in Figure 1—source data 1.

During homeostasis, planarian stem cells replenish organs by steady proliferation that drives cellular turnover (Pellettieri and Sánchez Alvarado, 2007). Within hours of any injury, a general increase in stem cell division occurs, along with vast transcriptional changes (Baguñà, 1976; Gaviño et al., 2013; Sandmann et al., 2011; Wenemoser et al., 2012; Wenemoser and Reddien, 2010). These changes are only sustained beyond the first day if tissue is removed, in what is referred to as the ‘missing tissue response’ (Baguñà, 1976; Gaviño et al., 2013; Wenemoser and Reddien, 2010). Activated by injury, the extracellular signal-regulated kinase (ERK) contributes to many of these wound-induced transcriptional changes, in addition to regulating stem cell proliferation, differentiation, and survival (Owlarn et al., 2017; Shiroor et al., 2020; Tasaki et al., 2011). Because these injury-induced changes have predominantly been characterized by analyzing broad stem cell behaviors, how they regulate the transition from homeostasis to regeneration of particular organs are key issues to resolve.

Most planarian organs extend throughout the entire body (Figure 1A), and injuries often cause simultaneous damage to multiple organs (Elliott and Sánchez Alvarado, 2013). The resulting complex regenerative response has limited our ability to decipher how stem cells respond to damage of particular organs. Unlike most planarian organs, except the eye, the pharynx is anatomically distinct (Adler and Sánchez Alvarado, 2015; Kreshchenko, 2009). Importantly, it can be completely and selectively removed without perturbing other tissues by brief exposure to sodium azide (Figure 1B; Adler et al., 2014; Shiroor et al., 2018). Because only a single organ is removed, pharynx amputation vastly simplifies the regeneration challenge posed to the animal. Previous work identified the forkhead transcription factor FoxA as an essential regulator of pharynx regeneration (Adler et al., 2014; Scimone et al., 2014a). Under homeostatic conditions, FoxA is expressed in the pharynx and a subset of stem cells. Pharynx amputation triggers an increase in FoxA+ stem cells, demonstrating that injury expands the pool of pharynx progenitors. These properties allow us to dissect how stem cells respond to loss of a specific organ and are regulated to restore it.

The ability of planarians to replace exactly the tissues that have been damaged or removed by injury remains one of the outstanding questions in regeneration (Mangel et al., 2016; Nishimura et al., 2011). Previous studies have suggested that stem cells selectively increase the output of specific progenitors of depleted organs, implying a targeted mode of regeneration (Figure 1C; Thi-Kim Vu et al., 2015). However, others have shown that stem cells respond indiscriminately to tissue removal, incorporating new cells into tissues regardless of whether they have been damaged, suggesting a non-targeted mode of regeneration (LoCascio et al., 2017). Based on these findings, the authors proposed that stem cells non-selectively increase production of any nearby progenitors, determined by the size and position of a wound, rather than the identity of missing tissues (Figure 1C). These two seemingly contradictory models introduce uncertainty into our understanding of the relationship between tissue loss and the stem cell behaviors that ultimately contribute to the regeneration of missing tissues.

By performing an in-depth analysis of specific populations of stem cells in response to different injuries, we show that stem cells can sense the identity of missing tissues. Inflicting various injuries to both the pharynx and body defines distinct contributions of stem cells to regenerated tissue depending on when they divide relative to injury. Planarian stem cells respond to organ loss by selectively increasing expression of organ-specific transcription factors required for subsequent regeneration. Amputation of non-pharyngeal tissues only amplifies non-pharyngeal progenitors, while removal of pharynx tissue selectively increases pharynx progenitors. This increase in pharynx progenitors, and subsequent pharynx regeneration, depends on stem cell division and the MAP kinase ERK, during defined times after tissue loss. Unlike the pharynx, eye regeneration following selective removal is not dependent on stem cell division or ERK signaling, suggesting that different injuries may require distinct regenerative mechanisms. We propose that, in addition to non-targeted and passive modes of regeneration, stem cell behavior can be altered by the loss of specific tissues, selectively channeling their output towards replacement of missing organs.

Results

Both targeted and non-targeted mechanisms contribute to pharynx regeneration

Planarian stem cells have distinct responses to injury depending on whether or not tissue has been removed. Any injury induces a proliferative response within hours, while tissue removal causes a sustained response for up to 4 days, leading to localized proliferation and differentiation (Baguñà, 1976; Wenemoser and Reddien, 2010). Therefore, it has been hypothesized that the initial injury response is a ‘general’ mechanism for repair, whereas the later ‘missing-tissue response’ may be tailored to target the replacement of lost tissue. To evaluate the outcome of stem cell proliferation during these specific injury responses, we altered the timing of stem cell labeling relative to different injuries and analyzed the prevalence of labeled cells in mature organs.

We labeled stem cells with the thymidine analogue F-ara-EdU (Neef and Luedtke, 2011) for 4 hr either immediately or 1 day after pharynx or head amputation. We then analyzed F-ara-EdU+ cells in the pharynx 7 days after amputation (Figure 1D). When F-ara-EdU was applied immediately after amputation (D0), we observed increased F-ara-EdU+ cells in the pharynx following either pharynx or head removal, as compared to intact controls (Figure 1E,F). The timing of this pulse, relative to injury, confirms a previous study showing that amputation stimulates general incorporation of newly generated cells into non-injured tissues (LoCascio et al., 2017). However, F-ara-EdU administration 1 day after amputation (D1) resulted in a specific increase in F-ara-EdU+ pharynx cells only after removal of the pharynx, but not the head (Figure 1E,F). To determine if increased tissue production requires tissue removal, prior to F-ara-EdU administration we performed incisions anterior to the pharynx, which damaged the body without removing any tissue (Figure 1F). However, the number of F-ara-EdU+ cells in the pharynx were comparable to controls, suggesting that tissue removal strongly stimulates production of new tissue, while injury alone does not.

To determine whether other tissues incorporate new stem cells in a time-dependent manner relative to injury, we analyzed the number of newly generated neurons in the brain by combining F-ara-EdU staining with FISH for the neuronal marker ChAT (Figure 1—figure supplement 1A; Wagner et al., 2011). In the newly regenerated brain, head amputation increased F-ara-EdU+ ChAT+ neurons after both F-ara-EdU pulse conditions, as compared to intact controls. However, the number of F-ara-EdU+ ChAT+ brain neurons were comparable to controls after either pharynx amputation or incisions, regardless of when the F-ara-EdU pulse was administered (Figure 1—figure supplement 1B,C). Because chemical pharynx removal does not increase production of neural tissue (Figure 1—figure supplement 1B,C), while its surgical removal does (LoCascio et al., 2017), non-targeted regenerative mechanisms may require injury to specific types of tissues, such as body-wall muscle, epithelia, or intestine. In fact, amputation-specific transcriptional changes important for regeneration have recently been identified within these tissues (Witchley et al., 2013; Lander and Petersen, 2016; Scimone et al., 2016; Benham-Pyle et al., 2020). These results suggest that while cells generated immediately after tissue removal can be broadly deployed to all surrounding tissues, those generated 1 day later are targeted toward only those that are missing.

Pharynx tissue loss selectively increases pharynx progenitors

Our data so far indicate that injury channels the output of stem cells towards missing tissues. If this targeted model is true, injuries that do not remove pharynx tissue, like head amputations, should not increase pharynx progenitors (Figure 1C). Alternatively, if regeneration is non-targeted, injury should non-selectively increase production of any nearby progenitors (LoCascio et al., 2017). If this is the case, head amputation, where pharyngeal tissue is not removed, should also stimulate an increase in pharynx progenitors (Figure 1C).

To test the response of organ-specific progenitors (Figure 1A) to loss of different tissues, we challenged animals with either head or pharynx amputation and analyzed changes in expression of organ-specific progenitor markers within piwi-1+ stem cells. First, we labeled pharynx progenitors with double fluorescent in situ hybridization (FISH) for piwi-1 and the pharynx-specific progenitor marker FoxA, 3 days after pharynx or head amputation. We then quantified pharynx progenitors in the same region, anterior to the pharynx (Figure 2A). As previously reported, we found that pharynx removal caused a significant increase in pharynx progenitors as compared to intact controls (Adler et al., 2014; Scimone et al., 2014a). By contrast, head amputation did not influence the number of pharynx progenitors, which were similar to intact animals (Figure 2A,B). To determine when pharynx progenitors emerge and how long they persist, we quantified the number of pharynx progenitors at various times after pharynx amputation and found that they significantly increased 3 days after amputation (Figure 2C). Because injury broadly influences stem cell behavior, we also analyzed the proportion of these pharynx progenitors relative to all other piwi-1+ stem cells at the same times after pharynx and head amputation and found a similar trend (Figure 2—figure supplement 1A,B). These data indicate that pharynx progenitors are selectively produced 3 days after pharynx loss, but not after loss of other tissue types.

Figure 2 with 3 supplements see all
Pharynx tissue loss selectively increases pharynx progenitors.

(A) Confocal images of double fluorescent in situ hybridization (FISH) for FoxA (green) and piwi-1 (magenta) in intact and injured animals, 3 days post-amputation (dpa). Images are partial projections of a portion of the area outlined by dashed boxes. DAPI = DNA (blue); arrows = double-positive cells; scale bar = 10 μm. (B) Number of FoxApiwi-1+ cells in the area outlined by dashed boxes in A. (C) Number of FoxA+ piwi-1+ cells at indicated times post-pharynx amputation in the area outlined by dashed boxes in A. (D) Number of FoxA+ piwi-1+ cells in intact and injured animals, 3 days after injury (red lines) in the area outlined by dashed boxes in cartoons. (E) Number of cells double-positive for piwi-1 and the indicated progenitor marker in intact and injured animals, 3 days after amputation in the area outlined by dashed boxes in A. For graphs, a 6000 μm2 region in the same location of the pre-pharyngeal region was analyzed over 20 z-sections, as represented by dashed boxes in A and D. Graphs represent mean ± SD; symbols = individual animals; shapes distinguish biological replicates and; **, p≤0.01; ***, p≤0.001; ****, p≤0.0001, one-way ANOVA with Tukey test. Raw data can be found in Figure 2—source data 1.

To determine which types of injuries stimulate an increase in pharynx progenitors, we inflicted various injuries to or around the pharynx (Figure 2D). We then labeled and quantified pharynx progenitors 3 days later. Incisions that damaged the pharynx without removing any tissue failed to stimulate an increase in piwi-1+FoxA + stem cells. However, partial removal of the pharynx (~50–80%) caused a significant increase in pharynx progenitors compared to intact controls. We also performed flank resections, which removed tissue from regions adjacent to the pharynx but did not damage the pharynx itself. Despite being previously shown to increase new cells into the uninjured pharynx with BrdU labeling (LoCascio et al., 2017), we observed comparable numbers of pharynx progenitors as in intact controls (Figure 2D). To determine if the increase in pharynx progenitors was localized, we analyzed the same-sized regions in tails, farther from the site of amputation. However, we did not detect an increase in pharynx progenitors in tails after either pharynx or head amputation as compared to intact controls (Figure 2—figure supplement 1C). These data suggest that stimulation of pharynx progenitor production is local and requires recognition of lost pharynx tissue, but not necessarily loss of the entire organ.

Based on our finding that pharynx progenitors increase only in response to missing pharynx tissue, we hypothesized that the specific pairing of organ loss and progenitor increase would be true for other organs. Besides the pharynx, the eye is the only other planarian organ that is anatomically restricted and thus can be fully removed without leaving any remaining tissue behind (Lapan and Reddien, 2012; LoCascio et al., 2017). Expression of the eye-specific transcription factor ovo is required for eye regeneration (Figure 1A; Flores et al., 2016; Lapan and Reddien, 2012; Rouhana et al., 2013; Scimone et al., 2011; Scimone et al., 2017; Scimone et al., 2014a) and ovopiwi-1+ eye progenitors increase after decapitation (Lapan and Reddien, 2012). Conversely, following pharynx amputation, this increase did not occur (Figure 2E, Figure 2—figure supplement 2A), indicating that pharynx loss does not stimulate the production of eye progenitors.

We also quantified the responses of organ-specific progenitors for muscle (myoD+), intestine (gata-4/5/6+), the excretory system (six-1/2+) and the nervous system (pax6a+) (Figure 1A; Flores et al., 2016; Scimone et al., 2011; Scimone et al., 2017; Scimone et al., 2014a) after either pharynx or head removal. With the exception of intestinal progenitors, all others showed a similar behavior, increasing within 3 days after head removal, but not pharynx removal (Figure 2E, Figure 2—figure supplement 2B–E). Analysis of the proportion of progenitors relative to all piwi-1+ stem cells in the same regions yielded comparable outcomes (Figure 2—figure supplement 1B). No changes in any of these organ-specific progenitors were observed in tail regions after pharynx or head amputation, indicating that it is a local response (Figure 2—figure supplement 1C). Together, these data indicate that planarian stem cells sense the loss of missing tissues to initiate their regeneration through the selective expansion of organ-specific progenitors.

All these organ-specific transcription factors, with the exception of pax6a, are required for regeneration of their cognate organ (Adler and Sánchez Alvarado, 2017; Flores et al., 2016; Lapan and Reddien, 2012; Pineda et al., 2002; Scimone et al., 2011; Scimone et al., 2017). Therefore, to test whether or not these transcription factors regulate pharynx regeneration, we knocked them down with RNAi and assayed feeding ability 7 days after pharynx amputation (Adler et al., 2014; Ito et al., 2001). Unlike FoxA(RNAi), knockdown of other organ-specific progenitor markers did not impact the recovery of feeding ability (Figure 2—figure supplement 3A), despite efficient knockdown and manifestation of known phenotypes (Figure 2—figure supplement 3B, data not shown). Because pax6a is not required for brain regeneration, we performed RNAi of coe, a neural progenitor marker that is required for brain regeneration (Cowles et al., 2013). However, coe knockdown did not affect the recovery of feeding ability (Figure 2—figure supplement 3A). Therefore, it is unlikely that any of these other progenitor markers contribute to pharynx regeneration, despite the presence of muscle and neural tissue within the pharynx.

Pharynx loss selectively induces mitosis of pharynx progenitors

The increase in pharynx progenitors following pharynx amputation suggests that stem cells may divide in response to pharynx loss to selectively amplify pharynx progenitors. Because stem cells are the only dividing cells in planarians (Morita and Best, 1984), we can visualize stem cell division with antibody staining for histone H3Ser10 phosphorylation (H3P). Using this mitotic marker, we verified that stem cell division increased near wounds beginning 1 day after either pharynx or head removal (Figure 3A; Adler et al., 2014; Baguñà, 1976). To determine if these dividing stem cells are pharynx progenitors, we combined antibody staining for H3P with FISH for FoxA. One day after pharynx removal, we observed higher numbers of H3P+ pharynx progenitors in regions adjacent to wounds as compared to intact animals (Figure 3B). To determine how soon after amputation these pharynx progenitors initiate division, and how long it persists, we quantified the coincidence of FoxA+ H3P+ cells at various times after pharynx amputation in the pre-pharyngeal region (Figure 3B). Division of pharynx progenitors increased within 6 hr of pharynx amputation, peaked within 2 days, and returned to homeostatic levels by 5 days after amputation (Figure 3C). Despite an overall increase in H3P+ stem cells 1 day after head amputation (Figure 3A; Baguñà, 1976), numbers of H3P+ pharynx progenitors did not correspondingly increase (Figure 3B,D). Analysis of H3P+ pharynx progenitors relative to all dividing stem cells in the same prepharyngeal region sustained this dichotomy (Figure 3—figure supplement 1A). Interestingly, we were able to detect what appeared to be instances of both symmetric and asymmetric distribution of FoxA in cells undergoing anaphase (Figure 3—figure supplement 1B). Together, these data show that pharynx progenitors are selectively stimulated to divide in response to pharynx loss.

Figure 3 with 4 supplements see all
Pharynx loss selectively induces mitosis of pharynx progenitors.

(A) Whole-mount images of animals stained with anti-phosphohistone H3 (H3P) in intact and injured animals, 1 day post-amputation (dpa). Dashed line outlines animal; arrows = areas of increased H3P; scale bars = 250 μm. (B) Confocal images of FISH for FoxA (green) and H3P antibody (magenta) in intact and injured animals, 1 day post-amputation. Images are partial projections of a portion of the area outlined by dashed boxes. DAPI = DNA (blue); arrows = double-positive cells; scale bar = 10 μm. (C) Number of FoxA+ H3P+ cells at indicated times after pharynx amputation in the area outlined by dashed boxes in B. (D) Number of FoxA+ H3P+ cells at indicated times after head amputation in the area outlined by dashed boxes in B. (E) Number of cells double-positive for H3P and the indicated progenitor marker in the area outlined by dashed boxes in B. For H3P quantification, the entire pre-pharyngeal region was analyzed over 30 z-sections, as represented by dashed boxes in B, and normalized to area. Graphs represent mean ± SD; symbols = individual animals; shapes distinguish biological replicates and; *, p≤0.05; **, p≤0.01; ***, p≤0.001; ****, p≤0.0001, one-way ANOVA with Tukey test. Raw data can be found in Figure 3—source data 1.

In addition to the selective division of excretory system progenitors that occurs after RNAi depletion of excretory tissues (Thi-Kim Vu et al., 2015), progenitors in the epidermal lineage have also been shown to divide following head amputation (van Wolfswinkel et al., 2014). Therefore, we tested whether non-pharyngeal progenitors are selectively stimulated to divide 1 and 2 days after head amputation. Although the kinetics of each differed slightly, excretory system (six1/2+), nervous system (pax6a+), and muscle (myoD+) progenitor division increased 2 days after head amputation, while intestinal (gata-4/5/6+) progenitors did not. Additionally, division of nervous system and muscle progenitors increased only after head but not pharynx amputation (Figure 3E, Figure 3—figure supplement 2A–D). Analysis of these dividing progenitors relative to all H3P+ stem cells recapitulated these results, with the exception of six-1/2+ excretory progenitors, which did not proportionally increase after either amputation (Figure 3—figure supplement 1C). Despite minor differences of each progenitor, the overall trend supports the notion that loss of non-pharyngeal tissues triggers division of stem cells expressing non-pharyngeal progenitor markers while in most cases, pharynx loss does not. This mitotic response appears again to be local, as we did not observe increased division of any of these organ-specific progenitors in tail regions, distant from wounds (Figure 3—figure supplement 3A–D). Even after head amputation, we were unable to detect any dividing eye progenitors (ovo+H3P+) (Figure 3—figure supplement 4A), unless we stalled mitotic exit with the microtubule destabilizing drug nocodazole. Following nocodazole treatment, we detected rare instances of H3P+ eye progenitors; however, they did not increase after head amputation (Figure 3—figure supplement 4B). Therefore, while eye progenitors do divide, their division dynamics do not seem to be affected by injury, similar to intestinal progenitors (Figure 3E, Figure 3—figure supplement 2D). This finding suggests that there may be other eye and intestinal progenitors upstream of those expressing ovo or gata-4/5/6, or that expansion of these progenitors may occur via transcriptional upregulation.

Stem cell division within a critical window is required for pharynx regeneration

In planarians, pulse-chase experiments using thymidine analogs have shown that cell division contributes to the production of regenerated tissues (Cowles et al., 2013; Eisenhoffer et al., 2008; Forsthoefel et al., 2011; LoCascio et al., 2017; Newmark and Sánchez Alvarado, 2000; Wagner et al., 2011). Our results above identified an elevation in pharynx progenitor division within 2 days after pharynx removal (Figure 3C) that correlates with cell cycle entry of stem cells destined for missing pharynx tissue 1 day after amputation (Figure 1E,F). Together, these results define a window of 1–2 days after amputation in which pharynx progenitor division may selectively contribute to pharynx regeneration. Because this timeframe directly precedes the expansion of pharynx progenitors 3 days after pharynx amputation (Figure 2C), we hypothesized that stem cell division increases to specifically generate the progenitors that are necessary for pharynx regeneration.

To test this possibility, we blocked stem cell division with nocodazole, which induces a metaphase arrest with as little as 24 hr of exposure, resulting in the accumulation of mitotic (H3P+) stem cells (Figure 4A; Grohme et al., 2018; Molinaro et al., 2021; van Wolfswinkel et al., 2014). To specifically inhibit mitosis 1–2 days after pharynx amputation, we soaked animals in nocodazole for 24 hr, beginning 1 day after pharynx amputation (Figure 4B). We then assayed pharynx regeneration via recovery of feeding behavior. Animals treated with nocodazole for 24 hr had drastic delays in recovery of feeding, compared to DMSO-treated controls, with only 50% of worms regaining the ability to eat within 20 days, and 100% within 32 days of amputation (Figure 4C). To verify that nocodazole treatment under these conditions delayed pharynx regeneration, we examined pharynx anatomy with the marker laminin, which is strongly expressed in the mouth and pharynx, and weakly expressed in the body where the pharynx attaches (Adler et al., 2014; Cebrià and Newmark, 2007). As expected, while residual laminin expression was retained within the body, animals treated with nocodazole 1–2 days after pharynx amputation completely lacked a pharynx with its characteristic layered structure 7 days after amputation and sustained severe defects even up to 14 days (Figure 4D). Treatment with nocodazole for a full 48 hr, beginning immediately after amputation (Figure 4B), did not exacerbate the delay in feeding ability or defects in pharynx anatomy (Figure 4C,D). These findings suggest that stem cell division outside the 1–2 day window has a minor contribution to pharynx regeneration. To verify this, we soaked animals in nocodazole for 24 hr increments surrounding this 1–2 day window, beginning either immediately, or 2 days after pharynx amputation, which recovered feeding behavior at a similar rate as controls (Figure 4—figure supplement 1A,B). Further, animals treated from 0 to 1 days after amputation had only minor defects in pharynx anatomy, while those treated 2–3 days after amputation were normal (Figure 4—figure supplement 1C). Therefore, we conclude that stem cell division within a critical window of 1–2 days after amputation fuels the majority of pharynx regeneration.

Figure 4 with 2 supplements see all
Stem cell division within a critical window is required for pharynx regeneration.

(A) Whole-mount images of intact animals stained with H3P after treatment with DMSO or nocodazole for 24 hr. Dashed line outlines animal; scale bars = 250 µm. (B) Schematic of nocodazole treatment relative to pharynx amputation for graph in C and images in D. (C) Proportion of animals capable of feeding after pharynx amputation, treated as indicated in B and assayed daily. Error bars represent ± 95% confidence intervals. n ≥ 54 animals from three independent experiments. (D) Whole-mount FISH for the pharynx marker laminin 7 and 14 days post-pharynx amputation (dpa) in animals treated as indicated in B. Dashed blue line outlines pharynx; dashed yellow line outlines mouth; scale bars = 100 µm. n ≥ 23 animals from three independent experiments. (E) Confocal images of FISH for FoxA (green) and piwi-1 (magenta) 3 days post-pharynx amputation in animals treated with DMSO or nocodazole, 1 day after amputation for 24 hr. DAPI = DNA (blue); arrows = double-positive cells; scale bar = 10 µm. (F) Number of FoxApiwi-1+ cells quantified in animals after indicated treatments (schematic) in the area outlined by dashed box in cartoon. Graph represents mean ± SD; symbols = individual animals; shapes distinguish biological replicates and; ****, p≤0.0001, unpaired t-test. Raw data can be found in Figure 4—source data 1.

Figure 4—source data 1

Raw data for feeding assay (Figure 4C) and quantification of piwi-1+ cells (Figure 4F).

https://cdn.elifesciences.org/articles/68830/elife-68830-fig4-data1-v1.xlsx

To test whether stem cell division within this critical window generates pharynx progenitors, we exposed animals to nocodazole 1–2 days after amputation, and analyzed the impact on FoxA+ stem cells. First, we verified that this treatment caused an extensive increase in H3P+ pharynx progenitors 2 days after amputation (Figure 4—figure supplement 2A,B), illustrating that they were arrested in mitosis. Second, we performed double FISH for FoxA and piwi-1 3 days after pharynx removal and found that nocodazole treatment caused a dramatic decrease in pharynx progenitors compared to controls (Figure 4E,F). Importantly, intact animals treated similarly with nocodazole showed no difference in the abundance of pharynx progenitors compared to controls (Figure 4—figure supplement 2C,D). Therefore, perturbing stem cell division during this brief window specifically impacts the production of pharynx progenitors during regeneration. To determine if stem cell division during other times contributed to pharynx progenitor production, we again exposed animals to nocodazole for 0–1 and 2–3 days after pharynx amputation. While we observed some defects in the production of pharynx progenitors, they were more subtle than those in animals treated for 1–2 days after amputation (Figure 4—figure supplement 2E). Together, our data show that stem cell division in a critical window of 1–2 days after amputation produces pharynx progenitors that are likely essential for pharynx regeneration.

ERK phosphorylation is required to produce pharyngeal progenitors

The mitogen activated protein (MAP) kinase pathway drives proliferation and differentiation during development and regeneration in many organisms (Ghilardi et al., 2020; Patel and Shvartsman, 2018). In planarians, phosphorylation of the MAP kinase ERK is the earliest known injury-induced signal required for regeneration. ERK activity regulates broad stem cell proliferation and is required for transcriptional changes that drive axial repatterning (Owlarn et al., 2017; Tasaki et al., 2011). To determine whether pharynx loss also induces ERK phosphorylation, we performed a western blot with an antibody against phosphorylated ERK (pERK). pERK increased 15 min after pharynx amputation and returned to baseline levels within 6 hr (Figure 5A). Therefore, similar to other injuries (Owlarn et al., 2017), pharynx amputation also activates ERK by phosphorylation soon after injury.

Figure 5 with 3 supplements see all
ERK phosphorylation is required to produce pharyngeal progenitors.

(A) Western blot for phosphorylated ERK (pERK) and tubulin (loading control) in intact animals and at the indicated times after pharynx amputation. (B) Schematic of PD0325901 (PD) exposure relative to pharynx amputation for graph in C and images in D. (C) Proportion of animals capable of feeding after pharynx amputation, treated as indicated in B and assayed daily. Error bars represent ± 95% confidence intervals. n ≥ 47 animals from three independent experiments. (D) Whole-mount FISH for the pharynx marker laminin 7 days post-pharynx amputation (dpa) in animals treated as indicated in B. Dashed blue line outlines pharynx; dashed yellow line outlines mouth. Scale bars = 100 µm. n ≥ 18 animals from two independent experiments. (E) Confocal images of FISH for FoxA (green) and piwi-1 (magenta) 3 days post-pharynx amputation in animals treated with DMSO or PD (schematic). DAPI = DNA (blue); dashed box = region imaged; arrows = double-positive cells; scale bar = 10 µm. (F) Number of FoxApiwi-1+ cells 3 days post-pharynx amputation after indicated treatments (schematics E and F). (G) Confocal images of FoxA FISH (green) and H3P antibody (magenta) 2 days post-pharynx amputation in animals treated with DMSO or PD, 1 day after amputation for 24 hr. DAPI = DNA (blue); arrows = double-positive cells; scale bar = 10 μm. (H) Number of FoxA+ H3P+ cells 2 days post-pharynx amputation in animals treated with DMSO or PD (schematic). Bar graphs represent mean ± SD; symbols = individual animals; shapes distinguish biological replicates and; ***, p≤0.001; ****, p≤0.0001; one-way ANOVA with Tukey test (F), unpaired t-test (H). Raw data can be found in Figure 5—source data 1.

Figure 5—source data 1

Original, uncropped images of western blots in Figure 5A.

https://cdn.elifesciences.org/articles/68830/elife-68830-fig5-data1-v1.zip
Figure 5—source data 2

Raw data for feeding assay (Figure 5C), quantification of piwi-1+ cells (Figure 5F) and H3P+ cells (Figure 5H).

https://cdn.elifesciences.org/articles/68830/elife-68830-fig5-data2-v1.xlsx

Exposing animals to PD0325901 (PD), an inhibitor of the upstream ERK-activating MEK kinase, blocks ERK phosphorylation and permanently inhibits regeneration following substantial anterior tissue removal (Owlarn et al., 2017). To determine whether ERK is also required for pharynx regeneration, we exposed animals to PD for 5 days immediately after pharynx amputation and then assayed feeding behavior (Figure 5B,C). While DMSO-treated control animals regained the ability to feed within 7 days, animals treated with PD from 0 to 5 days after amputation had substantial delays in feeding, with 50% of worms feeding by day 13 and all worms feeding by day 29 (Figure 5C). Depending on the timing, delaying administration of MEK inhibitors relative to amputation partially or completely rescues anterior regeneration, and suggests that ERK acts within the first day of regeneration (Owlarn et al., 2017). To pinpoint when ERK signaling is important for pharynx regeneration, we delayed PD exposure for 1 or 2 days after pharynx amputation, and again assayed feeding (Figure 5B,C). Animals exposed 1 day after pharynx amputation (1–6 days) had delayed feeding ability, similar to those treated immediately after amputation (0–5 days), suggesting that ERK activity within the first day of amputation is dispensable for pharynx regeneration. Animals exposed 2 days after amputation (2–7 days) regained the ability to feed at rates similar to controls (Figure 5C), indicating that ERK is essential for regeneration within the first 2 days after pharynx amputation. Therefore, ERK likely acts primarily between 1 and 2 days after pharynx amputation. This timing occurs after the increase in pERK following injury has already subsided (Figure 5A), suggesting that pharynx regeneration may be facilitated by homeostatic levels of ERK signaling instead of its injury-induced high level activation.

To verify that the inability of ERK-inhibited animals to feed was caused by defects in regeneration, we analyzed pharynx anatomy 7 days after pharynx amputation with FISH for laminin. Animals exposed to PD for 5 days, beginning 0 or 1 day after amputation, lacked a pharynx, while pharynges in animals exposed beginning 2 days after amputation were comparable to controls (Figure 5D), mirroring the results of our feeding assay (Figure 5C). PD-exposed animals eventually regenerated a pharynx within about 2 weeks (Figure 5C, Figure 5—figure supplement 1A), suggesting that ERK inhibition does not permanently block pharynx regeneration. Therefore, we confirmed the effects of ERK inhibition by repeating pharynx regeneration experiments with UO126 (UO), another potent, but structurally independent, MEK inhibitor and observed the same outcomes (Figure 5—figure supplement 1B–D). Furthermore, exposure to PD or UO eliminated pERK after pharynx amputation on a western blot (Figure 5—figure supplement 1E) and also permanently blocked regeneration after extensive anterior tissue removal, for up to 70 days (Figure 5—figure supplement 1F). Together, these results define a window 1–2 days after amputation in which ERK activity is required for pharynx regeneration.

Soon after amputation, ERK signaling is required for upregulation of several genes including follistatin (fst) (Owlarn et al., 2017), which accelerates regeneration by inhibiting activin-1 and -2 (Gaviño et al., 2013; Roberts-Galbraith and Newmark, 2013; Tewari et al., 2018). Like ERK inhibition, fst(RNAi) prevents regeneration following substantial anterior tissue removal (Owlarn et al., 2017; Tewari et al., 2018). However, if less tissue is removed, the requirement for fst diminishes (Tewari et al., 2018). Likewise, when we amputated animals pre-pharyngeally and maintained them to PD for 5 days, head regeneration was initially delayed in 100% of animals 7 days after amputation, but eventually occurred in 88% of animals within 2 weeks (Figure 5—figure supplement 1G), on a similar timeline as pharynges (Figure 5C,D, Figure 5—figure supplement 1A). Therefore, ERK may mediate pharynx regeneration via fst expression, resulting in a short-lived block of pharynx regeneration. To test this, we analyzed fst expression, which increases within 6 hr of head amputation (Gaviño et al., 2013), but not until 24 hr after pharynx amputation (Figure 5—figure supplement 2A), suggesting that injury-induced ERK activation (Figure 5A) may not always coincide with fst upregulation. Despite upregulation of fst expression after pharynx amputation, fst(RNAi) animals regained the ability to feed at a normal rate (Figure 5—figure supplement 2B and C), indicating that fst is not required to accelerate pharynx regeneration. Therefore, although pharynx loss eventually induces fst expression, regulation of pharynx regeneration via ERK is independent of fst.

Our data suggests that ERK activity is required 1–2 days after pharynx amputation, just prior to the emergence of pharynx progenitors 3 days after amputation. We hypothesized that ERK may promote the production of these progenitors, which we tested by maintaining animals in PD for 3 days following pharynx amputation (Figure 5E). PD exposure significantly inhibited the increase in FoxApiwi-1+ cells typically observed after pharynx amputation (Figure 5E,F). Importantly, intact animals treated with PD for 3 days showed no difference in the abundance of pharynx progenitors as compared to controls (Figure 5—figure supplement 3A), indicating that ERK activity for this duration is not necessary for maintaining pharynx progenitors during homeostasis. Therefore, the decrease in pharynx progenitors following pharynx amputation and PD treatment is likely due to their reduced production, rather than altered survival. To determine when ERK promotes pharynx progenitor production, we exposed animals to PD in 24 hr increments during this 3-day window (Figure 5F). While PD treatment starting 0 or 2 days after pharynx amputation had no effect on pharynx progenitors, treatment starting 1 day after amputation significantly inhibited pharynx progenitor increase, similar to those treated for three full days (Figure 5F). Similar experiments with UO yielded the same outcomes (Figure 5—figure supplement 3A,B), demonstrating that ERK activity 1–2 days after pharynx amputation promotes the production of pharynx progenitors during regeneration.

To test whether PD-mediated inhibition of pharynx progenitor production following pharynx loss has long-term consequences on pharynx regeneration, we exposed animals to PD for 0–3 days and 1–2 days after amputation, and assayed feeding behavior. These animals had delays in feeding, although less severe than those treated for five full days (Figure 5—figure supplement 3C), presumably because animals start recovering after drug washout. Meanwhile, those treated for 0–1 day after pharynx amputation recovered feeding ability at normal rates (Figure 5—figure supplement 3C), illustrating that PD exposure that does not affect the production of pharynx progenitors does not delay regeneration. Together, these results indicate that ERK activity 1–2 days after amputation promotes pharynx regeneration by contributing to the increase in pharynx progenitors 3 days after amputation.

Because ERK signaling regulates broad stem cell division associated with missing tissue in planaria (Owlarn et al., 2017), the decrease in pharynx progenitor production after drug exposure could be a result of reduced stem cell division. To determine if amputation-induced pharynx progenitor division depends on ERK activity, we exposed animals to MEK inhibitors 1–2 days after pharynx amputation, and analyzed FoxA+ H3P+ stem cells. Neither PD or UO exposure substantially impacted the number of dividing pharynx progenitors as compared to controls (Figure 5G,H, Figure 5—figure supplement 3D), despite an overall decrease in H3P+ stem cells in the same animals (Figure 5—figure supplement 3E). These data indicate that ERK signaling is not required for the selective increase in pharynx progenitor division induced by pharynx loss. Because the timing of ERK’s requirement for increasing pharynx progenitors overlaps with when stem cell division is required (Figure 4C–F) 1–2 days after pharynx removal, ERK signaling is unlikely to regulate initiation of pharynx progenitor division. Instead, ERK likely promotes pharynx progenitor production and regeneration by regulating FoxA expression and stem cell differentiation.

Stem cell division and ERK phosphorylation are not required for eye regeneration following selective removal

Unlike pharynx removal, selective removal of the eye does not increase stem cell division or expansion of eye-specific progenitors. Instead, following resection, the eye regenerates by passive homeostatic turnover that is not regulated by the presence or absence of the eye (LoCascio et al., 2017). Therefore, we speculated that eye regeneration, following specific removal, may not have the same requirements as the pharynx. To test whether eye regeneration relies on stem cell division and ERK signaling, we performed eye-specific resections and immediately exposed animals to either nocodazole or MEK inhibitors (Figure 6A). We monitored eye regeneration with FISH for ovo and the eye-specific marker opsin Sánchez Alvarado and Newmark, 1999. We confirmed that eye tissue was successfully removed by the absence of ovoopsin + photoreceptors immediately after surgery (Figure 6B). Exposure to either nocodazole for 2 days, or MEK inhibitors for 5 days, did not impact eye regeneration (Figure 6B,C), despite a complete block of pharynx regeneration under these conditions (Figure 4D, Figure 5D, Figure 5—figure supplement 1D). Our results show that, unlike pharynx regeneration, eye regeneration does not require stem cell division or ERK activity after selective removal. Instead, much like the maintenance of pharynx progenitors in intact animals (Figure 4—figure supplement 2D, Figure 5—figure supplement 3A), eye regeneration is unaffected by drug treatments. Therefore, it is possible that pre-existing eye progenitors, or more likely, the continued production of homeostatic levels of eye progenitors (LoCascio et al., 2017) is sufficient for eye regeneration following selective removal.

Stem cell division and ERK activation are not required for eye regeneration following selective removal.

(A) Schematic of drug treatment relative to injuries for B-E. (B) Confocal images of FISH for ovo (green) and opsin (magenta) immediately (0dpa), or 7 days post-eye resection in animals treated as in A. Dashed box in cartoon represents area imaged. Blue dashed line outlines anterior edge of worm; red arrows = missing eyes; yellow arrows = regenerated eyes; scale bars = 50 µm. (C) Proportion of ovoopsin+ eyes that regenerated 7 days post-eye resection. n ≥ 34 animals from three independent experiments. (D) Confocal images of FISH for ovo (green) and opsin (magenta) 7 days post-head amputation in animals treated as in A. Dashed box in cartoon represents area in top images. DAPI = DNA (blue); blue dashed line outlines anterior edge of worm; red arrows = missing eyes; yellow arrows = regenerated eyes; scale bars = 50 µm. Bottom = zoom of ovo+ cells from the regions outlined by gray boxes in top images. Scale bars = 5 µm. (E) Proportion of ovoopsin+ eyes that regenerated 7 days post-head amputation. n ≥ 32 animals from three independent experiments. In graphs, error bars = ± 95% confidence intervals and; ****, p≤0.0001; Fisher's Exact Test. Raw data can be found in Figure 6—source data 1.

Previous studies have shown that eye regeneration may occur in alternative ways depending on the context of the wound. While eye-specific resections do not invoke typical injury responses, more severe injuries stimulate the expansion of ovo+ eye progenitors (Lapan and Reddien, 2012; LoCascio et al., 2017). Because wounds generated from eye resection do not stimulate the same response as more severe injuries, their repair may not depend on the same mechanisms facilitated by proliferation and ERK signaling. Therefore, we tested whether stem cell division and ERK activity are required for eye regeneration after head amputation (Figure 6A). In controls, ovoopsin + photoreceptors re-emerged 7 days after amputation (Figure 6D,E; Lapan and Reddien, 2011). By contrast, animals treated with either nocodazole for 2 days or ERK inhibitors for 5 days failed to regenerate eyes, despite the presence of ovo+ eye progenitors (Figure 6D,E). We conclude that unlike pharynx regeneration, eye regeneration only requires stem cell division and ERK signaling in the context of more severe injuries.

Discussion

In this study, to evaluate the contribution of stem cells to tissue regeneration, we challenged stem cells by inflicting different types of injuries to planarians. Analysis of organ-specific progenitors after various wounds highlighted shifts in the abundance of these stem cell populations based on the presence or absence of specific organs. We uncovered a mechanism that targets regeneration towards missing tissues, fueled by the division and subsequent expansion of organ-specific progenitors required for regeneration. In particular, we found that pharynx loss induces a selective increase in pharynx progenitor division, 1–2 days after amputation, followed by an ERK-dependent expansion of pharynx progenitors that are channeled towards pharynx regeneration (Figure 7). These findings suggest that in many cases, stem cells can sense the identity of missing tissues to launch their targeted regeneration.

Model for targeted pharynx regeneration.

Soon after pharynx loss, stem cells recognize the pharynx is missing and target regeneration toward the pharynx by selectively inducing division of existing FoxA-expressing stem cells (red), or expression of FoxA in division-competent stem cells, within 1–2 days. This division drives an ERK-dependent increase in pharynx progenitors 3 days after pharynx loss, which is required for pharynx regeneration.

Modes of regeneration: targeted or non-targeted?

Previous work has suggested two potential models underlying planarian regeneration. One group proposed a non-targeted model, in which stem cells broadly incorporate into both damaged and undamaged tissue, dependent on indiscriminate amplification of progenitors triggered by nearby wounds (LoCascio et al., 2017). On the other hand, a targeted model suggests that stem cells amplify organ-specific progenitors in response to perturbations to organ tissues (Thi-Kim Vu et al., 2015). By labeling proliferating stem cells with F-ara-EdU at different times after amputation, we have uncoupled the contribution of these two mechanisms to the production of regenerated tissues. Depending on the type of injury, cells generated soon after amputation are channeled into all nearby tissues, even undamaged ones. However, those generated 1 day after amputation are selectively targeted for missing tissues. Importantly, the timing of this targeted mechanism overlaps with an essential peak in pharynx progenitor division, revealing that this mechanism generates organ progenitors necessary for the replacement of missing tissues.

The cells that regenerate the pharynx are primarily generated through this targeted mechanism, mediated by stem cell division and ERK signaling between 1and 2 days after amputation (Figure 7). Smaller lesions, such as eye resections, do not stimulate a proliferative wound response (LoCascio et al., 2017), nor do they depend on stem cell division and ERK signaling for subsequent regeneration. A previous study proposed that regeneration initiation requires signals regulated by both injury and tissue loss (Owlarn et al., 2017). If eye removal is not detected as an injury, and is insufficient to trigger a ‘missing tissue signal’, this could explain why eye regeneration, following resection, happens passively via homeostatic cellular turnover in the regenerating eye (LoCascio et al., 2017). Further, while surgical removal of the pharynx does increase production of neural tissue (LoCascio et al., 2017), selective chemical pharynx removal does not, suggesting that non-targeted regeneration may require injury to specific types of tissues, such as body wall muscle or epithelia, in addition to tissue loss. Therefore, multiple avenues lead to regeneration: passive homeostatic turnover in the absence of typical injury responses, and both targeted and non-targeted mechanisms that increase cellular production when tissue is lost.

Our work highlights several key features that are important to consider for future analysis of regeneration. In particular, depending on the size and severity of the wound, initial injury signals may vary, triggering a differential requirement for stem cell division and regulatory signaling pathways. Our results underscore the distinct requirements for proliferation and ERK activity in regeneration of eyes and the pharynx. Also, the timing of experimentation and analysis should be deliberate, as we show that F-ara-EdU-labeled stem cells are differentially incorporated into regenerating and non-regenerating tissues depending on when they are labeled relative to injury. Another key consideration is the intrinsic heterogeneity of cell populations that are analyzed. Stem cells identified by piwi-1 and H3P staining encompass cells with different potencies and differentiation states (van Wolfswinkel et al., 2014; Zeng et al., 2018), which may respond uniquely to the changing environment of a regenerating animal. Therefore, broad analysis of stem cells lacks the resolution required to tease apart the intricacies involved in coordinating regeneration. Restricting our analysis to organ progenitors and even further to those that are actively dividing narrows this focus. However, our observation of the asymmetric and symmetric segregation of FoxA in dividing cells suggests that heterogeneity exists even within these subsets of stem cells. Resolving the specific stem cells responsible for driving different modes of regeneration, and when cell fate is established in them, will be an exciting area for future work.

ERK signaling plays multiple roles during regeneration

Phosphorylation of ERK promotes regeneration in many animals (DuBuc et al., 2014; Wan et al., 2012; Yun et al., 2014). In planaria, ERK has been implicated as a regeneration trigger, as it is briefly activated by phosphorylation within minutes of injury and is required for wound-induced transcription, stem cell differentiation and broad stem cell proliferation (Owlarn et al., 2017; Tasaki et al., 2011). ERK also functions to re-establish axial patterning during regeneration (Owlarn et al., 2017; Umesono et al., 2013), which depends on a network of positional cues that are expressed in muscle cells throughout the body (Lander and Petersen, 2016; Scimone et al., 2016; Witchley et al., 2013). Because tissue removal from the body requires re-establishment of these positional cues for regeneration to proceed (Rink, 2018), it has been difficult to distinguish ERK’s roles in organ regeneration.

Unlike amputations to the body, pharynx removal does not broadly disrupt positional cues, which has allowed us to pinpoint a distinct role for ERK in organ regeneration. Both ERK activity and stem cell division act simultaneously, but independently, 1–2 days after pharynx loss, to drive the expansion of pharynx progenitors 3 days after amputation. These events occur after the injury-induced increase in pERK has subsided. Also, ERK activity is dispensable for pharynx progenitor division. Taken together, these results suggest that an ERK-independent signal triggers division of FoxA+ stem cells, and that ERK acts later during organ regeneration to facilitate stem cell differentiation or maintain cell fate.

Among the ERK-dependent wound-induced genes is follistatin (fst) (Owlarn et al., 2017), which promotes regeneration in ways similar to ERK (Tewari et al., 2018). Interestingly, both ERK and fst are absolutely essential for regeneration following substantial anterior tissue removal, but become less so if smaller amounts of tissue are removed. This variability may be due to the extent of axial patterning disruption induced by different amputations. In fact, the inability of fst(RNAi) animals to regenerate is entirely dependent on their failure to reset positional information, as inhibition of wnt signals that restrict head formation rescue these regeneration defects (Tewari et al., 2018). Therefore, it is likely that injury-induced fst expression and ERK phosphorylation primarily regulate regeneration initiation by establishing repatterning rather than triggering stem cell behaviors that directly contribute to organ regeneration. ERK inhibition reduces broad stem cell division after tissue removal (Owlarn et al., 2017), but not the specific increase in division of pharynx progenitors that accompanies pharynx loss, suggesting that not all injury-induced stem cell behaviors may be critical for regeneration. Further, rescuing axial repatterning, and thus regeneration after fst knockdown, does not rescue defects in missing tissue-induced stem cell proliferation and apoptosis (Tewari et al., 2018). Therefore, the ‘missing-tissue response’ may encompass multiple events including patterning, broad stem cell division, and the generation of organ-specific progenitors that contribute independently to different aspects of regeneration.

Receptor tyrosine kinases such as the epidermal growth factor receptor (EGFR) and the fibroblast growth factor receptor (FGFR) have been shown to play critical roles in signaling upstream of ERK in many organisms (Patel and Shvartsman, 2018), making them intriguing candidates for potential regulators of regeneration. In planarians, egfr-3 is required to activate ERK during regeneration (Fraguas et al., 2017) and is also involved in stem cell differentiation (Fraguas et al., 2011; Lei et al., 2016). Other studies have highlighted roles for the ligand egf-4 and the receptors egfr-1 and egfr-5 in the differentiation of stem cells into brain, intestinal and excretory tissues, respectively (Barberán et al., 2016; Fraguas et al., 2014; Rink et al., 2011). Further, some FGFRL-Wnt circuits restrict pharynx formation to the trunk region, possibly through regulation of FoxA expression (Lander and Petersen, 2016; Scimone et al., 2016). Whether any of the planarian EGF or FGF ligands or receptors similarly regulate the production of pharyngeal progenitors remains to be determined (Cebrià et al., 2002; Ogawa et al., 2002).

Shifts in stem cell heterogeneity contribute to regeneration

By studying the dynamics of FoxA expression in stem cells after pharynx or head removal, we have uncovered shifts in stem cell heterogeneity that depend on the presence or absence of a particular organ. Intriguingly, we find that both the overall number of FoxA+ pharynx progenitors, as well as those that are actively dividing, increase only after pharynx removal. Therefore, stem cells sense the absence of the pharynx and channel their proliferative output toward the population of stem cells required to regenerate it. Combined with our analysis of non-pharyngeal progenitor dynamics after different amputations, our results suggest that the heterogeneity of the stem cell population can be differentially deployed depending on the severity of the injury and the particular tissues that need repair.

Interestingly, removal of non-pharyngeal tissues like the head, does not increase pharynx progenitor proliferation but nevertheless results in increased F-ara EdU+ cells within the pharynx, raising a conundrum regarding the source of these cells. One possibility is that other, non-FoxA+ progenitors which have yet to be identified, may contribute to pharynx regeneration. However, no other tissue-specific transcription factors or proposed pharynx progenitor markers (meis, twist, or dd_554) appear to be required for pharynx regeneration (Cowles et al., 2013; Scimone et al., 2014a; Zhu et al., 2015). Alternatively, a recent study has suggested that planarian stem cells, even those expressing organ-specific transcription factors, may harbor a large degree of plasticity that allows fate switching between stem cell and progenitor types (Raz et al., 2021). While this hypothesis has not been tested in the context of injury, it is possible that stem cells generated soon after tissue loss could adopt a pharynx progenitor fate at various times over the course of regeneration, which would not necessarily generate a detectable increase in pharynx progenitors at any one time. It will be interesting to explore the potential of these cells in more detail when true lineage-tracing becomes possible in planarians.

Surprisingly, stem cell division in a narrow window, 1–2 days following pharynx amputation, is absolutely essential for pharynx regeneration, and coincides with the elevation of pharynx progenitor division that directly precedes their increase 3 days after pharynx amputation. The requirement for division in this brief moment after amputation suggests that stem cells detect tissue loss through transient signals regulated by injury. Indeed, a recent study has identified a population of potentially slow-cycling stem cells, reminiscent of reserve stem cells in mammals, that may be specifically induced to enter the cell cycle by tissue loss (Bankaitis et al., 2018; Molinaro et al., 2021). Whether this distinct population of stem cells contributes to the regeneration of particular organs is not known. Regulatory signals could either be produced upon injury to promote regeneration, or released from inhibitory cues that might emanate from organs when they are present (Rink, 2018; Ziller-Sengel, 1967; Ziller-Sengel, 1965). Intriguingly, a recent study characterizing transient amputation-induced transcriptional changes revealed that the majority of these changes occur within differentiated cell types (Benham-Pyle et al., 2020). The possibility of transient signals customized to particular organs and the ability of stem cells to readily sense them may explain how planarians exhibit such rapid and robust regeneration of all organs.

Cell fate acquisition can occur throughout the cell cycle (Fichelson et al., 2005; Pauklin and Vallier, 2014; Soufi and Dalton, 2016). The increase in FoxA expression in both actively dividing stem cells, and those outside of M-phase, does not pinpoint a particular time in the cell cycle where fate acquisition during regeneration might occur. Tissue loss could generate fleeting signals sensed by stem cells that influence them to adopt a specific cell fate during division to compensate for missing tissue. Alternatively, stem cells already expressing organ-specific markers may be poised to divide upon receiving such a signal, allowing them to quickly initiate regeneration upon exit of the cell cycle. Indeed, studies in human hepatoma cell lines have shown that FoxA1 remains attached to chromatin during mitosis, contributing to rapid activation of downstream targets following mitosis during liver differentiation (Caravaca et al., 2013). It will be interesting to explore these possibilities in future studies.

Mammalian homologs of FoxA were the first identified ‘pioneer’ transcription factors, characterized by their ability to engage closed chromatin and drive organogenesis (Hsu et al., 2015; Iwafuchi-Doi and Zaret, 2016; Lam et al., 2013; Zaret and Mango, 2016). This raises the possibility that pioneer factors may be viable in vivo targets for achieving regeneration of entire organs. In fact, overexpression of a related mammalian transcription factor, FoxN, is sufficient to drive regeneration of the thymus in mice (Bredenkamp et al., 2014). The increased proliferation of stem cells expressing FoxA after pharynx removal suggests that activation of pioneer factors may also drive organ regeneration in planaria. Other pioneer factors, including gata-4/5/6, soxB1-2 and FoxD, are also expressed in planarian stem cells and are required for regeneration of the intestine (Flores et al., 2016; González-Sastre et al., 2017), sensory neurons (Ross et al., 2018), and anterior pole (Scimone et al., 2014b; Vogg et al., 2014), respectively. Therefore, upregulation of pioneer factors in stem cells may be a general strategy used to initiate organ regeneration. Identifying the regulatory mechanisms responsible for the selective activation of pioneer factors in stem cells may be an ideal approach to understanding how organisms initiate regeneration of targeted organs in vivo. In conclusion, our work sheds light on the flexibility and dynamic responses of stem cells to different injuries, and highlights potential mechanisms to activate organ-specific transcriptional programs required for regeneration.

Materials and methods

Key resources table
Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional
information
AntibodyAnti-DIG-AP (sheep polyclonal)RocheCat#11093274910, RRID:AB_514497in situ: 1:3000
AntibodyAnti-DIG-POD (sheep polyclonal)RocheCat# 11207733910, RRID:AB_514500in situ: 1:1000
AntibodyAnti-DIG_FITC (sheep polyclonal)RocheCat# 11426346910, RRID:AB_840257in situ: 1:1000
AntibodyAnti-phosphohistone H3 (Ser10) (rabbit monoclonal)AbcamCat# Ab32107, RRID:AB_732930IF: 1:1000
AntibodyAnti-Oregon Green-HRP (rabbit polyclonal)Thermo FisherCat# A21253, RRID:AB_2535819IF: 1:1000
AntibodyAnti-tubulin (mouse monoclonal)Sigma/MilliporeCat# T5168, RRID:AB_477579WB: 1:1000
AntibodyAnti-Phospho-p44/42 MAPK (Erk1/2) (rabbit monoclonal)Cell Signaling TechnologiesCat# 4370S, RRID:AB_2315112WB: 1:1000
AntibodyGoat anti-mouse Alexa Flour 488 (polyclonal)Thermo FisherCat# A11029, RRID:AB_2534088WB: 1:4000
AntibodyGoat anti-rabbit IRDye 800CW (polyclonal)LI-CORCat# 926–32211, RRID:AB_621843WB: 1:20,000
Chemical compound, drugF-ara-EdUSigmaCat# T511293dilution: 0.5 mg/mL
Chemical compound, drugOregon Green 488 azideThermo FisherCat# O10180F-ara-EdU development: 100 µM
Chemical compound, drugProteinase KThermo FisherCat# 25530049F-ara-EdU development: 10 µg/mL in situ: 4 µg/mL
Chemical compound, drugRoche Western Blocking ReagentRocheCat# 11921673001dilution: 0.5%
Chemical compound, drugHorse serumSigmaCat# H1138-500mLdilution: 5%
Chemical compound, drugNocodazoleSigmaCat# M1404dilution: 50 ng/mL
Chemical compound, drugPD0325901EMD Millipore/CalbiochemCat# 4449685 MGdilution: 10 µM
Chemical compound, drugUO126Cell Signaling TechnologiesCat# 9903Sdilution: 25 µM
Chemical compound, drugWestern blot lysis bufferZanin et al., 2011PMID:22118282
Chemical compound, drugPierce Protease InhibitorThermo FisherCat# A32965
Chemical compound, drugPierce Phosphatase InhibitorThermo FisherCat# A32957
Chemical compound, drugBolt LDS sample bufferLife TechnologiesCat# B0007
Chemical compound, drugBolt 4–12% Bis-Tris polyacrylamide gelInvitrogenCat# NW04125BOX
Chemical compound, drugOdyssey blocking bufferLI-CORCat# 927–40000
Software, algorithmGraphPad PrismGraphPad Prism (https://graphpad.com)RRID:SCR_002798Version 9
Software, algorithmGraphPad QuickCalcsGraphPad QuickCalcs (https://graphpad.com/quickcalcs/)RRID:SCR_000306
Software, algorithmImageJImage J https://imagej.net/RRID:SCR_003070
OtherDAPI stain 5 µg/mLThermo Fisherdilution: 1:5000
OtherAqua-PolymountPolysciences IncCat# 18606
OtherPVDF Immobilon membraneMerck MilliporeCat# IPFL00010

Worm care

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Animals of Schmidtea mediterranea asexual clonal line CIW4 were maintained in a recirculating water system (Arnold et al., 2016; Merryman et al., 2018) containing Montjuïc salts (planaria water) (Cebrià and Newmark, 2005). Prior to experiments, animals were transferred to static culture and maintained in planaria water supplemented with 50 µg/mL gentamicin sulfate. Animals used for experiments were between 2 and 3 mm in length and starved for approximately 5–7 days.

Amputations, sodium azide treatment, and tricaine anesthetization

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Pharynx removal was performed by chemical amputation as previously described (Adler et al., 2014; Shiroor et al., 2018). Planarians (2–3 mm in size) were placed in 100 mM sodium azide diluted in planaria water. After 4–7 min, the pharynx extended out of the body and was plucked off using fine forceps (#72700-D; Electron Microscopy Sciences). Animals were kept in sodium azide for no longer than 10 min, rinsed three times, and then transferred into a fresh dish. For pharynx incisions and partial amputations, animals were soaked in tricaine solution (4 g/L in 21 mM Tris pH 7.5) diluted 1:3 in planaria water which causes the pharynx to extend but not detach. Pharynx incisions were created by using forceps to snip along the length of the pharynx. For partial pharynx amputations, the proximal end of the pharynx was snipped off with forceps or trimmed with a scalpel. To resect eyes, animals were immobilized on moist filter paper, and eyes were scraped out using the tips of fine forceps. All other amputations and injuries were performed with a micro feather scalpel (#72046–15 or #72045–45; Electron Microscopy Sciences). For direct comparisons to pharynx-amputated animals, head-amputated and intact animals were soaked in sodium azide for 2–3 min.

F-ara-EdU administration

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Animals were soaked in 0.5 mg/mL F-ara-EdU (Sigma T511293) in planaria water containing 3% DMSO for 4 hr either immediately or 24 hr after amputation and fixed 7 days after amputation.

In situ hybridizations and immunostaining

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Animals were fixed as previously described (Pearson et al., 2009) with minor modifications. Briefly, animals were killed in 7.5% N-acetyl-cysteine in PBS for 7.5 min and fixed in 4% paraformaldehyde in PBSTx (PBS + 0.3% Triton X-100) for 30 min. Worms were then rinsed twice with PBSTx and incubated in pre-warmed reduction solution (PBS + 1% NP-40 + 50 mM DTT + 0.5% SDS) at 37°C for 10 min. Worms were rinsed twice more with PBSTx, dehydrated in a methanol series and stored at −20°C.

For F-ara-EdU detection, following fixation, animals were rehydrated and bleached in 6% H2O2 overnight. Animals were then treated with proteinase K (10 µg/mL proteinase K and 0.1% SDS in PBSTx) for 15 min, and post-fixed in 4% formaldehyde in PBSTx for 10 min. A F-ara-EdU development solution was made containing PBS + 1 mM CuS04 and 100 µM Oregon Green 488 azide (Thermo Fisher O10180). Freshly made 100 mM ascorbic acid was added to this solution immediately before administering it to samples, which were then incubated for 30 min in the dark. Following a few rinses with PBSTx, animals were post-fixed, rinsed 2x in PBSTx, and put through in situ (see below). Following in situ, animals were placed in K block (5% inactivated horse serum, 0.45% fish gelatin, 0.3% Triton-X and 0.05% Tween-20 diluted in PBS) at room temperature for 4 hr or 4°C overnight. To detect F-ara-EdU, animals were incubated with 1:1000 anti-Oregon Green-HRP (Thermo Fisher A21253) and counterstained with DAPI in K block at 4°C overnight. Antibodies were washed off in PBSTx, pre-incubated with tyramide (1:2000 FAM) for 10 min and developed for 15 min.

Colorimetric in situ hybridizations were performed as described in Pearson et al., 2009 using anti-DIG-AP (Roche 11093274910) at 1:3000. Fluorescent in situ hybridizations were performed as in King and Newmark, 2013 with minor modifications. Briefly, animals were rehydrated and bleached (5% formamide, 1.2% H2O2 in 0.5x SSC) for 2 hr, then treated with proteinase K (4 µg/mL in PBSTx, Thermo Fisher 25530049). Following overnight hybridizations at 56°C, samples were washed 2x each in wash hybe (5 min), 1:1 wash hyb:2X SSC-0.1% Tween 20 (10 min), and 2X SSC-0.1% Tween 20 (30 min), 0.2X SSC-0.1% Tween 20 (30 min) at 56°C followed by 3 × 10 min PBSTx washes at room temperature. Subsequently, animals were placed in blocking solution (0.5% Roche Western Blocking Reagent and 5% inactivated horse serum diluted in PBSTx). Animals were then incubated with an appropriate antibody: 1:1000 anti-DIG-POD (Roche 11207733910) or 1:1000 anti-FITC-POD (Roche 11426346910) in blocking solution at 4°C overnight followed by several washes with PBSTx. For development with FAM (1:2000) or Cy3 (1:7500), animals were preincubated with tyramide in borate buffer for 30 min and then developed with 0.005% H2O2 in borate buffer for 45 min. For development with rhodamine, animals were pre-incubated with tyramide (1:5000) for 10 min and developed for 15 min. To inactivate peroxidases, animals were treated with 200 mM sodium azide or 4% H2O2 in PBSTx for 1 hr, then rinsed with PBSTx >6 times before application of the next antibody.

For H3P detection, following in situ, animals were incubated with anti-phosphohistone H3 (Ser10) antibody (Abcam, Cambridge, MA Ab32107) diluted 1:1000 in blocking solution (0.5% Roche Western Blocking Reagent and 5% inactivated horse serum in PBSTx) for 2 days at 4°C. Primary was washed off with PBSTx followed by incubation with goat anti-rabbit-HRP (Thermo Fisher 31460) diluted 1:2000 in PBSTx overnight at 4°C. Antibody was washed off with PBSTx and samples were pre-incubated and developed with rhodamine tyramide as described above.

For all in situ and immunostaining experiments, DAPI [5 µg/mL] (1:5000 dilution; Thermo Scientific) was added along with the last antibody (except for colorimetric in situ). After the final development, animals were soaked in ScaleA2 (4M urea, 20% glycerol, 0.1% Triton X-100, 2.5% DABCO) (Hama et al., 2011) for at least 3 days. Animals were mounted ventral side up except for those stained for ovo, which were dorsal side up, and embedded in Aqua-Polymount (Polysciences Inc 18606). To maintain consistent sample thickness, animals were mounted in wells cut from a double layer of double stick tape (Scor-Pal 6’ wide Scor-Tape 209).

Western blot

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Ten animals per condition were snap frozen in lysis buffer (50 mM Hepes pH 7.5, 1 mM EGTA, 1 mM MgCl2, 100 mM KCl, 10% glycerol, 0.05% NP40, and 0.5 mM DTT) (Zanin et al., 2011) containing Pierce protease and phosphatase inhibitors (Thermo Fisher A32965 and A32957). A cup horn sonicator (Branson Ultrasonics Corporation, Danbury, CT) chilled to 4°C was used to generate extracts by sonication for a total of 2 min with 1 s pulses at 90% amplitude. Total protein was quantified using a NanoDrop OneC (Thermo Fisher). After quantification, Bolt LDS sample buffer (Life Technologies B0007) was added to the extracts and 100 µg of each sample was run on a polyacrylamide gel (Bolt 4–12% Bis-Tris, Invitrogen NW04125BOX). The gel was transferred onto a PVDF Immobilon membrane (Merck Millipore IPFL00010) using the Pierce Power Blot Cassette system (Thermo Scientific), then treated with Odyssey blocking buffer (LI-COR 927–40000) for 1 hr at RT. Membranes were incubated overnight at 4°C with mouse anti-tubulin (Sigma/Millipore T5168) and rabbit anti-Phospho-p44/42 MAPK (Erk1/2) (Cell Signaling Technologies 4370S), diluted 1:1000 in blocking buffer. The blot was washed 3 × 10 min in TBST (TBS +10% Tween 20) and incubated for 1 hr at RT with goat anti-mouse Alexa Flour 488 (Thermo Fisher A11029) and goat anti-rabbit IRDye 800CW (LI-COR 926–32211) secondary antibodies (diluted 1:4000 and 1:20,000 respectively in Odyssey blocking buffer). Membranes were then washed as before and imaged using a Bio-Rad ChemiDoc MP. Western blots were repeated at least twice with comparable results.

Drug treatments

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Nocodazole (Sigma M1404) was administered in 24 or 48 hr increments at 50 ng/mL. PD0325901 (EMD Millipore/Calbiochem 4449685 MG) and UO126 (Cell Signaling Technologies 9903S) were administered at 10 µM and 25 µM, respectively. Drugs were diluted in planaria water containing 0.05% DMSO. Animals were rinsed three times after treatment and either fixed immediately, or transferred to a new dish and rinsed daily until further experimentation.

Feeding assay

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Animals were fed 20 µL of colored food (4:1 liver:milliQ water with 2% red food coloring) in a petri dish. After approximately 30 min, the number of animals with red intestines were scored. For time courses, feeding assays started at 4 days post-amputation and any animals that ate were removed from the dish. Feeding assay time courses were repeated at least three times with ~20 animals assayed per experiment.

RNA interference

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RNAi was performed as previously described (Rouhana et al., 2013), with in vitro-synthesized double-stranded RNA (dsRNA). dsRNA was diluted to a final concentration of 400 ng/µL in colored food. RNAi food was administered every 3 days, six times in total, except for gata-4/5/6 and six-1/2, which caused phenotypes after 1–2 feeds. C. elegans unc22 dsRNA was used as a control. Amputations were carried out 5–7 days after the last feed. All RNAi experiments were repeated at least twice with ~10 animals per experimental group.

Image acquisition, quantification, and statistical analysis

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Whole-mount colorimetric in situ hybridizations and live worms were imaged on a Leica M165F. Fluorescent in situ hybridizations were imaged on a Zeiss 710 confocal microscope using a 25x objective with 2.28 µm z-sections. ImageJ software was used for processing and quantification (Schindelin et al., 2012). All samples were quantified without blinding by manual examination of optical sections of overlaid fluorescence channels in pre-defined regions of animals as indicated in figures. Cells were identified as positive for markers if fluorescence coincided with DAPI signal and was easily distinguishable from background levels, as demonstrated in figures with images. Quantification was performed in a minimum of 6 animals per experimental group.

For piwi-1+ progenitor analysis, a 6000 μm2 region in the same location of the pre-pharyngeal area was captured at 1.3x zoom. Quantification included 20 z-sections (45.6 µm) beginning at the first piwi-1+ cell. For H3P analysis, the entire pre-pharyngeal region was imaged, captured at 0.6x zoom. Quantification included 30 z-sections (68.4 µm) beginning at the first H3P+ cell and was normalized to area. Representative confocal images are partial projections of ~5 z-sections from regions that were used for quantification, re-imaged at 4x zoom. For F-ara-EdU quantification, images were captured at 1x zoom. In most cases, the pharynges and brains of each experimental group were imaged from the same animal. All visible F-ara-EdU+ cells in the pharynx and all F-ara-EdU+ ChAT+ cells in the brain were quantified throughout the entire organ. Representative F-ara-EdU images are projections of the entire analyzed region. In bar graphs, symbols represent individual animals, and shapes distinguish biological replicates. Statistical analysis was performed using PRISM-Graphpad version nine or GraphPad Quick Calcs to perform one-way ANOVA with Tukey test, unpaired t-test or Fisher’s Exact test as indicated in figure legends. *p≤0.05; **p≤0.01; ***p≤0.001; and ****p≤0.0001.

Primers

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Sequences for all transcripts used in this study were cloned using the following primers:

GeneSmed IDForward primerReverse primer
piwi-1dd_Smed_v6_659_0_1gaccaagaagaggaggtctccgcgttcgcgaattctgtcatt
FoxAdd_Smed_v6_10718_0_4aacgacctcaacggaatgtttcatgcgccaaagttaaggata
ovodd_Smed_v6_48430_0_1aatgcccacagatttgtccataaagtgaattcgggtg
myoDdd_Smed_v6_12634_0_1ctattccggtccatactcagcactcttgatcaactttcctcg
gata-4/5/6dd_Smed_v6_4075_0_1gtccgtaagatccacgatccgtgattgaggaatagggcttcg
six-1/2dd_Smed_v6_9774_0_1ccttgtcagggatctaatccggtgaggatgataagttggg
pax6add_Smed_v6_17726_0_1ctgggcataaatcaaaccgccttgggggataaactgatcc
coedd_Smed_v6_9893_0_1cgaagagcagacaacagcacttttaccaacacccgattgc
laminindd_Smed_v6_8356_0_1agtcgctggcaaagtgcatctaatgatgcgtggtatccacag
fstdd_Smed_v6_9584_0_1cagtggtgtgcaatttagcgagttcgcaggtattcttggtttcgtaattcg
ChATdd_Smed_v6_6208_0_1tcggttgctgaaggtattgcaggcatatagcattctacacgg

Data availability

All data generated and analyzed in this study are included in the manuscript and supporting files. Numerical data used to generate all graphs are included in a single Source Data File with an individual tab containing the raw data for each figure panel.

References

  1. Book
    1. Ziller-Sengel C
    (1965)
    Inhibition de la régénération du pharynx chez les planaires
    In: Edwin G, editors. Regeneration in Animals and Related Problems. Amsterdam: North Holland Publishing Co. pp. 193–201.
    1. Ziller-Sengel C
    (1967)
    Investigation of the inhibition of the regeneration of the planarian pharynx. I. Evidence of an auto-inhibiting factor found in the regenerating pharynx
    Journal of Embryology and Experimental Morphology 18:91–105.

Decision letter

  1. Phillip A Newmark
    Reviewing Editor; Morgridge Institute for Research, United States
  2. Didier YR Stainier
    Senior Editor; Max Planck Institute for Heart and Lung Research, Germany

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

How stem cells respond to missing tissue is an important question for understanding regenerative processes. Here, Bohr et al. address this question by comparing how stem cells respond to loss of a single organ (the pharynx) versus loss of many tissues (after head amputation) in the planarian, an organism that can regenerate its entire body from a tiny piece of tissue. The authors find that the stem cells respond to loss of the pharynx by producing more pharynx progenitors; this increase is not observed after removal of non-pharyngeal tissues. Thus, the planarian's stem cells are able to "sense" when certain tissues are missing and target their fates accordingly.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Planarian stem cells sense the identity of missing tissues to launch targeted regeneration" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

All three reviewers recognized the potential significance of this work, but also shared the same concerns about sample sizes, lack of biological replicates, and insufficient details about cell quantification. Given the interest in the question being pursued, if these issues can be addressed satisfactorily, a revised paper could be considered here as a new submission. We have included the reviews below and hope you will find the comments helpful.

Reviewer #1:

This manuscript by Bohr et al. explores how planarian stem cells respond to the loss of a specific organ: the pharynx. The previously proposed "target-blind" model of planarian regeneration (LoCascio et al. 2017) posited that stem cells do not respond directly to missing tissues, but rather replace missing cell types based on their normal rates of homeostatic turnover. In contrast, Bohr et al. suggest that planarian stem cells can sense and respond to the loss of specific missing tissues, using the pharynx as a case study. The authors conclude that planarians may use more than one mode of regeneration, depending upon the target being regenerated (eye vs. pharynx).

The question explored in this paper is of fundamental importance, and providing an alternative model by which planarian stem cells regenerate missing tissues should be of interest to a broad readership. Unfortunately, in its current form, the manuscript presents enticing preliminary findings, rather than robust experimental observations. Currently, the manuscript has limited samples sizes and experimental replicates, which is unfortunate. Because this paper is attempting to refute a previously published model it is critical that the data are clear and convincing. If not, these findings could be summarily dismissed without appropriate debate. If the authors can show the robustness and rigor of their results, and address the major issues listed below, this manuscript would represent a significant contribution to our understanding of planarian regeneration.

1. Throughout the manuscript, experiments were either not repeated, or the number of biological replicates was not reported. In most cases, it appears that experiments were done only once (with the exception of the drug treatments). Numbers of biological replicates and sample sizes should be explicitly stated and the data from different replicates reported for Figures 1D-G, 2B-D, 3C, 3E-F, 4B, 4D-H, 5C-D, and 6A-E.

2. The authors do not sufficiently describe their methods for imaging and quantifying cells (Figures 1E, 1G, 2C-D, 3F, 4E-H, 5D, 6B). The size of the area covered to collect these data is unclear. High-magnification images are shown: are these the areas that were imaged? If so, their results could be biased by choosing small regions of interest. Ideally, the authors should quantify more than one region per animal. Also, they do not describe the depth of the z-stacks collected or how these stacks were normalized/standardized across conditions. All their conclusions hinge on the quantification of progenitor populations in response to different amputation paradigms or chemical treatments, so the standards for imaging and quantification must be clearly reported.

3. Inappropriate statistical tests were used throughout. The use of multiple t-tests amplifies the chance of a Type I error and is especially problematic when up to 7 comparisons were made! The authors should use one-way ANOVA with multiple comparison corrections for all experiments with more than two groups.

4. Figures 1D-E show that upon pharynx amputation but not head amputation, FoxA+ piwi+ pharynx progenitors increase. These data suffer from the quantification issues highlighted above: how the data were quantified is not sufficiently described, only 3 data points were taken (one per animal), the experiment appears to have been performed only one time, and the wrong statistical test was used. Rather than reporting the number of FoxA+ piwi-1+ cells counted, the authors should quantify the total number of doubly positive cells as a percentage of piwi-1+ cells, as was previously published (Adler et al. 2014). The authors also fail to specify whether the change observed between "3 dpa phx" and "3 dpa head" is significant, which is a material point.

5. Figure 2D also suffers from the inadequate quantification practices described above. Ideally, FoxA+ cells should be quantified as a percentage of the H3P+ cells observed.

6. The authors use "stem cell", "progenitor", "stem cell progenitor", and "progenitor stem cells" in a mixed and confusing way throughout the paper. For example, in lines 174-175 the authors state that "proliferation of FoxA+ stem cells precedes the increase in pharynx progenitors." This refers to FoxA+ H3P+ cells vs. FoxA+ piwi-1+ cells, but the only difference is that the former are stem cells in the act of mitosis. Is a distinction being made? Elsewhere in the paper, FoxA+ piwi+ cells are referred to as stem cells. The terminology used needs more clarity and consistency.

Along these lines, what is a FoxA+ PSC (Line 168)? Are the authors suggesting that FoxA is a pluripotency marker? Or are the authors saying that tgs-1 has a more expansive expression pattern and is co-expressed with progenitor markers? If double FISH was performed with the progenitor markers reported (ovo, myoD, gata-4/5/6, six-1/2, and pax6), would they all overlap with tgs-1? These experiments need to be performed to make any claims about FoxA expression in the context of pluripotency.

7. In Figure 3C, how does pharynx regeneration occur over such a long period of time after nocodazole treatment in the 1-2 day window? Does target-blind regeneration occur once this window is missed? The authors should repeat analysis of FoxA+ progenitors at later time points in this condition, and/or show rates of BrdU incorporation into the pharynx with and without nocodazole treatment in this window.

8. The use of the inhibitor PD in Figure 4 is problematic. No data are shown to verify that phosphorylation of ERK was inhibited in these experiments. A citation of previous use is not sufficient. The effect of PD on WT uncut animals with regards to FoxA+ cells is not shown and is a necessary control. To address questions of drug specificity, the authors should corroborate their findings with a second inhibitor of ERK signaling like U0126, which has already been shown to work in planarians (Owlarn et al. 2017).

9. The conclusion that ERK signaling functions in regulating differentiation but not proliferation is premature (line 249-264). Figures 4E-H should be quantified as percentages of piwi-1+ and H3P+ cells, especially since there is decreased proliferation overall with PD treatment.

10. The use of head fragments to compare eye regeneration vs. pharynx regeneration is inappropriate. Previous studies have already shown that the absence of eyes is not required to induce ovo+ progenitor amplification (LoCascio et al. 2017). Thus, this is not a surprising result (line 343-345) and the authors are mis-citing previous observations (in the earlier work, head fragments were never described). The region where ovo+ cells was quantified in Figure 6A is not justified or explained. The yellow box is placed in a medial region where ovo+ cells do not normally reside. The authors should image within the laterally positioned ovo+ streams that have been previously described (Lapan and Reddien 2012; LoCascio et al. 2017).

11. Rather than using head fragments, the authors should repeat the flank resection experiments shown previously (LoCascio et al. 2017). This previous study showed that increased BrdU incorporation into the pharynx occurred following flank resection even though the pharynx was present. That result may have been 1) an artifact of increased BrdU staining due to stimulation of proliferation upon injury, 2) caused by unintentional damage to cells associated with the pharynx, or 3) a response to the loss of FoxA+ progenitor populations that surround the pharynx rather than the loss of the differentiated organ. The authors have the opportunity to revisit this published observation by quantifying the FoxA+ progenitor response during flank resection +/- pharynx. Without these data, this story is incomplete and therefore the conclusion of a targeted regeneration response is not yet convincing.

12. The negative results that proliferation and ERK signaling are dispensable for eye regeneration in Figure 6 are weak and unconvincing. The regenerated eyes appear smaller; this should be quantified (number of PRNs per eye). If the small pharynges that form in Figures 3D and 4C are considered a deleterious phenotype, why is the same standard not applied to the eye? Also, existing eye progenitors could have been sufficient for eye regeneration in these drug treatments. Furthermore, eyes did not regenerate after nocodazole treatment 50% of the time. Is it not more likely that the observations reported are dosage and timing artifacts? How has proliferation been affected? These observations do not live up to the claims made.

13. The authors claim that ovo+ cells are not proliferative (H3P+) even in cases where there is eye progenitor amplification (head amputation), but the data are not shown (line 321). They should be. Indeed, previous publications have never shown that ovo+ cells proliferate. This might mean that there are proliferating eye progenitors that precede expression of ovo. The authors should discuss this alternative.

14. The authors claim that eye regeneration does not require proliferation or ERK signaling but pharynx regeneration does. This conclusion hinges on the gross observation that eyes can regenerate in the presence of nocodazole and PD (see point 12 above). These data are coarse and the interpretations are unconvincing. Instead, their model can be directly tested using BrdU pulse-chase experiments. According to the authors' model, one would predict that following pharynx amputation, the rate of incorporation of BrdU+ cells into the regenerating pharynx should be higher than in uninjured controls. Conversely, the rate of BrdU incorporation into the regenerating eye should remain unchanged between injured (i.e. eye-resected) and control animals (LoCascio et al. 2017). Once the authors have established the prediction above, they have the opportunity to show the effects of nocodazole, PD, and U0126 on BrdU incorporation in the regenerating eye vs pharynx following eye resection and pharynx amputation in the same animal. This way, the authors can directly test the requirement for proliferation and/or ERK signaling in both tissues.

Reviewer #2:

Bohr, Shiroor, and Adler investigate how stem cells respond to the loss of specific tissues in planarians. The planarian stem cell population (neoblasts) are distributed throughout the planarian body and include pluripotent stem cells and a wide range of lineage-committed progenitor cells. How this heterogenous pool of cells behave post-injury or amputation is incompletely understood. The discovery of markers to label stem cell progeny have opened the door to investigate stem cells respond to tissue loss. However, the anatomy of planarians makes it difficult to surgically remove or damage specific organs. The PI of this study developed an assay to remove the pharynx by "chemical amputation" to study the mechanisms underlying regeneration of this organ without drastically perturbing or injuring other tissues. Using this approach, this paper investigates how a well-defined population of FoxA+ progenitors respond to pharynx removal at early time-points during the regeneration. Their data suggest that stem cells are able to detect loss of the pharynx and respond by generating significantly more cells fated to become pharynx, whereas amputation of non-pharyngeal tissues does not have an obvious effect on pharynx progenitor specification dynamics. In addition, using pharmacological treatments, the authors show that cell proliferation and ERK signaling are required for the expansion of pharynx progenitors and cell differentiation. In contrast, other cell types in the planarian eye do not appear to require proliferation or ERK signaling, suggesting that stem cell responses "target blind" as suggested in a previous study, but are rather tuned to specific missing tissues.

This work has the potential to make a significant contribution to the field by advancing our understanding of how the planarian heterogenous stem cell population responds to the loss of a specific organ. However, the report is preliminary as presented. It appears that the authors performed many experiments a single time. In addition, description of the methods is insufficient. Consequently, before this work can be considered for publication, the authors need to chiefly demonstrate the reproducibility of the data and robustness of the observations.

1. The authors need to replicate experiments to increase the sample size for most experiments.

2. Details for imaging and quantification should be explicitly stated in the methods, and the reported cell count numbers should be normalized as appropriate for each set of experiments.

3. Although the authors mention "lineage-tracing" experiments, they do not perform DNA analog pulse-chase experiments to analyze a temporal progression and spatial localization of stem cells to FoxA+ progenitors after pharynx removal. The authors rely on PH3-staining in conjunction with FoxA, and supplementary experiments using the pluripotent stem cell marker tgs-1 (which was only examined at 1 dpa). Could the authors clarify what they think FoxA+ stem cells represent? Are these self-renewing pluripotent stem cells or lineage-committed progenitors? Can the authors get some insight by scanning their images of PH3+ cells expressing FoxA visibly undergoing metaphase? Are daughter cells uniformly FoxA+ as reflected in the model? At least in one of the cells shown in the nocodazole treated controls it appears that both daughter cells express FoxA (Figure 3). I suggest showing some higher magnification images to support the interpretations/conclusions. Others have posited (e.g., Rink, Chapter 2 of Planarian Regeneration: Methods and Protocols), that not every dividing cell may be a long-term self-renewing stem cell and whether a transient amplifying cells exist or contribute to regeneration in planarians is unknown. Adler and Sánchez Alvarado (2015) discuss the role of transient states and how the transcriptional profiles change in response to regeneration. It wasn't clear to me how the authors think about these cells based on the limited number of experiments and analysis, and there are a few places where the terminology is inconsistent, especially in reference to proliferating ovo+ progenitors (P. 14). The authors need to be clear, and it might be helpful to illustrate their model in one of the early figures or to include it in the final model, which omits tgs-1 due to the limited number of experiments performed with this marker gene (Figure 7).

4. The pharynx is complex and there is no data to assess what the contribution of other progenitor populations might be. I don't think or think it is unlikely that FoxA+ progenitors are solely responsible for reconstructing the pharynx. The authors should examine how other progenitor populations behave during the process of pharynx regeneration by extending the timeline of progenitor cell analysis. This would reveal if there is fluctuation in progenitor dynamics as animals regenerate the pharynx or re-scale proportions after pharynx regeneration. For example, can the authors test if they are able to detect a contribution of neural progenitors to regeneration of the pharyngeal nervous system? And if so, when during the regeneration process does it take place in the context of their study?

Reviewer #3:

In this manuscript, Bohr et al. examine how the pluripotent stem cell system of planarians responds to organ-specific damage. If and how the differentiation of specific cell types is dynamically regulated is a conceptually fascinating problem in planarians and in general stem cell research. The authors address this problem by comparing the stem cell response between a single-organ amputation (the pharynx) versus broad tissue loss (decapitation). Their findings indicate that only the removal of the pharynx triggers the increased differentiation of pharyngeal cell types, while the loss of non-pharyngeal tissues upregulates the differentiation of progenitors of multiple organs, but not the pharynx. Further, the authors implicate temporally restricted ERK signaling as a regulatory component in the differentiation of pharyngeal cell types. These observations are also important because they contrast with the previously proposed "target blind" model (LoCascio et al., 2017) that posits the differentiation of different cell types at constant relative proportions, with the rate of stem cell divisions as global production rate regulator. In contrast, the observations by Bohr et al. provide further evidence for more flexibility and specificity within the planarian stem cell system ("target consciousness") in the sense of lineage-specific adjustments in the production rates of specific cell types.

That said, the manuscript generally suffers from an overly narrow focus. Important questions remain regarding the specificity of the stem cell response to pharynx amputation and multiple experiments lack important controls (see below). Moreover, the authors have overlooked that a "target conscious" progenitor response has already been demonstrated by the selective proliferation of protonephridial marker expressing neoblasts in response to protonephridial damage by RNAi (Vu et al., 2015). As a result, the current manuscript would require substantial additional experimentation to consolidate its findings sufficiently for publication in eLife.

1. Specificity of the stem cell response

The central premise of the paper is the selective amplification of pharyngeal progenitors in response to pharynx amputation. This the authors conclude on basis of i) an increase in the absolute number of foxA+/piwi-1+ cells in a specific area, while ii) de-capitation has no effect on the absolute number of foxA+/piwi-1+ cells in the same area. This approach is an insufficient demonstration of specificity, as the known phenomenon of wound-induced stem cell activation might also change the absolute number of specific neoblast subclasses and might do so in an injury-dependent manner.

To account for this important caveat, the authors need to i) quantify RELATIVE proportions of foxA+/piwi-1+ cells out of total piwi cells (or of total H3P+ cells) and ii) they need to include other organ progenitors in the initial analysis. The latter is also critical because the pharynx is a complex organ comprising descendants of multiple lineages (e.g., muscle, neurons, epidermis) and it is not clear whether the foxA+/piwi-1+ cells indeed serve as a singular origin of all constituting lineages (as assumed by the authors), or if they only provide a subset of pharyngeal cells with a rate-limiting role in pharynx assembly (e.g., pharyngeal muscle). In face of such uncertainty, iii) the quantification of new cell incorporation into the pharynx versus other tissues via BrdU labeling would be necessary to address this caveat and to provide a global perspective on the specificity of the response independent of incompletely characterized marker genes.

In addition, the following experimental design problems need to be addressed or better documented, including:

• The authors provide insufficient methodological detail on progenitor quantifications, even though the entire manuscript rests on this assay. What are their criteria for scoring a piwi-1+ cell as double-positive for the often weak and noisy lineage labels? If done "by eye", was double-blind scoring used? Were all cells in a given Z-stack counted or only specific planes? If the latter, by which criteria were image planes selected for quantification? How were the specific specimens out of an experimental cohort selected for imaging/quantification? Though not necessarily a unique shortcoming of this particular study, these points simply need to be adequately addressed in order to rigorously support quantitative differences between experimental conditions (e.g., specificity).

• The authors appear not to distinguish at all between technical replicates (e.g., multiple specimens within an experimental cohort) versus biological replicates (independent experimental cohorts). This is significant, because (i), the use of the standard error of the mean (SEM) that the authors employ throughout is not really an appropriate measure for a single biological replicate with 3 animals – the standard deviation (SD) would seem a more appropriate measure in this context (SD). (ii), the number of worms quantified for each experiment is generally low (n=3 animals in Figure 1, 3, 5, 6; n=5 animals in the rest of the figures) given the observed variability in the data (e.g., ~25-30 foxA+/piwi-1+ cells 3d after pharynx amputation in Figure 1E versus 50 cells in figure 1H). Similarly, for kinetic experiments as in Figure 1H or 2C, it is simply crucial to ensure that the error bars include the variation in response dynamics between multiple replicates due to the drift in the baseline fraction of H3P+ cells or varying staining efficiencies (e.g., different batches of animals on different days), rather than the technical variation in a single experimental cohort only. Please address these concerns by adding more specimens and a thorough description of the experimental design.

2. Timeline of pharynx regeneration

The pharynx regeneration timeline and associated events that the authors present are insufficiently supported by experimental data. The conclusion in line 215 "that proliferation in a critical window of 1 to 2 days after pharynx amputation produces a population of progenitors that are likely essential for pharynx regeneration" rests (i) on the diagnosis of a "proliferative peak of FoxA+ stem cells that occurs after pharynx amputation (Figure 2C)" (line 202). However, rather than a "proliferative peak", Figure 2C shows a broad "proliferative plateau" of FoxA+ stem cells between 6h and 3 days after amputation. Similarly, the foxA+/H3P+ quantification after pharynx amputation in Figure 4G also displays a lack of a peak of foxA+/H3P+ from 1d to 2d after amputation. (Ii), the associated nocodazole experiments suffer from the fact that the authors did not quantify the impact of the drug on the abundance of foxA+/piwi-1+ cells during treatment intervals from 0-1d and 2-3d after pharynx amputation. Therefore, the authors cannot rule out that nocodazole treatment might have similar effects on the abundance of foxA+/piwi-1+ cells throughout the 1-3 d post-amputation time interval, with the more severe organ-level phenotype of the 1-2d treatment window being caused by some other effect of the drug (e.g., on the differentiation of another rate-limiting cell type for pharynx regeneration or, conceivably, inhibition of priming neuronal activity ). Similar concern apply regarding the statement in line 242, "… a window 1-2 days after amputation in which activation of ERK signaling is important for pharynx regeneration." Here, (i) the quantification of the end-stage phenotype of drug treatment during the 1-2d time interval (regain of feeding ability) is missing. (ii) Similarly, the examination of the consequences of PD treatment on foxA+ expression in piwi-1+ cells in panel 4D-H employs drug soaking for 3 days, yet the corresponding end-stage phenotype of 3-day drug treatment is not shown. (iii) the implications of ERK in pharynx regeneration are tentative. Even though the PD compound is initially correctly introduced as "MEK inhibitor", the authors subsequently switch to the factually wrong "ERK inhibitor" designation (e.g., line 358). Further, additional experimental evidence for the assumed Erk inhibition as the cause of the observed phenotypes would be desirable to rigorously support the conclusion.

These caveats need to be addressed if a cell biological timeline is to remain part of this manuscript.

3. Integration with the existing literature

The authors need to better integrate their findings with the literature. First, they need to cite the findings of Vu et al., which explicitly demonstrated a specific increase in the fractional abundance of piwi-1+/protonephridial marker+ cells in response to RNAi-mediated damage to protonephridia(Vu et al., 2015). As such, this study already demonstrates the main point of Bohr et al., namely that the planarian stem cell system is capable of "target conscious" progenitor provision. In the very least, the authors should credit these results as additional evidence for their model. A further finding that they should discuss is the demonstration by LoCascio et al. (LoCascio et al., 2017) that flank region cut-outs cause a significant increase in pharynx cell incorporation over baseline, despite the absence of injury to the pharynx. How do the authors reconcile the discrepancy between these data and their own? In general, the discussion would benefit greatly from a more explicit comparison between the "target blind" model versus their data, as well as a broader perspective on the regulation of stem cell homeostasis.

https://doi.org/10.7554/eLife.68830.sa1

Author response

All three reviewers recognized the potential significance of this work, but also shared the same concerns about sample sizes, lack of biological replicates, and insufficient details about cell quantification. Given the interest in the question being pursued, if these issues can be addressed satisfactorily, a revised paper could be considered here as a new submission. We have included the reviews below and hope you will find the comments helpful.

Reviewer #1:

This manuscript by Bohr et al. explores how planarian stem cells respond to the loss of a specific organ: the pharynx. The previously proposed "target-blind" model of planarian regeneration (LoCascio et al. 2017) posited that stem cells do not respond directly to missing tissues, but rather replace missing cell types based on their normal rates of homeostatic turnover. In contrast, Bohr et al. suggest that planarian stem cells can sense and respond to the loss of specific missing tissues, using the pharynx as a case study. The authors conclude that planarians may use more than one mode of regeneration, depending upon the target being regenerated (eye vs. pharynx).

The question explored in this paper is of fundamental importance, and providing an alternative model by which planarian stem cells regenerate missing tissues should be of interest to a broad readership. Unfortunately, in its current form, the manuscript presents enticing preliminary findings, rather than robust experimental observations. Currently, the manuscript has limited samples sizes and experimental replicates, which is unfortunate. Because this paper is attempting to refute a previously published model it is critical that the data are clear and convincing. If not, these findings could be summarily dismissed without appropriate debate. If the authors can show the robustness and rigor of their results, and address the major issues listed below, this manuscript would represent a significant contribution to our understanding of planarian regeneration.

We thank the reviewer for recognizing the importance of the questions we have pursued here. However, we would like to clarify that our intention was not to refute the previously published model. This confusion likely arose because of a lack of clarity in our writing. In the current version, we explicitly test the ‘non-targeted’ model with EdU incorporation (Figure 1E,F and Figure 1—figure supplement 1) and flank resection experiments (Figure 2D). Our data shows that the ‘targeted’ model is the primary mode by which the pharynx regenerates, but the ‘non-targeted’ model also operates immediately after wounding. We have added an entire section in the discussion to make it plainly apparent to readers that both mechanisms likely operate during regeneration.

1. Throughout the manuscript, experiments were either not repeated, or the number of biological replicates was not reported. In most cases, it appears that experiments were done only once (with the exception of the drug treatments). Numbers of biological replicates and sample sizes should be explicitly stated and the data from different replicates reported for Figures 1D-G, 2B-D, 3C, 3E-F, 4B, 4D-H, 5C-D, and 6A-E.

We agree that the manuscript suffered from a general lack of experimental replicates and small sample sizes, as addressed in the overview above. Taking into account these essential criticisms, we have repeated each experiment 2-3 times, increased overall sample sizes, and explicitly marked distinct biological and technical replicates on graphs, where possible. We have also included a table detailing biological and technical replicates for each relevant figure.

2. The authors do not sufficiently describe their methods for imaging and quantifying cells (Figures 1E, 1G, 2C-D, 3F, 4E-H, 5D, 6B). The size of the area covered to collect these data is unclear. High-magnification images are shown: are these the areas that were imaged? If so, their results could be biased by choosing small regions of interest. Ideally, the authors should quantify more than one region per animal. Also, they do not describe the depth of the z-stacks collected or how these stacks were normalized/standardized across conditions. All their conclusions hinge on the quantification of progenitor populations in response to different amputation paradigms or chemical treatments, so the standards for imaging and quantification must be clearly reported.

We agree that the previous version of the manuscript lacked clarity with regard to imaging and quantification. We have now addressed these important criticisms by clearly describing them in the figure legends and the Image acquisition, quantification and statistical analysis section of the methods (beginning on line 759). We now explicitly state the following standardized parameters for a given experiment: 1) the region analyzed within the animal, 2) the x-y area measured, 3) thickness (z-sections) for a given experiment, and 4) the z-section where quantification began.

3. Inappropriate statistical tests were used throughout. The use of multiple t-tests amplifies the chance of a Type I error and is especially problematic when up to 7 comparisons were made! The authors should use one-way ANOVA with multiple comparison corrections for all experiments with more than two groups.

As suggested, all figures in which multiple comparisons are made now use one-way ANOVA to determine statistical significance.

4. Figures 1D-E show that upon pharynx amputation but not head amputation, FoxA+ piwi+ pharynx progenitors increase. These data suffer from the quantification issues highlighted above: how the data were quantified is not sufficiently described, only 3 data points were taken (one per animal), the experiment appears to have been performed only one time, and the wrong statistical test was used. Rather than reporting the number of FoxA+ piwi-1+ cells counted, the authors should quantify the total number of doubly positive cells as a percentage of piwi-1+ cells, as was previously published (Adler et al. 2014). The authors also fail to specify whether the change observed between "3 dpa phx" and "3 dpa head" is significant, which is a material point.

We have addressed the concerns with biological replicates, sample sizes and statistical analysis as described in our responses to points 1 and 3. We have also attempted to indicate when relevant differences are not statistically significant, as is the case for the comparison in question above.

As for representing our data as the absolute number of cells rather than as a proportion, the reviewer is correct in noting that this strategy is a departure from our previous paper, where FoxA+piwi-1+ cells were represented as a relative percentage (Adler et al., 2014). However, similar quantification of ovo+ eye progenitors is represented as absolute numbers rather than percentages (Lapan et al. and LoCascio et al.). Without clear quantification standards in the literature, we initially evaluated both strategies, but found the results to be very similar. We have included two figures of side-by-side comparisons of these two quantification strategies for key panels from our manuscript (Author response images 1 and 2). Given the complex dynamics of the stem cell population, we feel that representing the data as relative percentages oversimplifies our analysis. Because our progenitor analysis initially compares multiple injury scenarios, we are confident that the changes we see in progenitors occur independently of broad injury responses. Further, representing the data as means of absolute numbers of cells in a standardized area/thickness allows us to indicate individual animals and biological replicates on graphs, providing a more transparent view of the data to readers. Based on these criteria, we have maintained our strategy of representing the data as absolute values, along with extensive additions to the manuscript clarifying where and how the data were obtained.

Author response image 1
Pharynx loss selectively increases pharynx progenitors in proportion to stem cells.

(A) Proportion of cells double positive for the indicated progenitor marker and piwi-1+ relative to all piwi-1+ stem cells in the area outlined by dashed boxes in cartoons. Cartoons depict different amputation conditions. n ≥ 790 cells per experimental group from 3 independent experiments. (B) Average number of FoxA+ piwi-1+ cells in the same animals and regions as A. Same data as is in Figure 2B, E of manuscript. (C) Proportion of FoxA+ piwi-1+ cells at indicated times post-pharynx amputation relative to all piwi-1+ stem cells in the area outlined by dashed boxes in A. n ≥ 631 cells per experimental group from 3 independent experiments. (D) Average number of FoxA+ piwi-1+ cells in the same animals and regions analyzed as C. Same data as in Figure 2C of manuscript. For all graphs a 6000μm2 region in the same location of the pre-pharyngeal region was analyzed over 20 z-sections, as represented by dashed boxes in A. Graphs represent a proportion ± 95% confidence intervals (A, C) or the mean ± SD with symbols = individual animals; shapes distinguish biological replicates (B, D). *, p ≤ 0.05 **, p ≤ 0.01; ***, p ≤ 0.001; ****, p ≤ 0.0001, Fisher’s Exact Test (A, C) or one-way ANOVA with Tukey test (B, D).

Author response image 2
Pharynx tissue loss selectively increases mitotically active pharynx progenitors.

(A) Proportion of FoxA+ H3P+ cells relative to all H3P+ stem cells at indicated times after pharynx or head amputation in the area outlined by dashed boxes in E. n ≥ 515 cells per experimental group from 2 independent experiments. (B) Average number of FoxA+ H3P+ cells quantified in the same animals and regions as A. Same data as is in Figure 3C, D of manuscript. (C) Proportion of cells double-positive for the indicated progenitor marker and H3P+ relative to all H3P+ stem cells in the area outlined by dashed boxes in E. n ≥ 472 cells per experimental group from 2 independent experiments. (D) Average number of cells double-positive for the indicated progenitor marker and H3P+ quantified in the same animals and regions as C. Same data as in Figure 3E of manuscript. (E) Cartoons depicting different amputation conditions. For A-D, the entire pre-pharyngeal region was analyzed over 30 z-sections, as represented by dashed boxes. Graphs represent a proportion ± 95% confidence intervals (A, C) or the mean ± SD with symbols = individual animals; shapes distinguish biological replicates (B, D). *, p ≤ 0.05 **, p ≤ 0.01; ***, p < 0.001; ****, p ≤ 0.0001, Fisher’s Exact Test (A, C) or one-way ANOVA with Tukey test (B, D)

5. Figure 2D also suffers from the inadequate quantification practices described above. Ideally, FoxA+ cells should be quantified as a percentage of the H3P+ cells observed.

We have addressed this concern as described to points 1-4 above.

6. The authors use "stem cell", "progenitor", "stem cell progenitor", and "progenitor stem cells" in a mixed and confusing way throughout the paper. For example, in lines 174-175 the authors state that "proliferation of FoxA+ stem cells precedes the increase in pharynx progenitors." This refers to FoxA+ H3P+ cells vs. FoxA+ piwi-1+ cells, but the only difference is that the former are stem cells in the act of mitosis. Is a distinction being made? Elsewhere in the paper, FoxA+ piwi+ cells are referred to as stem cells. The terminology used needs more clarity and consistency.

Along these lines, what is a FoxA+ PSC (Line 168)? Are the authors suggesting that FoxA is a pluripotency marker? Or are the authors saying that tgs-1 has a more expansive expression pattern and is co-expressed with progenitor markers? If double FISH was performed with the progenitor markers reported (ovo, myoD, gata-4/5/6, six-1/2, and pax6), would they all overlap with tgs-1? These experiments need to be performed to make any claims about FoxA expression in the context of pluripotency.

We agree that our language, particularly with regard to terminology surrounding stem cells, progenitors, and organ progenitors lacked consistency. We have now modified the language in the following ways: 1) we refer to “stem cells” as piwi-1+ or H3P+ cells, 2) we use the term “organ-specific progenitors” to refer to cells double-positive for progenitor markers and either H3P or piwi-1 as labeled in Figure 2 and 3, and 3) we removed all instances of the confusing terms “stem cell progenitor” and “progenitor stem cells” throughout. In addition, we removed the tgs-1 data and any claims about pluripotent stem cells, which we agree were preliminary and inconclusive.

7. In Figure 3C, how does pharynx regeneration occur over such a long period of time after nocodazole treatment in the 1-2 day window? Does target-blind regeneration occur once this window is missed? The authors should repeat analysis of FoxA+ progenitors at later time points in this condition, and/or show rates of BrdU incorporation into the pharynx with and without nocodazole treatment in this window.

We agree that it would be valuable to know how animals eventually regenerate after perturbation of pharynx progenitors and whether non-targeted mechanisms take over to slowly regenerate this organ. However, we have struggled to devise an experiment to clearly define these parameters. Nocodazole treatment during the 1-2 day window likely results in culling of pharynx progenitors by apoptosis. Because progenitors are not undergoing mitosis at equivalently high rates in intact animals, nocodazole treatment would not perturb them similarly, making comparisons difficult. Therefore, administration of BrdU during this window could potentially label completely distinct stem cell populations.

8. The use of the inhibitor PD in Figure 4 is problematic. No data are shown to verify that phosphorylation of ERK was inhibited in these experiments. A citation of previous use is not sufficient. The effect of PD on WT uncut animals with regards to FoxA+ cells is not shown and is a necessary control. To address questions of drug specificity, the authors should corroborate their findings with a second inhibitor of ERK signaling like U0126, which has already been shown to work in planarians (Owlarn et al. 2017).

We have addressed all the concerns raised above. 1) All MEK inhibitor experiments have been repeated with UO126, with comparable results (Figure 5—figure supplement 1C-F, Figure 5—figure supplement 3B-E, Figure 6). 2) A western blot verifies that MEK inhibitor treatment for as little as 24 hours prevents ERK phosphorylation (Figure 5—figure supplement 1E). 3) We now include data showing that exposure to either inhibitor blocks regeneration in tail fragments, indicating that the pharynx regeneration defect is not due to inefficient inhibition (Figure 5—figure supplement 1F). 4) We now include quantification of FoxA+piwi-1+ in intact animals after treatment with PD and UO (Figure 5—figure supplement 2A).

9. The conclusion that ERK signaling functions in regulating differentiation but not proliferation is premature (line 249-264). Figures 4E-H should be quantified as percentages of piwi-1+ and H3P+ cells, especially since there is decreased proliferation overall with PD treatment.

In the current version of the manuscript we have made every effort to dissect the role of ERK signaling in stem cell division versus differentiation. We have now extensively tested the timeline for ERK requirement in pharynx regeneration. Based on either PD or UO exposure (Figure 5C-H, and Figure 5—figure supplement 1 and 3), our data now clearly shows that ERK activity is not required prior to the stem cell division that occurs 1-2 days after pharynx amputation. Further, even though it is true that PD and UO exposure do reduce overall stem cell division, these drug treatments do not impact division of pharynx progenitors (Figure 5H and Figure 5—figure supplement 3D,3E). Because analysis of FoxA+ H3P+ cells and total H3P+ cells are done in the same animals, one can extrapolate that the proportional data would look similar. In fact, a decrease in overall H3P would inflate the relative numbers of FoxA+ H3P+ cells. We have added text to clarify this in the Results section (line 426-429).

10. The use of head fragments to compare eye regeneration vs. pharynx regeneration is inappropriate. Previous studies have already shown that the absence of eyes is not required to induce ovo+ progenitor amplification (LoCascio et al. 2017). Thus, this is not a surprising result (line 343-345) and the authors are mis-citing previous observations (in the earlier work, head fragments were never described). The region where ovo+ cells was quantified in Figure 6A is not justified or explained. The yellow box is placed in a medial region where ovo+ cells do not normally reside. The authors should image within the laterally positioned ovo+ streams that have been previously described (Lapan and Reddien 2012; LoCascio et al. 2017).

We agree that this experiment did not add much to the overall findings, nor did we adequately explain our rationale for including it. It has been removed from this version.

11. Rather than using head fragments, the authors should repeat the flank resection experiments shown previously (LoCascio et al. 2017). This previous study showed that increased BrdU incorporation into the pharynx occurred following flank resection even though the pharynx was present. That result may have been 1) an artifact of increased BrdU staining due to stimulation of proliferation upon injury, 2) caused by unintentional damage to cells associated with the pharynx, or 3) a response to the loss of FoxA+ progenitor populations that surround the pharynx rather than the loss of the differentiated organ. The authors have the opportunity to revisit this published observation by quantifying the FoxA+ progenitor response during flank resection +/- pharynx. Without these data, this story is incomplete and therefore the conclusion of a targeted regeneration response is not yet convincing.

We appreciate the reviewer’s suggestion to include this important experiment. We now include quantification of FoxA+piwi-1+ cells after flank resection (Figure 2D), which proves that injuries outside of the pharynx do not trigger an increase in pharynx progenitors. Additionally, we administered EdU at different times (0hr or 24hr) after head or pharynx amputation (Figure 1E,F). We found that cells generated immediately (0hr) after amputation are incorporated broadly, reinforcing the (LoCascio et al. 2017) result that flank resection increases BrdU labeling into the uninjured pharynx. Importantly, by administering EdU 1 day after amputation, we found that cells generated at this time were channeled specifically into regenerating organs, providing evidence supporting a targeted mechanism for regeneration that selectively produces progenitors of missing tissues.

12. The negative results that proliferation and ERK signaling are dispensable for eye regeneration in Figure 6 are weak and unconvincing. The regenerated eyes appear smaller; this should be quantified (number of PRNs per eye). If the small pharynges that form in Figures 3D and 4C are considered a deleterious phenotype, why is the same standard not applied to the eye? Also, existing eye progenitors could have been sufficient for eye regeneration in these drug treatments. Furthermore, eyes did not regenerate after nocodazole treatment 50% of the time. Is it not more likely that the observations reported are dosage and timing artifacts? How has proliferation been affected? These observations do not live up to the claims made.

We have addressed potential issues with dosage and timing by standardizing the exposure times for nocodazole experiments for 2 days, increasing animal numbers, and improving the quantification. Proliferation and ERK activity are only required for eye regeneration in the context of larger amputation, but not after eye resection alone (Figure 6). Regarding the pharynx, laminin is expressed strongly in the pharynx and mouth, and weakly in the body where the pharynx attaches. Even though there is residual laminin staining 7 days after amputation with PD, UO, or nocodazole exposure (Figure 4D, 5D, Figure 5—figure supplement 1D), this is not at all a ‘pharynx’. The normal architecture, which is apparent in the DMSO controls and in the supplemental figures associated with Figures 4 and 5, is completely lost after these treatments. We have clarified this in the writing (line 290-296). Conversely, eye regeneration in drug-treated animals is undoubtedly more comparable to controls (Figure 6B).

Eye regeneration is mediated by homeostatic turnover, but whether existing eye progenitors are sufficient to regenerate these small structures after drug treatments remains unclear. We have added text in this section to address this possibility (line 450-453).

13. The authors claim that ovo+ cells are not proliferative (H3P+) even in cases where there is eye progenitor amplification (head amputation), but the data are not shown (line 321). They should be. Indeed, previous publications have never shown that ovo+ cells proliferate. This might mean that there are proliferating eye progenitors that precede expression of ovo. The authors should discuss this alternative.

We have now included images for ovo+ H3P+ cells in intact animals and after head amputation with or without nocodazole treatment to enrich for cells in mitosis (Figure 3—figure supplement 3). We were only able to detect dividing ovo+ cells after nocodazole treatment, but did not observe any increase after head amputation as compared to intact nocodazole treated controls. As suggested, we added discussion of the possible explanations to this section (line 265-267).

14. The authors claim that eye regeneration does not require proliferation or ERK signaling but pharynx regeneration does. This conclusion hinges on the gross observation that eyes can regenerate in the presence of nocodazole and PD (see point 12 above). These data are coarse and the interpretations are unconvincing. Instead, their model can be directly tested using BrdU pulse-chase experiments. According to the authors' model, one would predict that following pharynx amputation, the rate of incorporation of BrdU+ cells into the regenerating pharynx should be higher than in uninjured controls. Conversely, the rate of BrdU incorporation into the regenerating eye should remain unchanged between injured (i.e. eye-resected) and control animals (LoCascio et al. 2017). Once the authors have established the prediction above, they have the opportunity to show the effects of nocodazole, PD, and U0126 on BrdU incorporation in the regenerating eye vs pharynx following eye resection and pharynx amputation in the same animal. This way, the authors can directly test the requirement for proliferation and/or ERK signaling in both tissues.

We agree with the reviewer that this is an important experiment, and we have now addressed the different contributions of EdU+ cells into either the regenerating pharynx or the brain (Figure 1E,F). However, we have reserved the pharmacological perturbations for future work.

Reviewer #2:

Bohr, Shiroor, and Adler investigate how stem cells respond to the loss of specific tissues in planarians. The planarian stem cell population (neoblasts) are distributed throughout the planarian body and include pluripotent stem cells and a wide range of lineage-committed progenitor cells. How this heterogenous pool of cells behave post-injury or amputation is incompletely understood. The discovery of markers to label stem cell progeny have opened the door to investigate stem cells respond to tissue loss. However, the anatomy of planarians makes it difficult to surgically remove or damage specific organs. The PI of this study developed an assay to remove the pharynx by "chemical amputation" to study the mechanisms underlying regeneration of this organ without drastically perturbing or injuring other tissues. Using this approach, this paper investigates how a well-defined population of FoxA+ progenitors respond to pharynx removal at early time-points during the regeneration. Their data suggest that stem cells are able to detect loss of the pharynx and respond by generating significantly more cells fated to become pharynx, whereas amputation of non-pharyngeal tissues does not have an obvious effect on pharynx progenitor specification dynamics. In addition, using pharmacological treatments, the authors show that cell proliferation and ERK signaling are required for the expansion of pharynx progenitors and cell differentiation. In contrast, other cell types in the planarian eye do not appear to require proliferation or ERK signaling, suggesting that stem cell responses "target blind" as suggested in a previous study, but are rather tuned to specific missing tissues.

This work has the potential to make a significant contribution to the field by advancing our understanding of how the planarian heterogenous stem cell population responds to the loss of a specific organ. However, the report is preliminary as presented. It appears that the authors performed many experiments a single time. In addition, description of the methods is insufficient. Consequently, before this work can be considered for publication, the authors need to chiefly demonstrate the reproducibility of the data and robustness of the observations.

1. The authors need to replicate experiments to increase the sample size for most experiments.

We have addressed this concern, as detailed in the overview above. All experiments have been repeated, increasing the sample sizes across the board. We have also included a table detailing biological and technical replicates for each relevant figure.

2. Details for imaging and quantification should be explicitly stated in the methods, and the reported cell count numbers should be normalized as appropriate for each set of experiments.

We have addressed this concern, as detailed in the overview above. Briefly, imaging and quantification methods are now described in detail within the figure legends and methods section entitled Image acquisition, quantification and statistical analysis (beginning on line 759). Regions analyzed were either of standard area or normalized to area, and for a given confocal imaging experiment, the numbers of z-sections were kept consistent.

3. Although the authors mention "lineage-tracing" experiments, they do not perform DNA analog pulse-chase experiments to analyze a temporal progression and spatial localization of stem cells to FoxA+ progenitors after pharynx removal. The authors rely on PH3-staining in conjunction with FoxA, and supplementary experiments using the pluripotent stem cell marker tgs-1 (which was only examined at 1 dpa). Could the authors clarify what they think FoxA+ stem cells represent? Are these self-renewing pluripotent stem cells or lineage-committed progenitors? Can the authors get some insight by scanning their images of PH3+ cells expressing FoxA visibly undergoing metaphase? Are daughter cells uniformly FoxA+ as reflected in the model? At least in one of the cells shown in the nocodazole treated controls it appears that both daughter cells express FoxA (Figure 3). I suggest showing some higher magnification images to support the interpretations/conclusions. Others have posited (e.g., Rink, Chapter 2 of Planarian Regeneration: Methods and Protocols), that not every dividing cell may be a long-term self-renewing stem cell and whether a transient amplifying cells exist or contribute to regeneration in planarians is unknown. Adler and Sánchez Alvarado (2015) discuss the role of transient states and how the transcriptional profiles change in response to regeneration. It wasn't clear to me how the authors think about these cells based on the limited number of experiments and analysis, and there are a few places where the terminology is inconsistent, especially in reference to proliferating ovo+ progenitors (P. 14). The authors need to be clear, and it might be helpful to illustrate their model in one of the early figures or to include it in the final model, which omits tgs-1 due to the limited number of experiments performed with this marker gene (Figure 7).

We appreciate the suggestion to clarify our language surrounding stem cells, pluripotent stem cells, and progenitors. To address the important point regarding the temporal progression of pharynx progenitors, we now include DNA analog (EdU) pulse-chase experiments to analyze the output of stem cells generated either immediately after amputation, or 1 day later when we see a specific increase in pharynx progenitors. This experiment (Figure 1E,F) shows that stem cells produced 1 day after amputation are selectively incorporated into the regenerating pharynx. As addressed in the overview above, point 4, we have made our terminology throughout the manuscript more consistent.

We agree with the reviewer regarding the complexity of the stem cell population, which includes both long-term self-renewing cells and progenitors. Because we could not make clear conclusions based on the tgs-1 data, we removed this figure and any claims about pluripotent stem cells from this version of the manuscript. Although we do not have data to definitively indicate the potency of FoxA+ progenitors, we now include images of anaphase cells that suggest that FoxA can segregate into daughter cells both symmetrically and asymmetrically (Figure 3—figure supplement 1A). We have added a section addressing these issues to the discussion, because we agree that investigating these questions is important (lines 511-528).

4. The pharynx is complex and there is no data to assess what the contribution of other progenitor populations might be. I don't think or think it is unlikely that FoxA+ progenitors are solely responsible for reconstructing the pharynx. The authors should examine how other progenitor populations behave during the process of pharynx regeneration by extending the timeline of progenitor cell analysis. This would reveal if there is fluctuation in progenitor dynamics as animals regenerate the pharynx or re-scale proportions after pharynx regeneration. For example, can the authors test if they are able to detect a contribution of neural progenitors to regeneration of the pharyngeal nervous system? And if so, when during the regeneration process does it take place in the context of their study?

The reviewer is correct in noting that we still lack definitive proof of whether FoxA+ progenitors are the sole source of pharyngeal tissue or only generate a subset of critical cells. The pharynx is indeed a complex organ and we would love to know whether FoxA is a master regulator of them all. To address this point, we have knocked down every progenitor included in this paper and every proposed pharynx progenitor from the literature (dd554, meis, twist) and none of them seem to play any role in pharynx regeneration (Figure 2—figure supplement 3 and data not shown). The lack of a detectable phenotype from these markers has made us reluctant to draw conclusions from their dynamics during pharynx regeneration.

Reviewer #3:

In this manuscript, Bohr et al. examine how the pluripotent stem cell system of planarians responds to organ-specific damage. If and how the differentiation of specific cell types is dynamically regulated is a conceptually fascinating problem in planarians and in general stem cell research. The authors address this problem by comparing the stem cell response between a single-organ amputation (the pharynx) versus broad tissue loss (decapitation). Their findings indicate that only the removal of the pharynx triggers the increased differentiation of pharyngeal cell types, while the loss of non-pharyngeal tissues upregulates the differentiation of progenitors of multiple organs, but not the pharynx. Further, the authors implicate temporally restricted ERK signaling as a regulatory component in the differentiation of pharyngeal cell types. These observations are also important because they contrast with the previously proposed "target blind" model (LoCascio et al., 2017) that posits the differentiation of different cell types at constant relative proportions, with the rate of stem cell divisions as global production rate regulator. In contrast, the observations by Bohr et al. provide further evidence for more flexibility and specificity within the planarian stem cell system ("target consciousness") in the sense of lineage-specific adjustments in the production rates of specific cell types.

That said, the manuscript generally suffers from an overly narrow focus. Important questions remain regarding the specificity of the stem cell response to pharynx amputation and multiple experiments lack important controls (see below). Moreover, the authors have overlooked that a "target conscious" progenitor response has already been demonstrated by the selective proliferation of protonephridial marker expressing neoblasts in response to protonephridial damage by RNAi (Vu et al., 2015). As a result, the current manuscript would require substantial additional experimentation to consolidate its findings sufficiently for publication in eLife.

We appreciate the reviewer’s recognition of our work and their thoughtful criticisms. In the revised version we have added new controls, included the important reference of Vu et al., 2015, and extensively discussed the conflicting models for regeneration (referred to in our manuscript as ‘targeted’ vs ‘non-targeted’). Addressing these criticisms has resulted in a substantially more clear manuscript that adds to our understanding of the mechanisms underlying regeneration.

1. Specificity of the stem cell response

The central premise of the paper is the selective amplification of pharyngeal progenitors in response to pharynx amputation. This the authors conclude on basis of i) an increase in the absolute number of foxA+/piwi-1+ cells in a specific area, while ii) de-capitation has no effect on the absolute number of foxA+/piwi-1+ cells in the same area. This approach is an insufficient demonstration of specificity, as the known phenomenon of wound-induced stem cell activation might also change the absolute number of specific neoblast subclasses and might do so in an injury-dependent manner.

To account for this important caveat, the authors need to i) quantify RELATIVE proportions of foxA+/piwi-1+ cells out of total piwi cells (or of total H3P+ cells) and ii) they need to include other organ progenitors in the initial analysis. The latter is also critical because the pharynx is a complex organ comprising descendants of multiple lineages (e.g., muscle, neurons, epidermis) and it is not clear whether the foxA+/piwi-1+ cells indeed serve as a singular origin of all constituting lineages (as assumed by the authors), or if they only provide a subset of pharyngeal cells with a rate-limiting role in pharynx assembly (e.g., pharyngeal muscle). In face of such uncertainty, iii) the quantification of new cell incorporation into the pharynx versus other tissues via BrdU labeling would be necessary to address this caveat and to provide a global perspective on the specificity of the response independent of incompletely characterized marker genes.

In addition, the following experimental design problems need to be addressed or better documented, including:

• The authors provide insufficient methodological detail on progenitor quantifications, even though the entire manuscript rests on this assay. What are their criteria for scoring a piwi-1+ cell as double-positive for the often weak and noisy lineage labels? If done "by eye", was double-blind scoring used? Were all cells in a given Z-stack counted or only specific planes? If the latter, by which criteria were image planes selected for quantification? How were the specific specimens out of an experimental cohort selected for imaging/quantification? Though not necessarily a unique shortcoming of this particular study, these points simply need to be adequately addressed in order to rigorously support quantitative differences between experimental conditions (e.g., specificity).

• The authors appear not to distinguish at all between technical replicates (e.g., multiple specimens within an experimental cohort) versus biological replicates (independent experimental cohorts). This is significant, because i), the use of the standard error of the mean (SEM) that the authors employ throughout is not really an appropriate measure for a single biological replicate with 3 animals – the standard deviation (SD) would seem a more appropriate measure in this context (SD). ii), the number of worms quantified for each experiment is generally low (n=3 animals in Figure 1, 3, 5, 6; n=5 animals in the rest of the figures) given the observed variability in the data (e.g., ~25-30 foxA+/piwi-1+ cells 3d after pharynx amputation in Figure 1E versus 50 cells in figure 1H). Similarly, for kinetic experiments as in Figure 1H or 2C, it is simply crucial to ensure that the error bars include the variation in response dynamics between multiple replicates due to the drift in the baseline fraction of H3P+ cells or varying staining efficiencies (e.g., different batches of animals on different days), rather than the technical variation in a single experimental cohort only. Please address these concerns by adding more specimens and a thorough description of the experimental design.

We thank the reviewer for highlighting these important caveats with our interpretation of our data. In point 2 of the introduction to this response, and in Author response figures 1 and 2, we have included a side-by-side comparison of absolute vs relative quantification and show that for key figures in our manuscript, the two strategies yield very similar results. In addition, because we are comparing different wounding paradigms that induce general wound responses, our experiments are internally controlled for wound-induced proliferation.

The reviewer is correct in noting that we still lack definitive proof of whether FoxA+ progenitors are the sole source of pharyngeal tissue or only generate a subset of critical cells (also addressed in reviewer 2’s major concern 4 above). To address this point, we have knocked down every progenitor used in this paper and every proposed pharynx progenitor from the literature (dd554, meis, twist). None of them cause defects in pharynx regeneration (Figure 2—figure supplement 3 and data not shown). In the current version of the manuscript, we also include pulse-chase labeling with EdU to demonstrate that stem cells are indeed selectively incorporated into the regenerating pharynx, depending on when they proliferate relative to tissue removal (Figure 1E,F). However, we acknowledge the caveat that FoxA may only control the production of a subset of cells. Ongoing work not included here is aimed at addressing this caveat.

We have also corrected all of the flaws the reviewer has pointed out related to replicates and statistical analysis in this version as outlined in the introduction to this response, point 1. Briefly, these include: 1) clarifying and standardizing methods for imaging and quantification, 2) distinguishing biological vs technical replicates in all experiments, 3) replaced error bars representing SEM with error bars representing standard deviation, and 4) increased number of animals in every experiment.

2. Timeline of pharynx regeneration

The pharynx regeneration timeline and associated events that the authors present are insufficiently supported by experimental data. The conclusion in line 215 "that proliferation in a critical window of 1 to 2 days after pharynx amputation produces a population of progenitors that are likely essential for pharynx regeneration" rests i) on the diagnosis of a "proliferative peak of FoxA+ stem cells that occurs after pharynx amputation (Figure 2C)" (line 202). However, rather than a "proliferative peak", Figure 2C shows a broad "proliferative plateau" of FoxA+ stem cells between 6h and 3 days after amputation. Similarly, the foxA+/H3P+ quantification after pharynx amputation in Figure 4G also displays a lack of a peak of foxA+/H3P+ from 1d to 2d after amputation. Ii), the associated nocodazole experiments suffer from the fact that the authors did not quantify the impact of the drug on the abundance of foxA+/piwi-1+ cells during treatment intervals from 0-1d and 2-3d after pharynx amputation. Therefore, the authors cannot rule out that nocodazole treatment might have similar effects on the abundance of foxA+/piwi-1+ cells throughout the 1-3 d post-amputation time interval, with the more severe organ-level phenotype of the 1-2d treatment window being caused by some other effect of the drug (e.g., on the differentiation of another rate-limiting cell type for pharynx regeneration or, conceivably, inhibition of priming neuronal activity ). Similar concern apply regarding the statement in line 242, "… a window 1-2 days after amputation in which activation of ERK signaling is important for pharynx regeneration." Here, i) the quantification of the end-stage phenotype of drug treatment during the 1-2d time interval (regain of feeding ability) is missing. ii) Similarly, the examination of the consequences of PD treatment on foxA+ expression in piwi-1+ cells in panel 4D-H employs drug soaking for 3 days, yet the corresponding end-stage phenotype of 3-day drug treatment is not shown. iii) the implications of ERK in pharynx regeneration are tentative. Even though the PD compound is initially correctly introduced as "MEK inhibitor", the authors subsequently switch to the factually wrong "ERK inhibitor" designation (e.g., line 358). Further, additional experimental evidence for the assumed Erk inhibition as the cause of the observed phenotypes would be desirable to rigorously support the conclusion.

These caveats need to be addressed if a cell biological timeline is to remain part of this manuscript.

We thank the reviewer for their insightful criticisms. While we agree that the FoxA+ H3P+ peak may more closely resemble a shield volcano than a Matterhorn, regardless of terminology, there is an obvious and significant elevation in proliferation that occurs within two days of pharynx amputation. This conclusion has been strengthened by reviewers’ suggestions to boost sample size and alter statistical analysis methods. We now include additional characterization of FoxA+piwi-1+ pharynx progenitors after nocodazole treatment 0-1 and 2-3 days after pharynx amputation (Figure 4—figure supplement 2E), reinforcing our claims that proliferation occurring 1-2 days after amputation is critical for pharynx progenitor production. Additionally, this 1-2 day window is concurrent with the timing of selective incorporation of EdU+ into the regenerating pharynx, supporting our conclusion that this 1-2 day proliferative window is essential for pharynx progenitor production and subsequent regeneration.

Regarding ERK experiments, we have addressed every point raised by the reviewer by expanding our MEK inhibitor experiments to more thoroughly test when ERK is required during pharynx regeneration. This includes standardizing the length of MEK inhibitor treatments and performing many additional controls to show that multiple MEK inhibitors both prevent phosphorylation of ERK and lead to the same outcomes in regeneration. To bolster evidence supporting our timeline, we have done the following:

We now include data for the end-stage phenotype following both 1-2 day and 0-3 day PD exposure (Figure 5—figure supplement 3C), which both show a delay in regeneration. We also exposed animals to PD or the alternative MEK inhibitor U0126 for 24 hour intervals from 0-1, 1-2 or 2-3 days after pharynx amputation, and analyzed pharynx progenitor abundance. These experiments show that ERK inhibition between 1-2 days after amputation specifically impacts the increase of pharynx progenitors (Figure 5F, Figure 5—figure supplement 3B). Together, these experiments have strengthened our argument that ERK is required for pharynx progenitor production and pharynx regeneration.

We have modified our language to refer to the drug inhibitors as MEK inhibitors, but continue to refer to ERK inhibition, as application of MEK inhibitors does inhibit ERK activity (Owlarn et al. 2017) and phosphorylation, as shown in our western blot (Figure 5—figure supplement 1B).

3. Integration with the existing literature

The authors need to better integrate their findings with the literature. First, they need to cite the findings of Vu et al., which explicitly demonstrated a specific increase in the fractional abundance of piwi-1+/protonephridial marker+ cells in response to RNAi-mediated damage to protonephridia(Vu et al., 2015). As such, this study already demonstrates the main point of Bohr et al., namely that the planarian stem cell system is capable of "target conscious" progenitor provision. In the very least, the authors should credit these results as additional evidence for their model. A further finding that they should discuss is the demonstration by LoCascio et al. (LoCascio et al., 2017) that flank region cut-outs cause a significant increase in pharynx cell incorporation over baseline, despite the absence of injury to the pharynx. How do the authors reconcile the discrepancy between these data and their own? In general, the discussion would benefit greatly from a more explicit comparison between the "target blind" model versus their data, as well as a broader perspective on the regulation of stem cell homeostasis.

We thank the reviewer for this suggestion, and apologize for this oversight of our colleagues’ work. In the revised version, we have more thoroughly described and contextualized our findings with the existing literature. We added sections in the introduction, discussion, and figures that specifically address the contrast between existing models for regeneration (referred to in our manuscript as targeted vs non-targeted).

We have also included experiments analyzing FoxA+piwi-1+ cells after flank resection (Figure 2D), but do not observe any increase. We have reconciled this finding with the existing literature by performing EdU labeling after different amputations, somewhat similar to those performed in (LoCascio et al. 2017), but at different times. These experiments reveal that a targeted mechanism channels cells towards missing tissues one day after amputation (Figure 1E,F), but that a non-targeted mechanism also contributes to regeneration, potentially if small amounts of tissues have been damaged (like eye resections). The first section of the discussion extensively describes these two models and how they may be utilized in planarian regeneration.

https://doi.org/10.7554/eLife.68830.sa2

Article and author information

Author details

  1. Tisha E Bohr

    Department of Molecular Medicine, Cornell University College of Veterinary Medicine, Ithaca, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2336-407X
  2. Divya A Shiroor

    Department of Molecular Medicine, Cornell University College of Veterinary Medicine, Ithaca, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9085-5675
  3. Carolyn E Adler

    Department of Molecular Medicine, Cornell University College of Veterinary Medicine, Ithaca, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    cea88@cornell.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3883-0654

Funding

National Institute of General Medical Sciences (R01GM139933)

  • Carolyn E Adler

College of Veterinary Medicine, Cornell University

  • Divya A Shiroor

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

Acknowledgements

We thank the Cornell University Biotechnology Resource Center for assistance with data collection on the Zeiss LSM 710 Confocal which is supported by the NIH (NIH S10RR025502). We would also like to thank Kuang-Tse Wang and Justin Tapper for helpful comments on the manuscript and Melanie Issigonis for assistance with F-ara-EdU-labeled FISH. DAS was supported by a GRA Fellowship from the Cornell College of Veterinary Medicine. This work was supported by Cornell University startup funds and R01GM139933 awarded to CEA.

Senior Editor

  1. Didier YR Stainier, Max Planck Institute for Heart and Lung Research, Germany

Reviewing Editor

  1. Phillip A Newmark, Morgridge Institute for Research, United States

Publication history

  1. Received: March 26, 2021
  2. Accepted: May 22, 2021
  3. Version of Record published: June 22, 2021 (version 1)

Copyright

© 2021, Bohr 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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