# On the hormonal control of posterior regeneration in the annelid Platynereis dumerilii

Patricia Álvarez-Campos, Anabelle Planques, Loïc Bideau, Michel Vervoort, Eve Gazave

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Patricia Álvarez-Campos, Anabelle Planques, Loïc Bideau, Michel Vervoort, Eve Gazave. On the hormonal control of posterior regeneration in the annelid Platynereis dumerilii. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution, 2023, 340 (4), pp.298-315. 10.1002/jez.b.23182. hal-03851548

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**On the hormonal control of posterior regeneration in the annelid *Platynereis dumerilii***

Patricia Álvarez-Campos<fnref n="1" />,<fnref n="2" />, Anabelle Planques<fnref n="1" />, Loïc Bideau<fnref n="1" />, Michel Vervoort<fnref n="1" />,*, and
Eve Gazave<fnref n="1" />,.*

*Co-senior and corresponding authors: michel.vervoort@ijm.fr; eve.gazave@ijm.fr

**Abstract**

Regeneration is the process by which many animals are able to restore lost or injured
body parts. After amputation of the posterior part of its body, the annelid *Platynereis dumerilii* is able to regenerate the pygidium, the posteriormost part of its body that bears the anus, and a subterminal growth zone containing stem cells that allows the subsequent addition of new segments. The ability to regenerate their posterior part (posterior regeneration) is promoted, in juvenile worms, by a hormone produced by the brain and is lost when this hormonal activity becomes low at the time the worms undergo their sexual maturation. By characterizing posterior regeneration at the morphological and molecular levels in worms that have been decapitated, we show that the presence of the head is essential for multiple aspects of posterior regeneration, as well as for the subsequent production of new segments. We also show that methylfarnesoate, the molecule proposed to be the brain hormone, can partially rescue the posterior regeneration defects observed in decapitated worms. Our results are therefore consistent with a key role of brain hormonal activity in the control of regeneration and growth in *P. dumerilii*, and support the hypothesis of the involvement of methylfarnesoate in this control.

**Key words:** regeneration, brain hormone, *Platynereis*, annelid, methylfarnesoate, hormonal regulation

<footnote n="1">¹Université Paris Cité, CNRS, Institut Jacques Monod, F-75013, Paris, France</footnote>

<footnote n="2">²Present address: Centro de Investigación en Biodiversidad y Cambio Global (CIBC-UAM) & Departamento de Biología (Zoología), Facultad de Ciencias, Universidad Autónoma de Madrid, Spain</footnote>
## Introduction

Regeneration, the ability to restore lost or injured body parts (Brockes & Kumar 2008; Poss 2010), is a widespread phenomenon in Metazoa (Bely & Nyberg 2010; Bideau et al. 2021). However, regeneration in animals is also highly variable as some species are able to regenerate only specific cell types or tissues, while others can regenerate complex structures such as limbs, and even their whole body from a small piece of tissue (Bely & Nyberg 2010; Grillo et al. 2016; Bideau et al. 2021). Segmented worms (Annelida) have been widely used to study this amazing process and its evolution in animals (e.g., Zattara & Bely 2016, Özpolat & Bely 2016). Many annelids (with the noticeable exception of leeches) display substantial regeneration abilities, which however may differ from one species to another. In fact, even in closely-related species, a huge variability is observed, in particular with respect to the possibility of regenerating their anterior body region upon amputation (Özpolat & Bely 2016, Nikanorova et al. 2020). Moreover, regeneration in some semelparous annelids (i.e., species with a single reproduction episode followed by death), such as nereids (Nereidae), is intimately linked to its sexual reproduction mode, given that these species lose their growing and regenerative abilities when sexual maturation starts (e.g., Clark & Ruston 1963; Clark & Scully 1964; Schenk et al. 2016).

Nereids, such as *Hediste diversicolor* (formerly *Nereis diversicolor*), *Alitta virens* (formely *Nereis virens*) and *Platynereis dumerilii*, are indeed able to grow during most of their life (juvenile phase), by adding segments in their posterior body region one by one, a process known as posterior growth or posterior elongation (e.g. Clark & Scully 1964; de Rosa et al. 2005; Gazave et al. 2013). During this growth phase, these worms are also able to successfully regenerate the posterior part of their body upon amputation, a process known as posterior regeneration (e.g., Clark & Ruston 1963; Özpolat & Bely 2016; Kozin & Kostyuchenko 2015; Planques et al. 2019). Interestingly, both posterior growth and posterior regeneration abilities stop when the worms undergo sexual maturation, a process known as epitoky, which involves dramatic modifications of the morphology, physiology, and behaviour of the worms to become reproductive gonochoristic individuals (e.g. Chatelain et al. 2008). During this sexual metamorphosis, the benthic tubicolous juvenile worms turn into pelagic reproductive individuals, undergoing striking morphological and physiological modifications and producing massive amounts of gametes (e.g., Fauvel 1959; Clark 1961; Fischer 1999). Sexually mature animals are then ready to spawn, and they start a nuptial dance (females and males
rapidly swim together in a circle) to deliver their gametes and die shortly after spawning.

A large series of experiments have shown that sexual maturation in nereids is under an endocrine control and identified the brain or supraesophageal ganglion as the source of this hormonal activity (e.g., Durchon 1948; Hauenschild 1964; Bertout 1983; Fischer 1984; Schenk et al. 2016). In juvenile worms, the brain hormone, traditionally termed nereidin, was shown to repress sexual metamorphosis, that is only triggered when the worms become older and during the breeding season, by the decrease of the effective concentration of this brain hormone (e.g., Clark & Ruston 1963; Scully 1964; Golding 1983). In addition to repressing epitoky, the brain hormone was also suggested to promote both posterior growth and posterior regeneration. In fact, if the supraesophageal ganglion is totally removed from nereid worms before posterior amputation, posterior regeneration and subsequent posterior growth are impaired (e.g., Clark & Bonney 1960; Clark & Evans 1961; Clark & Ruston 1963; Durchon & Marcel 1962; Hauenschild 1966; Golding 1967a; Hofman 1976). The molecular identity of the brain hormone has remained elusive for many years. While initially believed to be a neuropeptide (Müller 1973; Hofmann 1976; Cardon et al. 1981; Durchon 1984), it was recently found that nereidin activity in *P. dumerilii* involves methylfarnesoate (MF), a sesquiterpenoid hormone related to arthropod juvenile hormones (Schenk et al. 2016). MF was indeed shown to repress vitellogenesis in coelomic cells, which was used as a read-out of sexual maturation, and its concentration drops in the course of maturation, as expected for nereidin (Schenk et al. 2016). MF treatment also increased the expression of *Hox3*, a gene expressed in the growth zone in both growing and regenerating *P. dumerilii* worms (Pfeifer et al. 2012; Gazave et al. 2013; Planques et al. 2019), which led to the hypothesis that MF might also promote posterior growth and regeneration (Schenk et al. 2016).

In this article, we further studied the hormonal control of posterior regeneration in *P. dumerilii*, which has become an important model for comparative neurobiology and developmental biology in recent years (e.g., Williams and Jékely 2016; Schenkelaars & Gazave 2021; Özpolat et al. 2021). Life cycle of this small-sized nereid annelid can vary from around 3 months to a much longer time period depending on culturing conditions (Özpolat et al. 2021). Embryonic and larval development gives rise to small juvenile worms that enter a posterior growth phase which ends when the worms metamorphose into sexually mature individuals (Supp. Fig. 1 A to C). This posterior growth relies on the presence and activity of a subterminal growth zone, located immediately in front of the pygidium (terminal body part that bears the anus and a pair of appendages, the anal cirri),
which contains putative stem cells whose sustained divisions allow the serial addition of
new segments in the posterior body region (de Rosa et al. 2005; Gazave et al. 2013).

During this juvenile growth phase, after posterior amputation of several segments,
the growth zone, and the pygidium, *P. dumerilii* worms are able to quickly regenerate
their pygidium and their growth zone. The new growth zone will then produce new
segments as in normal posterior growth. Regeneration and post-regenerative posterior
growth are thus sequential processes. We recently showed that posterior regeneration in
*P. dumerilii* is a rapid process that includes five well-defined stages in a reproducible
timeline (Planques et al. 2019). As shown in Planques et al. 2019, and illustrated in
supplementary Fig. 2, stage 1, reached one day post posterior amputation (1 dppa),
corresponds to the completion of wound healing. At stage 2 (2 dppa), the anus opening is
reformed and a small blastema is observed, where intense cell proliferation occurs. The
size of the blastema increases at stage 3 (3 dppa) and the regenerated pygidium starts to
differentiate as shown by the formation of small anal cirri and pygidial muscles. At stage
4 (4 dppa), a well-differentiated pygidium with long anal cirri is observed and at stage 5
(5 dppa), the boundary between the pygidium and the new developing segments becomes
clearly visible through the appearance of lateral segmental grooves. At this stage,
posterior regeneration is finished and the worms enter a post-regenerative posterior
growth phase. While the timeline of regeneration stages is highly reproducible in worms
of a given age and size, the subsequent addition of segments is much more variable
(Planques et al. 2019).

Taking advantage of our previous morphological and molecular detailed
characterizations of posterior regeneration in *P. dumerilii*, we here re-address the role of
the brain hormonal activity in this process. By combining anterior and posterior
amputations and performing both morphological and molecular characterizations on
precisely staged worms, we found that both posterior regeneration and post-regenerative
posterior growth are severely affected in the absence of the brain, and characterized these
defects at the morphological and molecular levels. We also observed that posterior
regeneration defects in decapitated worms can be partially rescued by applying
exogenous MF hormone, pointing out the key role of MF in the control of posterior
regeneration.
Materials and methods

Worm culture, amputation procedures and production of the biological material

Individuals of *P. dumerilii* were obtained from the laboratory culture established at the Institut Jacques Monod (France), following Dorresteijn et al. (1993) protocol (Vervoort & Gazave 2022). A total of 782 individuals were used to perform the different experiments (see details on results section). In all but one experiment, 3-4-month-old worms with 30-40 segments were used. In addition, in the sexual maturation experiment, 1-month-old worms with 10–15 segments and 1-year-old worms with 85–95 segments were also used. After anaesthesia (with a solution of MgCl₂ 7.5% and sea water, 1/1), amputations of the anterior part of the worms were done using a microknife (SharpointTM) at the anterior boundary of the first segment in order to remove the prostomium (including the supraesophageal gland, Supp. Fig 1D). Amputations of the posterior-most segments of 3-4-month-old worms were also performed in order to remove the last 5-6 segments, the growth zone and the pygidium (Planques et al. 2019). After amputation, all worms were let to recover in natural fresh sea water (NFSW) for a couple of minutes and were then placed individually in 6-well plates in 10 ml of NFSW.

Methylfarnesoate treatments

Following Schenk *et al.* (2016) experimental design, (E,E)-methylfarnesoate (MF; Echelon Biosciences) 100 nM stock was prepared by adding 1 µl of pure hormone in 33.6 ml of DMSO in a glass container to prevent potential hydrophobic binding of the hormone to plastic. MF stock was then aliquoted in 10-12 glass vials that were frozen at -20°C, using only one aliquot per day of experiment in order to avoid MF alteration after several freeze-thaw cycles. Five to six amputated worms (see above) were placed in 150 ml glass beakers. MF-treated worms were maintained in a solution of 100 ml filtered NFSW and 100 µl of MF (from 100 nM stock solution in 0.1% DMSO). Worms used as controls (non-MF-treated, with or without anterior amputation) were maintained in 100 ml filtered NFSW and 0.1% DMSO.

Scoring of the worms and statistical analyses

To monitor the morphological changes expected for sexual maturation following
anterior amputation, as well as the posterior regeneration stages following anterior and/or
posterior amputations, daily observations of the worms were done after amputation and/or
hormonal treatments (see the “Results” section for the different timing of scoring/imaging procedures) using a ZEISS dissecting scope (Stemi SV11). Scoring was done according to the staging system established in Planques *et al.* (2019): worms were scored either by the regeneration stage that has been reached (stage 1 to 5) or, after regeneration completion, by the number of new segments that has been produced during the post-regenerative posterior growth process. During regeneration, worms showing a morphologically intermediate stage were coded as 1.5, 2.5, 3.5, and 4.5. Worms that died in the middle of the experiment were counted but removed from the statistical analyses, and those with abnormal morphology were analysed separately in all the experiments, to avoid biased results, as they cannot be scored properly due to their aberrant shape. Statistical analyses and graphic representation of the results of the amputation experiments were done using Prism 7 for Mac OS (GraphPad software) (Ivashchenko et al. 2017). Detailed results of all the statistical analyses are provided in Supp. Table 1.

*Whole-mount in situ hybridizations (WMISH) and imaging*

Single nitro blue tetrazolium chloride/5-bromo-4-chloro-3'-indolymphosphate (NBT/BCIP) WMISH on regeneration stages were performed as previously described (e.g., Tessmar-Raible et al. 2005; Gazave et al. 2017; Planques et al. 2019; Vervoort & Gazave 2022). Bright field pictures for NBT/BCIP WMISH were taken on a Leica microscope DM5000B. Editing and compilation of Z projections were achieved using ImageJ and Adobe Photoshop CS5.1. The final figure panels were then compiled using Adobe Illustrator CS5.1.

*Cell proliferation assays*

Proliferating cells are labelled by incubating worms (either control, only posteriorly amputated or C2 conditions – anterior amputation being performed two days before posterior amputation at 3 and 5 dppa) with the thymidine analog 5-ethynyl-2'-deoxyuridine (EdU) (1h in sea water, EdU 5 µM) and labelled using the Click-It EdU imaging kit (555, Life technology) as described in Vervoort & Gazave (2022). Samples were then counterstained with Hoechst (1 µg per ml), mounted in 87% glycerol/DABCO (2.5 mg/ml) and imaged on Zeiss LSM 980 confocal microscope (n= 4 to 6 per condition). Hoechst and EdU positive cells were thus counted using the Imaris software (Imaris 9.5 by BitPlane). Statistical analyses were performed using the Prism software (Prism 7 by GraphPad): proportions of EdU+ cells between conditions were compared using non-
parametric tests.

## Results

*Timeline of events following anterior amputation*

In order to be able to identify the effects of brain removal on *P. dumerilii* posterior regeneration, we briefly characterized their consequences on worm survival and sexual maturation, as decapitation has indeed been shown to lead to accelerated sexual maturation followed by the death of the animals (e.g., Hauenschild 1960, 1964, 1966). Effective concentration of brain hormones is thought to decrease from younger to older animals (e.g., Golding 1967a, 1983), raising the possibility that decapitation could have age-dependent effects (or timelines) due to different hormone concentrations in the worms' tissues. We therefore did a series of decapitation experiments using 40 worms with supposedly high, intermediate and low hormone levels (1-month, 3-4-month, and 1-year-old worms, respectively). Worms were observed at many different time points after anterior amputation and compared to 10 non-amputated control worms of the same age (Supp. Fig. 3). Of note, some 1-year-old control worms became sexually mature during the course of the experiment.

First of all, we found that the vast majority of decapitated worms survived for a long period after amputation, at least for 12 days (Supp. Fig. 3). In addition, regardless of their age, all decapitated worms showed some signs of sexual maturation such as the presence and accumulation over time of large cells which could be maturing germ cells, as they are expressing the germ cell marker *Vasa* (Supp. Fig. 3A1-2, B1-2, C1, 4A to C') as well as the enlargement of parapodia (Supp. Fig. 3A2-3, B2, C2). In addition, more than a week after amputation, 1-year-old decapitated worms will start to habor the typical color differences between males (reddish individuals) and females (yellowish individuals) (Supp. Fig. 3C3, C4). They also show an accumulation of large cells, that are very probably gametes as they are expressing *Vasa* and are morphologically similar to oocytes and sperm observed in non-amputated worms that underwent sexual maturation (Supp. Fig. 4C-C'). Later, those 1-year-old decapitated animals showed the typical *P. dumerilii* courtship dancing, yet none of them managed to release their gametes and perform fertilization.

In summary, anterior amputation does indeed induce sexual maturation leading to
sexual maturity in 1-year-old worms while they were however unable to reproduce. In 1-month-old and 3-4-month-old worms, sexual maturation was induced but never lead to maturity (Supp. Fig. 3A, B). Importantly, decapitated worms were able to survive for more than 10 days, which made possible the combination of anterior and posterior amputations to assess the effects of the absence of the brain on posterior regeneration.

*Morphological characterization of posterior regeneration after both anterior and posterior amputations*

Seminal older studies convincingly showed that the removal of the brain impaired posterior regeneration in nereid annelids, but these studies were somehow limited as animals were not always precisely staged and, importantly, no distinction was generally made between regeneration *per se* and post-regenerative posterior growth. To precisely reassess the influence of the absence of the brain hormone on posterior regeneration in *P. dumerilii*, we performed experiments combining anterior amputation followed by posterior amputation. We used a total of 336 3-4-month-old worms able to survive 14 days after anterior amputation (see previous section). We performed anterior amputation (Fig. 1A, black diamond) at different time points before posterior amputation (Fig. 1A, red diamond), as we did not know for how long the brain hormone is present in the animal after decapitation. Seven experimental conditions (316 amputated individuals in total) plus one control experiment (only posterior amputation) were designed for a total duration ranging from 7 to 14 days (Fig. 1A). Day 0 always corresponds to the day when posterior amputations were performed. In condition C0 (n= 43), both anterior and posterior amputations were performed the same day (day 0). In condition C1 (n= 49), anterior amputation was performed one day before posterior amputation (day -1). C2 to C7 experiments (n= 43 for C2, n= 43 for C3, n= 37 for C4, n= 43 for C5, n= 36 for C6, n= 32 for C7) were similarly designed and worms were scored every day for seven days post posterior amputation (dppa) (Fig. 1A). Morphology of the regenerated region was also monitored and 131 individuals with abnormal morphologies, as impossible to assign to a precise stage were analyzed separately (Fig. 2, Supp. Fig. 5).

The overall results of these experiments are depicted in Fig. 1B. Detailed statistics can be found in Supp. Table 1. Stage 1 (wound healing achieved), was reached at 1 dppa in all conditions, like in the control, and as no morphological abnormality was observed, this indicates that wound healing is not affected by the absence of the head. Worms in all conditions also reached stage 2 (i.e. presence of a regenerated anus and a small blastema).
At 3 dppa, worms' posterior regeneration is significantly delayed in all conditions and none of them were able to complete it (i.e., to reach stage 5) and produce segments at the end of the experiment (at 7 dppa), in contrast to the controls. The importance of the posterior regeneration delay was correlated to the moment when anterior amputation is performed. Indeed, C0 and C1 worms were the less affected ones and some reached stage 4 at 7 dppa, unlike worms from all the other experimental conditions. In addition to a delay in posterior regeneration, various abnormal morphologies were also observed in all experimental conditions from 3 dppa onwards (Fig. 2, Supp. Fig. 5), while not found in the control worms. From 3 dppa onwards, several worms displayed an enlarged regenerated region (Fig. 2, red arrowheads) and some showed one or two reduced or misplaced outgrowths at the posterior end of the pygidium (Fig. 2, pink and blue arrows, respectively). In some worms, additional tiny outgrowths often located next to two well-developed anal cirri, were also observed (Fig. 2, green arrows). C5 was the only condition in which worms with abnormal morphologies (~5% of the total number of worms) were observed at 3 dppa (Supp. Fig. 5). At 4 dppa, worms with abnormal morphologies were found in C3 to C7 conditions (Supp. Fig. 5), with a proportion ranging from ~3% to ~13% (Supp. Fig. 5). Abnormalities were also observed in C2 at 5 dppa (~18%), and in C0 and C1 at 6 dppa (~9% and ~18%, respectively) (Supp. Fig. 5). At the end of the experiment (7 dppa), the conditions that resulted in the highest number of abnormal regenerated worms were C2, C4, and C5 (Supp. Fig. 5A), which corresponds to ~60% of the total observed specimens (Supp. Fig. 5B).

In summary, our data show that, while not affecting early steps of regeneration, the absence of the head prevents regeneration from proceeding beyond stage 3 and leads to morphological abnormalities in the regenerated region. The severity in the delay in regeneration increases with the time period between anterior and posterior amputations. The number of worms with altered morphologies of the regenerated region increases with time after posterior amputation.

*Molecular characterization of posterior regeneration after both anterior and posterior amputations*

To further understand defects in *P. dumerilii* posterior regeneration induced by anterior amputation, we performed whole-mount *in situ* hybridizations (WMISH) with nine molecular markers whose expression during posterior regeneration was previously characterized (Planques et al. 2019) on C2 (anterior amputation two days before posterior
amputation) and control (posterior amputation only) worms. We chose C2 since worms
in this condition were strongly delayed in their regeneration and, while displaying some
abnormalities, had regenerated regions with morphologies that are not too severely
altered, to be comparable with the controls. Representative images of the expression of
the genes at five time points after posterior amputation are shown in Figs. 3 and 4.

We first studied four genes whose expression allowed us to monitor the
reformation of different structures and tissues during posterior regeneration. *Pdum-hox3*,
a growth zone marker, is first expressed in the wound epithelium at 1 dppa (Fig. 3A1,
blue arrows) and, from 2 dppa onwards, in the growth zone stem cells (Fig. 3A2-A5, blue
arrowheads). A similar expression was found in C2 worms at 1 and 2 dppa (Fig. 3B1-2,
blue arrows and arrowheads), but at 3 and 5 dppa, a slightly extended and thicker
expression on the lateral sides of the regenerated region was observed (Fig. 3B3, B4; blue
arrowheads and asterisks). A large asymmetrical expression, completely different to that
found in controls, was observed at 7 dppa (Fig. 3B5; white asterisk). These data therefore
suggest that, while probably initially regenerated in a more or less normal fashion, the
growth zone is not properly maintained at later stages in C2 worms, which may explain
why these worms do not produce new segments. *Pdum-wnt1*, a marker of the terminal
structures of the worms (anus and pygidium), is expressed at 1 dppa in the wound
epithelium (Fig. 3C1; blue arrows), in the posterior-most part of the gut (Fig. 3C2-C5;
brown arrows) and in the pygidial epidermis (Fig. 3C2-C5; brown arrowheads) from 2
dppa onwards, and in faint ectodermal stripes at 5 and 7 dppa (Fig. 3C4, C5; black
arrowheads) which corresponds to segment primordia. In C2 worms, at 1 dppa, we found
*Pdum-wnt1* expression to be restricted to the lateral part of the wound epithelium (Fig.
3D1; faint blue arrows), while at 2 dppa, a strong expression in the wound epithelium was
observed (Fig. 3D2; blue arrows), and no expression in the gut can be detected (in contrast
to control). At 3 dppa, the gene is expressed in a large extended posterior expression
domain and includes both epidermal and posterior gut cells (Fig. 3D3; brown arrowheads
and arrows). At 5 and 7 dppa (Fig. 3D4, D5), *Pdum-wnt1* is expressed both in the posterior
gut cells (brown arrow) and in the prospective pygidium, in an extended fashion (brown
arrowheads). No epidermal segmental stripes of *Pdum-wnt1*-expressing cells were
observed, confirming the impairment of segment production in C2 worms. *Pdum-wnt1*
expression also suggests that the overall patterning of the regenerated region, notably the
pygidium, is altered in C2 worms.

*Pdum-ngn* is a nervous system marker expressed in neural progenitors, first in a
few lateral cells, at 1 and 2 dppa (Fig. 3E1, E2; purple arrowheads), and, from 3 dppa onwards, in many neural precursors of the developing ventral nerve cord (Fig. 3E3-E5, purple arrows) and in sensory cells in the anal cirri (Fig. 3E3, E4, purple arrowheads). In C2 worms, *Pdum-ngn* expression was not detected at 1 dppa (Fig. 3F1) but was roughly normal at 2 dppa (Fig. 3F2, purple arrowheads). The number of *Pdum-ngn*-expressing cells was strongly reduced, as compared to controls, in later stages (Fig. 3F3-F5), strongly suggesting that nervous system formation is impaired in C2 worms in which some neurogenesis does however occur both in the anal cirri (Fig. 3F3, F4, purple arrowheads) and in the ventral nerve cord (Fig. 3F5, arrows). *Pdum-twist* is a muscle marker expressed in muscle progenitors and their differentiating progeny. It is weakly expressed in internal cells at 1 dppa (Fig. 3G1; red asterisks), then in a ring of posterior cells at the origin of the pygidial muscles from 2 to 5 dppa (Fig. 3G2-G4; red arrowheads), and in segmental somatic muscles from 3 dppa onwards (Fig. 3G3-G5; red arrows). Expression of *Pdum-twist* appeared roughly normal in C2 worms at 1 dppa (Fig. 3H1, red asterisks). Expression in putative pygidial muscle precursors was observed at the next stages, in a pattern that however differs from that of control animals (absence of ring shape and/or extended expression, Fig. 3H2-H4, faint red arrowheads), suggesting that pygidial muscles are produced in an abnormal way. Expression of *Pdum-twist* was also found in cells located more anteriorly in the regenerated region (Fig. 3H3-H5, faint red arrows), which may be somatic segmental muscles. However, no segmental stripes of *Pdum-twist* were observed, suggesting that while some myogenesis occurs, muscle formation is nevertheless strongly altered in C2 worms.

We next studied the expression of two genes linked to cell proliferation, *Pdum-pcna* and *Pdum-cycB1* (Fig. 4A-D). At 1 dppa, *Pdum-pcna* is expressed in two large groups of cells close to the wound epithelium (Fig. 4A1; orange arrows), while *Pdum-cycB1* is only expressed in a few cells in this same region (Fig. 4C1; orange arrows). From 2 dppa onwards, the two genes become largely expressed in the blastema (Fig. 4A2, A3, C2, C3, green arrows) and subsequently in the developing segments (Fig. 4A4, A5, C4, C5, green arrowheads) as well as in the growth zone stem cells (Fig. 4A2, A3, C3-C5, green asterisks). Until 3 dppa, no convincing alterations of the expression patterns of those two genes were found in C2 worms (Fig. 4B1-B3, D1-D3). From 5 dppa onwards, in contrast, the number of cells expressing the two genes was strongly reduced in C2, and their overall expression patterns are strikingly altered (Fig. 4B4, B5, D4, D5, faint green arrowheads) with no expression in the growth zone stem cells. In addition, we performed
EdU labelling experiments, to precisely quantify the proportion of cells in S phase, for both C2 and control worms at 2 stages, 3 and 5 dppa (Supp. Fig. 6A, B, D and E, respectively). At 3 dppa, the percentage of EdU positive cells is around 30% in both C2 and control worms, even if the morphology of the regenerated structures are not similar. In contrast at 5 dppa, the percentage of EdU positive cells is significantly lower in C2 worms (around 10%) compared to controls (around 30%) (Supp. Fig. 6C and F). These data show that in C2 worms head amputation does not prevent cell proliferation during early stages of regeneration but at later stages, proliferation is likely strongly reduced in link with the failure to properly form new segments as shown in the previous sections.

We lastly studied the expression of three genes expressed in the stem cells of the growth zone, as well as in proliferating cells of the developing segments: *Pdum-piwiA*, *Pdum-vasa* and *Pdum-myc*. *Pdum-piwiA* becomes expressed at 2 dppa in two bilateral groups of internal cells (Fig. 4E2, orange arrows), its expression extends to most blastemal cells at 3 dppa (Fig. 4E3, green arrows) and is detected in cells of the mesodermal part of the growth zone (Fig. 4E3, green asterisk). At 5 and 7 dppa, *Pdum-piwiA* continues to be expressed in the mesodermal growth zone (Fig. 4E4, E5, green asterisk), as well as in the mesodermal cells of the developing segments (Fig. 4E4, E5, green arrowheads) while no expression is found in the pygidium (Fig. 4E5). In C2 worms, no major abnormalities in the expression pattern of *Pdum-piwiA* were found until 3 dppa, although, as reported for *Pdum-pcna*, the expression level seemed to be reduced (Fig. 4F1-F3). At 5 and 7 dppa, *Pdum-piwiA* expression pattern is strikingly different from controls, with no segmental expression, a broad intense expression including in the pygidium (Fig. 4F4, F5, double faint green arrowheads) and no expression in mesodermal growth zone stem cells was detected. At 1 dppa, *Pdum-vasa* expression is restricted to two lateral patches of ectodermal cells (Fig. 4G1, orange arrows) in control worms and from 2 dppa onwards, its expression is similar to that of *Pdum-piwiA* (Fig. 4G2-G5). In C2 animals, *Pdum-vasa* expression was found to be roughly normal at 1 and 2 dppa (Fig. 4H1, H2). At 3 dppa, no clear expression in the mesodermal growth zone stem cells is observed (Fig. 4H3), while a broad expression in the blastema, similar to that in controls is present (Fig. 4H3, green arrows). From 4 dppa onwards, the expression of the gene in C2 worms was strongly reduced, with no clear expression in the mesodermal growth zone stem cells (Fig. 4H4, H5, faint green arrowheads). In control worms, *Pdum-myc* is first expressed in a few scattered cells (at 1 dppa, Fig. 4I1, orange arrows), then strongly expressed at 2 dppa in two groups of posterior cells, where anal cirri will form (Fig. 4I1,
orange arrows) and weakly in more anterior blastemal cells (Fig. 4I2, green arrows). From 3 dppa onwards (Fig. 4I3-I5), *Pdum-myc* is expressed in the growth zone stem cells (Fig. 4I3-I5, green asterisk), at basis of the anal cirri (Fig. 4I3, I4, purple arrowheads), and both in mesodermal and ectodermal cells of developing segments (Fig. 4I4, I5, green arrowheads). In C2 worms, at 1 dppa, *Pdum-myc* was found to be expressed in two small groups of lateral cells (Fig. 4J1, orange arrows) and from 2 dppa onwards, we found that its expression pattern strikingly differed from that of controls (Fig. 4J2-J5). A ring of expressing cells was observed at 2 dppa (Fig. 4J2, blue arrows), a broad and diffuse expression at 3 dppa (Fig. 4J3, orange arrows), and at 5 and 7 dppa, an expression that seemed to be restricted to the growth zone and some adjacent cells (Fig. 4J4, J5, green asterisk). No segmental expression could be detected. The expressions of the three studied stem cell genes were therefore strikingly modified in decapitated worms suggesting defaults in growth zone regeneration and subsequent segment formation.

In summary, the molecular characterization of the defects observed during *P. dumerilii* posterior regeneration following an anterior amputation highlights how the overall patterning of the regenerated regions is altered, especially in late regeneration stages. While some neurogenesis and myogenesis occur, the pygidium does not differentiate properly and the growth zone, while probably initially regenerated, does not function normally. This is in line with the fact that cell proliferation is highly reduced and stem cells markers are mis- and/or under-expressed. Altogether, this leads to the non-production of new segments and therefore to an impairment of the post-regenerative posterior growth process.

## Effects of anterior amputation on post-regenerative posterior growth

We wanted to further evaluate the effects of brain removal on post-regenerative posterior growth in *P. dumerilii*, and performed a set of experiments combining posterior amputation followed by anterior amputation (Fig. 5A, red diamond and black diamond, respectively) after the regeneration of the growth zone (i.e. after 3 dppa for 3-4-month-old worms (Planques et al. 2019)). Three experimental conditions (total of n=92) plus one control experiment (n=38; only posterior amputation) were designed (Fig. 5A) and as before, posterior amputation were performed on day 0. In condition C3, anterior amputation was performed 3 days post posterior amputation (3 dppa), conditions C4 and C5 were designed similarly. Worms were monitored and scored every day for ten days and the overall results of these experiments are depicted in Fig. 5B (detailed statistics can
be found in Supp. Table 1). Morphology of the regenerated region was also monitored and abnormal individuals with abnormal morphologies, as impossible to assign to a precise stage were analyzed separately (Supp. Fig. 7).

As expected, control worms show no sign of segment addition between 1 to 5 dppa (Fig. 5B and Supp. Table 1) but at 6 dppa, the great majority of them (84,6%) do present new segment(s) produced by the regenerated growth zone (Fig. 5B). From 7 dppa onwards, all control worms have added at least one new segment (Fig. 5B). C3, C4 and C5 worms show no sign of segment addition between 1 to 5 dppa (Fig. 5B) but in striking contrast to control worms, they showed a much-delayed posterior growth in subsequent stages. For the C3 condition, new segment start to be produced for a small proportion of worms from 8 dppa onwards. In contrast C5 worms present at least one segment from 6 dppa onwards (Fig. 5B) and the majority of them present post-regenerative posterior growth at 8-10 dppa (57.7% at 8 and 9 dppa, and 73.1% at 10 dppa). The proportion of worms that were able to produce new segments and the timing of the beginning of post-regenerative posterior growth is thus highly correlated with the moment when the anterior amputation was performed. We also observed some worms with abnormal morphologies from 6 dppa onwards in C3 and C4 conditions and for all conditions from 7dppa onwards (Supp. Fig. 7A, B) while no abnormalities were observed in the control worms.

We performed an additional experiment in which we assessed the number of new segments that have been produced after anterior amputation for the C5 condition (n=18), (or no additional amputation for the control worms n=28) (Fig. 5C) and scored them until 15 dppa. From 8 dppa onwards, significant differences between C5 and control worms were observed: C5 worms only added one more segment during the seven following days (until 15 dppa), while control animals produced an average of five segments during the same time period (Fig. 5C). Unexpectedly, no abnormal morphologies were found until 11 dppa for C5 worms (no abnormalities were observed in the control worms) (Supp. Fig. 7C, D)

Altogether, these results show that after anterior amputation post-regenerative posterior growth is drastically affected and that a very limited number of new segments is produced. To further characterize these defects, we performed WMISH on C5 and control worms at 10 dppa for *Pdum-pcna* to monitor cell proliferation, *Pdum-hox3* to visualize the stem cells of the regenerated growth zone, and *Pdum-en* (*Pdum-engrailed*) to follow the production of new segments and therefore the activity of the growth zone (Fig. 6). As expected, control and C5 worms harbor a quite different morphology, as very
few post-regenerative segments have been produced in C5. In control worms, *Pdum-pcna* is broadly expressed in the developing segments (Fig. 6A, green arrowheads) while in C5 ones a similar expression is observed, yet in a smaller region given the reduced number of segments (Fig. 6A', green arrowheads). The ring-shaped expression of *Pdum-hox3* in the growth zone located at the anterior margin of the pygidium, found in the control worms (Fig. 6B, blue arrowheads), is also present in the C5 worms (Fig. 6B', blue arrowheads), suggesting the presence of the growth zone in the C5 worms. Finally, while several segmental stripes of *Pdum-en* expression delineating the newly produced segments are found in the control worms (Fig. 6C, brown arrowheads), only one or two such stripes are present in the C5 worms (Fig. 6C', faint brown arrowheads), suggesting a defect in the production of segments by the growth zone and therefore alterations of the function of the growth zone stem cells.

In summary, we have shown that after anterior amputation post-regenerative posterior growth is deeply affected, and the production of post-regenerative segments is extremely limited. These drastic defaults are not due to the absence of the growth zone but are likely due to an altered functioning of those cells.

*Effects of exogenous methylfarnesoate (MF) on posterior regeneration after both anterior and posterior amputations*

Methylfarnesoate (MF) has been reported to likely be the putative brain hormone controlling sexual maturation and suggested to also be involved in the control of posterior regeneration and growth in *P. dumerilii* (Schenk et al. 2016). We have shown (see previous sections) that, while not affecting early steps of posterior regeneration, the amputation of the head prevents regeneration from being completed and leads to morphological abnormalities in the regenerated region. To assess whether the defects in posterior regeneration after head amputation are due to the absence of MF, we tested if these defects can be rescued by the exogenous addition of MF hormone, as it was the case when supraoesophageal ganglia from intact worms were implanted in decerebrated host (e.g., Scully 1964; Golding 1967b). As in the previous experiments, we used 3-4-month-old worms and compared three sets of worms. Worms of the “control 1” condition (n=29) were amputated only posteriorly at day 0 and display a normal posterior regeneration process; worms of the “control 2” condition (n=27) were posteriorly and anteriorly amputated at day 0, and present delay and defects in posterior regeneration (these worms correspond to the C0 condition in Figure 1); MF-treated worms (n=51) were similarly
posteriorly and anteriorly amputated at day 0 but were subsequently incubated in sea water containing 100 nM of MF hormone, renewed every 24 hours (Fig. 7A). The three sets of worms were scored for the regeneration stage that was reached every day for five days (until 5 dppa). The overall results of these experiments are depicted in Fig. 7B. Morphology of the regenerated region was also monitored (Supp. Fig. 8) and abnormal individuals with morphologies difficult to assign to a precise stage were excluded from the statistical analysis.

As expected, from 4 dppa onwards, control 2 worms were significantly delayed as compared to the control 1 condition (Fig. 7B and Supp. Table 1). Interestingly, at 4 dppa, MF-treated worms were not significantly different to neither control 1 nor control 2 worms. At 5 dppa, however, MF-treated worms were significantly delayed compared to the control 1 worms but also regenerated significantly faster than the control 2 worms (Fig. 7B and Supp. Table 1). These results therefore indicate that the addition of exogenous MF positively affect posterior regeneration in headless worms. However, as MF-treated worms did not regenerate as the control 1 worms, it also indicates that MF is not completely reverting the defect produced by head amputation. We observed a similar proportion of worms with abnormal morphologies both in control 2 (13%) and MF-treated worms (15%) (Supp. Fig. 8 and Supp. Table 1), further indicating that the treatment with exogenous MF cannot fully compensate for the absence of the head.

Discussion

*Brain hormonal activity is required for both posterior regeneration and post-regenerative posterior growth in the annelid P. dumerilii*

Studies mostly conducted in the fifties and sixties, pointed out the important role of the brain and its hormonal production in the reproductive and regenerative processes of nereid annelids (see Introduction for details and references). These studies were however performed on worms that were not raised in standardized laboratory conditions, often did not make a distinction between posterior regeneration and post-regenerative posterior growth (e.g., Golding 1967a – see discussion about the differences between the two processes in Planques et al. 2019), which may lead to biases in the conclusions drawn, and lack of molecular data, which clearly limited the understanding of the regeneration defects that were observed after head amputation or brain removal. The recent use of molecular and cellular markers has made an in-depth characterization of both normal
posterior regeneration and post-regenerative posterior growth possible (Gazave et al. 2013; Planques et al. 2019) and provides invaluable tools to further characterize the role of the brain during both processes. We therefore re-assessed and further analyzed the role of the brain hormonal activity on posterior regeneration and post-regenerative posterior growth in *P. dumerilii* (i) using homogenous worms in terms of size and age and raised in stereotypical laboratory conditions and (ii) combining morphological and molecular data to increase our understanding of this role. We performed anterior amputations that remove the brain, at different time points before or after posterior amputation, to assess their effects on posterior regeneration and post-regenerative posterior growth, respectively.

Effects on posterior regeneration:

To investigate the effect of anterior amputation on posterior regeneration, we first
performed anterior amputations either at the same time than posterior amputation
(condition C0) or at six different time points (from one day to seven days) before posterior
amputation (conditions C1 to C7; Fig. 1A). We found that in all conditions, in contrast to
control worms, no worm whose head was amputated was able to complete posterior
regeneration even 7 days post posterior amputation (dppa), in addition, many of them
show morphological abnormalities (Fig. 1B, 2). These results confirm the absolute
requirement of the head (and thus likely the brain) for proper posterior regeneration. A
second important observation is that the severity of the delay in posterior regeneration
increases with the time period between anterior and posterior amputations (Fig. 1). This
is consistent with the hypothesis that anterior amputation impairs posterior regeneration
by causing a decrease in the concentration of the brain hormone in the blood and/or tissues
of the worms, due to the absence of the brain where this hormone is produced. Significant
differences in the regeneration stages reached by the worms were observed between C0
and C1 conditions, and all other conditions (C2-C7) (Supp. Table 1), suggesting a rapid
decrease in the hormone concentration in the absence of the brain. A third compelling
observation is that in all conditions, headless worms were able to reach stage 1 (wound
healing achieved) and stage 2 (a small blastema is formed), suggesting that these early
steps of the regeneration process therefore likely occur irrespective of the presence or
absence of the brain hormone. Further progression through the following steps of
regeneration, in contrast, does not happen in the absence of the head, suggesting that these
later steps require the activity of the brain hormone.
The analysis of several genes' expression that are well-known markers of specific tissues/structures during posterior regeneration in C2 worms allowed us to further characterize the defects in the posterior regeneration process caused by anterior amputations (Fig. 3). Firstly, we showed that the normal expression of the cell cycle genes *Pdum-cycB1* and *Pdum-pcna* at early stages of posterior regeneration in C2 worms (until 3 dppa; Fig. 4 A1 to D2) suggests that head amputation does not block cell proliferation during these early stages. It also suggests that the defects caused by head amputation are not primarily due to the blocking of cell proliferation but rather to a patterning default. Accordingly, the expression patterns of *Pdum-hox3*, *Pdum-twist*, and *Pdum-neurogenin* in C2 worms clearly differ from those of these genes in worms that have been treated with the cell proliferation inhibitor hydroxyurea after posterior amputation (Planques et al. 2019), indicating that head amputation and the inhibition of cell proliferation have distinct effects on posterior regeneration. At 5 and 7 dppa, the expression of *Pdum-cycB1* and *Pdum-pcna* are strikingly reduced, showing a decrease in cell proliferation at these later stages, which is potentially due to the failure to produce segments (Fig. 4 A4 to D5). Abnormal expression patterns of three stem genes previously shown to be expressed in proliferative cells, *Pdum-piwiA*, *Pdum-vasa*, and *Pdum-myc* were also observed in C2 worms at these late time points, confirming major defects in the pattern of proliferative cells caused by head amputation.

Secondly and importantly, the posterior growth zone is initially regenerated in C2 worms as seen by the proper expression of *Pdum-hox3* (a growth zone marker) at early time points of posterior regeneration (1 to 2 dppa, Fig. 3 A, B). The regenerated growth zone is however not able to produce any segment as seen by the absence of visible segment or stripes of cells expressing *Pdum-wnt1* (a marker of segmentation) at 5 and 7 dppa (Fig. 3, C, D), which may be due to its improper maintenance in the absence of the head, as suggested by the increasingly modified expression of *Pdum-hox3* from 3 dppa onwards (Fig. 3A,B). The expression of *Pdum-wnt1*, *Pdum-twist* and *Pdum-neurogenin* highlights the fact that several other structures or tissues (e.g., wound epithelium, pygidium, muscles and nervous system) of the regenerating posterior part are also severely affected in C2 worms (Fig. 3C-H). Taken together, these observations strongly suggest that the brain hormonal activity is required for many aspects of posterior regeneration including the formation of a proper wound epithelium and the efficient regeneration of the growth zone (and its subsequent ability to produce segments).

*P. dumerilii*'s growth zone is a structure composed of very probably stem cells
with high potency, that can be regenerated (Gazave et al 2013; Planques et al 2019). Those stem cells express the so-called GMP signature (for Germline Multipotency Program), found in many other stem and germ cells in animals (Juliano et al., 2013; Rinkevich et al., 2022). While the presence of *Hox3* expression reveals that in the absence of brain hormone, the growth zone regeneration occurs, the misexpression of the GMP markers, in particular *myc*, highlights the fact that its molecular identity is probably altered. To which degree the growth zone stem cell molecular identity is modified is an interesting point that will require more investigation in the future. Indeed, with the democratization of single-cell RNAseq technology in various contexts, notably regeneration (Bideau et al., 2021), it will be soon possible to compare the molecular signature of those cell populations, under the brain hormone control, or in its absence.

## Effects on post-regenerative posterior growth:

To investigate the effect of anterior amputation on post-regenerative posterior growth, we first performed anterior amputations 3, 4, or 5 days after posterior amputation (Fig. 5A). We observed a striking reduction of the number of worms undergoing post-regenerative posterior growth from 6 dppa to 10 dppa in all three experimental conditions (Fig. 5B). Post-regenerative posterior growth is therefore clearly affected by anterior amputation. Two additional observations can be done: (i) the defects are more severe if the anterior amputation is done earlier; (ii) some of the decapitated worms were able to produce at least one new segment. These observations can be explained by the fact that the regenerated growth zone has already produced one segment primordium at 3 dppa and two at 5 dppa as shown by the expression of *Pdum-engrailed* (Planques et al. 2019). In the C3 condition, anterior amputation has therefore been done in worms that have not all produced a segment primordium or whose segment that has been produced is still in a very early step of its formation. In the two other conditions, especially C5, head amputation is performed in worms that have already one or two segment primordia produced at earlier time points, which would explain why more worms display post-regenerative posterior growth at later time points such as 10 dppa.

We further characterize the C5 condition and observed that these worms, form at most two segments (in 15 dppa, while control added an average of five segments) (Fig. 5C). This strongly suggest that the C5 worms do not produce any segments after head amputation and therefore that the head, and thus the brain hormone is fully required for
post-regenerative posterior growth. Accordingly, only one or two stripes of *Pdum-engrailed* were observed in C5 worms at 10 dppa (Fig. 6). Our data also suggest that segments whose primordium has been produced before head amputation may still differentiate in the absence of the head, suggesting that the brain hormone would be required to produce the segment and/or early steps of its development, but not for its full differentiation. Alternatively, the potential remaining of low quantities of brain hormone in the body during the few days that follow head amputation might be sufficient to allow the differentiation of segments produced by the regenerated growth zone before head amputation, but not to sustain the formation of new segments after head amputation. Importantly, the normal expression of *Pdum-hox3* in C5 worms at 10 dppa suggests that the defect in segment formation is not due to the disappearance or degeneration of the growth zone but rather in some alterations of its functioning.

How the brain hormone is regulating the growth zone stem cells functioning is still an open question. Accumulating evidence, from flies and mammals notably, support the idea that hormones can finely regulate the functions of various stem cells (germ cells, intestinal, neural, hematopoietic, embryonic stem cells ...) and their niches (Gancz and Gilboa, 2013). Hormonal cues control notably the maintenance, differentiation and proliferation of stem cells, often though downstream effectors, notably the key signaling pathways Wnt and Notch. Investigating the role of such pathways during posterior elongation, in the presence and absence of brain is an interesting avenue to follow in the future.

### Involvement of methylfarnesoate (MF) in *P. dumerilii* posterior regeneration

A wealth of old studies has led to the hypothesis that the brain hormonal activity, which would consist of a single hormone being continuously secreted by the brain until sexual maturation, would exert both a positive control on posterior growth and regeneration and a negative control of sexual maturation (see Introduction for details and references). While the molecular nature of the brain hormone has remained elusive and controversial for many years, a recent study has identified methylfarnesoate (MF) as a most likely candidate for being the *P. dumerilii* brain hormone (Schenk et al. 2016). While MF was convincingly shown to act as a repressor of sexual maturation, its potential role in mediating regeneration remains more elusive.

We therefore tested whether the exogenous addition of MF hormone might rescue
the impairment of posterior regeneration caused by the absence of the brain hormone. We
found that MF-treated worms regenerate significantly faster than decapitated animals not
treated with MF but significantly slower than worms that have not been decapitated (Fig.
7B), indicating that the exogenous addition of MF was able to partially rescue the defects
caused by the absence of the head. These results provide a clear support to the hypothesis
that MF can mediate the positive control exerted by the brain on posterior regeneration,
reinforcing the previous claim that MF is the brain hormone and that it can act on both
regeneration and sexual maturation (Schenk et al. 2016). The rescue of the regeneration
defects caused by head amputation that we obtained with MF treatments was however
only partial. This appears different from what has been obtained in previous studies
involving the implantation of supraoesophageal glands in decerebrated worms, which
were described as leading to a seemingly complete capacity to regenerate (e.g., Golding
1967b). This discrepancy could be due to the experimental design of the experiment, as
MF is provided from the sea water in which the worms are incubated and not directly in
the internal medium of the animals. We do not know what the stability of MF in sea water
is, nor what the efficiency of the uptake of the compound by the worm cells is. It is
therefore possible that the concentration of active MF in the internal medium of the
worms in our experimental conditions is inferior to that in non-decapitated worms or
decerebrated worms in which a supraoesophageal ganglion has been grafted. One
possible explanation would therefore be that the amount of MF present in the body might
be not high enough to completely revert the defects in posterior regeneration caused by
the lack of the brain. We used a concentration of MF which was shown to have a strong
negative effect on Vitellogenin production in whole worms (Schenk et al 2016). It is
however conceivable that different quantities of MF may be required for the different
functions in sexual maturation and regeneration that this hormone fulfills. We may also
not exclude that the brain hormonal activity may involve (an)other hormone(s) than MF
and therefore that supplementing MF alone is insufficient to compensate for the absence
of the brain and rescue the defects in the posterior regeneration process due to this
absence.

In addition, we should also stress out the fact, that worm decapitation induces the
removal of much more than the brain itself. Many sensory structures, such as antennae,
palps, cirri, eyes and many nerves are also eliminated from this procedure, in contrast to
the original decapitation performed in old studies on big individuals. Such structures and
in particular nerves, are known to play critical roles during many contexts of regeneration, notably in annelids (Sinigaglia and Averof, 2019). In *P. dumerilli*, removal of the ventral nerve chord induces the loss of dorso-ventral polarity in the regenerated region (Boilly et al., 2017). But above such example of local nerve-dependency, nerves could also provide distant support for other cell types, or systemic signals, important for regeneration to succeed. If such type of additional signals exists, in the case of a full head decapitation, we eliminated them, preventing the complete regeneration to proceed even in the presence of exogenous MF. In *P. dumerilli*, conditional and specific cell ablation is now available (Veedin-Rajan et al., 2013) and such technic could be a powerful approach, in the future, to specifically remove the supraoesophageal ganglion, as well as other sensory and nervous structures, in order to evaluate their respective importance for posterior regeneration and elongation.

*Hormonal control of regeneration – is MF playing a conserved role beyond Nereididae?*

While less studied than molecular cues, endocrine regulation of blastema-based regeneration is a key process to understand regeneration potential in animals (Easterling et al., 2019 a & b). Both in vertebrates and non-vertebrates, compelling evidence support their role in each step of regeneration *ie*: wound healing, blastema formation and morphogenesis. For instance, a steroid hormone, the ecdysone, induces the restriction of regenerative potential at a specific developmental stage of *Drosophila* (Narbonne-Reveau and Morange, 2019). In various vertebrates, notably amphibians, thyroid hormone inhibits blastema formation (Easterling et al., 2019 a & b). In the annelid *P. dumerilii*, we have shown here that the brain hormone, previous identified as methyl farnesoate (MF) – a sesquiterpenoid hormone (Schenk et al., 2016), not only regulates reproduction, but positively impacts regeneration as well. The sesquiterpenoid hormonal system is supposed to be ancestral to Metazoa, while showing variations between lineages (So et al 2022). More specifically, MF, already known as the non-insect juvenile hormone, has been proposed, to play a role in controlling development and reproduction in protostomes. MF receptor has been identified in the annelid *Syllis magdalena* and thus, MF was proposed to play a role in the initiation of its reproductive process by stolonization (Álvarez-Campos *et al*., 2019). MF has been also directly identified in head extracts of an earthworm (*Lumbriculus sp.*) (Schenk *et al*., 2016). Both species being able to posteriorly regenerate, it would be interesting to investigate if MF has a function on this
process. As *S. magdalena*, like *P. dumerilii*, pertains to one of the main group of annelid,
the Errantia, while *Lumbriculus*. *sp*. in contrast is a member of the Sedentaria, this would
help us to understand is MF could have an ancestral function in controlling regeneration
in annelids. Comparative analyses made on a couple of additional annelid species, as well
as some mollusks, will highlight in the near future, the importance of hormonal control
of regeneration in the yet understudied group of protostomes.

Conclusions

We re-addressed the role of the brain hormonal activity on posterior regeneration
and post-regenerative posterior growth in the annelid *P. dumerilii*. We show that the
removal of the head prevents posterior regeneration from being successful and impairs
multiple aspects of this regenerative process. We also show that post-regenerative
posterior growth is severely affected in the absence of the brain. Indeed, while the
posterior growth zone is apparently regenerated, its functioning is affected in decapitated
worms. Our results therefore strongly suggest a key requirement of brain hormonal
activity for regeneration and growth in *P. dumerilii*. We also show that exogenous
methylfarnesoate (MF), which was proposed to be the *P. dumerilii* brain hormone and
shown to act on worm reproduction, partially rescues posterior regeneration defects in
decapitated worms, supporting the hypothesis that MF mediates the positive control of
the brain on posterior regeneration. Our study therefore extends our knowledge about the
control of posterior regeneration by the brain in *P. dumerilii*, paving the way for future
mechanistic analysis of hormonal control of regeneration in this species and beyond.

Acknowledgements

We thank all Vervoort lab members for helpful discussions and feedback on the
manuscript. We are much grateful to Sven Schenk for his invaluable advice about the
treatments with methylfarnesoate. We thank Florian Raible and Alexander Stockinger for
their valuable comments on the manuscript. We thank Haley Flom for her diligent
proofreading of the manuscript. We thank Patrick Beckers for his useful advices to make
a proper diagram showing the internal anatomy of *P. dumerilii's* head. We acknowledge
the ImagoSeine imaging facility (member of the France BioImaging infrastructure
supported by the Agence Nationale de la Recherche, ANR-10-INSB-04, "Investments of
the future"). Animal facility members are thanked for their help with the worm culture.
This work was funded by a collaborative grant jointly attributed by the Agence Nationale
de la Recherche (ANR) and the Austrian Science Fund (FWF) to the Vervoort (Institut
Jacques Monod Paris), Balavoine (Institut Jacques Monod Paris), and Raible (Max Perutz
labs Vienna) teams (grant TELOBLAST no. ANR-16-CE91-0007), Labex ‘Who Am I?’
laboratory of excellence (No. ANR-11-LABX-007), Centre National de la Recherche
Scientifique, Université de Paris, Association pour la Recherche sur le Cancer (grant PJA
20191209482), and comité départemental de Paris de la Ligue Nationale Contre le Cancer
(grant RS20/75-20).

**Conflicts of interest**

The authors declare that there is no conflict of interests.

**Data availability statement**

All the data are included in the manuscript, figures, supplementary figures, and
the supplementary table.

**Figure legends**

**Figure 1. Influence of anterior amputation on *P. dumerilii* posterior regeneration.**

A) Schematic representation of the experimental design used to address the impact of anterior amputation (aa, black diamonds) on the regeneration that follows posterior amputation (pa, red diamonds). A timeline representing the whole duration of the experiment (14 days) and the timepoints for anterior amputation (aa) and posterior amputation (pa) for each condition, as well as the scoring days (green bars, from 1 to 7 days post posterior amputation, dppa), are provided. Control worms were only amputated posteriorly; for C0, both anterior and posterior amputations were done the same day (0 dppa); for C1 to C7, anterior amputation was made 1 to 7 days before posterior amputation (-1 to -7 dppa). B) Graphic representation of the regeneration stages reached by the worms, every day for 7 days after posterior amputation, for each condition (see inset for the color code and the number of worms used per condition). Asterisks above the bar indicate that this condition was significantly different to the other conditions
specified by the asterisk color. Statistics: 2-way ANOVA on repeated measures with
Tukey correction. * 0.0001 > p < 0.05 (for specific p-values, see Supp. Table 1). Error
bar: Standard Deviation (SD)
A

<figure><img src="image_2.png" /></figure>

<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /></figure>

B

<figure><img src="image_5.png" /></figure>
**Figure 2.** Abnormal morphologies observed during *P. dumerilii* posterior regeneration after anterior amputation. Bright-field microscopy images of the most striking abnormal morphologies observed on posterior regeneration at different time points (4 to 7 days post posterior amputation or dppa). For each abnormal morphology shown, the condition of anterior amputation performed (C0 to C7, see Figure 1 and text for details) are mentioned on the image. Black arrows = well-developed anal cirri; red arrowheads = enlargement of regenerated region; pink arrows = reduction of anal cirri; green arrows = additional tiny outgrowths; blue arrows = misplaced anal cirri.
3 dppa

<figure><img src="image_6.png" /></figure>

4 dppa

<figure><img src="image_7.png" /></figure>

5 dppa

<figure><img src="image_8.png" /></figure>

6 dppa

<figure><img src="image_9.png" /></figure>

7 dppa

<figure><img src="image_10.png" /></figure>

C3

<figure><img src="image_11.png" /></figure>

C2

<figure><img src="image_12.png" /></figure>

C0

<figure><img src="image_13.png" /></figure>

C1

<figure><img src="image_14.png" /></figure>

C4

<figure><img src="image_15.png" /></figure>

C3

<figure><img src="image_16.png" /></figure>

C3

<figure><img src="image_17.png" /></figure>

C2

<figure><img src="image_18.png" /></figure>

C5

C4

C4

C7
**Figure 3.** Effects of anterior amputation on the expression of genes involved in segment, organ or tissue patterning and differentiation during posterior regeneration. Whole-mount *in situ* hybridizations (WMISH) for the genes (n=4) whose name is indicated are shown for five time points (1, 2, 3, 5, and 7 days post-posterior amputation or dppa). Control worms are only posteriorly amputated. C2 worms are amputated anteriorly 2 days before posterior amputation. All panels are ventral views (anterior is up). Blue arrows = wound epithelium; blue arrowheads = posterior growth zone; white asterisks = extended or aberrant expression of *Pdum-hox3*; brown arrow = gut posterior-most part; brown arrowheads = pygidial epidermis; black arrowheads = segmental stripes; faint blue arrows = *Pdum-wnt1* restricted expression in the wound epithelium; purple arrowheads = putative sensory cell progenitors; purple arrows = developing ventral nerve cord; red asterisk = early expression in internal cells of *Pdum-twist*; red arrowheads = pygidial muscle cells progenitors; red arrows = segmental muscles; faint red arrowheads = misshapen expression domain of *Pdum-twist* in pygidial muscle cells progenitors; faint red arrows = misshapen expression domain of *Pdum-twist* in segmental muscles.
Pdum-hox3

Control

C2

<figure><img src="image_19.png" /></figure>

A1

<figure><img src="image_20.png" /></figure>

B1

<figure><img src="image_21.png" /></figure>

C1

<figure><img src="image_22.png" /></figure>

D1

Pdum-wnt1

Control

C2

<figure><img src="image_23.png" /></figure>

C2

<figure><img src="image_24.png" /></figure>

D2

Control

C2

<figure><img src="image_25.png" /></figure>

E1

<figure><img src="image_26.png" /></figure>

F1

Pdum-ngn

Control

C2

<figure><img src="image_27.png" /></figure>

G1

<figure><img src="image_28.png" /></figure>

H1

<figure><img src="image_29.png" /></figure>

H2

<figure><img src="image_30.png" /></figure>

H3

<figure><img src="image_31.png" /></figure>

G3

<figure><img src="image_32.png" /></figure>

H4

<figure><img src="image_33.png" /></figure>

G4

<figure><img src="image_34.png" /></figure>

H5

<figure><img src="image_35.png" /></figure>

G5

1 dppa

2 dppa

3 dppa

5 dppa

7 dppa

<figure><img src="image_36.png" /></figure>

A3

<figure><img src="image_37.png" /></figure>

B3

<figure><img src="image_38.png" /></figure>

C3

<figure><img src="image_39.png" /></figure>

D3

<figure><img src="image_40.png" /></figure>

E3

<figure><img src="image_41.png" /></figure>

F3

<figure><img src="image_42.png" /></figure>

A4

<figure><img src="image_43.png" /></figure>

B4

<figure><img src="image_44.png" /></figure>

C4

<figure><img src="image_45.png" /></figure>

D4

<figure><img src="image_46.png" /></figure>

E4

<figure><img src="image_47.png" /></figure>

F4

<figure><img src="image_48.png" /></figure>

A5

<figure><img src="image_49.png" /></figure>

B5

<figure><img src="image_50.png" /></figure>

C5

<figure><img src="image_51.png" /></figure>

D5
**Figure 4. Effects of anterior amputation on the expression of cell proliferation and stem cells genes during posterior regeneration.** Whole-mount *in situ* hybridizations (WMISH) for the genes (n=4) whose name is indicated are shown for five time points (1, 2, 3, 5, and 7 days post-posterior amputation or dppa). Control worms are only posteriorly amputated. C2 worms are amputated anteriorly 2 days before posterior amputation. All panels are ventral views (anterior is up). Orange arrows = proliferating internal cells below the wound epithelium; green arrow = large expression in the blastema; green arrowheads = large expression in the developing segments; green asterisks = growth zone stem cells; faint green arrowheads = reduced expression in the developing segments; double faint green arrowheads = aberrant expression in the pygidium; purple arrowheads = expression at the basis of the anal cirri; blue arrows = extended abnormal expression of *Pdum-myc*.
Pdum-pcna

Control

C2

Pdum-cycB1

Control

C2

Pdum-piwiA

Control

C2

Pdum-vasa

Control

C2

Pdum-myc

Control

C2

1 dppa

<figure><img src="image_52.png" /></figure>

<figure><img src="image_53.png" /></figure>

<figure><img src="image_54.png" /></figure>

<figure><img src="image_55.png" /></figure>

<figure><img src="image_56.png" /></figure>

<figure><img src="image_57.png" /></figure>

<figure><img src="image_58.png" /></figure>

<figure><img src="image_59.png" /></figure>

<figure><img src="image_60.png" /></figure>

<figure><img src="image_61.png" /></figure>

2 dppa

<figure><img src="image_62.png" /></figure>

<figure><img src="image_63.png" /></figure>

<figure><img src="image_64.png" /></figure>

<figure><img src="image_65.png" /></figure>

<figure><img src="image_66.png" /></figure>

<figure><img src="image_67.png" /></figure>

<figure><img src="image_68.png" /></figure>

<figure><img src="image_69.png" /></figure>

<figure><img src="image_70.png" /></figure>

<figure><img src="image_71.png" /></figure>

3 dppa

<figure><img src="image_72.png" /></figure>

<figure><img src="image_73.png" /></figure>

<figure><img src="image_74.png" /></figure>

<figure><img src="image_75.png" /></figure>

<figure><img src="image_76.png" /></figure>

<figure><img src="image_77.png" /></figure>

<figure><img src="image_78.png" /></figure>

<figure><img src="image_79.png" /></figure>

<figure><img src="image_80.png" /></figure>

<figure><img src="image_81.png" /></figure>

5 dppa

<figure><img src="image_82.png" /></figure>

<figure><img src="image_83.png" /></figure>

<figure><img src="image_84.png" /></figure>

<figure><img src="image_85.png" /></figure>

<figure><img src="image_86.png" /></figure>

<figure><img src="image_87.png" /></figure>

<figure><img src="image_88.png" /></figure>

<figure><img src="image_89.png" /></figure>

<figure><img src="image_90.png" /></figure>

<figure><img src="image_91.png" /></figure>

7 dppa

<figure><img src="image_92.png" /></figure>

<figure><img src="image_93.png" /></figure>

<figure><img src="image_94.png" /></figure>

<figure><img src="image_95.png" /></figure>

<figure><img src="image_96.png" /></figure>

<figure><img src="image_97.png" /></figure>

<figure><img src="image_98.png" /></figure>

<figure><img src="image_99.png" /></figure>

<figure><img src="image_100.png" /></figure>

<figure><img src="image_101.png" /></figure>
**Figure 5. Influence of anterior amputation on *P. dumerilii* post-regenerative posterior growth.** A) Schematic representation of the experimental design used to address the impact of anterior amputation (aa, black diamonds) on post-regenerative posterior growth (i.e., following posterior amputation or pa, red diamonds). A timeline representing the whole duration of the experiment (10 days) and the timepoints for anterior amputation and posterior amputation for each condition, as well as the scoring days (green bars, from 1 to 10 days post posterior amputation, dppa), are provided. Control worms were only amputated posteriorly; for C3, anterior amputation was done 3 days after posterior amputation; similarly, for C4 and C5, anterior amputations were done 4 and 5 days after posterior amputation, respectively. B) Graphic representation of the percentage of worms that have undergone post-regenerative posterior growth (addition of at least one new segment) every day for 10 days after posterior amputation, for each condition (the number of scored individuals is indicated next to the condition name). C) Graphic representation of the regeneration stages reached by the worms or the number of newly-added segments (1s = 1 segment, 2s = 2 segments, ...) every day from 6 dppa to 15 dppa in control (only posterior amputation) and C5 (anterior amputation five days after posterior amputation) worms (see inset for the color code and the number of worms used per condition). Mean + SD are shown. Asterisks indicate significant differences as defined by 2-way ANOVA on repeated measures with Dunnett correction. * 0.0001 > p > 0.01 (for specific p-values, see Supp. Table 1).
A

<figure><img src="image_102.png" /></figure>

<figure><img src="image_103.png" /></figure>

Anterior amputation (aa)

<figure><img src="image_104.png" /></figure>

Posterior amputation (pa)

<figure><img src="image_105.png" /></figure>

Scoring days

<figure><img src="image_106.png" /></figure>

Days post posterior amputation (dppa)

<figure><img src="image_107.png" /></figure>

B

<figure><img src="image_108.png" /></figure>

C

<figure><img src="image_109.png" /></figure>
**Figure 6.** Effects of anterior amputation on the expression of cell proliferation, growth zone and segment marker genes during post-regenerative posterior elongation. Whole-mount *in situ* hybridizations (WMISH) for the genes (n=3) whose name is indicated are shown at ten days post-posterior amputation (10 dppa) for C5 (anterior amputation five days after posterior amputation) and control (no anterior amputation) worms. All panels are ventral views (anterior is up). Green arrow = large expression in the regenerating structure; blue arrowheads = posterior growth zone; black arrowheads = segmental stripes; faint black arrowheads = abnormal segmental stripes in a reduced number; black arrows = expression at the border between the last non-amputated segment and the regenerated region.
Pdum-pcna

Pdum-hox3

Pdum-en

Control worms

A

<figure><img src="image_110.png" /></figure>

B

<figure><img src="image_111.png" /></figure>

C

<figure><img src="image_112.png" /></figure>

C5 worms

A'

<figure><img src="image_113.png" /></figure>

B'

<figure><img src="image_114.png" /></figure>

C'

<figure><img src="image_115.png" /></figure>
**Figure 7. Assessment of the effect of methylfarnesoate (MF) on *P. dumerilii* posterior regeneration.** A) Schematic representation of the experimental design used to address the impact of exogenous MF treatment on posterior regeneration. A timeline representing the whole duration of the experiment (5 days) and the timepoints for anterior amputation (aa) and posterior amputation (pa) for three conditions, as well as the scoring days (green bars, from 1 to 10 days post posterior amputation, dppa), are provided. Control 1: posterior amputation, no anterior amputation, no MF treatment; Control 2: posterior amputation, anterior amputation, no MF treatment; MF-treated: posterior amputation, anterior amputation, MF treatment (100nM in sea water, renewed every day). B) Graphic representation of the regeneration stages reached by the worms every day for 5 days after posterior amputation, for each condition (see inset for the color code and the number of worms used per condition). Mean + SD are shown. Significant differences (0.0001 > p < 0.01) are indicated (see inset). Statistics: 2-way ANOVA on repeated measures with Tukey correction.
A

<figure><img src="image_116.png" /></figure>

<figure><img src="image_117.png" /></figure>

B

<figure><img src="image_118.png" /></figure>
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**Supplementary data Legends**

**Supplementary Figure 1. Morphology of *P. dumerilii* worms.** Juvenile (A) and sexually mature (female, B and male, C) worms are shown. Scale bar = 500 µm. (D) Sagittal diagram of the worm showing the most important structures in the prostonium.
<figure><img src="image_119.png" /><figcaption>Prostomium, sagittal view</figcaption></figure>

A

<figure><img src="image_120.png" /></figure>

B

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C

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♀
**Supplementary Figure 2.** Schematic representation with drawings and bright-field microscopy images of posterior regeneration stages in *P. dumerilii* worms, as defined by Planques et al. 2019. 3-4-months-old juvenile worms are posteriorly amputated (removing around 5 segments, the growth zone and the pygidium). One day post posterior amputation (1 dppa), wound healing is completed; at 2 dppa, the formation of the blastema has started; at 3 dppa, a large blastema with small cirri is present; at 4 dppa, a large blastema containing a differentiated pygidium and long anal cirri is present; at 5 dppa, newly-added segments produced by the regenerated growth zone start to become clearly distinct from the pygidium through the appearance of lateral segmental grooves. At 5 dppa, the regeneration is completed and post regenerative posterior growth starts.
<figure><img src="image_123.png" /><figcaption>1dppa</figcaption></figure>

<figure><img src="image_124.png" /></figure>

<figure><img src="image_125.png" /><figcaption>2dppa</figcaption></figure>

<figure><img src="image_126.png" /></figure>

<figure><img src="image_127.png" /><figcaption>3dppa</figcaption></figure>

<figure><img src="image_128.png" /></figure>

<figure><img src="image_129.png" /><figcaption>4dppa</figcaption></figure>

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<figure><img src="image_131.png" /><figcaption>5 dppa</figcaption></figure>

<figure><img src="image_132.png" /></figure>

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**Supplementary Figure 3. Effects of anterior amputation on *P. dumerilii* worms' survival and sexual maturation.** Three categories of worms were studied: 1-month-old (A), 3-4-month-old (B) and 1-year-old worms (C). For each panel (A to C), a timeline in days post anterior amputation (dpaa), as well as specific morphological features observed (with a representative picture) and their duration, are provided. Controls for each panel are also provided in order to see that they do not show any of the morphological features observed in experimental worms. A to C) At 2-3 dpaa, large cells (lc), that may be germ cells, started to appear near the amputation site (aa site) (A1, B1, C1). These cells accumulated in the whole body at around 8-10 dpaa, especially in 1- and 3-4-month-old worms (A2, B2). At the same time, parapodia (p) started to enlarge (A2, A3, B2, C2), and the typical color differences exhibited between mature females (yellowish) and males (pink-reddish) were patent in 1-year-old individuals (C3, C4). At 12 dpaa, 1-year-old individuals exhibit accumulation of gametes (g) (C5, C6). The death of the decapitated worms occurs from 12 to 21 dpaa, depending on their age.
<figure><img src="image_134.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_135.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_136.png" /></figure>

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**Supplementary Figure 4. Expression of *Pdum-vasa* on anteriorly-amputated *P. dumerilii* worms.**
A) 3-months-old or 1-year-old juvenile worms were anteriorly amputated and subsequently fixed 10 days later. B, C) WMISH for *Pdum-vasa* in 3-months-old worms (B) and 1-year-old worms (C) show expression in clusters of cells located inside the truck of the animals. B', C') Close-up views of vasa+ cell clusters in B and C, respectively. All panels are ventral views (anterior is up).
A

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B

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vasa

B'

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vasa

C

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vasa

C'

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vasa
**Supplementary Figure 5. Proportion of abnormal morphologies observed during *P. dumerilii* posterior regeneration after anterior amputation.**

A) The number of worms showing abnormal morphologies during posterior regeneration in each different condition (C0-C7 and control, see Figure 1 and text for details) is provided for each scoring day (from 1 to 7 dppa). No worms with abnormal morphology were found in the control condition. The total number of worms per experimental condition is mentioned. B) Graphic representation of the proportion of worms showing abnormal morphologies on posterior regeneration for each condition of anterior amputation (C1 to C7, and control) from 1 to 7 dppa. Asterisks indicate significant differences in comparison to the control, as defined by 2-way ANOVA on repeated measures with Dunnett correction. \* 0.0001 > p < 0.05 (for specific p-values, see Supp. Table 1).
A

<table><thead><tr><th rowspan="2">Condition</th><th rowspan="2">Total number of worms per condition</th><th colspan="7">Number of worms with abnormal morphology per condition</th></tr><tr><th>1dppa</th><th>2 dppa</th><th>3 dppa</th><th>4 dppa</th><th>5 dppa</th><th>6 dppa</th><th>7 dppa</th></tr></thead><tbody><tr><td>Control</td><td>20</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td></tr><tr><td>C0</td><td>43</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>4</td><td>11</td></tr><tr><td>C1</td><td>39</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>7</td><td>16</td></tr><tr><td>C2</td><td>43</td><td>0</td><td>0</td><td>0</td><td>0</td><td>8</td><td>19</td><td>26</td></tr><tr><td>C3</td><td>43</td><td>0</td><td>0</td><td>0</td><td>4</td><td>9</td><td>16</td><td>16</td></tr><tr><td>C4</td><td>37</td><td>0</td><td>0</td><td>0</td><td>5</td><td>13</td><td>19</td><td>23</td></tr><tr><td>C5</td><td>43</td><td>0</td><td>0</td><td>2</td><td>2</td><td>3</td><td>14</td><td>23</td></tr><tr><td>C6</td><td>36</td><td>0</td><td>0</td><td>0</td><td>3</td><td>8</td><td>8</td><td>9</td></tr><tr><td>C7</td><td>32</td><td>0</td><td>0</td><td>0</td><td>1</td><td>1</td><td>2</td><td>7</td></tr></tbody></table>

B

<figure><img src="image_143.png" /></figure>
**Supplementary Figure 6. Effects of anterior amputation on cell proliferation during posterior regeneration monitored with EdU.** A, B, D, E) Each panel shows a confocal z-stack of a posterior regenerative part in ventral view (anterior is up). EdU+ nuclei are displayed in red. Dotted lines highlight the morphology of the regenerated structures as revealed by the Hoechst staining (not shown). 1-hour-long EdU pulses were performed just before collecting the posterior regenerative parts either at 3 dppa (A, B), or at 5dppa (D, E). Control worms are only posteriorly amputated (A, D). C2 worms are amputated anteriorly 2 days before posterior amputation (B, E). C, F) Comparison of the proportions of EdU+ cells between conditions A and B (n=4), and D and E (n=6) respectively. Both brackets correspond to a non-parametric test. n.s. p >0.05; ** p <0.01. Both error bars correspond to the Standard Deviation. Cell proliferation is hampered in C2 conditions at 5dppa.
Control

C2

3dppa

<figure><img src="image_144.png" /></figure>

<figure><img src="image_145.png" /></figure>

<figure><img src="image_146.png" /></figure>

5dppa

<figure><img src="image_147.png" /></figure>

<figure><img src="image_148.png" /></figure>

<figure><img src="image_149.png" /></figure>
**Supplementary Figure 7. Abnormal morphologies observed during *P. dumerilii* post-regenerative posterior growth after anterior amputation.** A) The number of worms showing abnormal morphologies during regeneration (1 to 5 days post posterior amputation, dppa) and post regenerative posterior growth (6 to 10 dppa) for conditions C3 to C5 and control is provided for each scoring day (from 1 to 10 dppa). No worms with abnormal morphology were found in the control condition. The total number of worms per experimental condition is mentioned. Control worms were only amputated posteriorly; for C3, C4 and C5, anterior amputation was made 3, 4 and 5 days after posterior amputation, respectively. See Figure 5 for more details on the experimental designed. B) Graphic representation of the proportion of worms showing abnormal morphologies in condition C3 to C5 from 1 to 10 dppa. Asterisks indicate significant differences in comparison to control, as defined by 2-way ANOVA on repeated measures with Turkey correction. \* p < 0.005 (for specific p-values, see Supp. Table 1). C) The number of worms showing abnormal morphologies during post-regenerative posterior growth (6 to 15 dppa) in condition C5 and control is provided for each scoring day. No worms with abnormal morphology were found in the control condition. The total number of worms per experimental condition is mentioned. D) Graphic representation of the proportion of control and C5 worms showing abnormal morphologies during post-regenerative posterior growth, from 6 dppa to 15 dppa. No significant differences were found (for specific p-values, see Supp. Table 1).
A

<table><thead><tr><th rowspan="2">Condition</th><th rowspan="2">Total number of worms per condition</th><th colspan="10">Number of worms with abnormal morphology per condition</th></tr><tr><th>1 dppa</th><th>2 dppa</th><th>3 dppa</th><th>4 dppa</th><th>5 dppa</th><th>6 dppa</th><th>7 dppa</th><th>8 dppa</th><th>9 dppa</th><th>10 dppa</th></tr></thead><tbody><tr><td>Control</td><td>38</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td></tr><tr><td>C3</td><td>32</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>6</td><td>6</td><td>7</td><td>9</td><td>10</td></tr><tr><td>C4</td><td>30</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>3</td><td>5</td><td>8</td><td>9</td><td>11</td></tr><tr><td>C5</td><td>30</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>2</td><td>3</td><td>3</td><td>4</td></tr></tbody></table>

B

<figure><img src="image_150.png" /></figure>

C

<table><thead><tr><th rowspan="2">Condition</th><th rowspan="2">Total number of worms per condition</th><th colspan="7">Number of worms with abnormal morphology per condition</th></tr><tr><th>6 dppa</th><th>7 dppa</th><th>8 dppa</th><th>9 dppa</th><th>10 dppa</th><th>11 dppa</th><th>13 dppa</th><th>15 dppa</th></tr></thead><tbody><tr><td>Control</td><td>28</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td></tr><tr><td>C5</td><td>18</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>3</td><td>6</td><td>9</td></tr></tbody></table>

D

<figure><img src="image_151.png" /></figure>
**Supplementary Figure 8.** Abnormal morphologies observed during *P. dumerilii* posterior regeneration after anterior amputation and addition of exogenous methylfarnesoate (MF). A) The number of worms showing abnormal morphologies during posterior regeneration (1 to 5 days post posterior amputation, dppa) after anterior amputation and addition of MF (and controls) is provided for each scoring day. Control 1: posterior amputation, no anterior amputation, no MF treatment; Control 2: posterior amputation, anterior amputation, no MF treatment; MF-treated: posterior amputation, anterior amputation, MF treatment (100nM in sea water, renewed every day). No worms with abnormal morphology were found in the control condition. The total number of worms per experimental condition is mentioned. See Figure 7 for more details on the experimental design. B) Graphic representation of the proportion of worms showing abnormal morphologies during posterior regeneration for Control 1, Control 2 and MF-treated conditions from 1 to 5 dppa. No significant differences were found (for specific p-values, see Supp. Table 1).
A

<table><thead><tr><th rowspan="2">Condition</th><th rowspan="2">Total number of<br>worms per<br>condition</th><th colspan="5">Number of worms with abnormal<br>morphology per condition</th></tr><tr><th>1 dppa</th><th>2 dppa</th><th>3 dppa</th><th>4 dppa</th><th>5 dppa</th></tr></thead><tbody><tr><td>Control 1</td><td>29</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td></tr><tr><td>Control 2</td><td>31</td><td>0</td><td>0</td><td>0</td><td>0</td><td>4</td></tr><tr><td>MF-treated</td><td>60</td><td>0</td><td>0</td><td>0</td><td>0</td><td>9</td></tr></tbody></table>

B

<figure><img src="image_152.png" /></figure>
**Supplementary Table 1.** Results of the 2-way ANOVA statistical analyses performed for each experiment, with the specific correction used for each of them, detailed p-values and significant differences found for the multiple comparisons.