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. 2013 Sep 10;2:e00247. doi: 10.7554/eLife.00247

Tissue absence initiates regeneration through Follistatin-mediated inhibition of Activin signaling

Michael A Gaviño 1,†, Danielle Wenemoser 1,†, Irving E Wang 1, Peter W Reddien 1,*
Editor: Marianne E Bronner2
PMCID: PMC3771573  PMID: 24040508

Abstract

Regeneration is widespread, but mechanisms that activate regeneration remain mysterious. Planarians are capable of whole-body regeneration and mount distinct molecular responses to wounds that result in tissue absence and those that do not. A major question is how these distinct responses are activated. We describe a follistatin homolog (Smed-follistatin) required for planarian regeneration. Smed-follistatin inhibition blocks responses to tissue absence but does not prevent normal tissue turnover. Two activin homologs (Smed-activin-1 and Smed-activin-2) are required for the Smed-follistatin phenotype. Finally, Smed-follistatin is wound-induced and expressed at higher levels following injuries that cause tissue absence. These data suggest that Smed-follistatin inhibits Smed-Activin proteins to trigger regeneration specifically following injuries involving tissue absence and identify a mechanism critical for regeneration initiation, a process important across the animal kingdom.

DOI: http://dx.doi.org/10.7554/eLife.00247.001

Research organism: Other

eLife digest

Most animals can respond to injury with some form of tissue regeneration. In mammals, this is limited to wound healing, whereas other vertebrates—such as salamanders and zebrafish—can regenerate parts of internal organs and even entire appendages. The planarian, a flatworm, is even more remarkable, being able to regenerate its head or tail following amputation, and even a whole animal from just a small body fragment. This is particularly impressive given that planarians have a complex internal anatomy, which includes muscles, intestines, a system similar to kidneys, and a central nervous system with a brain.

How is such regeneration accomplished? Why are planarians able to regenerate their bodies so extensively, whereas humans cannot? To what extent are the mechanisms of planarian regeneration common to other animals? These questions have driven the study of planarian regeneration for more than a century, but it is only in recent years that the tools needed to address these questions at the molecular level have become available.

Planarian regeneration proceeds over several days and involves multiple processes, including gene expression, cell division and cell death. Importantly, it has recently been shown that planarians activate different responses depending on whether an injury results in significant tissue loss—and therefore requires regeneration for repair—or if simple wound healing will be sufficient. The mechanisms behind these different responses to injury have, however, remained a mystery.

Now, Gaviño et al. have identified a key mechanism in the initiation of regeneration following tissue loss. This is centered on the gene follistatin, which is expressed following wounding. When genetic techniques are used to disrupt the expression of follistatin, regeneration is completely blocked. However, the animal’s ability to routinely replace old cells via a stem-cell mediated mechanism is unaffected. This indicates that follistatin is specifically required for the replacement of cells lost through injury. Gaviño et al. further demonstrate that the protein encoded by follistatin likely initiates tissue regeneration upon substantial tissue loss through inhibition of proteins called Activins.

Activin and Follistatin proteins are broadly conserved in evolution, and are also expressed in mammals, raising the possibility that similar molecular circuits may govern regenerative responses in many species.

DOI: http://dx.doi.org/10.7554/eLife.00247.002

Introduction

Regeneration occurs in widespread contexts and species. Invertebrates such as Hydra are capable of whole-animal regeneration from tissue fragments, and many vertebrates can regenerate appendages or repair damaged organs (Sánchez Alvarado, 2000). Despite this widespread relevance, the central mechanisms that drive regeneration are poorly understood.

Planarians are flatworms capable of regeneration following an almost limitless variety of injuries and have emerged as a powerful model for exploring the molecular underpinnings of regeneration (Newmark and Sánchez Alvarado, 2002). New tissues are formed at planarian wound sites in a process called blastema formation, and pre-existing tissues are reorganized after amputation to accommodate reduced animal size and further generate missing tissues (Morgan, 1901; Reddien and Sánchez Alvarado, 2004). The source of regenerated tissue in planarians is a population of adult dividing cells called neoblasts (Reddien and Sánchez Alvarado, 2004), which include pluripotent stem cells called clonogenic neoblasts (cNeoblasts) (Wagner et al., 2011). Neoblasts are the only somatic cycling cells in adult animals and can be specifically ablated by gamma irradiation, allowing for dissection of the requirements for neoblasts in regenerative processes (Reddien and Sánchez Alvarado, 2004). Recent work has described the earliest molecular and cellular events that occur following injury (Pellettieri et al., 2010; Wenemoser and Reddien, 2010; Sandmann et al., 2011; Wenemoser et al., 2012). One finding to emerge from this work is that animals initiate distinct cellular and molecular responses to ‘major injuries’ that remove significant amounts of tissue (e.g., head amputation) and to ‘simple injuries’ that require only minimal healing for repair (wounds that do not elicit blastema formation, such as punctures or incisions). Following simple injury, for example, animals display an increase in mitotic numbers 6 hr after injury before returning to baseline levels (Wenemoser and Reddien, 2010), and expression of numerous wound-induced genes becomes undetectable by 24 hr after injury (Wenemoser et al., 2012). Following a major injury, these same initial responses are observed, but subsequent responses are also activated: the 6 hr increase in mitotic numbers is followed by a second increase 48 hr after amputation (Wenemoser and Reddien, 2010), and wound-induced gene expression persists beyond 24 hr and is refined over several days (Wenemoser et al., 2012). These responses are referred to as the ‘missing-tissue response’ (Wenemoser and Reddien, 2010; Wenemoser et al., 2012). How animals distinguish between injuries involving varying amounts of tissue loss and regulate these distinct wound response programs remains unknown.

We identified Smed-follistatin as required for molecular and cellular ‘missing-tissue’ responses during regeneration. Specifically, Follistatin-mediated inhibition of Activin signaling is required for regeneration to occur, with Smed-follistatin expression at wounds controlled by the extent of tissue absence following injury. These results suggest a mechanism by which regenerative responses can be initiated.

Results

Smed-follistatin is a wound-induced gene required for regeneration

To identify genes mediating regeneration-specific wound responses, we inhibited recently identified wound-induced genes (Wenemoser et al., 2012) with RNA interference (RNAi). Inhibition of Smed-follistatin (follistatin or fst), a gene encoding a Follistatin-like TGF-β-superfamily inhibitor, completely blocked regeneration (Figure 1A, Figure 1—figure supplement 1). No brain regeneration or anterior pole regeneration was observed in fst(RNAi) animals (Figure 1A, Figure 1—figure supplement 2). The anterior pole phenotype is consistent with a described role for follistatin in anterior regeneration (Roberts-Galbraith and Newmark, 2013). fst(RNAi) animals, however, also failed to produce a blastema following either tail amputation or the excision of lateral tissue wedges that left anterior and posterior poles intact (Figure 1B). These data demonstrate that fst is required broadly for regeneration.

Figure 1. fst is wound induced and required for regeneration.

(A) fst(RNAi) animals did not form blastemas after amputation (left, arrowheads, n > 100) and did not regenerate a brain as assayed with an RNA probe for choline-acetyltransferase (chat, middle, arrowhead, n = 9/9). (B) fst(RNAi) animals did not form blastemas 8 days following excision of a wedge of lateral tissue (arrowhead, n = 14/14). (C) fst(RNAi) animals displayed no phenotype in the absence of amputation (right, n = 30/30, 123 days RNAi). (D) fst(RNAi) animals still produced new neurons 20 days after failing to regenerate (n = 8/8; cells that are SMEDWI protein+ and chat+ are newly differentiating neurons [Wagner et al., 2011]), demonstrating ongoing tissue turnover in old tissues. Scale bar 10 µm. (E) fst was expressed sparsely throughout the intact animal and at the anterior pole. Following head and tail amputation, robust fst expression occurred at both anterior and posterior wound sites, with peak expression 12 hr post-amputation. Scale bars = 100 µm for left of (A), (B), and (C); 200 µm for right of (A) and (E). Anterior up.

DOI: http://dx.doi.org/10.7554/eLife.00247.003

Figure 1.

Figure 1—figure supplement 1. Specificity of the fst RNAi phenotype.

Figure 1—figure supplement 1.

Animals injected with dsRNA corresponding to non-overlapping sequences from either the 5′ half or the 3′ half of the fst gene failed to regenerate (n = 6/14 and 4/16 normal, respectively, compared to 16/16 normal for control), day 8. Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.004
Figure 1—figure supplement 2. Anterior pattern defects in fst(RNAi) animals.

Figure 1—figure supplement 2.

fst(RNAi) animals displayed normal anterior sfrp-1 expression 24 hr after amputation (n = 11/11, top), but by 8 days displayed none (n = 11/12, middle). fst(RNAi) animals also failed to regenerate anterior ndk expression (bottom, n = 7/8). Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.005
Figure 1—figure supplement 3. Efficacy of fst RNAi.

Figure 1—figure supplement 3.

fst expression was reduced both prior to and following amputation in animals treated with fst dsRNA as assayed by quantitative PCR (qPCR). For qPCR, total RNA was isolated from asexual Schmidtea mediterranea animals, and cDNA was prepared using SuperScript III (Invitrogen) with oligo-dT primers. qPCR was performed using SYBR Green (Applied Biosystems), and data were normalized to the expression of clathrin as previously described (Eisenhoffer et al., 2008). fst (left: CCAGGCGAAAGAAATCCAG and right: GAGGTGGGGCATTTGATACA) primers were used to evaluate expression.
DOI: http://dx.doi.org/10.7554/eLife.00247.006
Figure 1—figure supplement 4. Wound-induced fst expression persists for several days after amputation.

Figure 1—figure supplement 4.

fst is expressed at wound sites throughout regeneration, with additional expression in the brain at later timepoints (arrows, 6 days and 8 days). Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.007
Figure 1—figure supplement 5. fst is required after amputation for normal regeneration.

Figure 1—figure supplement 5.

Animals were amputated and injected four times with 30 nl fst dsRNA immediately after amputation and a second time 6 hr after amputation. Animals injected with fst dsRNA developed aberrant brains as labeled by chat expression, and in some cases produced no blastemas (n = 7/10 aberrant, 1/10 no blastema), while animals injected with control dsRNA regenerated normally (10/10 normal). Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.008
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Planarians constantly maintain adult tissues through cell turnover involving neoblasts (Reddien and Sánchez Alvarado, 2004). Consequently, most genes required for regeneration are also required for tissue turnover because of an involvement in neoblast biology (Reddien et al., 2005). Strikingly, unamputated fst(RNAi) animals did not shrink or lose structures, as is typically seen in animals with neoblast dysfunction, even after several months of significant expression reduction with RNAi (Figure 1C, Figure 1—figure supplement 3). Furthermore, amputated animals—despite failing to regenerate—displayed ongoing long-term neoblast-based tissue turnover of remaining tissue (Figure 1D). Together, these data suggest that the requirement for fst in tissue replacement is specific to regeneration, as it is not detectably required for neoblast-mediated tissue turnover. Because of the rarity of genes required for regeneration but not tissue turnover, fst was a good candidate for specifically mediating the processes that occur following injury to bring about regeneration.

fst expression was induced at wounds by 6 hr following amputation (Wenemoser et al., 2012; Roberts-Galbraith and Newmark, 2013) and persisted for several days, with maximal expression around 12 hr post-amputation (Figure 1E, Figure 1—figure supplement 4). In unamputated animals, fst was expressed sparsely throughout the animal, including ventrally, in a thin peripheral domain, and at the anterior pole (Figure 1E, Figure 1—figure supplement 4; Roberts-Galbraith and Newmark, 2013). Injection of fst dsRNA only after amputation caused poor blastema formation and regeneration defects (Figure 1—figure supplement 5), consistent with a requirement for wound-induced fst expression in regeneration. We conclude that fst is a wound-induced factor required for regeneration.

follistatin is required for the regeneration-specific neoblast response

To characterize the defects underlying regeneration failure in fst(RNAi) animals, we first investigated whether fst regulates neoblast function in regeneration. Neoblasts can be visualized by detecting neoblast-specific transcripts through whole-mount in situ hybridization (Reddien et al., 2005) and quantified using flow cytometry (Hayashi et al., 2006). fst(RNAi) animals displayed normal neoblast numbers prior to amputation, indicating that the observed regeneration failure is not caused by neoblast loss (Figure 2A). We next assessed whether neoblasts respond to injury in fst(RNAi) animals. The neoblast response to injury involves two peaks (6 hr and 48 hr post-amputation) in mitotic cell numbers, in between which neoblasts migrate to wounds (Wenemoser and Reddien, 2010). The first peak is generically induced by all injury types and is spatially widespread (Wenemoser and Reddien, 2010). The second peak occurs specifically following major injuries (removing tissues) and is biased toward wound sites (Wenemoser and Reddien, 2010). Amputated fst(RNAi) animals displayed a normal 6 hr mitotic peak, indicating that a normal generic injury response was present (Figure 2B). By contrast, these animals failed to display a 48 hr mitotic peak (Figure 2B). fst(RNAi) animals did however display localization of mitoses toward wound sites 48 hr after amputation (Figure 2—figure supplement 1), and neoblast enrichment at wound sites 18 hr after injury (Figure 2C), indicating that neoblast migration occurred normally.

Figure 2. fst is required for the neoblast response to missing tissue.

(A) fst RNAi did not affect neoblast number or distribution as assayed with an RNA probe for smedwi-1 (n = 5/5) and flow cytometry (percentage of live cells that were X1 cells). (B) fst(RNAi) tail fragments displayed reduced mitoses 48 hr and 72 hr after amputation (right, p<0.01 and p<0.001, two-tailed t-test). (C) Neoblasts migrated to wounds in fst(RNAi) animals as assayed for the presence of smedwi-1+ cells at wounds (NB.21.11E+ cells mark pre-existing tissue). (D) fst(RNAi) animals lacked eye progenitors following head amputation as assayed with RNA probes for ovo+/smedwi-1+ cells (p<0.001, two-tailed t-test). Scale bars = 100 µm. Anterior up.

DOI: http://dx.doi.org/10.7554/eLife.00247.009

Figure 2.

Figure 2—figure supplement 1. Neoblasts migrate to wounds normally in fst(RNAi) animals.

Figure 2—figure supplement 1.

Mitoses were enriched toward wound sites 48 hr after amputation in fst(RNAi) animals but were fewer in number (bottom right, p<0.001 at 200 µm, two-tailed t-test).
DOI: http://dx.doi.org/10.7554/eLife.00247.010
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Given that fst(RNAi) animals displayed a defective proliferative response to missing tissue, we tested whether these animals produced regenerative progenitor cell types. Head amputation normally induces neoblasts to produce ovo+ eye progenitors (Lapan and Reddien, 2012), but this process failed in fst(RNAi) animals (Figure 2D). From these data taken together, we conclude that fst is required for several aspects of the regeneration-specific neoblast response to injury.

follistatin is required for responding to tissue absence following injury

The abnormal missing-tissue-specific mitotic response of fst(RNAi) animals raised the possibility that other missing tissue responses could also require fst. Apoptosis increases following injury in planarians (Pellettieri et al., 2010), and, like the mitotic response, this increase involves a generic injury phase and a missing-tissue-specific phase. First, a local apoptosis burst occurs at wound sites 4 hr following any injury; second, a body-wide apoptosis burst occurs 72 hr after injury, but only in cases involving missing tissue (Pellettieri et al., 2010). The apoptosis level in this latter phase scales with the amount of missing tissue (Pellettieri et al., 2010). Planarians possess a centrally located pharynx used for feeding and defecation (Reddien and Sánchez Alvarado, 2004); measuring apoptotic cell numbers by TUNEL within the pharynx is an established assay for quantifying the body-wide increase in apoptosis that occurs 72 hr post-amputation (Pellettieri et al., 2010). Strikingly, fst(RNAi) pharynges displayed little increase in apoptotic cell numbers 72 hr post amputation, whereas a roughly 20-fold increase from pre-amputation levels occurred in control pharynges (Figure 3A). fst(RNAi) animals had a normal 4 hr apoptosis burst, indicating that fst is not generally required for apoptosis (Figure 3B). The 72 hr apoptotic response occurs in animals that have had their neoblasts ablated and cannot regenerate (Pellettieri et al., 2010). Therefore, the failure of fst(RNAi) animals to produce this response cannot be explained as a non-specific result of regeneration failure.

Figure 3. fst is required broadly for missing-tissue responses and morphallaxis.

(A) fst(RNAi) animals displayed no increase in pharyngeal TUNEL+ cells 3 days post-amputation (p<0.001, two-tailed t-test). Dotted white line = pharynx outline. (B) fst(RNAi) tail fragments displayed normal TUNEL+ cell numbers 4 hr post-amputation (n = 6/6). (C) fst(RNAi) animals displayed normal wound-induced gene expression 3 hr and 6 hr after amputation (jun-1: n = 20/20, nlg1: n = 5/5) but reduced expression compared to controls 24–48 hr after amputation (arrows; jun-1: 17/19 correctly scored blindly, p<0.01 Fisher’s exact test, nlg1: 22/27 correctly scored blindly, p<0.01, Fisher’s exact test). (D) fst(RNAi) animals had increased wound-induced expression of delta-1 24 hr after amputation (n = 12/12). (E) fst(RNAi) animals did not rescale expression of wntP-2 48 hr after amputation (n = 18/21). (F) fst(RNAi) animals did not reduce the number of cintillo+ cells in head fragments following amputation (p<0.001, two-tailed t-test). Scale bars = 100 µm. Anterior up in (A–C), (E), (F). Anterior left in (D).

DOI: http://dx.doi.org/10.7554/eLife.00247.011

Figure 3.

Figure 3—figure supplement 1. The fst(RNAi) phenotype is not a non-specific result of regeneration failure.

Figure 3—figure supplement 1.

Left: Following lethal irradiation and amputation, animals displayed higher expression of nlg1, lower expression of jun-1, and higher expression of delta-1 (n > 5 for each). smad1(RNAi) animals did not form blastemas but displayed normal delta-1 expression. Right: control tail fragments produced a pharynx de novo by 8 days after amputation, while fst(RNAi) tail fragments did not (arrowhead, n = 6/7). smad1(RNAi) animals failed to produce blastemas following amputation (arrowhead) but produced a pharynx normally (n = 5/5). The marker madt labels intestine and outlines pharynx (white area). Anterior up. Scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.012
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In addition to the cellular responses to missing tissue described above, persistence of wound-induced gene expression is another aspect of the planarian missing-tissue response (Wenemoser et al., 2012). We observed less expression of two wound-response genes in fst(RNAi) animals than in controls 24–48 hr post-amputation, despite expression levels being indistinguishable at earlier timepoints (Figure 3C). Notably, some wound-induced genes display expression that inversely scales with missing tissue amount; for example, Smed-delta-1 displays higher expression after an incision or puncture (simple injuries) than after amputation (a major injury) (Wenemoser et al., 2012). Amputated fst(RNAi) animals displayed a higher, rather than lower, level of Smed-delta-1 expression than did controls 24 hr after amputation (Figure 3D). Therefore, the lower expression levels observed for other wound-induced genes in fst(RNAi) animals do not reflect generically lower gene expression at wounds, but instead a specific requirement for fst for missing-tissue-specific gene expression.

Irradiated animals (which cannot regenerate) can display either higher or lower levels of wound-induced expression, depending on the gene examined (Wenemoser et al., 2012). Indeed, some wound-induced genes were similarly affected between irradiated and fst(RNAi) animals, while others were oppositely affected (Figure 3—figure supplement 1). As was the case for the failed apoptotic response of fst(RNAi) animals, the missing-tissue gene expression defects of fst(RNAi) animals cannot therefore be explained as a side-effect of regenerative failure.

In addition to producing a regeneration blastema, amputated animals must reorganize and rescale remaining tissue in a process termed morphallaxis (Morgan, 1901; Reddien and Sánchez Alvarado, 2004). Some aspects of this process do not require blastema formation. For example, wntP-2 (also known as wnt11-5 [Gurley et al., 2010]) is normally expressed in planarian tails (Petersen and Reddien, 2009; Gurley et al., 2010) and its expression domain restricts posteriorly within 48 hr of amputation whether regeneration proceeds or not (Gurley et al., 2010). fst(RNAi) animals did not rescale the wntP-2 expression domain 48 hr following amputation, further supporting a model in which fst is required for responding to missing tissue (Figure 3E). Following head amputation, head fragments not only produce posterior-specific cell types but also reduce numbers of anterior-specific cell types (which are overabundant for the new fragment dimensions). This process failed in fst(RNAi) head fragments (Figure 3F). Finally, fst(RNAi) fragments did not produce pharynges de novo (which normally occurs in pre-existing head and tail fragment tissue) (Figure 3—figure supplement 1). By contrast, RNAi of a different gene that blocked blastema formation (smad1) did not block pharynx formation, indicating this defect is not a simple consequence of blastema formation failure (Figure 3—figure supplement 1). We conclude that fst is required broadly for missing-tissue-specific wound responses, and that these defects likely underlie the inability of fst(RNAi) animals to regenerate.

Smed-activin-1 is required for the follistatin regeneration phenotype

Because Follistatin proteins are well-characterized extracellular inhibitors of TGF-β ligands (Nakamura et al., 1990; Hemmati-Brivanlou et al., 1994), we sought to identify putative TGF-β ligands that Smed-Follistatin might regulate to promote regeneration. Seven putative TGF-β superfamily members exist in the Schmidtea mediterranea genome (Figure 4—figure supplement 1 and Molina et al., 2007; Orii and Watanabe, 2007; Reddien et al., 2007; Gaviño and Reddien, 2011; Molina et al., 2011; Wenemoser et al., 2012; Roberts-Galbraith and Newmark, 2013). If Fst regulates one of the proteins encoded by these genes, then RNAi of that gene might suppress the fst RNAi phenotype. We tested this possibility (see Figure 4—figure supplement 2 and ‘Materials and methods’ for details) and found that RNAi of either of two genes, Smed-activin-1 (act-1 in short) or Smed-activin-2 (act-2), strongly suppressed the blastema formation defect (Figure 4A), the failure to regenerate a brain (Figure 4A), and the failed missing-tissue apoptotic response of fst(RNAi) animals (Figure 4B); RNAi of act-2 can also restore anterior pole regeneration in fst(RNAi) animals (Roberts-Galbraith and Newmark, 2013). Given that Follistatin proteins can directly regulate Activin proteins in other organisms (Nakamura et al., 1990; Hemmati-Brivanlou et al., 1994), these data suggest that Follistatin promotes missing tissue responses by inhibiting the function of Activin proteins.

Figure 4. act-1 and act-2 are required for the fst RNAi phenotype.

(A) fst(RNAi) animals treated with control dsRNA did not produce blastemas or a brain after amputation (n = 17/22), whereas fst(RNAi) animals treated with act-1 or act-2 dsRNA produced normal blastemas and brain (act-1: n = 19/23, p<0.0001; act-2: n = 16/26, p<0.001; Fisher’s exact test for both). Inhibition of other candidate genes did not suppress the fst RNAi phenotype (n > 9 for all other conditions). Aberrant animals were scored as having greatly decreased or absent brain. (B) fst(RNAi) animals treated with control dsRNA displayed a reduced apoptotic response 3 days after amputation, whereas fst(RNAi) animals treated with act-1 dsRNA displayed a normal apoptotic response (p<0.001 between control RNAi and fst;ctrl RNAi; p<0.01 between fst;ctrl RNAi and fst;act-1 RNAi, two-tailed t-test for both). (C) act-1(RNAi) animals displayed a greater induction of ovo+ eye progenitors compared to controls 2 days after head amputation (n = 24, p<0.0001, two-tailed t-test). Scale bars = 100 µm. Anterior up.

DOI: http://dx.doi.org/10.7554/eLife.00247.013

Figure 4.

Figure 4—figure supplement 1. Phylogeny of planarian activin homologs.

Figure 4—figure supplement 1.

The Smed-act-1 sequence was aligned with other TGF-β sequences using CLUSTALW (Higgins, 1994; Thompson et al., 1994). The alignments were trimmed using GBlocks (Castresana, 2000), allowing for smaller final blocks, gap positions within the final blocks, and less strict flanking positions. Bayesian analysis was performed using MrBayes (Huelsenbeck and Ronquist, 2001; Ronquist and Huelsenbeck, 2003). The Bayesian inference tree is shown with support values above 0.5 for each branch. Smed-Act-1 and Smed-Act-2 are shown in red. The phylogenetic positions of Smed-Act-1 and Smed-Act-2 support orthology with Activin proteins. Xl = Xenopus laevis, Mm = Mus musculus, Hs = Homo sapiens, Gg = Gallus gallus, Bf = Branchiostoma floridae, Dm = Drosophila melanogaster, Dr = Danio rerio, Sm = Schmidtea mediterranea.
DOI: http://dx.doi.org/10.7554/eLife.00247.014
Figure 4—figure supplement 2. RNAi controls for fst suppression experiments.

Figure 4—figure supplement 2.

Left: Animals treated with both fst dsRNA and act-1 dsRNA display no fst RNAi phenotype even though expression of fst is greatly reduced (top). Animals treated with both fst dsRNA and act-2 dsRNA displayed a similar reduction of the fst RNAi phenotype even though expression of fst is greatly reduced (bottom). Animals treated with fst dsRNA and another candidate dsRNA display the fst RNAi phenotype even though expression of candidate genes are greatly reduced (n > 6 for all). Anterior left, scale bars = 100 µm. Right: fst expression was reduced in fst(RNAi) animals concomitantly treated with control dsRNA or with act-1 dsRNA as assayed by quantitative PCR (performed as described in Figure 1—figure supplement 1) with two independent sets of fst primers (primer set 2 = left: TGGTGTGCAATTTAGCGAGT; right: ATTCACTGCAGCCTCTTGTG).
DOI: http://dx.doi.org/10.7554/eLife.00247.015
Figure 4—figure supplement 3. Efficacy of act-1 RNAi.

Figure 4—figure supplement 3.

act-1 expression was reduced in act-1(RNAi) animals both prior to and 24 hr after amputation as assayed by quantitative PCR. qPCR was performed as described in Figure 1—figure supplement 4. act-1 (left: GCGAGCTACCTTTCAATGCT, right: ACGGGAGTGCAACAGTTTTT) primers were used to evaluate expression.
DOI: http://dx.doi.org/10.7554/eLife.00247.016
Figure 4—figure supplement 4. act-1(RNAi) animals appear normal following regeneration.

Figure 4—figure supplement 4.

act-1(RNAi) animals formed blastemas and regenerated following head and tail amputation (n > 200). Scale bars = 100 µm, anterior up.
DOI: http://dx.doi.org/10.7554/eLife.00247.017
Figure 4—figure supplement 5. act-1(RNAi) animals display normal neoblast numbers.

Figure 4—figure supplement 5.

Intact act-1(RNAi) animals displayed normal numbers of X1 cells as counted by flow cytometry.
DOI: http://dx.doi.org/10.7554/eLife.00247.018
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activin-1(RNAi) animals display excessive progenitor production following amputation

Given that activin expression is required for the fst(RNAi) phenotype, we investigated the consequences of act-1 RNAi on regeneration. Although act-2(RNAi) has been reported to produce posterior regeneration defects (Roberts-Galbraith and Newmark, 2013), act-1(RNAi) animals were capable of regenerating (Figure 4—figure supplement 3, Figure 4—figure supplement 4) and, as with fst(RNAi), displayed normal neoblast turnover during homeostatic growth (Figure 4—figure supplement 5). act-1(RNAi) survived after amputation as well as controls did (observed more than a month following injury, n = 10/10). act-1(RNAi) animals did however display some abnormalities. Although act-1(RNAi) animals displayed normal ovo+ eye progenitor numbers prior to amputation, increased numbers as compared to controls were present following amputation (Figure 4C). By contrast, fst RNAi caused the opposite phenotype of reduced ovo+ eye progenitor formation. These data raise the possibility that act-1 regulates responses to injury, with some aspects of regeneration overactive following act-1 inhibition.

The amount of missing tissue regulates follistatin expression following injury

Because fst is required for regeneration but not for normal tissue turnover, we reasoned that fst expression might be high following amputation, an injury type requiring significant tissue regeneration, but low following incision or puncture, injuries requiring only wound healing. We therefore assessed fst as compared to act expression at wounds following either incision or excision of a tissue wedge. Increased act-1 expression was not detected following either type of wound, with expression detected throughout the intestine of uninjured animals, suggesting an intestinal source of Activin-1 protein (Figure 5A). act-2 expression was similar to act-1 in intact animals, but unlike act-1 is wound-induced (Figure 5B, Figure 5—figure supplement 1; Roberts-Galbraith and Newmark, 2013). Indeed, act-2 was wound-induced following either incision or tissue wedge excision, with expression persisting for several days irrespective of injury severity (Figure 5C, Figure 5—figure supplement 2). By contrast, fst expression was induced at both wound types by 6 hr after injury, but by 48 hr after injury was present only at wedge excision wound sites (Figure 5C, Figure 5—figure supplement 2). These results indicate that fst expression persists longer at wounds that result in tissue absence. Furthermore, fst expression was greater at wounds involving a large amount of missing tissue (assessed at 48 hr) than at wounds with little missing tissue (Figure 5—figure supplement 3). Together, these data are consistent with a model in which wound-induced fst expression levels are regulated by the amount of missing tissue. In this model, fst promotes regenerative responses by inhibition of act-1 and act-2 following major injury (Figure 5D).

Figure 5. fst induction is regulated by tissue absence following injury.

(A) act-1 was expressed broadly throughout the intestine and was not induced by amputation. (B) act-2 was expressed in the intestine and pharynx in intact animals and induced following amputation at wound sites (6 hr and 24 hr, arrows), eventually spreading throughout the body (48 hr, arrows). (C) Animals displayed wound-induced expression of fst 6 hr after either incision or tissue wedge removal, but expression persisted only in cases of tissue wedge removal (n > 5, p<0.01, white arrowheads = injury site). (D) A proposed genetic model for fst and act-1/2 in regeneration. Wounds induce fst expression. If there is missing tissue following injury, fst induction is high, Act-1/2 signaling is inhibited, and regeneration-specific responses are initiated. If there is no missing tissue following injury, fst expression is low, Act-1/2 signaling is not inhibited, and regeneration-specific responses are repressed. Scale bars = 100 µm. Anterior up.

DOI: http://dx.doi.org/10.7554/eLife.00247.019

Figure 5.

Figure 5—figure supplement 1. Wound-induced act-2 expression persists for several days after amputation.

Figure 5—figure supplement 1.

act-2 expression persisted at high levels throughout the animal during regeneration until at least 8 days after amputation. Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.020
Figure 5—figure supplement 2. fst and act-2 expression is negligible at wound sites prior to injury.

Figure 5—figure supplement 2.

Lateral, post-pharyngeal expression of fst and act-2 is minimal prior to injury (act-2 signal present is intestinal). Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.021
Figure 5—figure supplement 3. The amount of missing tissue regulates wound-induced fst expression.

Figure 5—figure supplement 3.

fst expression was higher in level 48 hr after an amputation resulting in a large amount of missing tissue than after an amputation resulting in little missing tissue, as measured by quantifying fluorescent in situ hybridization signal intensity (p<0.01, two-tailed t-test). Anterior up, scale bars = 100 µm.
DOI: http://dx.doi.org/10.7554/eLife.00247.022
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Discussion

Regeneration initiation

All long-living animals face the prospect of injury and require regenerative mechanisms. Planarians are an exceptional example of the regenerative potential of animals. Distinct cellular and molecular programs for responding to simple injury vs missing tissue exist in planarians. In the case of injuries involving substantial missing tissue, animals mount unique mitotic and apoptotic responses and produce an extended program of wound-induced gene expression (Pellettieri et al., 2010; Wenemoser and Reddien, 2010; Wenemoser et al., 2012). These events represent the earliest described divergent behaviors following major injuries requiring regeneration vs simple injuries requiring only wound healing. A central question has therefore become how these distinct responses are mediated.

We identified a gene encoding a homolog of the TGF-β inhibitor, follistatin, that is required for regeneration and for regeneration-specific cellular and molecular responses to injury. Our data suggest that inhibition of Activin signaling by Fst is required for initiating a regenerative response at wounds following major injury. Finally, fst is wound-induced, with the level of fst expression persisting at high levels longer following a major injury than following a simple injury. We propose that wound-induced fst expression allows for regenerative responses to be initiated specifically as a consequence of tissue absence.

The nature of the planarian missing-tissue response

fst is the first gene known to be required for regeneration-specific responses in planarians. Not all missing-tissue responses are abolished following fst inhibition, however. For example, neoblast migration to amputation sites occurred normally in fst(RNAi) animals, despite the absence of a normal proliferative response. Similarly, although expression of act-1 and act-2 are required for the fst(RNAi) phenotype, inhibition of activin expression in the absence of amputation does not affect homeostatic tissue turnover or induce a regeneration-like state, demonstrating that the suppression of Activin alone is not sufficient to induce missing-tissue responses. Therefore, some aspects of the missing-tissue response to injury require an as yet unknown ‘missing-tissue’ signal or signals that operate independently of fst and Activin signaling. Identifying and characterizing these processes will be important for understanding how the decision to mount a regenerative response occurs.

TGF-β signaling across regenerative contexts

Our findings describe a system in which suppression of Activin signaling is required for regeneration. The possibility therefore exists that Activin signaling may serve similar functions in other organisms. Indeed, TGF-β signaling has been implicated as a negative regulator of regeneration in a variety of contexts, including following partial hepatectomy (Russell et al., 1988; Kogure et al., 1995; Romero-Gallo et al., 2005), in embryonic chick retinas (Sakami et al., 2008), in renal regeneration following ischemia/reperfusion injuries (Kojima et al., 2001), and for mouse skeletal muscle regeneration (Zhu et al., 2011). Given the relevance of these systems to human medicine, it will be important to investigate to what extent regenerative regimes recapitulate the mechanisms observed in planarians. Interestingly, a number of systems use TGF-β signaling to promote rather than suppress regeneration: TGF-β signaling is involved in axolotl limb and Xenopus tail regeneration (Lévesque et al., 2007; Ho and Whitman, 2008), activin expression can be induced by wounding and exogenous TGF-β can speed healing in mammals (Mustoe et al., 1987; Hübner et al., 1996; Sulyok et al., 2004), TGF-β signaling can promote regeneration following mouse ear hole-punching (Liu et al., 2011), and wound-induced activin promotes cell proliferation and migration following zebrafish fin amputation (Jaźwińska et al., 2007). Despite these contextual differences, TGF-β signaling plays a major role in many forms of regeneration studied. Therefore, uncovering ‘missing-tissue’ signals in planarians, describing how these signals interact with Activin signaling, and identifying the key factors regulated by these signals will inform a broad understanding of core regenerative mechanisms.

Materials and methods

Gene cloning

For RNA probes, genes were cloned into pGEM and amplified with T7-promoter-containing primers. For RNAi, genes were cloned into pPR244 as described (Reddien et al., 2005). activin-1 was cloned with primers 5’-TCAACTGAAACGGAAGTTGG-3’ and 5’-TGGTGGATCCTTACTTGCAG-3’, activin-2 with primers 5’-ACCAATTATGGCCAATCCAG-3’ and 5’-CCGGCTAATTGTGAACAAAC-3’, and follistatin with 5’-CACAAGAGGCTGCAGTGAAT-3’ and 5’-CATTCAGAAGGCATTGTCCA-3’.

RNAi

The control dsRNA for all RNAi experiments was unc-22 from Caenorhabditis elegans. RNAi experiments were performed by feeding a mixture of liver and bacteria expressing dsRNA (Reddien et al., 2005). 20 ml of bacterial culture was pelleted and resuspended in 60 μl of liver. For fst and act-1 RNAi regeneration experiments, animals were fed on day 0, day 4, day 8, and day 12, amputated on day 16/17 and either soaked for 6 hr in 1 µg/µl dsRNA (TUNEL experiments), soaked for 2 hr in dsRNA (gene expression experiments), or not soaked in dsRNA. For suppression experiments, totals from two separate experiments were pooled: (1) animals were fed fst dsRNA on day 0, day 4, day 8, and day 12, fed candidate gene dsRNA on day 16, day 20, and day 23, and amputated on day 24. (2) Animals were amputated and injected four times with a 30 nl equimolar mixture of fst and candidate gene dsRNA on day 0, injected without amputation on day 1, amputated and injected on day 4, and injected only on day 5. Animals were scored and fixed 8 days after the final amputation.

In situ hybridizations, immunostaining, and TUNEL

Whole-mount in situ hybridizations and fluorescence in situ hybridizations (FISH) were performed as described (Pearson et al., 2009). For double/triple labeling, HRP-inactivation was performed between labelings (4% formaldehyde, 30 min). Immunostainings were performed as previously described (Reddien et al., 2005) using tyramide signal enhancement. TUNEL was performed as previously described (Pellettieri et al., 2010).

γ-irradiation

For elimination of neoblasts, planarians were exposed to 6000 rad (6K, ∼72 min) using a cesium source (∼83 rad/min).

Flow cytometry

Animals were amputated in cold CMFB, and cells prepared as described (Scimone et al., 2011). For quantification of X1 cells, five animals were used per RNAi condition in triplicate. Analyses and sorting were performed using a Moflo3 FACS sorter (Dako-Cytomation, Carpinteria, CA) and FlowJo.

Imaging and analyses

For quantifying cell numbers expressing a marker or an area of positive cells, equal numbers of optical stacks were taken per specimen, collapsed, and quantified using Automeasure in AxioVision (Zeiss, Jena, Germany) or manually. For quantification of fluorescence intensity, 7 optical stacks were acquired from the ventral surface of animals, collapsed, and values determined using the Automeasure module (Densitometric sum) in AxioVision (Zeiss). Images were acquired using an AxioImager with Apotome (Zeiss) or an LSM 700 (Zeiss).

Acknowledgements

We thank Reddien Lab members for comments and discussion. PWR is an early career scientist of the Howard Hughes Medical Institute and an associate member of the Broad Institute of Harvard and MIT. We acknowledge support from the NIH (R01GM080639) and the Keck Foundation.

Funding Statement

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

Funding Information

This paper was supported by the following grants:

  • National Institutes of Health R01GM080639 to Peter W Reddien.

  • Howard Hughes Medical Institute to Peter W Reddien.

  • Keck Foundation to Peter W Reddien.

Additional information

Competing interests

The authors declare that no competing interests exist.

Author contributions

MAG, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article.

DW, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article.

IEW, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article.

PWR, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

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eLife. 2013 Sep 10;2:e00247. doi: 10.7554/eLife.00247.023

Decision letter

Editor: Marianne E Bronner1

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for choosing to send your work entitled “Tissue absence initiates regeneration through Follistatin-mediated inhibition of Activin signaling” for consideration at eLife. Your article has been evaluated by a Senior editor and 3 reviewers, one of whom, Marianne Bronner, is a member of our Board of Reviewing Editors.

The Reviewing editor and the other reviewers discussed their comments before we reached this decision, and the Reviewing editor has assembled the following comments based on the reviewers' reports. Our goal is to provide the essential revision requirements as a single set of instructions, so that you have a clear view of the revisions that are necessary for us to publish your work.

This is a thorough and interesting paper that makes an excellent contribution to the body of knowledge in this field. Gaviño, et al. demonstrate a novel role for the Activin branch of TGFβ signaling in planarian regeneration. The paper is potentially appropriate for eLife after the authors attend to the following concerns:

1) The major findings revolve around two TGFβ family members, follistatin and activin-1, and their role in regeneration, effects on cell proliferation, etc. The story is interesting but fairly simple and straightforward. Therefore, the paper would be improved by significant shortening to concentrate on the major findings. The authors are encouraged to rewrite the paper in shorter format and reduce the number of figures accordingly.

2) The interpretation that activin-1(RNAi) significantly speeds up the normal regenerative response is not convincing. An alternative interpretation is that the mechanisms regulating homeostasis and regeneration may have become uncoupled in activin-1(RNAi). The increased ovo expression in animals 20 days after amputation is intriguing, suggesting that there is an elevated level of tissue homeostasis after a single round of RNAi and regeneration. Are there survival disadvantages for animals that have undergone long term activin-1(RNAi) coupled with amputation? The claim of speeding regeneration is semantic in nature and should be toned down.

3) The authors argue that activin is upstream of Smad4 by showing that Smad4(RNAi) causes an increase in apoptosis like activin(RNAi). Since Smad4(RNAi) causes a profound regeneration defect, the apoptotic increase could be secondary to this defect. Furthermore, it has been shown that activin can signal independent of Smad4 (Suzuki et al., Biochemical and biophysical research communications, 2010.) Are there any other upstream R-Smads capable of recapitulating the Activin defect or also rescue the Follistatin phenotype?

4) The authors show adult homeostatic expression patterns for follistatin and activin, yet they have no apparent phenotype under homeostasis. One concern is that the RNAi conditions may not be effective enough to knock down these homeostatic levels of these factors. This would prevent them from making the conclusion that these factors have no effect normally. Activin especially seems like a good candidate to regulate the neoblast homeostatic niche. In order for the authors to conclude that knockdown of these factors has no effect in homeostasis, they should provide in situ data to demonstrate that the adult expression of these factors is indeed repressed by RNAi to a reasonable degree.

5) The generic controls for RNAi seem inadequate. The authors should perform rescue experiments and lack of off-target effects.

eLife. 2013 Sep 10;2:e00247. doi: 10.7554/eLife.00247.024

Author response

1) The major findings revolve around two TGFβ family members, follistatin and activin-1, and their role in regeneration, effects on cell proliferation, etc. The story is interesting but fairly simple and straightforward. Therefore, the paper would be improved by significant shortening to concentrate on the major findings. The authors are encouraged to rewrite the paper in shorter format and reduce the number of figures accordingly.

We have reworked the paper as suggested. We reduced the number of figures from seven to five, removing data from multiple experiments from the main figures. Focus is increased on the follistatin RNAi phenotype and we removed roughly 1/3 of the text from the manuscript.

2) The interpretation that activin-1(RNAi) significantly speeds up the normal regenerative response is not convincing. An alternative interpretation is that the mechanisms regulating homeostasis and regeneration may have become uncoupled in activin-1(RNAi). The increased ovo expression in animals 20 days after amputation is intriguing, suggesting that there is an elevated level of tissue homeostasis after a single round of RNAi and regeneration. Are there survival disadvantages for animals that have undergone long term activin-1(RNAi) coupled with amputation? The claim of speeding regeneration is semantic in nature and should be toned down.

We have simplified the manuscript and it now focuses more on the follistatin phenotype. With activin RNAi, we present suppression of the follistatin RNAi phenotype and increased ovo+ progenitor presence at amputation sites. The issue of speeding regeneration will be a good target for future work.

We also investigated the long-term survival of activin-1(RNAi) animals following amputation as suggested. We found that there was not an increased level of animal death over the period observed (∼1 month) in activin-1(RNAi) animals as compared to controls. This is now referenced in the main text.

3) The authors argue that activin is upstream of Smad4 by showing that Smad4(RNAi) causes an increase in apoptosis like activin(RNAi). Since Smad4(RNAi) causes a profound regeneration defect, the apoptotic increase could be secondary to this defect. Furthermore, it has been shown that activin can signal independent of Smad4 (Suzuki et al., Biochemical and biophysical research communications, 2010.) Are there any other upstream R-Smads capable of recapitulating the Activin defect or also rescue the Follistatin phenotype?

We removed the smad4 phenotype from the manuscript, as we now are leaving the apoptosis responses to wounding in activin deficient animals for more detailed future work.

4) The authors show adult homeostatic expression patterns for follistatin and activin, yet they have no apparent phenotype under homeostasis. One concern is that the RNAi conditions may not be effective enough to knock down these homeostatic levels of these factors. This would prevent them from making the conclusion that these factors have no effect normally. Activin especially seems like a good candidate to regulate the neoblast homeostatic niche. In order for the authors to conclude that knockdown of these factors has no effect in homeostasis, they should provide in situ data to demonstrate that the adult expression of these factors is indeed repressed by RNAi to a reasonable degree.

We performed the RNAi controls in homeostatic conditions as suggested. We used quantitative PCR to measure fst expression levels in unamputated fst(RNAi) animals and also in fst(RNAi) fragments 24h after amputation. In these two cases, fst expression was inhibited by RNAi to a similar extent. We performed the same experiment in unamputated activin-1(RNAi) animals and 24h activin-1(RNAi) regenerating fragments. Here as well, expression was inhibited by RNAi to a similar degree in both contexts. These data are now included in Figure 1–figure supplement 3 and Figure 4–figure supplement 3, respectively.

To further demonstrate that the effect of fst RNAi is specific to regeneration, we labeled amputated fst(RNAi) animals that had failed to regenerate with a combination of markers that identify differentiating neurons 20d after amputation. Strikingly, in these animals that produced no blastemas, new neurons were still being made in old tissues. Furthermore, irradiated animals (irradiation kills neoblasts and blocks tissue turnover) die within roughly 2 weeks of irradiation, whereas these amputated animals survive long-term despite failing to regenerate. These data demonstrate that fst(RNAi) animals that fail to produce new tissues at wound sites continue to replace aging cells for tissue turnover, indicating the specificity of requirement for fst under these experimental conditions for regeneration. These data are now included in Figure 1.

5) The generic controls for RNAi seem inadequate. The authors should perform rescue experiments and lack of off-target effects.

We performed and now present quantitative PCR experiments to measure the extent of both fst and activin-1 inhibition by RNAi in unamputated and regenerating animals (see point 4 above).

Furthermore, (in addition to previously presented in situ controls) we measured the extent of fst and activin-1 inhibition in fst; activin-1 double RNAi animals and observed that fst is inhibited at least as strongly in these double RNAi animals as in fst; control double RNAi animals. These data are presented in Figure 4–figure supplement 2.

To address the specificity of fst RNAi, we produced two additional RNAi constructs for inhibiting fst expression, with each construct spanning a non-overlapping portion of the fst gene. Both of these constructs caused a regeneration phenotype. These data are now presented in Figure 1–figure supplement 1.

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