**Methods in Molecular Biology 2450**

# Simon Blanchoud Brigitte Galliot *Editors*

# Whole-Body Regeneration

Methods and Protocols

# M ETHODS IN M OLECULAR B IOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651 For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

# Whole-Body Regeneration

# Methods and Protocols

Edited by

# Simon Blanchoud

Department of Biology, University of Fribourg, Fribourg, Fribourg, Switzerland

# Brigitte Galliot

Dept Génétique et Evolution, Université de Genève, Genève, Geneva, Switzerland

Editors Simon Blanchoud Department of Biology University of Fribourg Fribourg, Fribourg, Switzerland

Brigitte Galliot Dept Ge´ne´tique et Evolution Universite´ de Gene`ve Gene`ve, Geneva, Switzerland

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2171-4 ISBN 978-1-0716-2172-1 (eBook) https://doi.org/10.1007/978-1-0716-2172-1

© The Editor(s) (if applicable) and The Author(s) 2022

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Cover Caption: The drawing visible on the cover of this book was created by Rosalie Fortabat, 10 year-old CM2 student from E´ cole du Port de la Ville de Nice, in the context of the multidisciplinary project "WORKSTATION NICE. MARINE MODELS. DRAWING REGENERATION" by the artist Irene Kopleman. Irene Kopleman's project was a collaboration between the artist, the Muse´e d'Art Moderne et d'Art Contemporain in Nice (MAMC, FR), the Tiozzo Lab at the Institute de la Mer de Villefranche-sur-Mer (FR), and the Ro¨ttinger Lab at the Institute for Research on Cancer and Aging in Nice (FR). The project allowed the public to experience, to participate, and to contribute to the progress of research on the regenerative capacities of marine invertebrates. May such multidisciplinary projects continue to flourish to stimulate the curiosity of young generations for art, for science and for regeneration research in particular.

This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature.

The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

# Preface

Whole-body regeneration (WBR) is the ability of an adult organism to restore a complete, functional body from a fragment of itself. Since Trembley's pioneering work in 1744, this amazing process has intrigued scientists who continue to characterize the mechanisms underlying spontaneous regeneration, with the hope that understanding them will open up avenues to human therapies. Although it is one of the oldest areas of experimental research in the life sciences, WBR retains enormous appeal as well as much of its mystery. One of the most puzzling aspects of WBR is the wide diversity of forms and species it involves. This phylogenetic dispersion is a fantastic asset for piecing together the puzzle of WBR origins and evolutions through comparative analyses. Indeed, each WBR context appears to be a mixture of evolutionarily conserved processes and species-specific innovations. From this perspective, the study of WBR in these aquatic invertebrates, which are generally not established model systems, is a major experimental and conceptual challenge.

This book aims to provide a comprehensive overview of the many tools available to scientists to study the numerous facets of WBR. The first part of this book provides information on the diversity of WBR, on the main challenges of this research, and on the variety of approaches used to address this topic over time. The second and third parts present a series of zoological contexts where WBR is well established and can be studied in the lab with appropriate cellular tools (Fig. 1). By including as many phyla as possible and

Fig. 1 Whole-body regeneration (WBR) across metazoan phyla. Taxa written in black are exemplified in the current issue, those in orange are not represented in this book, and the ones in purple do not have known examples of WBR. The phylogenetic tree is overlaid with key developmental innovations highlighted by red boxes. Debated phylogenetic relations are indicated by dashed lines. Phylogeny of the depicted taxa and the estimated times of divergence were adapted from (1) dos Reis et al., 2015, Current Biology 25, 2939–2950, DOI: 10.1016/j.cub.2015.09.066 and (2) Marle´taz et al., 2019, Current Biology 29, 312–318, DOI: 10.1016/j.cub.2018.11.042

Table 1 Techniques and taxa covered by every chapter


giving a focus to rather uncommon organisms, including a protist one, we have tried to foster diversity and interdisciplinarity for WBR research. Table 1 determines which species is covered by which technique. In Parts IV, V, and VI, we have selected what we believe to be the future of WBR research with cutting-edge techniques from established model organisms that open to broad and integrated molecular and systems biology approaches. These parts include genetics, omics, and synthetic techniques, which according to the species are or will become instrumental to address some of the central questions in WBR.

Thanks to the contributions of all the authors, whom we warmly praise here, this book will provide a source of reference laboratory protocols for WBR research, essential for both experienced scientists and those new to the field. With the new conceptual and technical tools described here, a new impetus is given to this nearly 300-year-old field of research to shed unprecedented light on the biological and biophysical mechanisms underlying this fascinating developmental process.

We believe that Open Access is essential for the whole community to benefit from the contributions made by the authors of this book. Open Access to the entire book was made possible by the support of the authors as well as a substantial contribution from the MARISTEM Cost Action CA16203. We are grateful for this important support, which makes science accessible to everyone.

Fribourg, Switzerland Simon Blanchoud Geneva, Switzerland Brigitte Galliot

# Contents






#### Matthias Christian Vogg and Brigitte Galliot

#### xii Contents


# Contributors


SIMON BLANCHOUD • Department of Biology, University of Fribourg, Fribourg, Switzerland


FILIPA REINOITE • European Research Institute for the Biology of Ageing, University of Groningen, University Medical Center Groningen, Groningen, The Netherlands


# Part I

Historical Overviews

# Chapter 1

# The Hazards of Regeneration: From Morgan's Legacy to Evo-Devo

### Chiara Sinigaglia , Alexandre Alie´, and Stefano Tiozzo

#### Abstract

In his prominent book Regeneration (1901), T.H. Morgan's collected and synthesized theoretical and experimental findings from a diverse array of regenerating animals and plants. Through his endeavor, he introduced a new way to study regeneration and its evolution, setting a conceptual framework that still guides today's research and that embraces the contemporary evolutionary and developmental approaches.

In the first part of the chapter, we summarize Morgan's major tenets and use it as a narrative thread to advocate interpreting regenerative biology through the theoretical tools provided by evolution and developmental biology, but also to highlight potential caveats resulting from the rapid proliferation of comparative studies and from the expansion of experimental laboratory models. In the second part, we review some experimental evo-devo approaches, highlighting their power and some of their interpretative dangers. Finally, in order to further understand the evolution of regenerative abilities, we portray an adaptive perspective on the evolution of regeneration and suggest a framework for investigating the adaptive nature of regeneration.

Key words Whole-body regeneration, Evolution, Phylogeny, Adaptation, Homology, Character, Ecology, Development

#### 1 Introduction

Thomas Hunt Morgan is considered one of the fathers of modern genetics. He is best known for demonstrating that chromosomes carry the mechanical basis of heredity, the genes. He also has the merit of introducing and developing a successful laboratory model for genetic studies, the fruit fly Drosophila. Yet, in his early career, while working at the Bryn Mawr women's college (1891–1904), Morgan devoted a significant amount of time to studying the problem of regeneration, focusing on a diverse array of regenerating animals (Fig. 1). Morgan's experimental and theoretical findings are synthesized in his now-classic book Regeneration [1]. Despite his extensive experiments and the diversity of the organisms studied, Morgan failed to identify a universal mechanism

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_1, © The Author(s) 2022

Fig. 1 Example of regenerating animal models reported in Morgan's Regeneration (1901). (a) Hydra viridis, (b) Planaria maculata, (c) Gonionemus vertens, (d) Linckia multiformis, (e) Stentor coeruleus, (f) Eupagurus longicarpus, (g) Allolobophora fœtida, (h) Ciona intestinalis. (Modified from Morgan (1901) [1])

governing regeneration. Probably in a lighter moment, he allegedly said that since he had been unable to solve the problem of regeneration, he had decided to try something easier such as the problem of heredity [2]. The fascination and the struggle of understanding regenerative phenomena and their evolution remain as alive today as it was then.

Over the last two decades, new cell and molecular biology tools have become available, allowing the exploration of a broader range of metazoan regenerative mechanisms and prompting a (re) expansion of the field of regenerative biology [3, 4]. A unifying theory of regeneration is nevertheless still lacking. Why do not all species regenerate? Does regeneration have a single or multiple (evolutionary) origin? Are the mechanisms of regeneration co-opted from other developmental phenomena (i.e., embryogenesis)? To what extent asexual reproduction, coloniality, cancer, and regeneration can be seen as different facets of the same phenomenon? Can we decipher the mechanisms of regeneration and reenable them in nonregenerating species? Such compelling questions are still waiting for satisfactory answers.

Morgan's book [1] is as relevant today as it was in the previous century, as, besides providing a historical perspective on regeneration studies across the nineteenth and the twentieth century, it lays down the conceptual and theoretical framework guiding our current research on regenerative phenomena.

### 2 The Legacy of Morgan's Regeneration

In Regeneration, Morgan synthesized and critically revised the work of his colleagues and predecessors. By analyzing classical studies, including the work of Trembley, Spallanzani and Bonnet, and the ongoing work of his contemporary scholars, such as Roux, Barfurth, and Driesch, Morgan realized how the results diverged significantly in relation to the organism studied and the methodology adopted, often leading to controversial interpretations. Through his exercise of synthesis, Morgan first attempted to group organism-specific processes into a general phenomenon of regeneration, framing his comparative approach into general questions concerning growth and differentiation, and eventually providing new insights to a theory of development. Indeed, one of the most important contributions of Morgan's book was the idea that regeneration should be considered as a growth property, and therefore approached as a developmental phenomenon. This approach to regeneration actively opposed the adaptationist view endorsed by August Weismann [5, 6], who considered regeneration as a phenomenon of adaptation and not a primary quality of the organism [7], and supported the existence of a causal relationship between the tendency to be injured and the capacity to "re-grow." With the filter of time, the inflamed debate between the two scientists was most likely rooted on methodological and epistemological grounds, with Morgan criticizing Weismann for his adherence to a "theory," instead of starting from a purely experimental approach [5]. These originally discordant approaches are not mutually exclusive, and studying regeneration today as a form of development does not mean that this process has to be considered irrespectively of its adaptive value [8].

Morgan advocated and emphasized the importance of comparing the widest diversity of organisms in order to recast the questions about development in terms of experimentally testable hypotheses. His view of regeneration was supported by a striking array of experiments that he and his students performed on a substantial number of vertebrate and invertebrate species (Fig. 1). Undeniably, the tenet that emerges in Regeneration and that is still acutely pertinent 120 years later is to challenge any general hypothesis about regenerative phenomena by performing comparative experiments using different model organisms [1, 6, 9].

2.1 Partial Versus Whole-Body Regeneration In the pursuit of a coherent explanation of regenerative phenomena, one of the priorities in Morgan's work was to introduce a clearer and more consistent terminology, able to reflect the variety of regenerative processes and to compare the many models that he and his students were describing. Even if Morgan's most famous dichotomous subdivision of regeneration based on cellular rearrangements (morphallaxis) and cell proliferation (epimorphosis) turned out to be too restrictive [10], some of his terminology and classifications are still relevant today. For instance, Morgan classified regenerative ontogenies according to the new anatomical structures that resulted from regeneration [1]. Another general classification provided by Morgan is based on the causality of the regenerative process. He distinguished between "restorative regeneration," which include post-traumatic regeneration and is the result of some exogenous injury to the organism, and "physiological regeneration," which occurs during body homeostasis, such as the turnover cycle of epithelial dermal cells in mammals, or during the "life cycle of the individual," like for example during budding, molting or feather replacements.

> To our knowledge, the expression whole-body regeneration (WBR) was not used in Morgan's work. It has been introduced relatively recently and spread widely in the scientific literature [11– 18]. The term WBR has been loosely used to describe regenerative processes that involve a "large" portion of an animal body, without adhering to a strict definition. According to Cary and colleagues, an organism undergoes WBR when it "[...] can re-grow all body parts following amputation," which is opposed to "partial regeneration," when regeneration is restricted to only some body structure [16]. Bely and colleagues also define WBR as the ability to

regenerate "all body parts," and considered that regeneration of the primary body axis is not by itself sufficient to define WBR [4]. When using WBR most authors refer to restorative regeneration but it has also been used for physiological regenerative processes [19, 20]. The expression is also employed regardless of the stages of an organism's life cycle [16, 17].

While venturing into a clearer definition of WBR we run into some classical philosophical problems. WBR brings to the forefront the problem of biological individuality and, more specifically, the issue of establishing criteria for the persistence over time of biological individuals [21, 22]: to which and how many changes an organism can go through and still be considered the same individual? When WBR leads to two or more individuals how regeneration can be considered different from reproduction, and which one is the original individual? Indeed, the expression "WBR" is rather idiomatic since, if an injury leaves some cells or tissues behind, the regeneration then cannot be "whole." It appears that the amount of regenerated material is the main property defining WBR, but what is the threshold above which regeneration can be labeled "whole"? We could consider, for example, that at least half of the original individual has to regenerate. Following this rule, in a beheaded Planaria maculata the head reforming the body would be a case of WBR, but not the body reforming the head [23] (Fig. 1b). Yet such a threshold would be clearly arbitrary, leading to conclusions that would need to be justified.

The term "whole-body regeneration" has become popular only in the last few decades. Just like the use of "regeneration," it is rich in emphasis, but not accurate and nor fully definable. Regardless of the criteria to define it, WBR in different species clearly refers to different processes.

While attempting to introduce a language that accommodates the various regenerative phenomena that had been studied so far, Morgan used the term regeneration to indicate diverse and heterogeneous phenomena of organ renewal, replacements of body parts, or asexual development [6]. He wrote that "regeneration" could constitute an umbrella term encompassing "not only the replacement of a lost part, but also the development of a new, whole organism, or even a part of an organism, from a piece of an adult, or of an embryo, or of an egg," and even including instances of imperfect regeneration: "[...] must include also those cases in which the part replaced is less than the part removed, or even different in kind" [1]. This broad definition of regenerative phenomena is still applied today. Just like WBR, it should however be regarded as a "working definition," encompassing a heterogeneous class of events, not necessarily shared among taxa [24, 25]. Despite the complexity of the phenomena considered and the blurriness of definitions, often there has been a tendency to map regeneration as a character on

#### 2.2 Regeneration: Function Versus Process

phylogenetic trees. However, regeneration cannot be reduced to a single trait, and plotting onto an existing phylogeny its presence or absence has no more value than charting the capacity of animals to fly instead of focusing on the mechanisms and structures that allow the flight. Indeed, functions can arise convergently by multiple means rather than by historical continuity [26]. Instead, regeneration must be considered as a spatiotemporal organized process, or assemblies of processes into modules [27, 28] that can be used as individual evolutionary characters [29, 30]. Then, only characters on which we can do a reasonable hypothesis of primary homology [31], for example morphological, cellular, or molecular characters associated with regeneration, can be plotted on a tree.

To identify characters associated with regeneration it may be convenient to move toward a more reductionist approach, and break down the regenerative process along its ontogenetic and evolutionary paths. In the first case, each regenerative process could be split into conserved subprocesses such as wound-healing (when present), precursor(s) mobilization, and morphogenesis [32]. The latter involves comparing these artificial ontogenetic steps between closely related phylogenetic clades, for example class, order, or family, minimizing divergence time [25, 33, 34]. The definition and the breakdown of components, and the identification of which, if any, descend from a common ancestor are among the key interests of the field of evo-devo.

2.3 Help from Evo-Devo Theoretical Tools If, as Morgan firstly suggested, regenerative phenomena can be considered as developmental processes, then the conceptual and methodological approaches developed by evo-devo research are valuable also to explore the evolution of regenerative processes [3, 19, 25, 35]. First, the use of an extended concept of homology, such as "process homology" [29] or "character identity networks" [26], which links characters from different biological hierarchies (e.g., gene, GRN, morphological characters), and, for instance, can help to describe relationships between homologous proteins and homologous molecular pathways, even if they do not necessarily lead to homologous anatomical structures [29, 36]. This more nuanced concept of homology is a powerful tool to refine comparisons of apparently unrelated regenerative processes, potentially also among phylogenetically distant and divergent species.

Second, another useful concept that captures the different levels and types of heterogeneity of an organism is the notion of modularity [27, 37, 38]. Regeneration, just like development can be divided into discrete and interacting modules, which can be tissues, fields (i.e., cells committed to forming the same structure), elements of gene enhancers, parts of gene regulatory networks, or any other "basic structural entities or regulatory phenomena necessary to assemble a complex morphological structure" [39]. The concept of modules also helps to distinguish the processes occurring during regeneration from the function of regeneration itself [25].

Third, conjointly with modularity comes the concept of developmental constraint, which restrains phenotype production due to a limited interaction among existing modules [29, 40]. For example, a limited or restrained propagation of morphogens, or bioelectric signals through voltage gradients, due to the increased histological and cytological complexity could prevent regeneration [41, 42]. The possible inhibitory effect of the immune system on regeneration is also another little-studied potential constraint [43– 45]. The existence of developmental constraints should also be taken into account when comparing regenerative processes across different species.

The conceptual tools that regenerative biology can borrow from the field of evo-devo are powerful. Comparative approaches however entail interpretive caveats, as illustrated in the following examples.

#### 3 The Difficult Task of Reconstructing WBR Evolution

The evolutionary questions concerning regeneration ultimately provide a complete narrative of the phenomenon. They are far from being just theoretical, and they can change the approach to the mechanistic study and guide the experimental design on a given model organism [8]. The three following examples illustrate the power of evo-devo experimental approaches to infer the evolution of regeneration—and of WBR in particular—but also point out some possible interpretive caveats.

3.1 Far from Basal: Diversity of Regeneration in Sponges Sponges are emblematic organisms to study the early evolution of regeneration because they have excellent regenerative abilities [46] and likely represent the monophyletic sister group of all other metazoans [47, 48]. Sponges are often considered as basal metazoans, or ancestral representative of animals. However, they are not more basal to eumetazoans than eumetazoans are basal to sponges (Fig. 2a), and there is no fossil evidence that their body plan represents an ancient state [49, 50]. As any organism, modern sponges are nothing but a mosaic of characters in their ancestral or derived state. This holds true for their regenerative mechanisms that show great inter-species variations. For instance, the proverbial ability of cell aggregates to generate a functional sponge varies even between closely related species [51–54]: Halisarca dujardini can reconstruct its body from cell suspension, whereas Halisarca panicea is unable to do so [53]. Whether or not cell reaggregation is ancestral to Porifera will remain unsolved without phylum-level comparative studies.

Fig. 2 Phylogenetic relationships between species cited in the text, and cell types involved in WBR, in sponges (a), xenacoelomorphs (b), and ascidians (c). The species that are reputed for their extensive ability to regenerate are represented in red. The cells drawn represent the cell types known to supplement more tissues during regeneration, by proliferation and/or differentiation. On branches are shown ancestral reconstruction regarding the role of each cell type in WBR based on parsimonious optimization

The mechanisms of WBR from body fragment also varies between the four sponges classes. Many demosponges use massive proliferation and migration of archaeocytes with the participation of dedifferentiated choanocytes, which all together form a regenerative blastema [55, 56]. In some other demosponges (e.g., Halisarca dujardini and Aplysina cavernicola) the cell plasticity is even greater, with dedifferentiation of various cell types that also participate in blastema formation [57, 58]. In contrast to demosponges, neither archaeocytes nor tissue regeneration has yet been observed in their sister group, the Hexactinellida [59]. Calcareous sponges, who also do not possess archaeocytes, regenerate through epithelial morphogenesis by spreading and transdifferentiation of pinacocytes and choanocytes (e.g., in Leucosolenia complicata) with minor cell proliferation and no blastema formation [60]. Finally, among the homoscleromorphs, the sister group of calcareous sponges, only Oscarella lobularis has been reported to regenerate [61, 62]. As in Calcarea, it involves choanocyte transdifferentiation and tissue rearrangement, without blastema formation or local proliferation. Due to this phylum-level variability in regenerative capability and mechanisms, reconstructing the origin and evolution of WBR in sponges is far from being a straightforward task (Fig. 2a).

Nevertheless, choanocyte dedifferentiation and/or transdifferentiation seem to be a common theme in regenerative species, which may be in line with the suspected stem cell nature of choanocytes [63]. Comparative investigations focused on choanocyte dynamics (e.g., time series of single-cell RNAseq) could unravel fundamental sets of genes regulating WBR potentially inherited from the last common ancestor of Porifera.

3.2 Acoels and Planarians: Lessons from Faraway Cousins Recent work on acoels and Platyhelminthes has provided fresh insights on the possible ancestral mechanisms of WBR in the last common ancestor of Bilateria. Acoels are flatworms belonging to a larger clade named Xenacoelomorpha, together with Xenoturbellids and Nemertodermatids (Fig. 2b). Some authors consider Xenacoelomorpha as the sister group of all other Bilateria [64, 65] and others the sister group of Ambulacraria. Despite being distantly related, acoels share a superficial morphological resemblance with Platyhelminthes, a group of lophotrochozoan flatworms. Their regenerative mechanisms also show extensive similarities. In acoels and planarians, regeneration involves the proliferation-dependent formation of a regenerative blastema by mesenchymal multipotent and totipotent stem cells, the neoblasts, which express homologous genes such as Piwi paralogs and other members of the Germline Multipotency Programs [66–68]. In both acoels and planarians, muscles play a contraction-independent role by secreting position control proteins (e.g., wnt and bmp ligands), thus providing positional information for correct body plan restoration upon WBR [69–72]. These shared characters suggest ancestral features inherited from the last common ancestor of Bilateria. However, proposing the homology of regenerative processes at such a large phylogenetic scale remains risky. For instance, while neoblast-like stem cells are present in several bilaterian lineages [68], their phylogenetic distribution is much more parsimoniously explained by convergent acquisition, rather than as an ancestral presence with multiple losses. Transcriptomic and genomic characterization of neoblasts in various animals may additionally reveal shared molecular signatures that also result from convergent acquisition. Also, the orthology of the position-control genes expressed by muscles during planarian and acoel regeneration has not been established [69], and therefore it's not clear if their role in regeneration is inherited from a common ancestor or not.

> To date, regeneration studies on acoels have been mainly done in species belonging to the Bursalia suborder (e.g., Hofstenia miamia, Isodiametra pulchra). But, to our knowledge, regeneration power is not yet reported in the ca. other 400 acoel species nor in other Xenacoelomorphs (Xenoturbellids and Nemertodermatids) [73]. The example of sponges clearly demonstrates the intraphylum plasticity of WBR and highlights the importance of studying more related models. This may be the case for acoels too, as

they are known to evolve relatively fast [74] and to harbor many derived characters among Xenacoelomorphs, such as the organization of body muscles, or the presence of epidermal eyespots [75, 76]. Consequently, acoels alone cannot be taken as a proxy for Xenacoelomorpha and ancestral reconstruction of bilaterian WBR will not be possible without exploring anatomical, cellular and molecular diversity across Xenacoelomorpha.

Despite these caveats, the comparison between acoels and planarians is highly relevant to reconstruct the ancestral mechanisms of WBR in Bilateria. It is important to note that this holds regardless of the position of acoels as the sister group of Nephrozoa or Ambulacraria, since in both cases the last common ancestor of acoels and planarians is the ancestor of all Bilateria (Fig. 2b).

Increasing the phylogenetic resolution and comparing multiple closely related species is crucial to assign confidently the directionality of evolutionary transitions. Tunicates include so-called solitary species, where regeneration is limited to some tissue and organs [77] and colonial species, which are all able to undergo WBR via different types of budding [78]. Tracking WBR evolution in tunicates benefits from numerous anatomical studies on many species combined with well-resolved and robust phylogenies that allowed to infer multiple independent acquisitions of WBR in the whole subphylum [34, 78, 79]. For example, the evolution of budding in the family of Styelidae remained largely speculative until recently. Berrill [80] considered that all colonial species belonging to this family should be unified as a natural group because he assumed that they all perform the same kind of budding. In contrast, Kott suspected that budding modes may be more diverse than expected and advocated for "accurate resolution of their taxonomy [and] information on the process of vegetative reproduction" [81]. Recent phylogenetic reconstruction of Styelidae [34], as well as a closer look at the budding tissues in the species Polyandrocarpa zorritensis [82] showed that the fundamental differences in the mechanisms of bud formation, as well as their phylogenetic distribution, are more parsimoniously explained by convergent acquisition [34]. Thus, according to these data, three modes of WBR have been independently acquired (Fig. 2c) from a solitary, nonbudding, ancestor of Styelidae. Therefore, the question is to know whether homologous modules (e.g., GRN made of orthologous genes) have been convergently deployed in these three nonhomologous budding modes. The discovery of such shared GRN or budding cell types between the different budding modes in Styelidae will be interpreted as independent co-options, as long as the phylogenetic topology makes the convergent acquisition of budding the most parsimonious hypothesis.

#### 3.3 Plastic Families: Convergent Acquisition of WBR in Tunicates

#### 3.4 A Roadmap to Investigate WBR Evolution

These three examples clearly show that, in the attempt to infer the evolution of regenerative phenomena, the phylogenetic relationships between the considered organisms must be used as an interpretative framework to formulate hypotheses on evolutionary trajectories. Then, each defined character should be first considered independently (presence/absence of neoblasts, expression of Wnt orthologs, a given morphogenetic movement, etc.) in order to reconstruct the mosaic of derived and ancestral states that make up the regenerative process and its phylogenetic distribution. Combining several lines of evidence such as histology, morphology, molecular signatures (e.g., by RNAseq) and phylogenetic analyses of genes of interest is, therefore, an informative way to refine homology hypotheses. When possible, multiple species must be considered in parallel to cover the diversity of the regenerative mechanisms (including absences) across the studied taxa. Finally, a particularly informative ontogenetic step to collect characters related to regeneration may be the earliest steps after the injury, at the interface between the wound healing (when present) and the mobilization of the precursors (i.e., stem cells or dedifferentiating cells). For instance, recent RNA-seq and ATAC-seq analyses on fine-grained time series have shown that several species of bilaterian and cnidarians overexpress immediate-response genes such as EGR or Runt homologs, and establish Wnt signaling centers at the onset of regeneration [11, 83–86]. However, Wnt genes expressed in different regenerative contexts across species are not orthologous and are likely under the control of nonhomologous mechanisms [83]. This and the patchy distribution of WBR may point toward an evolutionary scenario where WBR arose multiple times independently during metazoan evolution, often reusing similar modules co-opted from embryogenesis (e.g., Wnt canonical pathway) while also assembling original modules specific to each regenerative strategy.

#### 4 What Is the Significance of WBR? An Integrative and Practical Approach

Regardless of the phylogenetic context—single or multiple acquisitions/losses of regenerative capacities—the advantages of regenerating a large portion of the body, or of multiplying individuals by budding, might seem self-evident. These advantages were largely assumed by early scholars, as Reaumur [87] and Bonnet [88], long before any theorization of evolution by means of natural selection. Yet, trade-offs between costs and benefits of regeneration might exist—and sometimes the benefits themselves might be difficult to identify, as in the case of the constant cycles of zooids destruction and regeneration in the colonial ascidian Botryllus schlosseri [89]. The challenges in understanding the evolution of WBR among metazoans depend thus also on the difficulties in answering an apparently elementary question: what are the consequences of regeneration on the survival and/or reproductive fitness of an individual? In other words, is regeneration, or the loss of it, adaptive?

Following Darwin's work [90], Weismann explicitly regarded regeneration as an adaptive phenomenon "the degree to which it is present is mainly in proportion to the liability of the part to injury" [7]. Morgan, who was skeptical of untested theoretical explanations, set out to validate experimentally this prediction. In order to test whether the regenerative potential of a body part correlated with its risk of being injured in nature, he chose as a study model the hermit crab Pagurus longicarpus (Fig. 1f), as its anterior appendages were exposed to damage, while its posterior ones were "naturally protected" by the host gastropod shell. All appendages proved to regenerate well, which led Morgan to reject any adaptive value for regeneration. Morgan's experimental setup was however criticized for oversimplifying the parameters of the problem. Needham, in particular, argued that for a correct estimation of the evolutionary pressures, the "indispensability" of each appendage had to be considered. After recapitulating the experiments on Pagurus [91] and other crustaceans [92], Needham remarked that, (1) the frequency of regeneration in posterior, more protected, appendages was indeed lower (in Pagurus it was 21% vs. 83%), and that (2) each pair of posterior appendages was essential to locomotion (and thus for survival of the crab). Thus, not only there was a correlation between risk of injury and regenerative potential, but the maintenance of a complete pair of posterior appendages was likely under strong selective pressure, supporting the old idea that regenerative abilities had an adaptive value [92]. The question was thus far from being settled because if purely adaptive interpretations could explain the patchy distribution of regenerative potential among metazoans, it remained difficult to account for the similarities among regenerative processes [8]. Goss crystallized this idea and argued that if regeneration was truly an adaptive phenomenon, it must have arisen (and been positively selected) from nonregenerating ancestors multiple times, which would entail substantial differences between developmental mechanisms [8]. Shared features between diverse regenerative processes had instead been demonstrated, such as the requirement for innervation [93, 94]. Previous research had further highlighted a certain degree of similarity between embryonic and regenerating limbs, notably concerning patterning [95, 96] and morphogenesis [97]. Goss, like Morgan, favored a scenario where regeneration would be an inherent feature of metazoan life, and most likely a derivative of a core embryonic developmental program [98].

In his view, the modern phylogenetic pattern of regenerating taxa could be interpreted as the result of repeated losses of potential—themselves the consequence of other adaptive processes, for instance, the evolution of better brains in vertebrates [99], or the transition from aquatic to terrestrial habitats [8]. While some similarities among regenerative processes do exist, for example with regard to wound healing [100], it is today clear that the comparison is far from being trivial, as also concluded by Morgan, and that the answer cannot derive from the "mere" addition of further, diverse types of data. The previous examples on sponges, flatworms, and tunicates show that the identification of the relevant comparisons, at all the different scales, is key. Regarding the shared features of regeneration and embryogenesis, for example, recent transcriptomic approaches have indeed highlighted some degree of conservation in sequential gene usage between embryonic processes and regeneration [101–104]. On the other hand, regeneration is broadly thought to display specific features, such as an involvement of the immune response [105], of the nervous system [106], and perhaps of muscle cells [69].

4.1 The Puzzle of "Restriction and Absence" of WBR: Eco-Evo-Evo Perspectives Representatives of sponges, acoels, planarians, tunicates but also cnidarians, ctenophores, annelids, echinoderms, and placozoans display different WBR capacities. The ability to regenerate large portions of the body is conversely lacking in arthropods, which nevertheless can regenerate their appendages until they reach a terminal molting stage—suggesting a possible trade-off between a protective cuticle and WBR, probably emerging at the origin of Ecdysozoa [107]. The problem with the "restriction and absence" [108] of regenerative potential among taxa remains central to the study of the evolution of regeneration [4]. As highlighted in the previous sections, the fragmentary taxonomic sampling is a major limit in understanding the evolutionary trajectories of WBR. The absence of regeneration is particularly difficult to address, and any explanatory research would need to take into account three parameters:


Habitat, body size, reproduction modes, anatomy, and defense mechanisms might all be factors to consider. The intersection of ecological, developmental, and phylogenetic parameters poses a methodological challenge, and an eco-evo-devo approach has the potential for providing a common framework for tackling the issue [110].

Recent works have extensively discussed the ultimate causes of a reduced regenerative potential [4]. These works argue either that some selective pressure could play against the preservation of regenerative capacities, or that no particular pressure would maintain it, so that it becomes a neutral trait. The studies directly addressing the ultimate causes of regeneration are rare. A famous example is the loss of regenerative capacity in some groups of spiders, including the black widow (Latrodectus mactans). Spiders usually regenerate well their injured legs [111], with the notable exception of few orb-weaving genera, where it has been hypothesized that a regenerated appendage could impair web-making more than a missing one [112]. In this case, a strong pressure, the need for a geometrically accurate spider-web, selected against the maintenance of regenerative capacities. Conversely, if no particular pressure maintains regenerative capacities, for example, if predation is low [113, 114], these could be lost. Neutrality could also emerge if regenerative phenomena were essentially a by-product (an epiphenomenon) of other developmental processes under selective pressure and if the molecular link between modules was lost, for instance due to the activity of selfish genetic elements [115]. Continued tissue growth [116], agametic reproduction, or core embryonic mechanisms [117] have all been proposed as processes from which regeneration might have derived.

A taxon-restricted loss of regenerative capacities does not necessarily imply an elimination of the genetic program for regeneration. Are there any latent or inhibited regenerative capacities in taxa that usually do not display them—and which could thus be reactivated? In naidine annelids, both comparative regeneration experiments and phylogeny indicate multiple events of loss of head regeneration. Interestingly, in one species, amputation during asexual fission within a small proliferative region harboring activated stem cells could elicit regeneration of a normal head [118]. This indicates that, despite the loss of regeneration, the capacity remained latent in these annelids, and could be reactivated. This study is a further reminder that a comparative experimental approach is essential for understanding the evolutionary trajectories of regeneration.

The problems with the loss of regenerative capacities, its significance for the fitness of organisms, and the question of whether regeneration is an attribute of all organisms are not purely theoretical. Indeed, our hopes of inducing regeneration where it does not occur, for example in adult humans, ultimately rests on the assumption that potential for regeneration might remain latent in organisms who are currently unable to do it [119].

#### 4.2 Questions and Approaches to Investigate WBR Evolution

Regardless of the evolutionary scenario, WBR constitutes a particular category of regenerative phenomena, whose links to physiology and reproduction are blurred. Here we have considered WBR in its most inclusive sense, including physiological regeneration and asexual reproduction, and effectively adopting the functional definition of regeneration that—by replacing essential body parts significantly delays an organism's death. But how to practically study WBR, placing this phenomenon in its evolutionary, developmental, and ecological context? The questions raised through the Weismann vs. Morgan adaptive/innate debate are still highly relevant today. The criteria and strategies then proposed can represent today the starting points for practically shaping an integrative research program on the complex issue of whole-body regeneration.


invertebrates undergoing WBR or recruiting new larvae, have a key impact on the dynamics of the benthic community. With regard to the second point, annelids have an important biogeomorphic impact on marine sediments, and regeneration negatively impacts their reworking of sediments [127]. Additionally, as WBR is tightly linked to the production of new individuals, it might represent a dispersal strategy [128], allowing organisms to colonize rapidly a novel or changing habitat, as it has been shown for forest recovery after fires [129]. The consequences of WBR on the invasiveness of a species and perhaps on the emergence of new species following reproductive isolation have been poorly studied, but constitute an interesting avenue for future research.

The extreme nature of WBR poses unique challenges, in particular when we try to investigate and measure the ecological and physiological implications. The resources required during WBR cannot be made available to other processes [130] This suggests important trade-offs for the organisms concerned, which need to be identified and quantified. These trade-offs concern the regenerative events, but also the loss of a body part itself. With regard to the cost-benefits of the regenerative process itself, regeneration subtracts resources from growth and reproduction, the so-called regenerative load [131]. In sponges and corals, injuries inflicted when food is scarce or when the animal had been previously injured regenerate less well, showing that resource allocation is critical [132]. On the other hand, besides the obvious benefit in avoiding looming death, WBR might provide some specific advantages, for example, a rapid adaptation to changing environments [125, 133]. In heteromorphic colonies of hydrozoans and bryozoans, changing environmental conditions could cause the regression of existing individuals and the generation of a different type of specialized zooid [125, 134]. Interestingly, given the colonial nature of these organisms, the costs of the process would be reduced by the reutilization of regressing individuals [20] as a source of materials and energy for the growing ones.

The loss of body parts is more difficult to quantify. Energy loss is a multifaceted variable, but the dry weight of the removed body part has been used as an estimate [135]. Short-term, acute, costs include the loss of foraging or motility, of body mass, risk of infection, behavioral disruption, and impaired self and nonself recognition, while lower fecundity or growth (due to loss of germ cells or energy storage) might be seen in the long term [136]. The loss of an arm, for example, has a greater cost for asteroids than for crinoids or ophiuroids, as they bear gonads [135]. As for the eventual benefits, it might seem difficult to imagine any advantage in losing a body part. Yet autotomy, the active breaking of a body part along a predetermined "plane," suggests a possible scenario: crustaceans, annelids, holothurians and other animals shed body parts as a defense mechanism, in order to escape predators or to isolate infected or malfunctioning body parts (reviewed [137]).

#### 5 Conclusions

When, later in his life, Morgan heard that a 24-year-old Norman John Berrill was working on marine worms and ascidian development and regeneration, he reproached him saying, "You are being very foolish [...] At your age you cannot waste your time. We will never understand the phenomena of development and regeneration." [138]. Perhaps, if he had access to the theoretical tools of ecoevo-devo and to the technological resources available today, he would have thought otherwise. Morgan's emphasis on exploring the vast diversity of both developmental and regenerative phenomena, and experimenting with testable hypotheses in models, represents the assets of his legacy. The very same modus operandi could help to avoid hasty interpretation and to remove anthropomorphic biases in how we interpret natural phenomena. Luckily, the young Berrill did not take Morgan's advice and "[...] continued watching in wonder to my heart's content and I am even more bewildered, though more sophisticated, by what I see" [138].

#### Acknowledgments

The authors want to thank Evelyn Houliston, Lucas Leclere, and Elena Casetta for their valuable comments on the manuscript. This work was supported by ANR (ANR-14-CE02-0019-01).

#### References


regeneration. J Hist Biol 46:511–541. https://doi.org/10.1007/s10739-012- 9341-9


https://doi.org/10.1016/j.ydbio.2012. 11.006


evolvability of complex organisms. Nat Rev Genet 12:204–213. https://doi.org/10. 1038/nrg2949


Exp Zool Part B Mol Dev Evol 334:37–58. https://doi.org/10.1002/jez.b.22919


https://doi.org/10.1038/s41467-017- 01148-5


redeployment of the embryonic gene network. bioRxiv. https://doi.org/10.1101/ 658930


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# Studying Regeneration in Ascidians: An Historical Overview

### Virginia Vanni , Loriano Ballarin, Fabio Gasparini, Anna Peronato , and Lucia Manni

#### Abstract

Ascidians are sessile tunicates, that is, marine animals belonging to the phylum Chordata and considered the sister group of vertebrates. They are widespread in all the seas, constituting abundant communities in various ecosystems. Among chordates, only tunicates are able to reproduce asexually, forming colonies. The high regenerative potentialities enabling tunicates to regenerate damaged body parts, or the whole body, represent a peculiarity of this taxon. Here we review the methodological approaches used in more than a century of biological studies to induce regeneration in both solitary and colonial species. For solitary species, we refer to the regeneration of single organs or body parts (e.g., siphon, brain, gonad, tunic, viscera). For colonial species, we review a plethora of experiments regarding the surgical manipulation of colonies, the regeneration of isolated colonial entities, such as single buds in the tunic, or part of tunic and its circulatory system.

Key words Colonial circulatory system, Evisceration, Gonad, Neural complex, Partial regeneration, Siphon, Thorax, Tunic, Whole body regeneration

#### 1 Introduction

Within the phylum Chordata, which includes the three subphyla Vertebrata, Cephalochordata, and Tunicata (Fig. 1), the latter exhibits the more striking regenerative abilities. This feature, widely recognized by the scientific community since the end of the nineteenth century, raised renewed interests in the last 15 years, thanks to the availability of new methodological tools enabling the dissection of its molecular and cellular bases [1, 2]. In tunicates, the regenerative ability shows remarkable differences in various clades, even in different tissues and organs of the same organism [3]. Nonetheless, in some species, it can extend to extremes of complete individuals formed, from a small group of stem cells in the case of whole-body regeneration (WBR) [4, 5].

Tunicates are filter feeding, marine organisms, widespread in all the seas, constituting abundant communities in various ecosystems.

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_2, © The Author(s) 2022

Fig. 1 Phylogenetic tree of chordates (modified from [7]) reporting the ascidian species studied for regeneration. Dots indicate if species are solitary or colonial. Types of regeneration induced in the various species include the following: regeneration of the colonial circulatory system (CCS), gonads (Go), neural complex (NC), partial regeneration (PR), regeneration of the siphons (Si), thorax (th), tunic (tu), viscera (Vi). WBR: whole body regeneration

> They are considered the sister group of vertebrates (Fig. 1) [6, 7], therefore representing, from an evolutionary point of view, an intriguing taxon for comparative studies on regeneration, a limited process in vertebrates [8, 9]. Tunicates include both solitary and colonial species, the latter representing the only chordates capable of asexual reproduction (also called budding) [10].

> Ascidians represent the main tunicate group, now considered paraphyletic by several authors. Their life cycle includes a swimming larva, which exhibits the typical chordate body plan and is the dispersal phase of the species. The larva is lecithotrophic and swims for a few hours to select an appropriate substrate on which

Fig. 2 Schematic representation of a solitary (left) and a colonial (right) ascidian (in ventral view). In the colonial tunicate, three adult zooids are represented, each one bearing one primary bud and one secondary bud

to adhere. Then, it undergoes a deep metamorphosis becoming a sessile individual, the juvenile, which possesses the capacity to regenerate.

Ascidian adult body is cylindrical, with an anterior inhalant (oral) siphon and a dorsal exhalant (atrial) siphon (Figs. 2 and 3). The brain (called cerebral ganglion) is located between the two siphons. The brain, together with the associated neural gland, forms the neural complex. The majority of the body is occupied by a large branchial chamber (pharynx) perforated by numerous ciliated stigmata [11]. The inhaled water enters the branchial chamber and, passing through the stigmata, is filtered by a mucous net produced by the endostyle. The latter is a glandular groove located in the ventral floor of the branchial chamber. The filtered seawater passes then to the atrial chamber and is expelled through the atrial siphon. Nutrients, entrapped in the mucous net, are agglutinated in a mucous cord that is conveyed to the U-shaped gut, located posteriorly, below the branchial chamber. The anus opens into the atrial chamber, so that the fecal pellets are removed by the exhalant water flow. Ascidians are hermaphrodites; gonads can be located in the posterior body, close to the gut, or in the lateral body wall.

In solitary ascidians, the cylindrical, tube-like body shape suggested a wide range of regeneration experiments involving mainly distal body parts, such as the siphons and the neural complex (Table 1) (Fig. 3b–f). In the solitary species of the genus Ciona, C. intestinalis and C. robusta, for which very advanced methodological tools are available, regeneration has been comprehensively

Fig. 3 (A–F). Oral siphon regeneration in Ciona robusta. (A) Individual showing the typical cylindrical, tube-like body shape of solitary ascidians. The oral siphon (os) individuates the anterior side, the atrial siphon (as) the dorsal side. (B–F). Details of siphons of an individual before (B), and after 0 (C), 1 (D), 3 (E), and 6 days (F) from the amputation of the oral siphon. The red line in B labels the level of amputation. One day after amputation (D), the wound is closed. In (E, F), black arrowheads mark the basal limit of the regenerating oral siphon. In (B) and (F), red arrowheads label some of the eight orange-pigmented sensory organs located in the notches between the lobes of the oral siphon rim. Six similar organs are also on the atrial siphon. Note that after 6 days from oral siphon ablation (F), the organs are present. (G–J). WBR in Botryllus schlosseri. In a colony at takeover phase (G; ventral view), regressing adult zooids (rz) are in form of dense masses at the center of the colony; primary buds (1b) are almost ready to open their siphons becoming the new generation of filtering individuals; small secondary buds (2b) are recognizable on primary buds. The colony is embedded by the tunic (t), where the colonial circulatory system is located. A marginal vessel (mv) extends all around the colony, connecting and coordinating the zooids. Blind ampullae (a) elongate from the marginal vessel toward the periphery. (H) colony (dorsal view) showed in G 2 days after the ablation of all the zooids (regressing zooids, primary and secondary buds). The circulation is restored. Asterisks in H individuate the position of the three primary buds marked by asterisks in G. (I–J) 3 days post ablation, two new vascular buds (arrows) are recognizable close to the marginal vessel. Square area in I is enlarged in J. (K–L) Tunic and colonial circulatory system regeneration in Botryllus schlosseri (ventral view). (K) a portion of the tunic (t) with its marginal vessel (mv) and blood ampullae has been removed (dotted line) in front of three adult zooids (az).


#### Table 1 Types of regeneration studies in solitary ascidians

Fig. 3 (continued) Arrowheads individuate the lateral cut edges. (L) the same colony showed in (K), 6 days after ablation: new tunic covers the previously exposed zooids. (L) in the regenerated tunic, the marginal vessel and a crown of new ampullae (a) are recognizable. Asterisks in K and L individuate the same zooid, as reference. Scale bar ¼ 1 mm in a, (G–J); 5 mm in (B–F); 100 μ in (K, L).

explored in terms of both its morphological and cellular/molecular aspects (Table 2) [1]. Among other emerging model species, Polyandrocarpa mytiligera is a solitary ascidian of the Red Sea that, for example, can regenerate the whole gut after its evisceration [12– 14].

In colonial ascidians, several individuals (zooids) are organized in large colonies exhibiting astonishing morphologies and colors (Fig. 3g) [10]. Usually, regeneration is seen as the ability of an organism to regrow or repair its cells, tissues and organs after their loss or severe injury. However, to some extent, asexual reproduction in colonial ascidians is considered an expansion of regeneration, as a nonembryonic development of new individuals. Indeed, the ability of colonial ascidians to activate unusual developmental pathways in both natural and/or induced conditions makes the border between asexual reproduction and a true, injury-induced, regeneration quite faint [2, 11].

In colonial ascidians, a plethora of regeneration experiments has been performed regarding, in general, the removal of single individuals from colonies (single buds, adult individuals, both buds and adult zooids) (Table 3; Fig. 3g–j) [2]. However, the regeneration of isolated colonial entities (e.g., isolated buds in the tunic, the whole tunic with its circulatory system without any zooid) and the regeneration of part of the tunic and its circulatory system have also been studied (Fig. 3k–l). The latter is a network of hemolymphatic vessels (Fig. 3g) connected to a marginal vessel that extends along the periphery of the colony. Several radial vessels emerge from each zooid and connect them to the marginal vessel. Blind, sausage-like ampullae elongate toward the tunic periphery. Among colonial ascidians, Botryllus schlosseri is one of the most studied species. However, several ascidians species can be maintained in laboratory culture throughout their life cycle and used for regeneration experiments (Table 3). Thanks to their recurrent budding, the ability to survive in aquaria also beyond their natural lifespan (useful to study ageing), the possibility to be split in fragments to analyze different conditions in the same genetic environment, their sequenced genome, and the availability of some molecular tools for unbiased results (Table 4), colonial ascidians provide valuable models for an integrated approach to regeneration.

On the whole, studies on both solitary and colonial species are shedding light in outstanding challenging topics of contemporary biological science, such as the connections between animal regeneration and regenerative medicine, stem cells biology, aging, and tissue homeostasis [1, 15].

Here, we briefly review the different types of regeneration experiments performed in ascidians in more than a century. Moreover, we present, in an historical perspective, the methodological approaches used to induce regeneration in both solitary and colonial ascidians.


#### Table 2 Experimental procedures used in the study of regeneration of solitary ascidians

(continued)

#### Table 2 (continued)


#### 2 Regeneration in Solitary Ascidians

The study of regeneration in solitary ascidians has a long story as it began at the end of the nineteenth and the early beginning of the twentieth century, with the experiments carried out by a series of German scientists working at the Stazione Zoologica, in Naples (Italy), newly founded by Anthon Dohrn [16–18]. With the establishment of a series of new French marine stations at the beginning of the last century, the contributions of the Belgian-French scientists appeared and, rapidly, acquired a predominance that was maintained throughout the first half of the century [19–24]. Then, the American school arose and, quickly, reached the visibility that it still has [25–27].

Solitary ascidians are capable of partial body regeneration (Table 1). In all the studied species, the epidermis can easily regenerate the tunic once the latter is removed [19, 20, 23]. When part of the body is removed, in most cases it can be reformed quite rapidly. C. intestinalis has been the reference model for the study of solitary ascidian regeneration for more than a century. In this species, when an animal is bisected, only the basal part can


#### Table 3 Types of regeneration studies in colonial ascidians

regenerate the missing one, provided that part of the pharyngeal basket is conserved; conversely, the distal part is unable to regenerate any basal structure. The regeneration includes organs such as the siphons, the neural complex, the gonads and the missing part of the digestive system.

#### Experimental procedures used in the study of regeneration of colonial ascidians


(continued)



The regeneration of the Ciona siphons attracted the attention of many researchers for the simplicity of the technique required: a scalpel and some anesthetic [28–30]. A full regeneration of the siphons has also been described in P. mytiligera, Styela plicata and Herdmania momus, whereas, Microcosmus exasperatus show a scrubby siphon reconstitution, suggesting an unequal distribution of the regenerative abilities among solitary species [31].

Jeffery and collaborators [32, 33] reinvestigated in detail the regeneration of the oral siphon in C. intestinalis (formerly called Ciona intestinalis type B) [34]. They demonstrated that both short-distance and long-distance processes are involved in oral siphon regeneration, the former based on the presence, in the remaining siphon stump, of a local pool of progenitor cells, the latter relying on the migration of progenitor cells from niches in the pharynx. Short-distance recruitment of progenitor cells is mainly involved in the formation of the orange-pigmented sensory organs (Fig. 3b, f) and is much more influenced by the depletion of the progenitor cell reservoir through repeated ablations [32]. When the siphon is amputated at its base, only long-distance recruitment of progenitor cells occurs [32].

A spectacular example of regeneration regards the two congeneric species P. mytiligera and P. tenera that can eject their viscera when subjected to stress conditions and rebuild them in less than 3 weeks [13]. Moreover, individuals of P. mytiligera can be separated in fragments by cutting along the longitudinal or transverse body axes. Each fragment then is able to regenerate completely the missing organs forming independent functional individuals [14].

As stated above, the methods used for studying regeneration in solitary ascidians were, in the past, quite simple: a scalpel or a razor blade to cut the animals or ablate the siphon(s), a dissection microscope to observe the anesthetized animals after the operation, eventually equipped with a camera lucida apparatus to record the regenerating steps in the recovering specimens (see the following "methodological approaches" section for a detailed historical overview on the methods used to induce regeneration). Today, the experimental procedure is not greatly different. The only relevant change is the introduction of in vivo imaging, which renders reporting much easier, and of electron microscopy analysis that allows detailed observations of the events at cellular levels. In addition, it must be stressed that, today, research on regeneration can exploit the abundance of biochemical and biomolecular toolkits offering the possibility to study in detail the events occurring during recovery. Table 2 reports the various approaches used for the study of regeneration and the attempt to elucidate the cells, genes, signaling pathways involved in the process.

#### 3 Regeneration in Colonial Ascidians

The first studies on regeneration in colonial ascidians are those of Giard [35] in the second half of the nineteenth century, cited by [36]. Driesh [36], using Clavelina lepadiformis, observed that, dividing the animals in two, each fragment was able to regenerate the missing parts. C. lepadiformis was also used as a model organism for regeneration studies by Della Valle [37, 38]. In 1921, Huxley was one of the first authors to study regeneration in Perophora viridis, by recording the regenerative process after splitting the zooids in two [39]. The same author also investigated the regenerative capability of Aplidium pellucidum by isolating small colonial fragments. More detailed studies were carried out on the regeneration processes of P. viridis by Deviney, in 1934 [25], and by Goldin, in 1948 [40].

In the second half of the twentieth century, new model organisms, such as B. schlosseri and Botrylloides leachii, became the main protagonists of regeneration studies in colonial ascidians. Even today these two organisms are widely used for regeneration studies. Over the years, studies have focused on three aspects (Table 3):

1. WBR


#### 3.1 WBR WBR was studied in B. schlosseri, B. leachii, Botrylloides violaceus and Botrylloides diegensis following the surgical removal of all zooids and buds from a colony (Fig. 3g–j) or isolating small fragments of the colonial vasculature. This type of regeneration closely resembles vascular budding, a spontaneous formation of new buds from the vessels of the vascular system, first described in botryllid ascidians more than 200 years ago [41] and observed and described again by Giard [35], Bancroft [42], and Herdman [43]. This type of budding is constitutive in Botryllus primigenus [44], but it can also be induced through the isolation of small vascular fragments containing part of the colonial circulatory system [45–47]. In WBR, a bud, that eventually reconstitutes the whole colony, develops in the colonial vasculature from the aggregation of hemoblasts [5, 48– 52]. Recently, Rosner and collaborators observed and studied an additional form of WBR in B. schlosseri, termed "budectomy induced WBR." When, in a colony at takeover (the phase in which the adult zooids are being resorbed and replaced by their primary buds), all the buds are surgically removed leaving only old zooids undergoing resorption, new budlets can develop from the latter [53]. In this view, even the experiments performed on B. schlosseri colonies by Majone in 1977 can be considered as WBR [54]. In this case, new budlets develop from anterior bud fragments connected to the colonial vasculature. These new budlets eventually grow further to actively filter-feeding adults [54].

In species in which zooids are connected by stolons, such as Polyandrocarpa zorritensis, C. lepadiformis and P. viridis, WBR has been induced through the isolation of part of the stolon from the rest of the colony [21, 25, 37–39, 55, 56]. The success and timing of regeneration depend on the dimension of isolated stolon fragments [25, 38, 40]. Full recovery can require a period of time ranging from a few days, as in P. zorritensis [56], to several weeks, as in the case of small pieces of stolon of C. lepadiformis [37]. In A. pellucidum and P. viridis, Huxley [39] isolated small fragments of colonies or even of zooids, and observed what he called the "dedifferentiation" of the latter to undifferentiated structures from which new buds eventually developed [39]. These processes were further studied by Deviney in 1934 [25] and Goldin in 1948 [40].

In B. schlosseri, it is also possible to observe the regeneration of bud residuals. When, in a colony, a single adult individual is left and its primary and secondary buds are removed, bud residuals can reverse the degeneration process and start regeneration. This phenomenon, described for the first time by Sabbadin in 1956 [57, 58], probably occurs for the absence of competition among buds, allowing the residuals to rescue development. In this case, a constitutive degeneration of budlets is reversed in a regenerative process induced by the new colony condition. This is an example in which it is difficult to mark the border between asexual reproduction and typical regeneration, and it stresses the high homeostatic capacity of colonial organisms to survive adverse conditions.

3.2 Partial Body Regeneration Partial body regeneration can also be observed in colonial species. In B. schlosseri [27], C. lepadiformis [21], Pycnoclavella neapolitana [59] and P. misakiensis [60, 61], the regeneration of the missing parts of amputated buds have been studied (Table 3). In these cases, no regression and development of new budlets are observed.

3.3 Tunic and Colonial Circulatory System Regeneration Colonial ascidians have also been studied for the ability to regenerate their tunic and the colonial circulatory system. In B. schlosseri, the full regeneration of the tunic and circulatory system occur in few hours when the peripheral matrix (i.e., the tunic and the enclosed portion of vasculature) is removed [62, 63] or days, when also marginal and radial vessels are ablated from the colony (Fig. 1k, l) [64, 65].

> As in solitary species, in more than a century of studies the surgical procedures employed to induce whole body or partial regeneration in colonial ascidians are roughly unchanged. However, the development of imaging and molecular tools allowed, in the last decades, the detailed study of the kinetics of regeneration, and the investigation of the molecular pathways involved in many aspects of WBR, as stem cells maintenance and differentiation (Table 4).

#### 4 Methodological Approaches to Induce Regeneration: An Historical Overview

4.1 Solitary Ascidians The first reports on induction of regeneration in solitary ascidians comes from Loeb (1892, cited in [36]), who observed siphon regeneration in Ciona, and Schultze [16], who documented siphon and brain regeneration and studied the process both in vivo and at the histological level. Almost 15 years later, Hirschler [18] cut individuals of Ciona transversely and obliquely through the thorax and utilized camera lucida drawings to record the regeneration process. Unfortunately, we do not have any information on the methodological approach used by these authors to induce regeneration. In 1930, Wermel published some methodological notes to study the regeneration of the oral siphon: he anesthetized animals of 4–7 cm in length (with MS222 or 10% chloral hydrate) and used fine dissection scissors to cut tissues [28].

Similar approaches for inducing regeneration were followed by later authors up to the more recently published reports, with some minor differences, such as the use of a scalpel to perform the ablations after the anesthetizing step [31].

Consecutive amputations of the oral siphon were used to study the recruitment of progenitor cells required for regeneration in Ciona [32].

In 1915, Se´lys-Longchamps described the evisceration in individuals of Polycarpa tenera kept in aquaria [12], also induced in the congeneric species P. mytiligera by Shenkar and Gordon [13], whereas Bourchard-Mandrelle [66] observed gonad regeneration in Ciona after their removal through a small hole in the body wall.

In 1992, Bollner and collaborators [67] set up the methodological approach for inducing brain regeneration in Ciona, used also in subsequent works [68]: after anesthesia with MS222 (0.02%) they first cut the epidermis and the nerves anterior to the neural complex to expose the anterior neural gland (ciliated funnel), and then proceeded through the epidermis toward the posterior part of the neural complex that was finally separated from the pharyngeal basket and the posterior nerves.

Dahlberg et al. [69] obtained better results than traditional microdissection in the induction of brain regeneration in Ciona by anesthetizing animals in MS222 (0.4 g/L) or propylene phenoxetol (0.06%) in seawater for 15–30 min before ablation, dissection or live imaging. For the ablation, they used fine forceps and biopsy punch tools (2 and 3 mm diameter). Animals were placed in silicone-coated Petri dishes and the entire cerebral ganglion was removed (with the associated neural gland, its ciliated funnel and the dorsal tubercle), in a single action to minimize the trauma. A different method to produce a brain lesion and induce its regeneration in S. plicata was described in 2015 by Medina and collaborators [70]: it consisted on the systematic injection in the pharyngeal region of the neurotoxin 3-acetylpyridine (3-AP; 65 mg/kg body weight), diluted in sterilized artificial seawater. This compound is a niacinamide antagonist that inhibits ATP synthesis, resulting lethal to the high metabolic rate of neurons. At selected time points following injection, they anesthetized, killed, and dissected the animals to collect their brains, which were then processed for their analysis.

Some general methodological approaches to induce regeneration in solitary ascidian can be summarized as follow:

4.2 Colonial Ascidians

	- (a) MS222 (e.g., 0.4 g/L in [69]) or
	- (b) 10% (w/v) chloral hydrate [28] or
	- (c) 0.06% (v/v) propylene phenoxetol [69] or
	- (d) menthol crystals (e.g., 0.4% (w/v) in [31]).
	- (a) For amputations (siphons, thorax, etc.)
		- <sup>l</sup> Dissection scissors and/or [28].
		- <sup>l</sup> Scalpels (see for example [31]).
		- <sup>l</sup> Forceps and biopsy punch tools (2 and 3 mm diameter) [69].
		- <sup>l</sup> Dissection on petri dishes (e.g., Sylgard®-coated) [69].
	- (b) For chemical induction of brain degeneration.
		- <sup>l</sup> Injection, in the pharyngeal region, of 3-acetylpyridine (3-AP; 65 mg/kg body weight) diluted in sterilized artificial seawater [70].
	- (c) For evisceration.
		- <sup>l</sup> Specimens kept alive in aquaria [12].
		- <sup>l</sup> Gently squeezing [13].

Up to the half of the nineteenth century, publications reporting experiments on colonial ascidians did not provide details on the methodological approaches used to induce regeneration. Publications simply report that individuals or stolons were cut and observed in vivo. Sometimes, regenerating fragments could be labeled with vital stains such as neural red, whereas fixed specimens were labeled with carminium for whole mount analysis [21]. Below, we report the available information on the three main kinds of regeneration in colonial ascidians.

One of the first studies on partial regeneration in B. schlosseri was that of Sabbadin, in 1956 [57]. In his study, he removed all the budlets (budectomy) of a colony except one with thin tungsten needles and razor blades under a dissecting microscope. The only remaining budlet was then removed when, after 72 h, it became a bud. The atrophied buds which are normally resorbed restarted their development and, eventually, became adults. Since 1956, numerous studies used B. schlosseri as a model organism to study WBR in colonial ascidians. In 1975, Sabbadin and Zaniolo [50] removed all the zooids and buds from colonies, with needles and razor blades under a dissecting microscope, leaving only the peripheral colonial matrix, that is, the tunic and its vasculature. They observed that, in a few days, a vascular bud developed from aggregation of blood cells and generated a new zooid. The same method was used by Rinkevich and collaborators [71] in B. leachii, Brown and collaborators [72] in B. violaceus, and Sunanaga and collaborators [45–47] in B. primigenus. Over the past 20 years, studies on regeneration using B. schlosseri and B. leachii have been implemented using molecular and biochemical techniques, studying in detail the involvement and function of some genes in regenerative processes of this two colonial ascidian [5, 53, 73–77].

Studies on partial regeneration in B. schlosseri were performed by Watkins [27], who damaged, with sharpened steel needles, the colonial buds and observed that about half of them regenerated the damaged part and reached maturity. The same method was used by Kaneko and collaborators [61] to induce partial regeneration in P. misakiensis.

Up to now, the regeneration of the vascular system was studied only in B. schlosseri. Zaniolo and Trentin [64] removed the entire colonial matrix (tunic, vessels and ampullae) around some zooids using a tungsten needle under a dissection microscope and observed the full regeneration of the peripheral vessels and tunic in 5 days. The same method was used in later studies [62, 63, 65, 78].

A general methodological approach to induce regeneration in colonial ascidian can be summarized as follow:


For WBR

1. Remove all the zooids, buds, and budlets from the colony, leaving only the colonial matrix [50].

For partial body regeneration

1. Remove the parts of interest of the animal from the colonies [27, 61].

For circulatory system regeneration

1. Remove the colonial matrix after cutting the radial vessels and the test all around the zooids [64].

#### 5 Concluding Remarks

As evidenced by the studies reported here, tunicates exhibit remarkable regenerative abilities, which have been studied from many points of view in over a century of researches. Besides their easy rearing and maintenance under laboratory conditions, the methods to induce regeneration are relatively simple and require few and cheap tools. Moreover, among chordates, only tunicates regenerate complete adult individuals from small tissue fragments. Despite these important characteristics, many molecular tools are still missing for this group of organisms. Transgenesis is one example: this technique would allow the in vivo study of gene expression during regeneration phases. However, it is still not developed especially for colonial ascidians and many solitary species as well. Today, the improvement of imaging techniques, and the broad application of some molecular biology tools, like RNA sequencing, can speed up the advancement of knowledge in the regenerative biology of ascidians. The studies summarized here will serve as a reference and a starting point for future researches aimed to uncover the biological basic properties of the astonishing regeneration in these chordates.

#### Acknowledgments

The authors wish to thank the COST Action MARISTEM 16203 community for supporting short term scientific missions and useful discussions.

#### References


animal models. Nat Rev Genet 7:873–884. https://doi.org/10.1038/nrg1923


intestinalis. J Zool Syst Evol Res 53:186–193. https://doi.org/10.1111/jzs.12101


https://doi.org/10.1111/j.1440-169X. 2010.01196.x


(Pallas) [Ascidiacea]. Arch Ital Anat Embriol 58:177–221


schlosseri. Sci Rep 4:1–11. https://doi.org/10. 1038/srep06460


expression network during oral siphon regeneration in Ciona. Development 144: 1787–1797. https://doi.org/10.1242/dev. 144097


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# Part II

Zoological Approaches

# Chapter 3

# Studying Protista WBR and Repair Using Physarum polycephalum

### Megan M. Sperry, Nirosha J. Murugan, and Michael Levin

#### Abstract

Physarum polycephalum is a protist slime mould that exhibits a high degree of responsiveness to its environment through a complex network of tubes and cytoskeletal components that coordinate behavior across its unicellular, multinucleated body. Physarum has been used to study decision making, problem solving, and mechanosensation in aneural biological systems. The robust generative and repair capacities of Physarum also enable the study of whole-body regeneration within a relatively simple model system. Here we describe methods for growing, imaging, quantifying, and sampling Physarum that are adapted for investigating regeneration and repair.

Key words Slime mould, Networks, Signaling, Extract, Regeneration, Injury

#### 1 Introduction

The protist slime mould Physarum polycephalum (which we will refer to as Physarum) exhibits generative capacities that permit the study of the basis of regeneration without the complexities of multi-cellular model systems. Although the multinucleated singlecelled slime mould lacks a fixed shape and nervous system, Physarum relays signals throughout its body using a network of branching cytoskeletal tubes that expand and contract to distribute biochemicals (Fig. 1) [1]. That intracellular communication method, known as shuttle streaming, acts as an intrinsic cellular oscillator that drives synchronization across the cell and allows for collective behavior of the organism (Table 1) [1]. The coordination of activities over short and long distances gives rise to more complex mechanosensing, problem solving, and decision-making capabilities that optimize the slime mould's ability to find food and avoid danger [2–6]. The coordinated oscillations also seem necessary for regeneration and repair capacities [7], however, the prior use of Physarum to study regeneration is limited.

Fig. 1 Physarum culture. (a) Plasmodial Physarum growing on agar substrate on day 3 (seed point out of frame). (b) Physarum undergoes plasmodial proliferation and builds branching networks to explore its environment

Relatively simple to culture, image, and sample, the single-cell Physarum is capable of regeneration in its most basic form [7]. Physarum can grow from discrete drops of liquid protoplasm to form a single plasmodial cell [7], develop an entire plasmodial network from a small seed point by plasmodial proliferation [8], fuse with separate plasmodial networks [9], and rapidly heal from injuries within its vein network (Table 1). Furthermore, Physarum takes multiple forms that allow hibernation under stressful conditions and growth when environmental conditions are optimal [10]. In suboptimal conditions (low humidity, lack of food, and in the presence of light), Physarum will reconfigure into a dormant, encrusted state known as sclerotia (Table 1) [10], where it can hibernate for months to years. Sclerotia rapidly transforms to a vegetative plasmodial state in the presence of humidity and food sources (Table 1), forming a branching slime mould network. In this dynamic state, Physarum uses shuttle streaming to distribute biochemicals throughout the plasmodial network and is capable of kinase signaling typically observed in eukaryotic organisms [11].

In contrast to regeneration programs that advance toward a large-scale target morphology [12, 13], Physarum has a strong generative capacity with consistent regeneration of hierarchical


#### Table 1 Definitions of Physarum States & Anatomy

branching patterns [14]. Similar to Physarum, plants possess a high degree of developmental plasticity, exhibiting consistent, reproducible shoot and root systems from leaf cuttings, with a shoot on one side and a root on the other and small variations in shape from its original body (regeneration in plants fully reviewed in [15]). In this chapter, we define whole-body regeneration as the development of an entire plasmodial network from a small seed point by plasmodial proliferation. In addition, we present methods for transforming sclerotia into a vegetative state and for regeneration of Physarum veins, which are methods for revival and tissue repair, but do not represent regeneration of the entire organism.

Also similar to plants, the large-scale morphology of Physarum is strongly driven by conditions in its local environment, such as attractants, repellants, and substrate stiffness [4, 16]. Physarum is extremely capable of growing and retracting in response to its surroundings and seemingly prioritizes efficient sampling of its surroundings and reinforcement of its beneficial branches over growing into a precise shape [16]. However, given the same environmental conditions, like a maze, Physarum will repeatedly identify the minimum-length solution by retracting veins that reach dead ends [17–19]. Therefore, in this chapter, we offer examples of fixed environmental conditions that can be used to assess repair and regrowth in the context of functional patterning, such as the Salt Bridge. Salt interferes with the ionic homeostatic mechanisms of Physarum and therefore exposure to salt will cause Physarum to generate unique whole-body morphologies in its presence [9].

The strong generative capacity of plants relies on stem cells that form at the cut site, elongate and proliferate, and produce new plant bodies [15]. Dissimilar to plants that possess a meristem region of undifferentiated cells, the entire plasmodial Physarum network exists as an undifferentiated single cell. Nuclei of plasmodial Physarum divide without passing through cytokinesis, giving rise to a large multinuclear syncytium [8]. Although plasmodial Physarum and its encrusted state, sclerotia, do not undergo cellular differentiation or even cytokinesis, when Physarum mature or food becomes limited, plasmodia will differentiate into sporangia in the presence of light [11]. The sporangia release haploid spores, which germinate, form flagellated myxamoebae, genetically recombine with myxamoebae from other plasmodia, and fuse into a zygote that develops into a new plasmodium [11].

In this chapter, we present methods for regenerating Physarum from its dormant sclerotia state, culturing plasmodia, and drying plasmodia to sclerotia. We describe methods for the modeling of injury and regeneration, particularly in complex environments like mazes and salt bridges. We also present methods for automated macrophotography to monitor Physarum growth and form. For more detailed analysis of Physarum form and function, we introduce microscopy approaches, including the injection of fluorescent polymers and beads prior to imaging. Those fluorescent materials can be used to observe the response of Physarum to localized nutrient or chemical stimuli via cytoplasmic shuttle streaming, which is the primary method of information transfer throughout the organism's body [1]. Methods to quantify both the structural and functional Physarum networks during growth and repair are also described. Finally, we outline techniques to sample Physarum networks and plasmodial slime for downstream use in mass spectrometry metabolomics or gene expression measurements like polymerase chain reaction or RNA sequencing [11, 20].

#### 2 Materials

Prepare all solutions using ultrapure water (0.2 μm-filtered and deionized). Prepare and store all reagents at room temperature (unless indicated otherwise).



#### 3 Methods

Carry out all procedures at room temperature unless otherwise specified. 3.1 Physarum Culture 1. Pour 2 mL of sterile water onto a 1 cm2 filter paper containing dehydrated Physarum sclerotia (see Note 9). 2. Place moistened filter paper, sclerotia-side down, at the center of a 1% agar-filled plate. 3. Arrange 4–6 oatmeal flakes 3–5 cm from the sclerotia. 4. Apply a single spray of sterile water on the plate to moisten, but not soak, the environment. 5. Maintain cultures at 22 C and 90% humidity in a dark incubator (see Note 5). 6. Wait 2 days for the Physarum plasmodia to start growing toward the oat flake food sources (see Note 10). 7. Add 10 to 12 oat flakes to cover the rest of the plate. 8. Apply a single spray of sterile water to the plate. 9. Return to incubator. 10. Add fresh oat flakes every 2 days (see Note 10). 11. Return to incubator. 12. Wait a total of 5 days for Physarum to completely cover the plate. 13. Cut a 1 cm2 cube of Physarum-coated agar using a sterile toothpick. 14. Transfer the piece of agar, Physarum-up, to the center of a fresh 1% agar plate (Fig. 2a, b) to sub-culture the Physarum to a new plate. 15. Spread 10 to 12 oat flakes across the agar. 16. Apply a single spray with sterile water. 17. Incubate at 22 C and 90% humidity in the dark. 18. Repeat steps 14–17 to maintain a growing culture of Physarum (see Note 11). 19. Allow plasmodium to grow over filter paper to begin transition to the sclerotia state for long-term storage. 20. Move the filter paper with plasmodium to dry, dark conditions

for at least 2 days to produce sclerotia.

Fig. 2 Physarum growth and repair. (a) Physarum grows from a single seed point (arrow) towards food sources (arrowheads) one day after sub-culturing. (b) After 3 days, the plasmodial Physarum covers the plate, forming a network that includes the seed point (arrow) and all food sources (arrowheads). (c) Before injury, a vein connects two food sources. The food source in the upper right corner is proximal to the Physarum seed point compared to the food source in the lower left corner. (d) An injury to the vein that leaves the underlying agar intact can be (e) rapidly repaired within 5 h

21. Keep the sclerotia in dry and dark conditions for long-term storage. The dry sclerotia can remain dormant for up a year.

The repair processes described here can be monitored using the imaging approaches outlined in Subheading 3.4.


3.2 Models of Vein Repair After Injury & Amputation


The generative processes described here can be monitored using the imaging approaches outlined in Subheading 3.4.

	- 2. Place the Physarum dish(es) in the field of view of the macrophotography set-up inside the incubator.
	- 3. Set the image acquisition rate and total imaging time to automatically acquire images (Fig. 4a) (see Note 14).
	- 4. Wait until the total imaging time has elapsed.

3.3 Generation in Complex Environments: Maze & Salt Bridge Experiments

3.4 Structural & Functional Imaging

Fig. 3 Physarum generation in complex environments. (a) Physarum grows within the confines of the maze (white arrows) toward the food source and retreats from areas lacking food (red arrow), building an optimized morphology based on the environmental conditions. (b) The oat flakes at the primary maze entrance points (blue arrows) are placed to encourage Physarum growth in three directions. Despite this, after 48 h Physarum only grows toward the maze endpoint (purple arrow; oatmeal flake removed). (c) The agar bridge connects Physarum samples to the 10% flan medium. Salt or other substances may be added to the bridge to induce morphological changes, such as (d) growth around the noxious salt bridge after 24 h


Fig. 4 Structural and functional visualization of Physarum plasmodium. Physarum structure visualized using (a) a scanner, (b) brightfield microscopy, or (c) darkfield microscopy. Brightfield imaging captures the outer architecture of the Physarum, whereas darkfield imaging shows compartments within each vein (insets). (d) Fluorescent dextran travels through the Physarum network, with preference for certain paths (arrow) over others (star). Dextran can be shuttled through the network and deposited in slime secretions (box). (e) Fluorescent beads travel through the Physarum network (shown at high magnification)


#### 3.5 Quantification of Growth & Repair This section outlines a general protocol for Physarum image processing and network analysis.


Fig. 5 Image processing and network analysis pipeline to quantify Physarum shape and connectivity. (a) Images saved from a scanner are imported and (b) filtered, (c) used to create a template, and (d) processed to enhance vein structure. The single pixel-wide skeleton is extracted and used to form a (e) network graph for evaluation using metrics, like (f) vein width. Scale bar ¼ 1 cm for all panels


#### 3.6 Sampling Physarum


#### 4 Notes


#### Acknowledgements

This research was also supported by the Allen Discovery Center program through The Paul G. Allen Frontiers Group (12171). We also gratefully acknowledge the Defense Advanced Research Projects Agency (DARPA) under Cooperative Agreement Number HR0011-18-2-0022. The content of the information does not necessarily reflect the position or the policy of the Government, and no official endorsement should be inferred. Approved for public release; distribution is unlimited. The authors also thank Melanie Chien for assistance with experiments and Cuong Nguyen for training in laser cutting during preparation of the maze.

#### References


biological cognition. Biosystems 165:57–70. https://doi.org/10.1016/j.biosystems.2017. 12.011

3. Whiting JGH, Jones J, Bull L et al (2016) Towards a Physarum learning chip. Sci Rep 6: 1–14. https://doi.org/10.1038/srep19948


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 4

# Studying Porifera WBR Using the Calcerous Sponges Leucosolenia

### Andrey I. Lavrov and Alexander V. Ereskovsky

#### Abstract

Sponges (Porifera), basal nonbilaterian metazoans, are well known for their high regenerative capacities ranging from reparation of a lost body wall to whole-body regeneration from a small piece of tissues or even from dissociated cells. Sponges from different clades utilize different cell sources and various morphological processes to complete the regeneration. This variety makes these animals promising models for studying the evolution of regeneration in Metazoa. However, there are few publications concerning the regenerative mechanisms in sponges. This could be partially explained by the delicacy of sponge tissues, which requires modifying and fine adjusting of common research protocols. The current chapter describes various methods for studying regeneration processes in the marine calcareous sponge, Leucosolenia. Provided protocols span all significant research steps: from sponge collection and surgical operations to various types of microscopy and immunohistochemical studies.

Key words Regeneration, Sponges, Protocols, Microscopy, Surgical operations, Ultrastructure, Proliferation, Apoptosis

#### 1 Introduction

Sponges (Phylum Porifera) are thought to be the sister group of all other animals and the earliest branching multicellular lineage of extant animals [1]. As such, they represent a key group for the understanding of the evolutionary history of animals, including the origin and evolution of regeneration mechanisms. The body shape of sponges is very diverse; they may be film-like, encrusting, lumpy or spherical, tubular, branching, flabellate, and so on. The body size of sponges varies as much as their body shapes: from 3–10 mm to 1.5–2 m [2]. Their organization is peculiar: they have no distinct gut, muscles, gonads, nervous system, or respiratory system. The surface of a sponge is covered by a simple singlelayered flat epithelium (called exopinacoderm), while the internal parts of the animal body are occupied by a highly complex mesenchymal tissue (called mesohyl) that comprises numerous mobile cell

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_4, © The Author(s) 2022

types embedded in its extracellular matrix [3]. The rigidity of the sponge body is ensured by the collagen and spongin fibrils (in some Demospongiae orders) and by the internal inorganic skeleton, consisting of either calcium carbonate (CaCO3) (Calcarea, some Demospongiae) or silica (SiO2) (Hexactinellida, Demospongiae, many Homoscleromorpha).

The mesohyl is penetrated by a complex system of canals and choanocyte chambers (termed the aquiferous system), which is the most characteristic feature of the poriferan anatomy. Sponges use this constant water pumping system to obtain food and oxygen and to remove metabolic wastes. Surrounding water is drawn into the inhalant canals via numerous pores (ostia) in the exopinacoderm. Water then circulates through choanocyte chambers, where it is filtered, before leaving the sponge via the system of exhalant canals that converge to large exhalant openings (osculum) [4]. The choanocyte chambers are lined by flagellated collar cells (called choanocytes). The constant beating of choanocytes' flagella generates the water flow through the whole aquiferous system, and their collars serves as an ultimate filter for retaining particles from the pumped water [2].

Five types of the body organization have been described in sponges: (1) asconoid, (2) solenoid, (3) syconoid, (4) sylleibid, and (5) leuconoid [3]. These types differ by the complexity of the aquiferous system and the extent of the mesohyl development. In the simplest asconoid sponges, ostia lead directly to a single cavity completely lined with the choanocytes, and the mesohyl is represented only by thin, mostly acellular layer. The more complex leuconoid sponges are characterized by an elaborated aquiferous system with highly developed canals and numerous choanocyte chambers and thick mesohyl with numerous specialized cell types.

A characteristic feature distinguishing sponges from other Metazoa is the high plasticity of cellular differentiation, anatomical, and tissue structures throughout their life cycle. Various differentiated cells of the sponge can move, transdifferentiate, and switch functions. The direction of the differentiation depends on the current needs of the organism. Thus, the sponge is constantly in the state of rearrangements of all its structures [5–9]. This "chronic morphogenesis" contributes to the growth of the animal, for instance, by reconstructing its somatic tissue after degradation during sexual and asexual reproduction, as well as during regeneration [10–13]. Besides, sponges are not equipped with protective tissues or structures like cuticles, scales, or shells, but are covered only by a single-cell layer. It has been suggested that this lack of protection against injury closely correlates with the high regenerative capacity of sponges [14].

Sponges are known to possess remarkable reconstitutive abilities ranging from restoration of a lost body part to a complete organism development from a small piece of tissue and even from the cell suspension. However, only few reliable data on the regeneration mechanisms (morphogenesis, cell behavior, and regulation) and their distribution among sponge clades currently exists [10, 12, 15–18].

We provided complex and detailed investigations of reparative regeneration in homoscleromorphs [10], calcareous sponges [13, 19], and demosponges [11, 12]. These studies included various approaches: transmission electron microscopy (TEM), scanning electron microscopy (SEM), epifluorescent and light microscopy, immunohistochemistry, and time-lapse recordings. The obtained results show a high diversity of morphogenesis, cell mechanisms, and cell turnover, accompanying the regeneration processes.

The model sponges presented in this chapter belong to the genus Leucosolenia, an abundant species broadly distributed in the White Sea and in the North of Europe, where they are accessible throughout the year. Leucosolenia are calcareous sponges, characterized by a calcium carbonate mineral skeleton and the asconoid aquiferous system (Fig. 1a).

The body wall of Leucosolenia has a thickness of 15–30 μm and is composed of three layers: an outer layer—the exopinacoderm, a central region—the loose mesohyl, and an inner layer—the choanoderm (Fig. 2b). Inhalant pores (ostia) are scattered throughout the exopinacoderm. They are formed by tubular cylindrical cells (porocytes), which connect the external milieu with the internal choanocyte cavity. The mesohyl of Leucosolenia contains a variety of

Fig. 1 Surgical operations in Leucosolenia variabilis. (a) sponge in vivo; (b) scheme of a sponge with different types of surgical operations: (1) body wall regeneration, (2) whole-body regeneration (WBR) from an amputated oscular tube, (3) WBR from an amputated diverticulum, (4) WBR from an amputated cormus tube, (5) WBR from a small fragment of the body wall, (6) cell reaggregation after mechanical tissue dissociation. WBR could be observed during the restorative process in amputated body tubes (2–4), small fragments of the body wall (5) and during cell reaggregation after dissociation (6). WBR from amputated body tubes requires minimal rearrangements of intact tissues, while cell reaggregation is accompanied by complete destruction of intact tissue structure. WBR from small fragments of the body wall represents an intermediate type. d diverticula, ot oscular tubes

Fig. 2 Various methods for investigations of Leucosolenia variabilis regeneration. (a) regenerative membrane growing from the periphery to the center of the wound orifice, stereomicroscope (in vivo), white arrowheads mark mesohyl cells inside the regenerative membrane. (b) Semithin section of regenerative membrane transformed into intact body wall (96 hpo). (c) TEM micrograph of complete regenerative membrane (24 hpo) consisting of the flattened exopinacocytes and endopinocytes transdifferentiated from choanocytes. (d) SEM micrograph of choanocytes transdifferentiating into endopinacocytes during growth of regenerative

cell types, including sclerocytes, rare amoeboid cells, symbiotic bacteria, spicules, as well as gametes and developing embryos during the reproduction season.

The rapid wound healing and high regeneration capacity after different surgical interventions were demonstrated, indicating that Leucosolenia are a promising model for sponge regeneration investigations [20]. Leucosolenia complicata and L. variabilis have been successfully used for different experiments concerning the study of restoration morphogenesis [13, 19, 21–26].

In a recent publication, we revisited various regenerative processes of L. variabilis from the White Sea using electron microscopy, laser confocal microscopy, epifluorescence microscopy, and time-lapse microscopy. These approaches allowed us to precisely address the issues of morphogenetic mechanisms, cell transdifferentiations, movements, and proliferation. Our study reveals the contributions of cell types to reparative regeneration in this species and demonstrates a central role of epithelial morphogenesis and transdifferentiations in the regeneration process [13].

Leucosolenia demonstrate high and diverse regenerative capacity after various surgical operations. In this chapter, we provide methods to study the reparative regeneration of the body wall, whole-body regeneration (WBR) from amputated tubes and small fragments of the body wall, as well as cell reaggregation and primmorph formation after tissue dissociation. We also provide protocols for cell proliferation, apoptosis, and immunohistochemical studies. Finally, we present approaches for functional analysis of cell proliferation and skeleton synthesis during regeneration.

#### 2 Materials


Fig. 2 (continued) membrane (24 hpo). (e) Cell proliferation is neither affected nor contributes to the regeneration at any stage of the process, for example, during transformation of regenerative membrane into intact body wall (36 hpo), white dashed line delimits regenerative membrane, orange arrowheads mark cells in S phase of cell cycle (EdU-positive cells), white arrowheads—cells in late G2/M-phase of cell cycle (pH3-positive cells). (f) Apoptosis during early stages (3 hpo) of regeneration, white dashed line marks wound surface, white arrowheads mark apoptotic (TUNEL-positive) cells. (g, h) Spicule secretion in the regenerative membrane during its transformation into intact body wall (72 hpo), (g) general view under bright field microscopy, (h) epifluorescence view under FITC filter set with newly synthesized spicules showing bright green emission, white arrowheads marks the same spicules in (g) and (h). (a, b, c, d, g, h) WBR from an amputated oscular tube; (e) WBR from an amputated cormus tube; (f) body wall regeneration. Scale bars: (a) 200 μm, (b, e, f, g, h) 50 μm, (c, d) 5 μm. ch choanocyte, en endopinacocyte of regenerative membrane, ex exopinacocyte of intact tissue, f flagellum, m mesohyl, mv microvilli of choanocyte, rm regenerative membrane, tch transdifferentiated choanocyte, wo wound orifice


solution, 0.04 mL 10% (w/v) ruthenium red aquiferous solution, 2.96 mL 0.1 M Na-Cacodylate buffer. Use freshly prepared.


#### 3 Methods



	- 1. Cut oscular tubes perpendicular to the main axis using scissors.
	- 2. Remove sponge from the Petri dish.
	- 3. Maintain the amputated oscular tubes in the Petri dish with FSW at 10–12 C.
	- 4. Change half of FSW with fresh one every 24 h.
	- 5. Inspect and photograph the amputated oscular tubes as indicated above.

3.2.3 Whole-Body Regeneration Excision and cultivation of fragments of the body wall (Fig. 1b 5) will allow for studying the whole-body regeneration (WBR) of the sponge.


#### 3.2.4 Cell Reaggregation This type of operation (Fig. 1b 6) (see Note 6) will allow for studying the WBR of the sponge:



buffers provide equal quality of fixations. However, the same one should be used during fixation and treatments of a single specimen. Unless specified, all incubations and rinses are performed at RT "with shaking," that is, with constant orbital shaking at 70 rpm.


The fixation and subsequent treatments for ECM and cell junction visualization should be done using 0.1 M Na-Cacodylate buffer, as phosphate ions block ruthenium red interactions with tissues.


After fixation, specimens should be dehydrated and embedded "with shaking" until step 14:


The fixation, postfixation, and dehydration of specimens should be done as detailed in steps 1–11 in Subheading 3.5 (see Note 13).

3.6 Methods of Tissue Fixation and Processing for Scanning Electron Microscopy (SEM)

3.7 Cell Proliferation and Immunohistochemical Studies


For cell proliferation studies, a combination of 5-ethynyl-2- 0 -deoxyuridine (EdU), labeling cells in S-phase of the cell cycle (DNA-synthesizing cells), and anti-phospho-histone H3 antibodies, labeling cells in lateG2/M-phase of the cell cycle (dividing cells), is used (Fig. 2e). Unless specified, all incubations and rinses are performed at RT "with shaking," that is, with constant orbital shaking at 70 rpm.

	- 2. Change media every 12–24 h, maintaining a constant concentration of a blocking agent, if prolonged blocking is required.
	- 3. Wash sponge tissues in a large volume of fresh FSW and incubate it for at least 24 h at 10–12 C to release cell proliferation.

3.9 Apoptosis Studies The following protocol is a slightly modified manufacturer's protocol for Click-iT TUNEL Imaging assay (Thermo Fisher Scientific) and In Situ Cell Death Detection Kit (Merck) (see Notes 21 and 22). Unless specified, all incubations and rinses are performed at RT "with shaking," that is, with constant orbital shaking at 70 rpm.


```
As spicules of Leucosolenia are composed of calcium carbonate
(CaCO3), their synthesis could be visualized in vivo, using Calcein
disodium salt solution (see Note 25).
```

3.10 Skeleton Synthesis Studies

#### 4 Notes


regeneration occurs after the excision of a small part of the body wall or amputation of an oscular tube. The process ends within 4–6 days post-operation (hpo) with complete restoration of the lost body wall. Three main stages could be distinguished in this type of regeneration: (1) internal milieu isolation (3—12 hpo), (2) wound orifice healing (regenerative membrane formation) (12—24 hpo) (Fig. 2a), and (3) transformation of the regenerative membrane into an intact body wall (48—144 hpo) [13].


The illumination is no less important. Not all light sources are equally suitable to observe living specimens, especially long term during time-lapse recordings. Avoid using heating light sources, as they will quickly raise the temperature of a specimen. Also, pay attention to the wavelength profile of a light source, as a light source with high intensities in the "blue" part of the spectrum may have deleterious effects on a living specimen. For time-lapse recording with a stereomicroscope, a lateral illumination gives a better contrast to the specimens. It also could be combined with transmission illumination. The light intensity should be set to a moderate level, as too bright illumination could negatively affect the viability of a specimen. The optimal recording period depends on the studied process: the quicker process—the shorter period should be used. Usually, a set of preliminary recordings are required to determine the optimal period.

	- (a) Negative control specimen (NCS), an alive regenerating specimen, which is incubated in DMSO solution in FSW at step 1 (instead of EdU solution, as in experimental specimens). Each batch of experimental specimens should be supplemented with NCS. Incubate NCS in a 30 mm plastic Petri dish with 5 mL of FSW supplemented with DMSO for 6 h at 10–14 C in parallel with the experimental specimens according to Subheading 3.7. The volume of added DMSO is equal to the volume of the EdU stock solution added to the experimental specimens. After incubation, treat NCS similarly to experimental specimens. The staining patterns in EdU-channel (555 nm) in NCS should be recognized as unspecific, and similar patterns in experimental specimens should not be considered.
	- (b) Positive control specimen (PCS), any alive specimen, which admittedly contains DNA-synthesizing cells. We recommend supplying each batch of experimental specimens with PCS. Treat PSC similarly and parallel to the experimental specimens, starting from step 1 of Subheading 3.7. If, during the study with a confocal microscope PCS shows, no EdU-positive cells, then experimental specimens should be discarded as some issues with EdU incubation or Click-reaction arose.

Immunohistochemical studies could be done in parallel with EdU cell proliferation studies (in this case, apply primary and secondary antibodies at steps 10 and 12 in Subheading 3.7, respectively) or separately (in this case, omit steps 1–2, 5, and 7–8 in Subheading 3.7).

At least at several first treatments, two negative control specimens should be used to control the quality of immunohistochemical staining and correctly interpret obtained Z-stacks, discriminating between specific and unspecific signals:


apoptosis. In Situ Cell Death Detection Kit is based on one-step labeling through the incorporation of dUTP conjugated with fluorescent dye in cell DNA at double-strand break sites. In turn, Click-iT TUNEL Imaging assay is based on two-step labeling: at the first step, dUTP modified with alkyne (EdUTP) is incorporated in cell DNA at double-strand break sites, at the second—Click-reaction visualizes EdUTP with Alexa Fluor azide. While In Situ Cell Death Detection Kit offers simpler and shorter treatments, Click-iT TUNEL Imaging assay gets an advantage of better penetration in tissue due to the small size of both EdUTP and Alexa Fluor azide.

	- (a) Negative control specimen (NCS), a specimen, in which the TdT reaction at step 6 is blocked. Treat NCS similarly to experimental specimens but incubate it TdT-cocktail devoid of TdT-enzyme. The staining patterns in TUNELchannel in NCS should be recognized as unspecific, and similar patterns in experimental specimens should not be considered.
	- (b) DNase positive control specimen (DNase-PCS), a specimen (intact sponge tissues), in which dsDNA breaks are artificially introduced by DNase I. Treat DNase-PCS similarly to experimental specimens but incubate it in DNase I solution containing 1–2 U of the enzyme for 30 min at 37 C immediately prior to step 5. DNase I incubation solution could be prepared by mixing commercially available DNase I enzyme, 10 DNase reaction buffer, and appropriate volume of MilliQ. After DNase treatment, rinse DNase-PCS three times with 1 PBS, 10 min each time, and proceed to step 5.
	- (c) As DNase I is highly volatile, to avoid contamination of experimental samples with it (which will generate pseudopositive staining) use a separate set of instruments for manipulations with DNase-PCS. If during the study with a confocal microscope, DNase-PCS shows no TUNEL-positive cells, then experimental specimens should be discarded as some issues with TdT-reaction or Click-reaction arose.

should try to change enzymes on fresh ones. Also, you could try to extend the duration of enzymatic reactions and raise reaction temperature to 37 C.


#### Acknowledgments

We gratefully thank our colleagues Fyodor Bolshakov, Veronika Frolova, Nikolai Melnikov, Ksenia Skorentseva, Alexandra Koynova, and Igor Kosevich (Moscow State University, Russia); Daria Tokina and Ilia Borisenko (Saint-Petersburg State University, Russia); and Emilie Le Goff and Stephen Baghdiguian (Universite´ de Montpellier, France) for their invaluable advices and help. The light microscopy studies were conducted using equipment of the Center of microscopy WSBS MSU, electron microscopy studies in the Electron Microscopy Laboratory of the Shared Facilities Center of Lomonosov Moscow State University sponsored by the RF Ministry of Education and the Common Service of morphology in IMBE, France. We are also indebted to the COST Action 16203 MARISTEM, which facilitated author discussions over the course of its periodic meetings. Studies were supported by the grants of the Russian Science Foundation no. 17-14-01089 (in vivo regeneration experiments), RFBR nos. 19-04-563 and 19-04-00545, and Fund of President RF no. MK-1096.2021.1.4.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 5

# Studying Ctenophora WBR Using Mnemiopsis leidyi

### Julia Ramon-Mateu , Allison Edgar, Dorothy Mitchell, and Mark Q. Martindale

#### Abstract

Ctenophores, also known as comb jellies, are a clade of fragile holopelagic, carnivorous marine invertebrates, that represent one of the most ancient extant groups of multicellular animals. Ctenophores show a remarkable ability to regenerate in the adult form, being capable of replacing all body parts (i.e., wholebody regeneration) after loss/amputation. With many favorable experimental features (optical clarity, stereotyped cell lineage, multiple cell types), a full genome sequence available and their early branching phylogenetic position, ctenophores are well placed to provide information about the evolution of regenerative ability throughout the Metazoa. Here, we provide a collection of detailed protocols for use of the lobate ctenophore Mnemiopsis leidyi to study whole-body regeneration, including specimen collection, husbandry, surgical manipulation, and imaging techniques.

Key words Ctenophore, Mnemiopsis leidyi, Wound healing, Whole-body regeneration, Husbandry, Surgeries, Live imaging, Time-lapse

#### 1 Introduction

While regenerative capabilities are common across the animal kingdom, the ability to regenerate all the structures of the body (i.e., whole-body regeneration) is a rather unique feature only found in some species. Ctenophores (comb jellies) are one such animal with impressive whole-body regenerative capabilities; they are holopelagic, carnivorous marine invertebrates that represent one of the oldest extant metazoan lineages [1]. Ctenophores have a unique body plan characterized by a biradial symmetry (with no planes of mirror symmetry) and one primary body axis (the oral–aboral axis) delimited by a mouth (oral) and an apical sensory organ (aboral). The ctenophore body is composed of two epithelial layers: the ectoderm—including the epidermis, apical organ, pharynx, nerve net, ctene plates (or comb plates), and tentacle sheath—and the endoderm primarily composed of a system of endodermal canals that distribute nutrients to the periphery of the animal. The

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_5, © The Author(s) 2022

ectodermal and endodermal tissues are separated by a thick mesoglea mostly composed of extracellular matrix, but also containing several types of individual muscle and mesenchymal cells [2]. The characteristics of the mesoglea differ between ctenophore species. For example, in Pleurobrachia species (the sea gooseberry), the mesoglea is rather rigid, while in lobate ctenophores the mesoglea is highly pliable, presumably due to differences in hydration characteristics. Ctenophores' main mode of locomotion is via the coordinated beating of their comb plates. They possess eight longitudinally oriented rows of locomotory ctene plates, each plate composed of thousands of laterally arranged cilia which they coordinately beat to propel through the water column. Ctenophores have been accurately described morphologically for over a century, with the first volume of the Flora and Fauna of the Statione de Napoli being dedicated to Ctenophora by one of the world's first experimental embryologists, Carl Chun [3].

One of the best-studied species of ctenophores in the regenerative field is the lobate ctenophore Mnemiopsis leidyi [4–10]. Like the majority of ctenophores [11], M. leidyi is a self-fertile hermaphrodite, meaning that a single animal carries both female and male gonads. The eggs and sperm are released freely into a common sinus under each comb row and fertilization takes place upon release into the water column [12]. Like most ctenophores, M. leidyi produces embryos which are optically clear and, like all ctenophores, it has a very stereotyped, clade-specific cleavage program where rounds of division occur every 20 min at room temperature and the juvenile cydippid stage hatches from the fertilization envelope within 18–24 h after the first cleavage [13– 15]. The cydippid is a feeding form characterized by a pair of long branching muscular tentacles that define the tentacular axis and bear specialized adhesive cells called colloblasts, used to capture prey [16, 17]. In lobate ctenophores like Mnemiopsis, the tentacles are progressively reduced and internalized during the transition to adulthood as the animal forms two large oral lobes that are extremely efficient at prey capture (Fig. 1). Under optimal conditions, the adult form can get sexually mature at ~4 weeks of age, though sexual reproduction at the juvenile morphological stage, termed "dissogeny," has been documented as early as 2 weeks [18, 19].

Our recent study shows that in M. leidyi cell proliferation is activated (after wound-healing) at the wound site and is indispensable for whole-body regeneration. EdU pulse and chase experiments after surgery together with the removal of the two main regions of active cell proliferation suggest a local source of cells in the replacement of missing structures. Time-lapse live imaging during M. leidyi wound healing shows evidence of cells forming actin-based protrusions while migrating to the wound site [10]. While lobate ctenophores show an outstanding capacity to

Fig. 1 The life cycle of Mnemiopsis leidyi. The adult body plan is referred to as "lobate," describing their preycapture tissues (oral lobes) that extend from the oral end. Adults produce both eggs and sperm. Embryos are ~150 μm in diameter and develop from single cell to hatching over ~24 h. The hatched, free-swimming feeding juvenile body plan is referred to as "cydippid" and characterized by a relatively shortened body and long prey-capture tentacles. The cydippids will start to transition into the lobate body plan as their tentacles retract, body lengthens and lobes form. Adult lobates will continue to grow until they reach a maximum size of around 6–18 cm

regenerate all body parts, another group of ctenophores, the Beroids, have lost the ability to regenerate [9]. Hence, the comparison of cellular and molecular responses after amputation between lobate ctenophores and Beroids provides an ideal system to elucidate the core cellular and molecular responses required for the process (and loss) of adult regenerative potential.

The many favorable experimental features provided by ctenophores (optical clarity, stereotyped cell lineage, multiple cell types, sequenced genome available [20], comparative and functional genomics [21–27], rapid regeneration (48–72 h) and their early branching phylogenetic position (potentially the earliest extant animal clade [1, 20, 28, 29]) make them a new powerful research organism for the study of regeneration at a cellular and evolutionary level. Here we provide a detailed protocol to use ctenophores to study the process of whole-body regeneration from the collection of specimens and husbandry in the laboratory to the deployment of basic techniques for the study and monitoring of regeneration at a cellular level.

#### 2 Materials

	- 2. 20 L plastic bucket.
	- 3. Ctenophore tank system: 200 L tank made of plexiglass (PMMA) walls consisting of an inner tank module enclosed between two outer compartments with drains both at the bottom and the top of the tank (Fig. 2B, C).
	- 4. 6<sup>00</sup> diameter glass bowls (Fig. 3).
	- 5. 2 L glass beaker (Fig. 3).
	- 6. 1 mL transfer plastic pipettes.
	- 7. Rotifer culture system: 2 L glass beaker, a 5 - 5 cm piece of rotifer floss (e.g., Reed Mariculture, Inc), air pump (e.g., Tetra Whisper Aquarium Air Pump) connected by plastic tubing to a 1 mL serological pipette.
	- 8. A coarse filter: 30-μm nylon mesh screening (e.g., Nitex) affixed to a section of pipe or plastic container with the bottom removed.
	- 9. Artemia hatching system: 1.5 L plastic cone with a stopcock (valve) at the bottom placed inside a support, aeration system made from a 1 mL plastic pipette attached at one end to a plastic tube connected to a standard aquarium air pump from the other end.
	- 10. Operating dish: 35-mm plastic petri dish, coated with a 2-mm thick silicon (SYLGARD-184) layer.
	- 11. Microburner: 16-gauge syringe needle inserted into latex tubing attached to a propane source (Fig. 4A).
	- 12. Pulled glass needles from Pyrex capillaries (Fig. 4A).
	- 13. Pair of fine forceps (e.g., World Precision Instruments, Cat#500341).
	- 14. Siliconized slide: microscope glass slide treated with a synthetic hydrophobic surface-applied product (e.g., Rain-X, Inc.).

Fig. 2 Ctenophore collection and culturing materials. (A) A "ctenophore dipper" constructed to collect ctenophores from the field. Ctenophores located close to the surface of the water are gently scooped into the beaker portion of the dipper. (B) The pseudo-kreisel tank system used to contain adult ctenophores. (C) Diagram of a pseudo-kreisel tank system for culturing adult M. leidyi at the lab. Two water inlets located in the tank bottom generate a continuous flow that pushes water up the sides of the tank. Drains on the top and bottom of each side displace the input of water, while two partially perforated plexiglass sheets contain the animals in the central space. The flow of water keeps the ctenophores toward the tank's center, while constant water flow through the system prevents fouling


#### UV-FSW: UV treated 1.0-μm filtered full strength seawater (e.g., 35 g/L).

2. 1- 0.2-μm UV-FSW: UV treated 0.2-μm filtered full strength seawater.

### 2.2 Reagents 1. 1-

Fig. 3 Husbandry and culture of Mnemiopsis leidyi. Culturing conditions for each stage of the M. leidyi life cycle. Spawning is induced in wild-caught or captive adults by manipulating their light exposure. Embryos are collected and placed in a 6<sup>00</sup> diameter glass finger bowl in 1- UV-FSW. Hatching occurs 18–24 h postfertilization (hpf). Hatched cydippids (M. leidyi juveniles) are grown in 2 L glass beakers to provide space for hunting behavior as their tentacles tend to get tangled with other individuals in smaller containers. Cydippids grow into adults in the next 3–4 weeks postfertilization as they grow in size, retract their tentacles, and develop oral lobes for prey capture


Fig. 4 Methods for ctenophore tissue regeneration assays. (A) Surgery instruments. Glass needles are handpulled using a microburner. (B) Puncture assay for M. leidyi cydippids. Tissue is punctured in a space that is clear of organs (tentacle bulb, comb rows, etc.) to assay wound healing of the epithelia. (C) Designs for multiple types of amputations. Oral–aboral bisection is performed by cutting tissue parallel to the esophageal canal and slightly to the side of the apical organ. This is so one half of the animal maintains an intact apical organ, as it is more likely to regenerate with this feature. Apical organ amputation includes cutting the space between the top of the comb rows and the base of the apical organ. Careful attention should be taken to ensure that the canals connected to the comb rows are not damaged. Tentacle bulb amputation requires cutting tissue between two adjacent comb rows, cutting out one or both tentacle bulbs on either side of the body and leaving behind 4 of the 8 comb rows. Make sure the tentacle bulbs are completely removed, including the dense cluster of cells at the base of the bulb


#### 3 Methods

#### 3.1 Sources and Collection We collect M. leidyi on the northeast coast of Florida, around the Saint Augustine area where the University of Florida's Whitney Lab for Marine Bioscience is located. The confluence of the Matanzas intercoastal river with the Atlantic Ocean creates a system of estuarine saline waters which favors the appearance of M. leidyi specimens all year long. M. leidyi can also be found in coastal waters along the Atlantic coast of North and South America and it has become an invasive species in European waters through ballast water introduction, most notably in the Black Sea, eastern Mediterranean, and Caspian Sea [30]. Collecting ctenophores is an art,

relatively easy to find them.

1. Choose optimal environmental conditions for ctenophore collection (see Notes 3 and 4).

but once an experienced collector has seen several in the wild it is


Different culturing systems have been optimized for each M. leidyi life cycle stage. Adult specimens prefer to move vertically on an hourly regime, so keeping them in a large volume, with a tall height aspect to cross sectional area is optimal. If adult animals are allowed to interact with the bottom surface of their container, they will erode their epidermal surface and die. For example, M. leidyi can be kept in 20 L buckets for short periods of time as long as the water is changed once or twice daily, but after several days animals will start to deteriorate. The ideal situation is to keep them in a tank installed in open sea water system room at ambient temperatures that allows continual sea water circulation keeping adults off tank surfaces. Hatchlings and juvenile cydippid stages can be grown and cultured in the laboratory (room temperature, 20–22 C) in a variety of different types of glassware, but low-density cultures (~1 embryo/ 5 mL) are preferred. Generally, we dilute embryonic cultures to 2 larger volumes every 2–3 days to give them more space to set their tentacles and feed (see Subheading 3.4).

We keep both M. leidyi and the atentaculate Beroe ovata in pseudo-kreisel tanks made of clear 2 cm thick plexiglass sheets connected with stainless steel screws and sealed with silicone, that generate a circular flow which keeps animals suspended in the water column. In the pseudo-kreisel depicted (Fig. 2C), the inner tank module where the animals are kept is 90 cm in diameter and has rounded corners with small perforations allowing water to exchange into two outer compartments (10 cm each) that have drains both at the top (that also serve as water level overflows) and the bottom of the tank. Upwelling inlets at the bottom of the inner tank introduce fresh sea water and direct it to the edges. By controlling the volume of water entering and exiting the tank, the overall position of the ctenophores can be controlled. For example, if the flow rate is too high, all of the animals are concentrated in the center of the tank and if it is too low the animals may sink to the bottom. The optimal flow rate allows animals to swim freely but prevents them from approaching the bottom because abrasion of

#### 3.2 Laboratory Setup for Mnemiopsis leidyi Culture

ctenophore epidermis on the bottom of the tank causes wounds that lead to death. Incoming seawater at Whitney is naturally sand filtered and devoid of zooplankton or phytoplankton.

Water quality is important for the successful culturing of ctenophore embryos and adults. We routinely rear embryos and early hatched cydippids in glass bowls (Fig. 3) filled with 1- UV-FSW until they reach ~1.5–2 mm diameter (1 week old). They are then transferred to 2 L glass beakers in order to give them more vertical space for swimming and feeding and are kept in this container until they transition into the lobate state (Fig. 3) (see Subheading 3.4).

3.3 Culturing Live Feed for Mnemiopsis leidyi Husbandry Juvenile and adult ctenophores feed on zooplankton present in the water column. Adults typically feed on copepods and other pelagic organisms (including larval fish). In captivity, the best first food source for M. leidyi is rotifers (e.g., Brachionus plicatilis) because of their small size and the ability of M. leidyi cydippids to catch them with their tentacles. We feed adults with Artemia and/or mysid shrimp daily. Note that feeding Mnemiopsis on Artemia is not sufficient to maintain reproductive ability, so mysids must be fed at least 1–2 times per week and ideally daily. Other labs use fish eggs and larvae rather than mysids. This nutritional requirement is currently a bottleneck in the rearing of reproductive colonies of Mnemiopsis in laboratory culture (see Notes 8–10).

A small-scale rotifer culture is sufficient to cover the feeding regimes of growing cydippids. We follow the instructions provided by the supplier (Reed Mariculture, Inc) except that we keep a smaller culture volume (see Note 11).


We use just-hatched Artemia to complement the diet of growing cydippids once they reach a certain size (Subheading 3.4) as well as to feed adult ctenophores.


We use mysid shrimps to feed adult ctenophores (see Subheading 3.4). We obtain the mysids directly from a shrimp farm and feed adult specimens with ~2 mysid shrimps per ctenophore twice a week (see Note 13).

#### 3.4 Spawning and Husbandry of Mnemiopsis leidyi

M. leidyi has a natural circadian rhythm and spawns according to the light-dark cycle. Our protocol for M. leidyi spawning at the Whitney Lab in St. Augustine, FL. has been modified from Pang and Martindale, 2008 [31]. Under normal summer conditions in Cape Cod, MA (Woods Hole), M. leidyi spawning is triggered by the onset of darkness and it normally occurs ~8 h after sunset. In northeast Florida, spawning occurs after 3–4 h of darkness. In order to get M. leidyi to spawn at any time of the day, we keep animals under constant light conditions and then place them in the dark to induce spawning. It takes 2–3 days of constant light exposure to erase the endogenous circadian rhythm of wild caught animals so they reliably spawn 3–4 h after putting them in the dark. When spawning freshly caught specimens, wild caught adult ctenophores are kept in a 20 L bucket filled with 1- UV-FSW in the laboratory under constant light for 48–72 h (more detailed protocol in [32]). Here we describe the protocol to spawn adult ctenophores cultured in captivity at the lab. All steps are performed at room temperature (20–22 C).


3.5 Animal Surgeries to Study Wound Healing and Whole-Body Regeneration

Although lobate stage adults have a high capacity to regenerate, we utilize M. leidyi cydippid stages due to their smaller size, speed of complete regeneration, and ease of visualization [5].


Follow the steps described below according to the type of operation. All operations are performed at room temperature (20–22 C).

Puncture assay (Fig. 4B):

3. Place one cydippid in a small drop of water on a siliconized microscope slide.


Whole-body regeneration studies (WBR):

7. Transfer cydippids using a plastic or glass pipette larger than the diameter of the specimen to an operating dish filled with just enough 1- 0.2-μm UV-FSW to cover the specimens (see Note 35).

We use three types of operations to recover all the structures/cell types of the cydippid's body.


3.6 Cell Proliferation Inhibitor Treatment with Hydroxyurea (HU) The role of cell proliferation in replacement of missing cell types was first proposed by TH Morgan more than 100 years ago [34]. Cell proliferation inhibitor experiments are a straightforward way to evaluate the requirement of cell proliferation in regeneration. We expose amputated cydippids to hydroxyurea (HU) treatments, a drug that inhibits cell proliferation by inhibiting the ribonucleotide reductase enzyme and thereby arresting cells in S-phase [35].


3.7 Fixation of Mnemiopsis leidyi Cydippids The gelatinous body of M. leidyi cydippids is mostly composed of mesoglea with varying osmotic proprieties, which makes standard fixation protocols challenging; standard fixative preparations in direct contact with the cydippid's body generate osmotic changes that cause the structural mesoglea to collapse and tissue to disintegrate. To preserve both the cellular and gross anatomic integrity, we use a fixation protocol based on embedding of specimens in a low melting point agarose [32].


3.8 Live Imaging During Wound Healing and Regeneration The remarkable optical clarity and small size of M. leidyi cydippids make them an ideal system for live-imaging experiments. Here we describe a combination of differential interference contrast (DIC) live-imaging and time-lapse techniques to monitor wound healing. M. leidyi wound healing involves cell migration and formation of actin-based cellular protrusions, resulting in a scar-less wound epithelium [10]. Under normal conditions wound healing is completed in around 30 min to 1 h after injury, depending on the size of the cut.


#### 4 Notes

	- (a) We use Brachionus plicatilis (L-type) rotifers (Reed Mariculture, Campbell, CA, USA).
	- (b) We recommend growing two asynchronous rotifer cultures at the same time and alternate harvesting between them. Excessive repeated harvesting (>50%) in one single culture could lead to the crash of the culture.
	- (c) In order to maintain the productivity, it is important to not let the rotifers run out of food. Maintain a detectable light green tint in the water between feedings (https:// reedmariculture.com/support\_rotifers.php).
	- (d) It is recommended to feed rotifers every day. If this is not possible, add the volume of food required for the days they will not be fed (ideally not more than 4). The accumulation of debris at the bottom could be a sign of overfeeding.
	- (e) The stability of the rotifer culture is based on finding the right balance between harvesting and feeding. In a healthy culture all or the majority of the rotifers will be females and will reproduce clonally. An increase in the proportion of males (smaller individuals) in the culture is indicative of an unbalanced and stressed culture.
	- (f) In case the rotifer culture cannot be maintained/harvested for longer than 4 days, there is a way to put rotifers in "hibernation mode" by setting up a backup culture. Harvest ~40% of your culture and transfer it into a 1 L container filled with 20 ppt FSW. Add a bit of extra algae concentrate to darken the culture. Let the culture uncovered or cover loosely to allow oxygen to enter and keep it at 4 C to slower the metabolism of rotifers. After 7 days 50% of your rotifers should be alive.

and kept free of detergents or any other harmful chemicals. Culturing materials should be cleaned exclusively with tap water followed by a final rinse with distilled water.


with methyl cellulose can suffice. For longer periods synthetic hydrogels [10] may be of use. We find 7.5% hydrogel as described the optimal concentration for our wound-healing time-lapse experiments, in terms of keeping a good balance between osmolarity of the medium and immobilization of the specimen. However, we recommend trying different hydrogel concentrations depending on the mobility of the animal and the length of the experiment. Lower hydrogel concentrations are more osmotically compatible with the animal but they also are less effective for immobilization while higher hydrogel concentrations allow for better immobilization but tend to dehydrate the specimens. Optimize this tradeoff for specific experiments.


#### References


evolution and development. Cold Spring Harb Protoc 3(11):1–11. https://doi.org/10. 1101/pdb.emo106

3. Chun C (1880) Fauna und flora des golfes von Neapel: Die ctenophoren des golfes von Neapel und der angrenzenden meeres-abschnitte. W. Engelmann, Leipzig. https://doi.org/10. 5962/bhl.title.10162


ctenophore Mnemiopsis leidyi. EvoDevo 5(1):4. https://doi.org/10.1186/2041- 9139-5-4


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Studying Placozoa WBR in the Simplest Metazoan Animal, Trichoplax adhaerens

Hans-Ju¨rgen Osigus , Michael Eitel , Karolin Horn, Kai Kamm , Jennifer Kosubek-Langer, Moritz Jonathan Schmidt , Heike Hadrys, and Bernd Schierwater

#### Abstract

Placozoans are a promising model system to study fundamental regeneration processes in a morphologically and genetically very simple animal. We here provide a brief introduction to the enigmatic Placozoa and summarize the state of the art of animal handling and experimental manipulation possibilities.

Key words Placozoa, Trichoplax adhaerens, Regeneration, Vital Staining, Transplantation

#### 1 Introduction

The phylum Placozoa [1] comprises flat (approx. 20–30 μm in height) discoid animals with a body size commonly less than 4 mm in diameter [2–5] (Fig. 1). One recently described species, Polyplacotoma mediterranea, can reach a size of up to 10 mm by adopting a highly ramified and highly flexible body shape [6]. In contrast, specimens of the other described species, Trichoplax adhaerens [2] and Hoilungia hongkongensis [7], as well as all other undescribed species never grow larger than 3–4 mm in diameter. The sandwich-like body of placozoans lacks any kind of symmetry but possesses a clear top-bottom polarity (Fig. 2a) [4, 8, 9]. The upper epithelium is facing the water column while the lower epithelium adheres to the substrate [2, 3, 10]. Upside-down flipped animals rotate and bring their lower epithelium back into contact with the surface quickly. Flipped animals perform this rotation by beating with the cilia of the upper epithelium. During this phase, the lower epithelium glides along itself until it regains contact with the ground [2, 3].

The three-layered placozoan bauplan consists of at least nine differentiated somatic cell types: upper and lower epithelial cells,

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_6, © The Author(s) 2022

Fig. 1 Different vital stages of placozoans. (a) Light microscopy image of the morphologically most simple metazoan animal, the placozoan Trichoplax adhaerens ("Grell clone"). (b) Degenerative stage of Trichoplax adhaerens ("Grell clone"). Under unfavorable conditions the upper epithelium of the animal lifts up and forms a hollow bubble (red arrow). In most cases the specimen will die shortly thereafter. (c) Degenerative stage of Trichoplax sp. H2 ("Vieste clone"). These thread-shaped stages can be frequently found in old placozoan cultures and are likely caused to some extend by unfavorable water chemistry

fiber cells, sphere cells, three types of gland cells, lipophil cells, and crystal cells [2, 3, 11–13] (Fig. 2a). Noteworthy, multiple other somatic (sub)cell types are awaiting their description [13, 14]. In addition to the so far identified differentiated somatic cell types, pluri- or omnipotent stem cells are found near the contact zone between lower and upper epithelium [15, 16]. The fiber cell layer, which is sandwiched between the upper and lower epithelium, plays a major role, for instance, in animal body contraction [3, 17]. The inter-connected fiber cells are the contractile elements and therefore also play a major role in animal locomotion. Lipophil cells secrete enzymes for extracellular digestion (Fig. 2b) and are exclusively found in the lower epithelium [11]. The lower epithelium also harbors three different types of gland cells (type 1, 2 and 3), which synthesize neuropeptides or mucus (in case of type 2 gland cells) [12]. Given that gland cells of type 1 and 3 possess a cilium, they have been suggested to be secretory sensory cells [12]. It is worth mentioning that type 3 gland cells can also be found in the upper epithelium [12]. The recently described sphere cells in the upper epithelium [13] include the shiny spheres, which might play a role in predator defense [18]. Finally, the so-called crystal cells are located at the margin of the animal body [11] and serve functions in gravity perception [19]. Besides morphological studies, a single-cell RNAseq study has indicated the existence of even more somatic cell types in placozoans [14].

A shared feature of all placozoans is the exceptionally high degree of body plasticity due to the absence of any kind of skeleton or other solid body parts [2, 4, 5]. Placozoans constantly change shape by contracting and relaxing their flat body which causes locomotion. Another mode of locomotion is mediated by ciliary beats of the lower epithelial cells, which is not accompanied, however, by shape changes [2]. It has recently been suggested that the

Fig. 2 Ultrastructure and feeding behavior of placozoans. (a) Schematic cross section of Trichoplax adhaerens. The typical placozoan bauplan consists of an upper epithelium, a lower epithelium and a fiber cell layer between both epithelia. The shown schematic bauplan and cell types are a synthesis of recent studies on the placozoan ultrastructure [11–13]. Please note that the actual number of placozoan somatic cell types is likely even higher and that each respective major cell type might summarize multiple sub-cell types. (b) Illustration of the typical placozoan feeding behavior. The animal glides over a food particle (green) to form an external digestion cavity. Digestive enzymes (yellow) are secreted into this extracorporeal feeding cavity and nutrition uptake is performed by the lower epithelium by means of phago- and likely pinocytosis. Panels a and b are modified after [40]

amino acid glycine induces fiber cell contractions and also the activation of ciliated locomotion in Trichoplax adhaerens [20]. The locomotion activity of placozoans mirrors the vitality of the animals and is correlated to food availability and food uptake [21–24]. Locomotion pauses for different reasons, for example, during the phase of extracellular digestion of food particles [21–24]. This extracellular digestion (Fig. 2b) is carried out by release of secreted enzymes of so far unknown composition (for further details, see [23]). Although the cells involved and the underlying physiological mechanisms are yet unknown, placozoans are able to perceive light [25].

Placozoans show exceptional regeneration capacities. For instance, partial mechanical disruptions will usually heal very rapidly within minutes [26] (see Note 1). This high regeneration capacity relates to the simple body architecture, which lacks any kind of organs or other complex morphological structures, even a basal membrane and a complex extracellular matrix are missing [17]. The high regenerative capacity of the animals allows placozoans to reproduce very efficiently by binary fission, which is the dominant mode of reproduction under laboratory conditions [2, 3, 5] (see Note 2). For this type of vegetative reproduction, animals constrict in the center region of the body to form two daughter individuals of approximately the same size. The plane of each fission is orthogonal to the previous division plane [27] and a certain ratio of inner vs. marginal cells is required to trigger fission [26]. Daughter individuals stay connected for several hours by a cellular thread, which mechanically breaks when the daughter individuals move in different directions at the very end of the fission process [2, 3]. The resulting wounds heal rapidly and the original wound borders cannot be traced back under the light microscope 30 min after regeneration. The critical step of the wound healing process of both the outer margin as well as the central region of the animal body is the contraction of the adjacent epithelial cells which brings the cells within the cell layers in close contact [26].

Comprehensive studies on whole body regeneration in Trichoplax adhaerens have been conducted by Ruthmann and Terwelp (1979) [28], and by Schwartz (1984) [26]. Both studies applied procedures outlined by Miller in 1971 [29], but without citing this original work. Miller has been a methodical pioneer for modern placozoan regeneration experiments, although some of his results contradict findings by Ruthmann and Terwelp [28] and Schwartz [26]. Ruthmann and Terwelp [28] tested different chemicals (using for example colchicine or trypsin–EDTA-supplemented calcium/magnesium-free ASW, see Note 3) to dissociate the placozoan body. Succeeding dissociation of animals into small cell clusters or even single cells, their reaggregation was studied after bringing the cells into close proximity via gentle centrifugation.

Studies from Kuhl and Kuhl (1963, 1966) [30, 31] reported first observations on the healing process of animals after cutting off a small piece from the animal body. The second key study on placozoan regeneration by Schwartz (1984) [26] focused on mechanical manipulation of the animal body. Schwartz studied the regeneration capacities of placozoan specimens after selective removal of marginal and/or center cells in a quantitative fashion and identified an approximately 20–25 μm thick circumferential marginal zone of presumably particular relevance for regeneration processes. This morphologically cryptic (i.e., macroscopically indistinguishable) zone consists of specific cell types, which cannot be found in the center of the animal. The differential cell type distribution from the edge to the center of the animal implies that cell type populations from both areas are needed to allow full recovery of animals after mechanical disruption. Schwartz also conducted a series of transplantation experiments (see Note 4). For example, parts of the marginal zone from a donor animal were transplanted into the central body region of an acceptor animal (Fig. 3). An

Fig. 3 Tissue transplantation in Trichoplax adhaerens. (a) From the "donor" animal, which has been stained with methylene blue, a piece consisting of marginal as well as center cells (lower left) has been cut off with an acupuncture needle. (b) The "acceptor" animal (unstained) has been prepared for the transplantation by means of cutting a hole into the center of its body. (c) The "donor" fragment is placed into the body hole of the "acceptor" animal. (d) Complete intergrowth of the "donor" fragment and the "acceptor" animal (18 h posttransplantation). Please note that the marginal cells of the "donor" fragment keep their state and form a new small margin in the central body region of the "acceptor" animal. (Picture taken from [33])

important result from these experiments was the observation that cells from the animal margin do not de- or redifferentiate when transplanted into the center of the animal. Successfully integrated marginal body fragments led to a hole in the central part of the body, resembling a doughnut-like appearance, with marginal cells lining the central hole. Eventually, such holes were shifted (by an unknown mechanism of active cell movement) toward the outer margin of the animal where the transplanted and the original marginal zones fused [26]. Due to the microscopic size of placozoans, only body parts comprising all three cell layers have so far been successfully transplanted into an acceptor animal. Body parts consisting just of the upper or lower epithelium alone, respectively, have not yet been successfully isolated and transplanted. Schwartz [26] also showed that even small animal fragments keep their top-bottom polarity after excision, that is, that these fragments tightly reattach to the substrate by means of the lower epithelium.

The following protocols are based on previously published animal handling and manipulation techniques and may serve as an updated basic guideline to perform whole-body regeneration experiments in placozoans.

#### 2 Materials


Filter at 4 μm, adjust to pH 7.8 and leave at least 24 h to settle (see Note 5). Autoclave if needed for downstream application. Store at room temperature.


#### 3 Methods

Placozoan specimens are extremely fragile which is highly relevant when working with live animals. The standard procedure for transferring placozoans from one place (e.g., culture dish) to another (e.g., microscopy slide) is to first detach the animal by means of a focused water jet gently pipetted from the side of the animal (see Note 7). Afterward, floating specimens can be pipetted into the designated place. Sometimes placozoans adhere very strongly to their substrate (the name giving feature for Trichoplax adhaerens, the "sticky hairy plate" [2, 3]), and cannot be detached without body disruptions. Another, even more disruptive, threat to the fragile animal body is the exposure to air—attached placozoans that fall dry for 1 s only get irreversibly disrupted. Therefore, the regular exchange of artificial seawater (ASW) in the culture dishes must be performed as rapidly and as cautiously as possible and an uninterrupted liquid film has to constantly cover the placozoans. Direct contact of placozoans to large underwater air bubbles, for example, as generated by algae in the culture dishes, is not harmful to the animals. Obviously, it is the rapid change of surface tension, and not the air itself, that is disruptive.

	- 1. Fill a glass petri dish (see Notes 8 and 9) 2/3 with ASW.
	- 2. Add 1 mL of food source (see Notes 10 and 11).
	- 3. Cover the petri dish to minimize evaporation.
	- 4. Detach placozoans from the wall of the shipment vessel (e.g., a 50 mL tube) by gently pipetting a focused water jet on their side (see Note 7).
	- 5. Transfer floating placozoans into the petri dish by pipetting.
	- 6. Store the culture at room temperature (ideally at 23 C, tolerable range from 20 C to 25 C) under a 12:12 h light–dark cycle (see Note 12).
	- 7. Every two weeks replace half of the ASW with fresh ASW to prevent the accumulation of waste products.
	- 8. Add 1 mL of food source after replacement of ASW.
	- 9. Repeat steps 7 and 8 until the animal density reaches 300 specimens per dish (see Notes 13 and 14).
	- 10. Follow step 4 to detach 50 animals.
	- 11. Pour 50% of the volume (including detached animals) to a new petri dish.
	- 12. Add fresh ASW to the original and the new culture dishes.
	- 1. Pipet 30 μL of the membrane dye stock solution onto a depression microscope slide.
	- 2. Wait for 1 h for the solution to dry completely at room temperature.
	- 3. Detach placozoans from their culture dish by gently pipetting a focused water jet on their side.
	- 4. Transfer the detached placozoans to the dried slide.
	- 5. Do not cover the slide since removal of the cover glass might disrupt the animals.
	- 6. Incubate at room temperature in the dark for 7 h in a wet chamber.
	- 7. Transfer stained animals into a petri dish filled with ASW to wash them.
	- 8. Proceed with mechanical manipulation experiments (see Subheading 3.4).
	- 1. Transfer placozoans into a 4 cm petri dish filled with 8 mL ASW.
	- 2. Add 200 μL MBSS to the dish for the liquid to become dark blue.
	- 3. Stain animals for 45–60 min.
	- 4. Transfer stained specimens by pipetting into a new petri dish containing fresh ASW to wash the animals.
	- 5. Proceed with mechanical manipulation experiments (see Subheading 3.4).

#### 3.4 Mechanical Manipulation The following protocol refers to the procedures described in Schwartz 1984 [26].

#### 1. Transfer live specimens into a glass petri dish prefilled 2/3 with fresh ASW to remove algae.


#### 3.5 Animal Fixation This protocol follows [35].


3.6 Animal Body Dissociation and Reaggregation

In general, the success of regeneration, with the eventual rebuilding of a vital whole animal, is highly dependent on the random aggregation of all cell types (potentially including pluri- or omnipotent stem cells). The protocol below refers to the procedures as described in Ruthmann and Terwelp [28] as well as Sebe-Pedros et al. [14].


#### 4 Notes


#### References


214(4):170–175. https://doi.org/10.1007/ s00427-004-0390-8


F.E. Schulze (Placozoa). Z. Naturforsch. C 39: 818–832


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# Collecting and Culturing Kamptozoans for Regenerative Studies

## Achim Meyer , Julia Merkel, and Bernhard Lieb

#### Abstract

Kamptozoa, also known as Entoprocta, are small aquatic filter-feeders that belong to the Lophotrochozoan superphylum, which also contains other acoelomate phyla including Annelida, Nemertea, and Mollusca. The study of Kamptozoa is thus of great interest to understand the early Lophotrochozoan evolution. Moreover, many kamptozoans have been shown to possess great regeneration capacities, including wholebody regeneration. In addition, and in particular for colonial cosmopolitan species such as Barentsia benedeni, kamptozoans are highly suitable as laboratory model organisms because of their simple culture, low space requirements, transparency and rapid life cycle. This chapter provides a brief introduction into field collection, culturing techniques for both the animals as well as the algae required for their feeding, fixation, staining, and sequencing.

Key words Entoprocta, Kamptozoa husbandry, Cryptomonas baltica, Filtration feeding

#### 1 Introduction

Kamptozoans are aquatic acoelomate filter-feeding invertebrates. These animals have a typically minute body, called zooid, ranging from 0.1 mm (Loxosomatidae) to 15 mm (Barentsiidae), composed of a stalk part containing the stolon, and a head part called calyx that contains the other organs (Fig. 1). The calyx of kamptozoans is crowned by solid tentacles, which covering cilia generate a flow of water that brings food particles into the atrium of the animal for feeding. Although most of the approximately 180 described kamptozoan species are found in the marine environment (25 Loxosoma spp., 118 Loxosomella spp., 50 species from the families Loxosomatidae, Pedicellinidae, and Barentsiidae [1, 2]), some species such as the cosmopolitan Barentsia benedeni can retreat into brackish water and two species are found exclusively in freshwater (Urnatella gracilis [3], and Loxosomatoides sirindhornae [4]). All Kamptozoa bear ovoviviparous trochophore larvae

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_7, © The Author(s) 2022

Fig. 1 Sketch of a colonial kamptozoan. A: Anus, B: Budding zooids, S: Stomach. Blue arrows indicate the aspiration current and filtration mode

which are similar to the mollusk and polychaete larvae. Additionally, all known species propagate by budding.

Kamptozoa, also currently known as Entoprocta, initial classifications (e.g., van Beneden 1845 [5]) described colonial species such as Pedicellina cernua as an ingroup of the "moss animals," or bryozoans, also currently known as Ectoprocta. Contradiction was firstly raised by Nitsche 1869 [6] pointing out some fundamental differences between these two groups of animals. One of the most readily observable difference is the location of the anus, which is position inside the atrium in Kamptozoa and outside of it in Bryozoa. Consequently, Nitsche suggested the names Entoprocta and Ectoprocta, and these names are still well accepted. However, the mode of filtration generated by cilia on the tentacles is opposed between Entoprocta and Ectoprocta, thus the name-giving position of the anus is inevitably a consequence of this divergent filtration feeding pattern. Related to the fundamentally divergent adult body plan is also the reorganization of the alimentary system during metamorphosis of the kamptozoan trochophora, when the gut becomes U-shaped, the mouth opening breaks through at a new position and the anus finally settles down within the crown of solid tentacles of the adult zooid [7]. The alternative name Kamptozoa was dropped by Cori 1936 [7] from the Greek κμπτω ¼ bent and refers to the curved stolon which is observed as reaction to external stimuli. Thus, kamptozoa describes a feature which is related to the unique behavior of the disturbed living animal that facilitates their identification both in the field as well as below the stereomicroscope. Because Kamptozoa provides a friendlier semantic for a researcher focusing on these taxa, we tend to favor the use of this more recent nomenclature.

Phylogenetically, Kamptozoa belong to the Lophotrochozoan superphylum, which contains other phyla including Annelida, Nemertea, and Mollusca. Kamptozoa are thus of great interest to understand the early Lophotrochozoan evolution. Bleidorn [8] summarizes the current knowledge about Lophotrochozoan systematics including the description of characters which define Mollusca as the sister group of Kamptozoa. The internal kamptozoan phylogeny (Fig. 2) agrees on the distinction of Solitaria (130 species) and Coloniales (50 species) [9, 10]. Many colonial species and one solitary species, Loxosomella antarctica, have been shown to be capable of whole-body regeneration [11–13]. Whole-body regeneration can take place both under natural conditions (e.g., low salinity leads to the dropping off the calyx during winter [14]) or after zooid loss due to predation of the calyx [15]).

Understanding the regenerative pattern of Kamptozoa could shed light on general developmental processes. Their pronounced regeneration capacity together with the transparent and minute body size makes Kamptozoa an ideal organism for research on whole-body regeneration using modern laboratory procedures such as advanced labeling and staining methods (Fig. 3).

In this chapter, we present protocols for the isolation, culture and feeding of the colonial species Barentsia benedeni. In addition, we provide protocols for the monitoring of WBR as well as for the fixation, staining, and DNA extraction of whole zooids. These protocols can easily be adopted for the study of other colonial species, as it works well for B. elongata, and will enable shortterm (several weeks) work with most solitary species.

Fig. 2 Phylogenetic relationships of recent kamptozoan families, modified after [8, 9]

#### 2 Material


Fig. 3 Confocal micrographs of regeneration stages of P. cernua (thesis J. Merkel 2014). Scale bars 20 μm. (a–e) lateral view. (f) Laterofrontal view. (a) First regeneration stage. (b) Second regeneration stage. (c) Third regeneration stage. (d) Fourth regeneration stage. (e, f) Fifth regeneration stage. Indicated organs are atrium (at), stomach (sto), esophagus (oes), midgut (mg), and hindgut (hg). Micrographs were gained with a Leica SP5 II confocal microscope

Fig. 4 Culturing bowl with tools for cleaning. Note the green cut-in air tubing

	- 2. Fixative solution: 4% (v/v) paraformaldehyde (PFA) in 0.1 M phosphate buffered saline. Long-term storage (>2 weeks) in plastic containers at 20 C and short term at 4 C.
	- 3. PBT: 0.2% (v/v) Triton X-100 in 0.1 M PBS. If stored at 4 C in the dark it is usable for months.
	- 4. Conjugated phalloidin: dissolve (usually comes as a powder; e.g., Alexa Fluor 488 phalloidin; Invitrogen, BODYPI R6G phalloidin, Oregon Green 514 phalloidin, BODIPY FL phallacidin) in methanol to yield a final concentration of 200 units/ mL. This stock solution is stable for at least 1 year when stored at 20 C.
	- 5. Staining solution (1:20 diluted): 50 μL conjugated phalloidin, 950 μL PBT. Store at 4 C in the dark for up to 1 week.

2.4 RNA Extraction and Target Gene Amplification


#### 3 Methods

#### 3.1 Collection and Identification There are pronounced species-specific settlement preferences such as epizoic growth on sponges, bryozoans, polychaetes, and sipunculids or attachment to macroalgae, and for a few species also inanimate surfaces such as rocks or shells. Known substrate preferences will guide the collection of animals in the field. For example, the cosmopolitan colonial freshwater Kamptozoa Urnatella gracilis prefers shells of Dreissena polymorpha in the area around Berlin [3] and the solitary kamptozoan Loxosomella murmanica live epizoically on the sipunculan Phascolion strombus which inhabits empty shells of the gastropod Turritella sp. and the scaphopod Antalis sp.. Such shells can be dredged from about 30 m depth from muddy and rocky bottom in the North Sea [18]. We here present our protocol for the collection of Barentsia benedeni colonies.




#### 3.2 Food Algae Culture

	- 1. Pour a 200 mL aliquot of the algae stock on a petri dish.
	- 2. Screen for obvious signs of contamination with a microscope.
	- 3. Discontinue contaminated cultures, start with a new stock in step 1.
	- 4. Transfer 150 mL NFSW into a 250 mL Erlenmeyer flask with a cotton plug.
	- 5. Heat up for 1 min at 600 W in a microwave. Do not let the water boil or salts will precipitate [17].
	- 6. Add 3 mL of freshly filter-sterilized 50X F/2 medium to the hot seawater (see Note 4).
	- 7. Mix gently by swirling without wetting the cotton plug.
	- 8. Wait 1 h for the medium to cool down to culturing temperature (RT or lower).
	- 9. Inoculate with up to 50 mL algae stock.
	- 10. Place the culture on an orbital shaker (120 rpm) in the light cabinet (see Note 5).
	- 11. Leave the culture to grow for 2 weeks before using it for feeding kamptozoans.
	- 12. Algae cultures can be kept up to 1 month.
	- 13. Renew the culture by starting with step 1 using the old culture as stock.
	- 14. Keep two 50 mL cultures at lower light, without shaking nor routine opening, for up to 2 months as backups.

In our culturing setup, collected colonies will not attach themselves again to the walls of the culture vessel but will become floating balllike structures.

3.3 Barentsia benedeni Culture

	- 2. Starve the colony for 1 week to fully isolate the zooid (see Note 10).
	- 3. Clean the colony and the culture vessel twice during the starvation.
	- 4. Amputate the tissue to be isolated using a fine pair of Vannas scissors.
	- 5. Transfer the colony back to a clean culture vessel, without feeding.
	- 6. Monitor the regenerating colony every 48 h.
	- 7. In the case of whole-body regeneration induced by a stalk cut, regeneration will proceed as follows (Fig. 3).
	- 8. 2 days postamputation (dpa): first regeneration stage, atrium and stomach have already formed.
	- 9. 4 dpa: second regeneration stage, the esophagus has elongated, midgut is formed, and atrium is bulged on the anal side.
	- 10. 6 dpa: third regeneration stage, developing hindgut and atrium converge.

#### 3.4 Tissue Isolation and Regeneration

3.5 Fixation and F-Actin Staining This protocol describes staining against F-actin but can be generalized to other conjugated probes by adapting the composition and incubation time of the staining medium. Thus staining with other primary and secondary antibodies could be applied to study different structures such as nuclei or keratin.


#### 3.6 Total RNA Extraction and Target Gene Amplification

Although RNA can be extracted from any piece of tissue, we obtained good yields by pooling together tissue from approximately 40 clonal zooids. The sample volume should not exceed 10% of the volume of the TRIzol used for lysis. You may wash your bench and pipettes with RNase erase solution before work. All steps at room temperature unless otherwise specified. Wear protective gloves and googles when working with liquid nitrogen or deepfrozen devices. Use a fume hood during processing of phenol. Wear disposable gloves while handling organic reagents such as TRIzol (contains phenol) and RNA samples to prevent RNase contamination from the surface of the skin; change gloves frequently, particularly as the protocol progresses from crude extracts to more purified materials.


#### 4 Notes

1. In our lab, we use commercially available FNSW which is shipped in 5 L containers. Natural seawater is mandatory for long term culture of the species, but not for the algae. For algal culture, it is thus possible to use artificial seawater instead.


Fig. 5 Simple algae culturing unit using natural light without shaker. Bottles are shaken manually once every working day because algae tend to settle on the bottom. Isochrysis sp. (left bottle) and Nannochloropsis sp. (right bottle)


#### Acknowledgments

We are grateful for the detailed and comprehensive help from Simon Blanchoud turning our first draft into the desired style and language of the final manuscript. We thank Andreas Wanninger for sharing his experience in labeling marine invertebrates with us.

#### References


of kamptozoan diversity in Australia and New Zealand. T Roy Soc South Aust 126:1–20


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 8

# Collecting and Culturing Bryozoans for Regenerative Studies

### Abigail M. Smith , Peter B. Batson , Katerina Achilleos , and Yuta Tamberg

#### Abstract

Among marine invertebrates, bryozoans are small, not well known, and complex to identify. Nevertheless, they offer unique opportunities for whole-body generation research, because of their colonial, modular mode of growth. Here, we describe detailed methods for collection of bryozoans from a range of environments, sample preparation and identification, culture and feeding, spawning and breeding, marking colonies for growth studies, and histological preparation.

Key words Bryozoans, Culture, Collection, Growth, Feeding, Whole-body regeneration, Budding, Regression, Brown bodies, Anatomy, Histology, Larvae, Spawning, Settlement

#### 1 Introduction

The Bryozoa (moss animals) is a diverse phylum of colonial aquatic invertebrates found in almost all freshwater and marine environments. The phylum comprises ~6000 living species [1] which grow into a bewildering array of colony types, including soft (weedy or gelatinous) and hard (calcified) forms, which may be moss-, sponge-, or coral-like in overall appearance. Numerous taxa grow as thin crusts or delicate lace-like encrustations over suitable substrates (Fig. 1) [2]. Although often overlooked, bryozoans are often among the most diverse and abundant members of marine communities, especially in the Southern Hemisphere. All bryozoans are suspension feeders, extracting small food particles from the water column, and colonies typically live attached to seafloor substrates (e.g., shells, rocks, algae) or on surfaces in freshwater ponds, rivers, and lakes [3]. Three of the main extant clades are the freshwater Phylactolaemata, the marine Stenolaemata, and the predominantly marine Gymnolaemata (Fig. 2). All three offer possibilities for the study of WBR and related phenomena.

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_8, © The Author(s) 2022

Fig. 1 Morphology of bryozoans. (a) An encrusting colony of the marine cheilostome bryozoan Watersipora subatra. (b) SEM image of the calcified autozooids of a Microporella discors colony (marine Cheilostomatida) showing ~6 polygonal autozooids; black arrowhead—autozooidal aperture; white arrowhead—avicularium (defensive polymorphic zooid). (c) Part of a living colony of the marine cheilostome Hippomenella vellicata showing feeding autozooids with extended lophophores (top); retracted autozooids with closed lid-like opercula (middle); and developing asexually budded autozooids at the colony margin (bottom). (d) Living colony of Hastingsia sp. This well-calcified continental shelf cyclostome was successfully grown in a laboratory culture system using natural seawater supplemented by cultured microalgae. (e) Two polypide regression products (brown bodies), indicated with white arrowheads; the adjacent zooidal chamber contains a developing polypide that will replace the previous polypide, which has degenerated (Hornera sp., marine cyclostomate, H&E stained). (f) Large colony of the gelatinous freshwater phylactolaemate Pectinatella magnifica. (g) Living colony of Cristatella mucedo, a freshwater phylactolaemate bryozoan; several rows of horseshoe-shaped lophophores line the periphery of the colony, which is capable of creeping along the

Bryozoan colonies are composed of iterated (mostly) submillimeter animals called zooids, which are budded as asexual clones from a single founder zooid, the ancestrula, itself derived from a free-swimming larva [4]. Depending on the species, a single colony may contain several to many hundreds of thousands of zooids. Autozooids are the zooid polymorphs responsible for feeding within a bryozoan colony; each has a lophophore bearing a crown of ciliated tentacles that captures microscopic food particles [5], typically microalgae. This feeding apparatus is normally extended into the water column on a flexible sheath, but can be retracted into a protective box-like or tubular zooid chamber, which may be gelatinous, leathery, or rigid in marine species that secrete a calcified skeleton [3]. The remaining parts of an autozooid include the polypide (comprising the lophophore, u-shaped unidirectional gut, a ganglion, and retractor muscles), and the cystid, the living and nonliving structural parts of the body wall (Fig. 3) [5]. Species identification of bryozoans often relies upon examination of the individual zooid architecture, and commonly requires the use of a dissecting microscope.

Zooids are physiologically interconnected via tissue strands (funiculus) which pass through pores in shared body walls, or via shared body cavities in budding zones [6, 7]. Autozooids possess variable degrees of physiological integration within the colony, while retaining a basic functional autonomy. Nonfeeding polymorphic zooids are common in marine bryozoans, and include


Fig. 2 Generalized phylogeny and relationships of the phylum Bryozoa (includes only extant taxa)

Fig. 1 (continued) substratum. (h) Statoblast (asexually produced resting propagule) of the freshwater bryozoan Plumatella cf. geimermassardi (fixed but unstained whole mount, imaged under compound microscope). Scale bars: a, 10 mm; b, 200 μm; c, 1 mm; d, 1 mm; e, 50 μm; f, 5 cm; g, 1 mm; h, 100 μm

Fig. 3 Generalized anatomy of a cheilostomate bryozoan individual zooid. Scale: zooids generally range from 0.1 to 1.0 mm in length

reproductive, defensive, and structural modules [8] that rely on autozooids for nutrition. Bryozoans may undergo seasonal sexual reproduction, while asexual budding occurs all year. Freshwater and a few marine bryozoans produce clonal resting propagules (statoblasts and hibernacula) containing stem cells [9].

This group offers unique opportunities for whole-body regeneration (WBR) research, but has been underutilized compared to other invertebrate models. Studies of WBR in this phylum could focus on zooid-scale processes in the context of the whole colony. Autozooids undergo agametic cloning to produce new zooids by budding, resulting in new colony growth, and subsequently undergo one or more polypide degeneration–regeneration cycles, which replace senescent polypides within existing zooids of the colony. The latter process occurs throughout the functional life of a zooid and involves full breakdown of the incumbent polypide into a residual "brown body," and development of a polypide replacement, which arises from a blastema on the cystid [10]. Individual polypides typically last 1–10 weeks before regression commences, and the regression phase lasts 3–20 days, depending on the species [10].

Little is known of bryozoan regenerative processes at the subzooidal scale, for example, following partial injury to a polypide, but both WBR and body organ/tissue (¼ "structure") regeneration has been reported for this phylum by Bely & Nyberg [11]. In Cristatella, at least, surgical damage to zooids is repaired rapidly without apparent damage. It is relatively easy to surgically separate living colonies into multiple ramets (clonal subcolonies), which will heal and continue to grow by budding [12]. Some bryozoans (e.g., Cupuladria exfragminis) are known to naturally autofragment as a clonal propagation strategy [13]. Bryozoans usually maintain budding along a given body axis during normal growth [14], but some taxa can undergo reversed-polarity budding and lateral budding during repair of individual zooids or during regeneration of mechanically broken branches [15]. At the colony scale, reversedpolarity budding can happen following breakage in branching forms. The precise extent of WBR in bryozoans remains to be determined.

In this chapter we present methods to study bryozoans, starting with how to find and collect bryozoans. Intertidal and shallow subtidal bryozoans can be scraped off rocks, picked from macroalgae or collected by divers, whereas shelf and deep-sea bryozoans are commonly collected via dredge or grab sampling. Preliminary on-board classification and sorting of bryozoans is achieved using overall colony form and colour, but proper species determination requires either light microscopy (difficult but can be nondestructive if done carefully) or scanning electron microscopy (lethal). Keeping living bryozoans in tanks requires careful preparation and maintenance of water quality, regular feeding of mixed phytoplankton cultures and, in some cases, regular gentle cleaning of colonies. Some encrusting bryozoans grow well in culture, but many are capable of surviving a long time without growing at all. It is possible to spawn at least some species of bryozoans, settle larvae, and raise colonies in a laboratory setting. Growth in bryozoans can be difficult to ascertain [16], but nontoxic marking of colonies using Calcein can be effective [17]. Deeper study into the structure of soft-parts (histology) and hard parts (micro-CT, SEM) allows for evaluation of life-cycle and effect of experimentation. In this contribution we provide the basic techniques for bryozoan collection, culture, and maintenance.

#### 2 Materials

Not all materials presented in this section are required for every study. Researchers will need to choose the required materials based on their project specifications.

	- 2. Airtight container(s) with lids, 1–3 L capacity.
	- 3. Frozen cold-packs.
	- 4. Thin wet protective layer (e.g., fresh seaweed, damp blotting paper).

#### 2.2 Identification of Bryozoans


#### 2.3 Culture and Feeding


#### 2.4 Marking 1. Living calcified bryozoans.


#### 2.5 Anesthetizing and Fixing for Histology

Solutions should be prepared using ultrapure water and analytical grade reagents. Solutions should be stored at room temperature unless otherwise stated. Chemicals used in this section, with a single exception of MgCl2, are toxic. Use fume hood for preparation and store tightly stoppered.


#### 3 Methods

#### 3.1 Onshore Collection

Bryozoan colonies are very diverse in terms of external appearance (Fig. 1). Encrusting bryozoans are generally small roundish patches, 1–3 cm in diameter, and variable in colour. When poked, they are predominantly hard to the touch, but some species are filamentous, weedy or gelatinous. It is easy to confuse these bryozoans with patches of algae—examination with a hand lens will show the regular openings (like pinpricks) on the surface of bryozoans, whereas most algae are smooth. Bryozoans prefer undersides of hard substrate, while algae need sunlight. Erect bryozoans tend to occur in deeper waters, and come in a multitude of shapes: fans, nets, fingers, trees, feathers. They can be confused with corals, macroalgae. Again, if they are hard/rigid and covered with small openings, found on undersides and overhangs, they are likely to be bryozoans.


#### 3.2 Offshore Collection


#### 3.3 Postcollection Sorting


#### 3.4 Identification Bryozoans are difficult to identify in the field and in hand specimen. There are many species which are superficially similar. Acquiring a bryozoan expert for confirming IDs is an excellent plan. If you need to send images for ID, scanning electron microscope images are best. Methodical comparison of SEM photos with species descriptions in specialist literature is, unfortunately, the only reliable way to identify bryozoans.



	- 2. Keep colonies in seawater, undisturbed at a constant temperature in the dark for at least 24 h prior to spawning.
	- 3. Induce spawning in marine bryozoans by sudden exposure to bright light; freshwater species will spawn overnight, but must be watched as the larvae can be very short-lived. The time until larval release can vary between 5 and 60 min depending on the species [19–25] (see Note 29).
	- 4. Collect larvae by gentle pipetting and transfer to prepared aquarium set-up or experiment (see Note 30). Provide many different substrate options (see Note 28), as they may settle fairly randomly when competent.
	- 1. Fill tank (leaving headspace) with 8 L Calcein in seawater solution (see Note 1).
	- 2. Adjust the water temperature to the one of the tank where bryozoans have been living.
	- 3. Add bubbler for aeration and circulation.

Fig. 4 Tank-based culture systems for bryozoans. (a) Short-term culture system using individual tanks. (b) Flow-through tank system. (c) Slow flow-through system. (d) Recirculating system. The option of a semirecirculating system is also shown with a red arrow. BF biological filter, F filter sock, H/C heating/ cooling system, L light source, MF mechanical filter, Phyt. R phytoplankton room/area, pH pH meter, P pump. Colors: red ¼ aeration system, blue ¼ sea water inflow, brown: sea water outflow or waste water, yellow and light blue: water purification steps, green: phytoplankton as food


The process of relaxing the colony in such a way that the lophophores can be fixed in a protruded position needs to be controlled under the stereomicroscope, with minimal disturbance to the colony. A typical relaxation interval should not exceed 2–2.5 h.


3.8 Anesthetizing and Fixing for Histology


#### 4 Notes


with shaking. Overheating can be avoided with the use of cold packs and thermo-isolating boxes. Over short travel times, oxygen depletion is not significant if the density of animals is low, so we recommend filling the containers almost to the brim with water. This makes the colonies more resistant to shaking and mechanical damage. For long travel times, however, leave some air above the water and take extra care not to shake the containers.


natural environment, then transfer the developing colonies, along with their substrates, to a culture system. If research objectives require tightly controlled conditions, or the absence of contaminating organisms, a closed aquarium system will be required. However, provision of artificial or sterilized natural seawater and a cultured microalgal diet adds a significant level of time investment to a project. Furthermore, some bryozoans have been shown to develop different and often-unusual colony morphology when kept in highly controlled laboratory conditions and fed microalgal monocultures [26].


sophisticated. Tanks and other components in contact with the water should be nonmetallic and should not contain natural rubber, as both can be toxic to bryozoans. It is good practice to "condition" immersed system parts, especially new plastics, by placing them under running (sea)water for 24 h; this removes soluble residues and encourages the establishment of biofilms. Recirculating, semirecirculating and flow-through tanks can be used for bryozoan culture. These options have different advantages and drawbacks. Flow-through tanks are preferable if using natural seawater as the food source. Cultured food, however, can also be used in the case of a slow flow-through system. Recirculating or semirecirculating systems are ideal if precise control of water quality conditions is needed. Depending on the objectives, a very simple system can be effective: for example, plastic buckets can be used as tanks, and manual water changes every ~24 h can work for short-term studies. A good general principle for culturing bryozoans is to replicate natural conditions as much as possible. Many marine and freshwater bryozoans prefer low-light levels and shaded environments, growing best on underhanging substrates. If culturing is taking place in a well-lit room, a light cover over tanks should be considered. Note that exposure to high-intensity light can induce spawning in some species (see Subheading 3.4). Water movement is another consideration. Linear or oscillating currents can be generated using various methods, including submersible pumps, wavemakers, aerators and mechanical stirrers. Some species grow well in still water, although a small amount of water movement is beneficial to ensure that food particles do not sink to the bottom. In our experience care should be taken to ensure microbubbles are not introduced into the system during aeration, as these can adhere to colonies, disrupting normal function. Similarly, high current speeds can be damaging. For many bryozoan species, management of water temperature is important. Nearshore marine and freshwater bryozoans tend to be eurythermal. For stenothermal taxa, a responsive temperature controller heating and/or cooling system should be used. Tanks can also be housed in a controlledtemperature environment.

25. Bryozoan colonies can be fed simply by the provision of natural seawater in flow-through systems, or by adding cultured food (usually microalgae) to tanks. For freshwater culturing, Wood [18] describes use of a closed mixed culture system using a tank containing well-fed goldfish and a rich microbiota; food-rich water from this reservoir is supplied to bryozoan culture tanks via an airlift pump. Marine bryozoan culture requires cultured microalgae, using commercially obtained strains developed for the aquaculture industry. Conveniently their nutritional content is usually well-documented. Many marine laboratories maintain a dedicated phytoplankton culture room, and requests for culture of specific strains should be made 1–2 months in advance to allow time for culturing. Common microalgal genera used for feeding bryozoans include Dunaliella, Rhodomonas, Tetraselmis, and Pavlova. It is important to supply appropriate cell concentrations of the cultured microalgae to the bryozoan tank, taking into account the dilution volume of the tank itself. Feeding can be done by manual daily additions of cultured cells, or ideally, by a drip feed system, which can be applied both to closed and flow-through systems. Microalgal monocultures are commonly used for experimental studies of growth and feeding, but mixed microalgal diets may be more appropriate for some studies. It should be noted that abrupt changes in diet from one monoculture to another is reported to induce colony-wide polypide regeneration in some taxa [15]. Optimal cell concentrations can be found in the literature for some commonly studied species, such as Electra pilosa [33] or Cryptosula pallasiana [27]. To calculate the volume of cultured algae required for feeding, cell counts can be made using a cavity slide or an automated cell counter.


alternative is to employ gentle puffs from a disposable plastic pipette that has had the tip cut off to enlarge the opening. In both cases great care should be taken to avoid damaging the colonies, and it is prudent to do some "test cleans," followed by examination of colonies several hours later to ensure they are undamaged (e.g., feeding normally). Some bryozoans, such as free-walled cyclostomes, are prone to having their membranous body walls scraped off, and individual brush hairs can easily enter zooidal tubes, damaging terminal membranes and polypides. Water filtration can reduce the buildup of mobile detritus in a closed or semirecirculating system; however, doing so also removes food particles. One method is to run a timer-activated power filter once a day for a short period (~1 h) before feeding or turn off the water filtration system for a short period of time (~2 h) while feeding.


a microscope camera, either by video recordings or photo time frames [36]. Settlement intervals vary, but planktotrophic cyphonautes larvae produced by the marine ctenostome species Amathia gracilis may swim up to 10 h before settlement [37].


gradually, or else place small menthol crystals on the water surface of the culture container. It is important to keep the container covered, because menthol is an irritant and evaporates easily. The lid should be transparent to allow microscopic observation and cover the container in such a way as to be removed without shaking the colony. A large upturned Petri dish is usually good for this purpose. Use of a benchtop extraction hood during relaxation with menthol is recommended. The same procedure of gradually adding the relaxant applies for chloral hydrate solution. Chloral hydrate: 20 g C2H3Cl3O2 in 1 L H2O, prepared 1 month before use to saturate properly.

35. 4% Formalin and Bouin's solution (150 mL filtered saturated solution of picric acid, 50 mL formalin, 10 mL glacial acetic acid, use within a few days of preparing) both work well for fixation of both marine and freshwater bryozoans for paraffin based histological sectioning. Samples can be stored in formalin for several months, but prolonged storage in Bouin's solution is not recommended, as it will dissolve skeletal carbonate. Histological handbooks such as [39] provide more background and details on specific fixatives and processing methods. Formalin-fixed material can also be processed for aceto-orcein staining. For immunocytochemical studies with the use of confocal laser scanning microscope, Triton X-100 is a common permeabilization agent, with goat or bovine serum albumin as common blocking solutions. Primary antibodies successfully used with bryozoans include rabbit anti-serotonin and mouse anti-acetylated α-tubulin; secondary antibodies include goat and donkey anti-rabbit as well as goat and donkey anti-mouse (e.g., [40]). For SEM examination of larvae, use 2.5% GA fixative, osmolarity-adjusted and buffered with 0.2 M Milloning's phosphate buffer (pH 7.4) for 1 h at 20 C. Freshwater and ctenostome bryozoans may require stronger concentrations. Postfix the animals in 2% osmium tetroxide and 1.25% sodium bicarbonate (pH ¼ 7.2 with 1 N HCL immediately before use) at 20 C, for 1–2 h, then follow standard protocols for rinsing, dehydration, critical-point drying and coating [37]. Fixation and processing for visualization of ovaries, oocytes and nuclei requires some very specialized stains. For DNA-specific fluorochrome Bisbenzimide H333342 use samples fixed with buffered 4% formalin, stain with Bisbenzimide H333342 (10 pg/mL) for 5 min or more at room temperature. Rinse three times with filtered seawater [20]. The specimens can be stored at 4 C until imaging. For aceto-orcein staining, specimens can be fixed either directly with 3:1 methanol–acetic acid for 30 min, or in two stages. For the latter method, first fix with 4% formalin with 0.2 M PBS for 20 min and rinse thoroughly with any phosphate buffer, then postfix with 3:1 methanol–acetic acid for 60 min. Use 45% solution of aceto-orcein to stain the samples for 30 min. Rinse with 20% acetic acid [20, 34].

36. Most marine bryozoan taxa are biomineralized, with skeletons comprising CaCO3 in the form of calcite and/or aragonite. For histological and EM sectioning purposes, the skeleton must be fully removed. Failure to properly decalcify can lead to wholesale destruction of the expensive diamond knives used for ultramicrotomy. Decalcification can have harsh effects on delicate bryozoan tissues, and lead to mechanical damage and extraction of cell contents, so gentler, longer-period protocols generally work best for heavily calcified taxa if excellent-quality sections are required. Immersion of samples in decalcification solution in the absence of CaCO3 is especially damaging to tissues, so close monitoring of progress is recommended. Typically, a weak acid or a chelating agent is used to remove the skeleton. Popular acids for decalcification include ascorbic acid and formic acid, and these are used in diluted form (~2–4%), in a seawater-isotonic solution; this process can take several weeks and requires regular changes of solution. Calcium chelation is a highly regarded method, especially for TEM, and the preferred agent is EDTA (in 5–20% range). As decalcification protocols are required for marine bryozoans only, decalcifying solutions should done in a seawater isosmotic environment, ideally similar to that used for fixation and washing stages, so buffers should be used during this procedure, for example, PBS/cacodylate buffer. An osmometer is useful, and the osmolarity of local seawater should be used as a target. In most cases decalcification will be undertaken after the initial fixation, but can be done either before or after postfixation, in the case of TEM. Following decalcification, samples should be washed several times in seawater-isotonic buffer. As a general rule, further processing of the decalcified material is best done by hand, rather than with a tissue processor. For the most delicate specimens, embedding in low- or ultralow temperature agarose before decalcification dramatically improves overall sample integrity.

#### References


DP (ed) New Zealand inventory of biodiversity, vol 1. Canterbury University Press, Christchurch, NZ, pp 271–279


Island sound ascidians and bryozoans. Connecticut Department of Environmental Protection, Sea Grant Connecticut, Groton, CT


nervous system function. Preprint. https://doi. org/10.1101/869792


transfer from paternal to maternal individuals of Membranipora membranacea. Biol Bull 206: 35–45


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Studying Annelida Regeneration in a Novel Model Organism: The Freshwater Aeolosoma viride

# Chiao-Ping Chen, Sheridan Ke-Wing Fok, Cheng-Yi Chen, Fei-Man Hsu, Yu-Wen Hsieh, and Jiun-Hong Chen

#### Abstract

Aeolosoma viride, a globally distributed freshwater annelid, has a semitransparent appearance with 10 to 12 segments, about 2 to 3 mm in length. It is easy to raise and handle in laboratory conditions. Due to its robust regenerative capacity and applicability of various molecular tools including EdU labeling, wholemount in situ hybridization (WISH), and RNA interference (RNAi), it rises as a promising model for studying whole-body regeneration.

Key words Annelid, Aeolosoma viride, Asexual reproduction, EdU labeling, Whole-mount in situ hybridization, RNA interference

#### 1 Introduction

Earthworms are known form their regenerative capabilities [1, 2]. However, after testing several earthworm species in Taiwan, we could not find a robust model for studying annelid regeneration. The possibility of interrogating annelid regeneration came true with the presence of Aeolosoma viride. The discovery of this worm was an unexpected event when we collected Daphnia sp. from water ponds at National Taiwan University. Since these annelids exhibit asexual fission, and regeneration is recognized as one type of asexual reproduction [3], we inferred that this annelid may have regenerative abilities, which, to our knowledge, was not systematically documented previously. We tested its regenerative capability by amputating at the foregut-midgut or midgut-hindgut junctions (Fig. 1). Surprisingly, we found that this annelid can regrow anterior and posterior segments within 1 week.

Chiao-Ping Chen and Sheridan Ke-Wing Fok have contributed equally to this work.

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_9, © The Author(s) 2022

Fig. 1 Morphology and paratomic fission in Aeolosoma viride. The intact worm has a prostomium and a peristomium with the mouth in the first segment. The enlarged midgut locates at the center of its alimentary canal, and the fission zone locates between the parental chain and zooid. The posterior growth zone is located in the last segment before pygidium. The red dashed lines indicate the amputation sites in the experiment of anterior regeneration. The yellow dashed lines indicate the amputation sites in the synchronization and experiment of posterior regeneration

A. viride is a semitransparent, freshwater annelid of length 2–3 mm, comprising 10–12 segments [4]. Phylogenetically, A. viride is defined as a "clitellate-like polychaete" [5, 6]. Consistent with previous reports, A. viride reproduces exclusively by paratomic fission under our laboratory conditions (Fig. 1) [3, 7]. Paratomy is a form of agametic reproduction that produces multiple zooids simultaneously by fission in posterior segment. Species reproducing by paratomy have different regenerative capacities to regenerate anterior and posterior segments [8, 9]. Given their small size and transparency, as well as their strong regenerative ability together with applicability of various molecular tools including EdU labeling, whole-mount in situ hybridization (WISH), and RNA interference (RNAi) [4, 10], we anticipate that A. viride will be informative in comparative studies focused on whole-body regeneration. In this chapter, we will provide detailed steps on how to manipulate and conduct this novel model in regenerative research.

#### 2 Materials

All solutions are prepared using analytical grade reagents and dissolved in deionized ultrapure water at room temperature (RT).

	- 2. 1 phosphate buffered saline (PBS): 40 g/L NaCl, 1 g/L KCl, 7.2 g/L Na2HPO4, 1.2 g/L KH2PO4. Sterilize before use.
	- 3. Ground oatmeal.
	- 4. Saturated menthol in ASW (see Note 1).
	- 5. 4% (w/v) paraformaldehyde (PFA) in saturated menthol (see Note 2).
	- 6. 4% (w/v) PFA in 1 PBS.
	- 7. Mounting solution (e.g., 100% glycerol).
	- 8. Sterile needles 27 G 1/200.
	- 9. Microscope glass slides (e.g., 8 well-slide).
	- 10. Culture plate (e.g., 6, 12 or 24 wells).
	- 11. 25 C incubator.
	- 12. Dissection microscope (e.g., WILD M8, Leica).
	- 13. DIC microscope.

2.2 Cell Proliferation Assay


#### 2.3 Whole-Mount In Situ Hybridization (WISH)


#### 3 Methods

2.4 RNA Interference

(RNAi)



#### 3.3 Animal Fixation 1. Place a maximum of 10 intact or amputated worms in 20 μL ASW per well on a 8 well-slide under a dissection microscope.


#### 3.4 Cell Proliferation Assay

Fig. 2 Anterior regeneration in A. viride. Morphology was observed in intact and regenerating worms after amputation. The protruding blastema becomes visible 24–48 h after amputation (hpa). Mouth formation can be detected around 96 hpa as indicated with black arrows. The amputation site is marked by a black dotted line. Scale bar: 50 μm


3.5 Whole-Mount In Situ Hybridization (ISH)

	- 2. Place a maximum of 50 intact or amputated worms in 200 μL ASW per tube.
	- 3. Wash twice with ASW, for 5 min each time.
	- 4. Remove the ASW and add 200 μL TRIzol.
	- 5. Vortex for 20–40 s until the tissue is completely liquefied without visible particles.
	- 6. Add 40 μL chloroform.
	- 7. Gently invert 10 times.
	- 8. Keep at 25 C for 15 min.

Fig. 3 EdU labeling of blastema cells at 48 h postamputation. Animals were incubated in EdU for 0, 6, or 12 h and then immediately fixed at 48 hpa. EdU-labeled nuclei are detected red and costained with Hoechst 33342 (blue). Amputation plane is on the left. Scale bar: 100 μm


#### 3.6 RNA Interference (RNAi) by Feeding Method


Fig. 4 Expression of the Avi-caspase X gene during anterior regeneration. Whole-mount ISH was performed on intact and regenerating worms with sense (upper row) or antisense (lower row) riboprobe. The amputation site is located on the left. Note the Avi-caspase X expressing cells (blue purple) observed from 24 hpa and at later time-points. Scale bar: 100 μm


Fig. 5 Avi-beta tubulin isoform 1 RNAi inhibited regeneration in A. viride. The inhibitory effect of regenerates by Avi-beta tubulin isoform 1 RNAi feeding or microinjection was observed at 7 days postamputation. The head morphology of regenerating worms was obviously affected by feeding or dsRNA microinjection method. The black and white arrow respectively indicated the mouth. Scale bar: 100 μm


#### 1. Produce control and target dsRNA from L4440 vector (follow steps 1 and 2 in Subheading 3.6 and steps 23–26 in Subheading 3.5).


#### 3.7 RNAi by Microinjection


#### 4 Notes


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 10

# Studying Annelida Body Regeneration Under Environmental Stress in Diopatra neapolitana

Adı´lia Pires

#### Abstract

The polychaete Diopatra neapolitana is a cosmopolitan annelid that can robustly regenerate both its anterior and posterior body part depending on the position of the amputation. Previous studies demonstrated that body regeneration represents a sensitive and unspecific response to environmental stresses, including contaminants and climate alterations.

The posterior body regeneration of D. neapolitana is thus a suitable, ecological and relevant biomarker in ecotoxicological and ecological risk assessment assays. Here we describe the amputation process, the monitoring of the regeneration process of the polychaete D. neapolitana and the quantification of the impact of environmental stresses on its regenerative capacity.

Key words Polychaetes, Posterior regeneration, Biomarker, Endpoint, Environmental alterations

#### 1 Introduction

Annelids are known for their efficient wound healing and their capacity to regenerate both anterior and posterior segments after loss by injury [1, 2]. This regenerative ability varies significantly within the phylum, and some species can regenerate an entire individual from a single segment while others are much more limited [3]. Previous works demonstrated that species of the genus Diopatra could regenerate anterior and posterior segments and prostomial structures [3–6]. This mechanism performs a critical role in survivorship after tissue loss due to sublethal predation and harvesting [5, 7]. Additionally, it can also aid in recovery from injuries due to physical alterations [8].

Several studies demonstrated that exposure to environmental stressors such as contaminants or abiotic alterations reduced the regenerative capacity of polychaetes [9–17], with organisms regenerating slower and usually fewer chaetigers (segments that have chaetae). Nusetti et al. [9] observed that the polychaete Eurythoe complanata exposed to crankcase oil took longer to regenerate a

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_10, © The Author(s) 2022

new region and regenerated fewer segments. Exposure to microand nanoplastics reduced the capacity of Perinereis aibuhitensis and Hediste diversicolor to regenerate their posterior ends [14, 15]. Diopatra neapolitana exposed to several contaminants, such as metals, pharmaceuticals, carbon nanotubes, and environmental enrichment presented a delay in posterior segments regeneration, taking longer to achieve complete regeneration, and regenerated fewer segments [10–13, 16, 18]. Moreover, exposure to abiotic alterations, including pH variations and salinity changes, also reduced the regenerative ability of D. neapolitana [17].

Although the majority of toxicity studies with polychaetes have been conducted using the species H. diversicolor [14, 19–23], most of those regarding the use of the regenerative ability as a biomarker were carried out with the species D. neapolitana due to this process being well documented for Diopatra species (e.g., [3, 5, 24]). Additionally, this species represents a wide spatial distribution, being reported in intertidal and shallow subtidal habitats, namely, in the Red Sea and Indian Ocean [25], Mediterranean Sea [26–28], and the Atlantic Ocean [6, 29–31]. Furthermore, Diopatra species play an important ecological and economic role. Their tubes stabilize the sediments, increasing their structural complexity and thus their biodiversity, by supplying refugia from disturbance and predation [32] and facilitating the settlement and the attachment of some algal species [33]. Moreover, this species is commonly harvested to be sold as fish bait [31, 34, 35]. Altogether, these studies suggest that the regenerative capacity of polychaetes is a suitable biomarker in ecotoxicological and ecological risk assessment assays since it is sensitive to environmental stressors, including organic and inorganic contaminants and climate alterations.

The mechanisms behind this sensitive yet unspecific response to environmental stresses in Diopatra remain to be elucidated. Some authors suggested that the delay in regenerative capacity could be related to exposure to oxidative stress [10, 11, 14, 17] since free radicals may damage the biochemical and cellular functions that underlie the regenerative process. Moreover, Soneja et al. [36] reported that oxidative stress prolonged chronic wound inflammation as it stimulates cells of the immune system. Delayed regeneration may impact the sexual reproduction of individuals, as organisms will canalize their reserves toward tissue regeneration rather than producing gametes [5–7]. Also, the delay of organisms in starting gamete production compromises population maintenance, with consequences for communities and ecosystems [8].

Consequently, due to this species' ecological importance, understanding the interplay between environmental stresses and regenerative capacity is particularly pertinent since delays in regenerative capacity may negatively impact population and ecosystem function.

This chapter presents a detailed protocol to study the impacts of environmental stressors in the posterior regenerative capacity in field-collected organisms of the polychaete D. neapolitana.

#### 2 Materials

All reagents should be prepared with sterile reverse osmosis water and stored at room temperature (RT) unless otherwise stated.


#### 3 Methods

3.1 Collection of Organisms and Acclimation


Fig. 1 (a) Tube of Diopatra neapolitana at sediment surface and (b) shovel with the inclination that should be used to catch D. neapolitana specimens


Fig. 2 Diopatra neapolitana anterior end ventral view (a) and dorsal view (b); (c) D. neapolitana specimen regenerating the posterior end, (d) D. neapolitana with posterior end regenerated. The newly regenerated chaetigers have a lighter color, being possible to observe the blood vessels through the body wall. 10 chaetiger 10, 60—chaetiger 60, P—Prostomium, Br—Branchiae, Pa—parapode, R—width of the regenerated chaetiger; NR—width of the not regenerated chaetiger (chaetiger 60); RS—specimen with posterior end fully regenerated


#### 3.2 Regeneration Assay Experiments should be carried out with acclimatized specimens of similar size. The impacts of environmental stresses are tested by exposing the regenerating organisms to contaminated sediments and/or contaminated water.


Fig. 3 Different levels of posterior regeneration of Diopatra neapolitana exposed to sediments contaminated with lead (0.0, 3.0, 9.0 mg/kg). Photographic record of the regeneration process 14 (left column) and 28 days (right column) after amputation at control (a and b), 3.0 mg/kg (c and d), and 9.0 mg/kg (e and f)

#### 4 Notes


alive. However, if it does not run into the tube, it means that it is dying. In this situation, remove the organism from the aquarium with its tube. Dead organisms should be immediately removed from the aquarium because they begin to decompose very quickly and contaminate water and sediment.


#### Acknowledgments

Thanks to FCT/MCTES for the financial support to CESAM (UIDP/50017/2020+UIDB/50017/2020+ LA/P/0094/2020) through national funds. This work was also financially supported by the project BIOGEOCLIM —PTDC/CTA-GQU/29185/2017 (POCI-01-0145-FEDER-029185) funded by FEDER, through COMPETE2020—Programa Operacional Competitividade e Internacionalizac¸a˜o (POCI), and by national funds (OE), through FCT/ MCTES. Adı´lia Pires was contracted by national funds (OE), through FCT—Fundac¸a˜o para a Cieˆncia e a Tecnologia, I.P., in the scope of the framework contract foreseen in the numbers 4, 5, and 6 of the article 23, of the Decree-Law 57/2016, of August 29, changed by Law 57/2017, of July 19.

#### References


https://doi.org/10.1016/j.marenvres.2015. 03.002


26(3–4):265–272. https://doi.org/10.1111/ j.1439-0485.2005.00055.x


Polychaetes of commercial and applied interest in Italy: an overview. Me`moires du Muse`um Natl dHistoire Nat 162(July):593–603


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Studying Annelida Regeneration Using Platynereis dumerilii

#### Michel Vervoort and Eve Gazave

#### Abstract

Regeneration, the ability to restore body parts after an injury or an amputation, is a widespread property in the animal kingdom. This chapter describes methods used to study this fascinating process in the annelid Platynereis dumerilii. During most of its life, this segmented worm is able to regenerate upon amputation the posterior part of its body, including its pygidium (terminal non-segmented body region bearing the anus) and a subterminal posterior growth zone which contains stem cells required for the formation of new segments. Detailed description of Platynereis worm culture and how to obtain large quantity of regenerating worms is provided. We also describe the staging system that we established and three important methods to study regeneration: whole mount in situ hybridization to study gene expression, 5-ethynyl-20 -deoxyuridine (EdU) labeling to characterize cell proliferation, and use of pharmacological treatments to establish putative roles of defined signaling pathways and processes.

Key words Regeneration, Annelid, Platynereis dumerilii, Whole mount in situ hybridization, Gene expression, EdU, Cell proliferation, Pharmacological inhibitors

#### 1 Introduction

Regeneration, the ability to restore a lost or damaged body part is a fascinating process that has intrigued scientists since the pioneering study of Hydra regeneration by A. Trembley during the 1700s [1]. While having been intensively studied during the first part of the twentieth century, reparative regeneration has been less investigated since the rise of genetic and molecular studies of development in the 70s. This is intrinsically linked to the limited regenerative potential of the main developmental biology models, with the noticeable exception of zebrafish [2], which have emerged at that time. These last years, there has been a strong revival of the interest for regeneration, in part driven by possible applications for regenerative medicine [3].

Annelida (annelids) constitute a major lineage of the Lophotrochozoa super phylum, a group of primary importance to understand animal and especially bilaterian evolution [4]. Annelids represent a quite large phylum, with over 22,000 species including

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_11, © The Author(s) 2022

ragworms, earthworms and leeches. They can live in various ecosystems, mostly in the sea, but also in fresh water and humid terrestrial environments. They present a diversity of forms and life history traits; some live in a tube, while others are burrowed deep in the sand, stuck on algae or even parasitic [4, 5].

Interestingly, annelids, with the noticeable exception of leeches, are among the Metazoa that show the most important regenerative abilities [6, 7]. Indeed, many annelids are able to regenerate, after an amputation or injury, the posterior part of their body, their anterior part (including the head), or both, as well as appendages (named parapodia) and all kind of tentacles and cirri [6]. While the capacity to regenerate their posterior parts is almost shared by all annelids, anterior or head regeneration is less widespread [6].

There is a quite long history of experimental and descriptive morphological studies of regeneration in many annelid species [8, 9]. Many of these studies notably investigated possible sources of the cells involved in regeneration [8, 10], as well as the importance of the nervous system to allow a proper regeneration [11]. More recently, cellular and molecular aspects of annelid regeneration have been studied in a couple of model species, Pristina leydyi, Capitella teleta, and Enchytraeus japonensis, all belonging to the same group of annelids, the Sedentaria (for review see [6, 12]). While these studies provided interesting information, there is, however, still a crucial need for additional annelid models that allow to address fundamental and mechanistic questions about regeneration.

One major model species that has been successfully developed for decades is the Nereididae Platynereis dumerilii, which was originally described by Audouin and Milne Edwards in 1834 (Fig. 1) [13], and belongs to the Errantia lineage. Platynereis dumerilii is a medium-sized marine annelid that is easily maintained in laboratories world-wide. Like many other marine animals, such as corals, sea urchins and fishes, Platynereis's life cycle is synchronized with the lunar cycle [14]. Each worm will reproduce only once in its life before dying, and the timing of this reproduction is tightly regulated by this circalunar life cycle. Platynereis has emerged as an intensely studied model organism for developmental, marine, neuro, and evolutionary biology, as well as to study regeneration [15, 16]. Platynereis worms have indeed extensive regenerative capabilities: after amputation of the posterior part of their body, which leads to the removal of the pygidium (terminal non-segmented body part of the worm), the stem cell–rich subterminal growth zone (responsible for the continuous growth of the worms [17]) and several segments, Platynereis worms are able to regenerate both pygidium and growth zone which will in turn produce new segments [18]. Platynereis is also able to regenerate various body outgrowths, such as tentacles and appendages

Fig. 1 Platynereis dumerilii. Pictures of juvenile and adult (male and female) worms. Males and females harbor specific morphological features linked to sexual metamorphosis, notably their color. While juveniles are mainly brownish, females are bright yellow, as they are full of oocytes. Males are white in their anterior part, as they are full of sperm, and red in their posterior part, due to extensive accessory blood capillaries. Morphological differences between juveniles and maturing worms are not limited to their color. Indeed, during sexual maturation, the whole intestine of the worm regresses and the trunk of the animal is progressively modified to become a bag full of gametes. In addition, mature worms harbor bigger and darker eyes compared to juveniles

(parapodia), but not its head. Platynereis worms can thus properly regenerate both complex differentiated structures which includes different types of tissues or organs (pygidium and parapodia, for example) and stem cells (posterior growth zone) [17, 18]. In this chapter, we will describe protocols routinely used to breed and maintain Platynereis in the laboratory and prepare biological materials required for regeneration studies. We will also introduce molecular biology and functional tools used to address key questions about regeneration.

#### 2 Materials

Prepare all solutions using ultrapure autoclaved water (H2O). Prepare and store all reagents at room temperature (unless indicated otherwise).

#### 2.1 Platynereis Worms Culture and Biological Material Production


#### 2.2 Whole Mount In Situ Hybridization and EdU Labelling


#### 3 Methods

3.1 Platynereis Worms Culture Platynereis dumerilii is a marine worm found worldwide in temperate seas [19]. Since decades researchers have no longer taken animals directly from the sea (except if information related to environmental cues are needed), as Platynereis's full life cycle is completed easily and successfully in laboratory settings (see Note 5) [20]. To raise Platynereis worms, always rinse glassware with distilled water and never use detergents.


3.2 Production and Fixation of Samples at Specific Stages of Regeneration

To minimize, as much as possible, variability, notably due to the age and the size of the animals, strict procedures for worm selection and amputation should be followed.


Fig. 2 Regeneration stages. On the top of the figure is drawn a growing juvenile worms with its posterior growth zone (orange line and arrowhead) and, anterior to the growth zone, developing segments (purple asterisks) and, posterior to the growth zone, the pygidium characterized by the presence of two specific outgrowths, anal cirri (green arrowheads) and two large glands (gray circles). Amputation plane is represented by red dotted lines. The five stages of regeneration are depicted. At stage 1, wound healing is achieved. At stage 2, a small blastema composed of proliferating cells is formed and its size increases during subsequent stages. At stage 3, small anal cirri can be observed. They strongly extend at stage 4 and some signs of pygidium differentiation become obvious (e.g., presence of glands). At stage 5, pygidium differentiation has pursued and a few segments delimited by faint segmental boundaries are observed. Growth continues and an increasing number of differentiating segments (with obvious segmental boundaries and developing parapodia) can subsequently be observed


#### 3.3 Whole Mount In Situ Hybridization

Whole mount in situ hybridization (WMISH) is the specific annealing of a labeled RNA probe to complementary sequence of a target mRNA in a fixed specimen, followed by detection and visualization of the nucleic acid hybrids [21] (see Note 22) (Fig. 3).


Fig. 3 Whole mount in situ hybridization. Whole mount in situ hybridization for the genes whose name is indicated are shown for two posterior regeneration stages, stage 3 (a–c) and stage 5 (d–f). All panels are ventral views (anterior is up). Amputation plane is represented by yellow dotted lines. In a and d, blue arrowheads point to expression of Pdum-pax6 in two longitudinal rows of neuroectodermal cells which will give rise to ventral neurons of the ventral nerve cord. In b is shown the expression of Pdum-neurogenin in a large number of neuronal precursor cells of the both the central and peripheral nervous system. In c, brown arrows point to the expression of Pdum-cdki1a in internal cells located in the anal region. In e, green arrowheads point to segmental ectodermal stripes of cells expressing Pdum-prdm3/16 which is also expressed in cells of the posterior growth zone (red arrowheads). These cells also express Pdum-evx (red arrowheads in f)


3.4 EdU Labelling for Investigating Cell Proliferation During Platynereis Regeneration

5-ethynyl-2<sup>0</sup> -deoxyuridine (EdU) is a nucleoside analog that is widely used to detect cells that are in the S-phase of their cell cycle in various species (see Note 34). In Platynereis, EdU labeling has been used to study cell proliferation in whole mount animals during development, postembryonic growth, and regeneration (e.g., [17, 18, 22]) (Fig. 4).


Fig. 4 EdU Labeling. Confocal image of the posterior part of a stage 3 worm (3 dpa) incubated in 5 μM EdU for 1 h before fixation. Hoechst counterstaining has been performed and allows to visualize all nuclei (in blue). EdU labeling is shown in red. Amputation plane is represented by yellow dotted lines. Anterior is up


3.5 Pharmacological Treatments for Functional Studies During Platynereis Regeneration

Performing functional studies during postembryonic developments used to be challenging for many organisms in which genetic tools are not easily or fully mastered. One way to alter or modify various molecular signaling pathways or cellular mechanisms is to soak regenerating animals in specific pharmacological inhibitors or activators (see Notes 38 and 39). An initial and crucial step consists in defining the efficient concentration that induces defects in the regeneration process (e.g., morphological abnormalities) and/or in its timing, without (or with minimal) toxic effects (i.e., with a minimal number of dead or autotomized animals (see Note 40)).


#### 4 Notes


occurs when animals are facing stressful conditions (or are mechanically damaged). Autotomy can happen in normal culture conditions, at a low rate. When scoring worms during treatments, autotomy is easily visible as worms are separated in at least two fragments, the one bearing the head may start to regenerate again, while the other will not.


48. For inhibitor treatments, perform two-way ANOVA multiple comparisons between control versus treated worms per scoring day. Comparisons between different inhibitor concentrations are also interesting to perform.

#### Acknowledgments

Work in our team is supported by funding from Labex "Who Am I" laboratory of excellence (No. ANR-11-LABX-0071) funded by the French Government through its "Investments for the Future" program operated by the Agence Nationale de la Recherche under grant No. ANR-11-IDEX-0005-01, Centre National de la Recherche Scientifique, INSB (grant Diversity of Biological Mechanisms), Agence Nationale de la Recherche (grant TELO BLAST no. ANR-16-CE91-0007). The authors warmly thank all current and past members of the "Stem cells, Development and Evolution" team at the Institut Jacques Monod, Paris, France.

#### References


de Zoologie expe´rimentale et ge´ne´rale 110(1): 127–143


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 12

# Collecting and Culturing Lineus sanguineus to Study Nemertea WBR

### Eduardo E. Zattara and Fernando A. Ferna´ ndez-Alvarez

#### Abstract

Whole-body regeneration, the ability to reconstruct complete individuals from small fragments, is rare among ribbon worms (phylum Nemertea) but present in the pilidiophoran species Lineus sanguineus. This species can regenerate complete individuals from a tiny midbody section, and even from a quarter of a piece, provided it retains a fragment of a lateral nerve cord. While a few other unrelated species of ribbon worms are also excellent regenerators, L. sanguineus is unique in having evolved its regenerative abilities quite recently and thus offers an exceptional opportunity to gain insight into the evolutionary mechanisms of regeneration enhancement. Interestingly, both its sister species Lineus lacteus and Lineus pseudolacteus, a third species derived from the recent hybridization of the other two, differ in their regeneration abilities: while L. lacteus is uncapable of regenerating a lost head, L. pseudolacteus is capable of anterior regeneration, albeit at a slower rate than L. sanguineus. L. sanguineus has a worldwide distribution in temperate shores of both hemispheres, is readily found at intertidal habitats, and can survive, feed and be bred through asexual replication with minimal effort in laboratory settings. All the above make this species a superb candidate for studies of regenerative biology. In this chapter, we present protocols to collect, identify and breed L. sanguineus to study the extraordinary whole-body regeneration abilities found in this species.

Key words Heteronemertea, Intertidal, Invertebrate rearing, Pilidiophora, Spiralia

#### 1 Introduction

Regeneration, the ability of organism to regrow lost body parts, is widespread across metazoan groups [1–3]. Regeneration varies broadly both in restorative potential and phylogenetic distribution: while many lineages are only capable of physiological tissue turnover or restoration of smaller amounts of lost tissues, others are capable of amazing regeneration feats, from restoring lost appendages to reconstructing whole new individuals from very small fragments. This later ability, known as whole-body regeneration, is well exemplified by planaria and other turbellarian flatworms, but is also commonplace in many cnidarians, ctenophores, sponges, xenacoelomorphs, and colonial tunicates. Some other groups, such as annelids and echinoderms, also show members with more

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_12, © The Author(s) 2022

limited but still exceptional regenerative powers [3]. Such broad array of regenerative potential across animals suggests that regenerative abilities have a rich evolutionary history that is mostly unexplored.

Understanding how regeneration evolves can help elucidate the cellular and molecular underpinnings of this developmental ability. One particularly informative approach is through comparative studies of regenerative ability in species that span an evolutionary transition in regenerative potential. Candidate mechanisms can be gleaned from comparing developmental and molecular genetic differences across such species and correlating them with changes in extent of regeneration. Mechanisms can then be experimentally assessed to test whether they inhibit or enhance regeneration. Studies on species spanning an evolutionary loss or reduction of regenerative ability can inform how the potential to regenerate might have become blocked or dampened, and eventually lead to strategies to lift or alleviate such blocks and allow for better regeneration and healing in systems that do not regenerate well (including most mammals in general, and humans in particular). In contrast, studies on species spanning an evolutionary gain or enhancement of regenerative ability can give insight on how organisms might be able to reboot embryonic developmental capabilities in a postembryonic context and inspire novel tools to induce regeneration after traumatic injury. While many evolutionary transitions leading to diminished or lost regenerative ability have been identified, there are very few examples of increased or gained regeneration [4, 5]. Thus, while we have many systems where we can study how regeneration is lost, we lack good models of how it is gained.

Ribbon worms (phylum Nemertea) are a phylum of about 1300 known species of elongated, primarily marine predatory worms [6–8]. While most species of nemerteans are capable of restoring a lost posterior end after a transverse amputation behind the brain, only a few have so far been shown to be able of restoring their anterior end after a similar injury [9]. The few species capable of anterior regeneration are taxonomically scattered across the phylum; ancestral trait reconstruction strongly suggests that lack of anterior regeneration is the ancestral and most common condition for nemerteans. Thus, species capable of regrowing a lost anterior end represent lineages that experienced evolutionary gains in regenerative ability.

Among anteriorly regenerating species, Lineus sanguineus (Rathke, 1799) stands out, unquestionably one of the champions of regeneration possessing some of the highest regenerative abilities known among animals [2]. A single worm of this species can be repeatedly amputated to obtain a complete regenerated worm just 1/200,000th of the volume of the original individual. Furthermore, a complete worm can regenerate not only from a thin transverse slice of the body, but even from just one quadrant of a thin slice [10]. Regeneration rate varies with the size and condition of the fragment, but a recognizable head and tail can be rebuilt in around a week or two.

Lineus sanguineus individuals tend to have a slender body, often a 100 times longer than their body width (Fig. 1a). They are slightly flattened dorsoventrally with a pair of long lateral grooves at the anterior end, followed by a reddish brain region. They have about 2 to 8 pairs of dorsolateral ocelli arranged in a bilateral pair of rows extending along the anterior half of the head, over the lateral cephalic slits. The mouth opens ventrally some distance behind the brain. Except for the frontal margin and lateral borders of the head, worms tend to be uniform in color. Color itself varies across the species' distribution, showing olive, green, brown or red hues. L. sanguineus inhabits sheltered stony regions, among algae, lurking within shellfish beds, or inside the fouling/encrusting community growing over natural and manmade substrates of the intertidal zones of the marine shore. It can be found on temperate seashores around the world [9, 11–15] (Fig. 1d). At the Southern Hemisphere, it has been found on the South Atlantic shores of Uruguay and Argentina, and the South Pacific shores of Chile and New Zealand. In the Northern Hemisphere, it has been reported from the North Atlantic shores of North America (from the Gulf of Mexico and Florida to Newfoundland, Canada), Europe (along the Bay of Biscay, English Channel and North Sea) and Eastern Asia (South and East China Seas, Yellow and Bohai Seas, Sea of Japan and eastern Japanese Atlantic shores). This widespread distribution makes it more readily available for collection by researchers near most temperate locations around the world.

Lineus sanguineus belongs to the class Pilidiophora, the nemertean group that contains the highest number of species with wholebody regenerative ability [9]. This cosmopolitan species [12] belongs to a mostly European species group that also includes Riseriellus occultus (described from NW Spanish and N Welsh shores), Lineus longissimus (found all along European Atlantic shores), two cryptic species known as Lineus lacteus (L. lacteus A, associated with the Bay of Biscay and English Channel shores, and L. lacteus M, found in the Mediterranean Sea), and the endemic Lineus pseudolacteus (found only at the French Atlantic shores near Roscoff, Bretagne) [12, 13]. Of the above species, only L. sanguineus and L. pseudolacteus are capable of anterior regeneration. The other species are limited to regenerate posterior ends [9].

Lineus sanguineus and L. lacteus A are sister species, estimated to have diverged about 10 My ago, while L. pseudolacteus likely emerged from a much more recent (12–25 Ky ago) single hybridization event between L. sanguineus and L. lacteus—likely after fertilization of an unreduced L. sanguineus oocyte by a L. lacteus sperm [13, 16]. Due to its triploid condition, L. pseudolacteus has

Fig. 1 Live and regenerating examples of Lineus sanguineus. (a) Live, extended individual of L. sanguineus; this specimen is relatively short—much longer specimens can be found. Anterior end at the left. The left inset shows a detail of the head in dorsal view. The right inset shows a detail of the head on lateral view (b) Live, coiled individual of L. sanguineus. Head toward upper right (c) Example of anterior regeneration from a posterior fragment, shown from 2 through 18 days postamputation (dpa); the red dashed line at 2 dpa shows the location of the healed anterior wound. Notice first ocelli appearing at 7 dpa, proboscis apparatus formed by 11 dpa and brain visible by 13 dpa (inset showing detail of head on lateral view); also note how the stump elongates and becomes slenderer to match the width of the regenerating anterior end. (d) Current known been reproducing exclusively asexually since that event, a regeneration-dependent strategy inherited from their maternal species. Lack of anterior regeneration in L. lacteus places an upper bound on how long ago did the L. sanguineus lineage evolve anterior regeneration. Both parent species have been shown to possess private alleles, that is, unique haplotypic variants found only in one of the species. This should give a considerable fraction of their genome a specific signature which could facilitate detection of allelic bias in genes differentially expressed during regeneration of L. pseudolacteus, complementing gene expression studies comparing postamputation responses between L. lacteus and L. sanguineus. Furthermore, populations of L. sanguineus display different morphotypes with corresponding differences in regeneration potential, that are not obviously correlated with genetic differences [9, 12, 13]. For example, two ecologically isolated morphotypes with the same genotypic structure are present in Iberian shores: smaller individuals (<5 cm of total length) can be found among algae in lower, mid and sub-tidal regions, while the larger ones (5–20 cm) are found among sand below boulders in the high intertidal region. Interestingly, larger individuals regenerate more slowly than the smaller ones, suggesting that regenerative potential might be modulated by physiological trade-offs and local adaptations. All the above make L. sanguineus, L. lacteus and L. pseudolacteus a unique and powerful three-species system to inquire into the molecular and developmental mechanisms that evolved to enable the spectacular whole-body regeneration currently found in L. sanguineus.

In this chapter, we present simple and inexpensive methods to collect, keep and experiment on Lineus sanguineus. Most of these methods are applicable also to L. lacteus and L. pseudolacteus (except of course for asexual propagation, which cannot be used in L. lacteus). Many of these methods might also apply to a variable degree to other nemertean species too.

#### 2 Materials

#### 2.1 Field Collection of Specimens


Fig. 1 (continued) distribution of Lineus sanguineus. Occurrence data obtained from the Global Biodiversity Information Facility and other sources [9, 11–14, 25] and curated by the authors. Occurrences are color-coded to show the different species under which the specimens had been originally described; all but Lineus pseudolacteus are now synonymized to Lineus sanguineus


#### 3 Methods

### 3.1 Field Collection

of Specimens

Lineus sanguineus inhabit rocky or pebbly areas of the intertidal zone, the region left exposed by receding waters during low tides (see Note 2, Fig. 2a). They often co-occur with other, sometimes similar looking species (see Note 3), like Lineus ruber, L. viridis, or L. clandestinus. In some places and locations, worms can be found and collected directly behind rocks and pebbles. This method is typically suitable for large specimens from the high intertidal zone (see Note 4). In most other occasions, however, they lurk within encrusting and fouling communities growing over large rocks and cannot be easily retrieved directly. In those cases, the most successful strategy is to cause hypoxia-induced migration (see Subheading 3.2).


Fig. 2 Field collection of nemerteans. (a) A typical intertidal area where L. sanguineus can be found. (b) Direct collection of specimens under pebbles, rocks and shells. (c) Large specimen of Lineus lacteus inside a mollusk shell while eating its owner; the anterior end is deep within the shell. (d) Removal of a sample of the fouling community encrusted on the pylons of a pier. (e) Removed rubble is placed in trays and covered with sea water. (f) Rubble is spread out on the bottom of the tray, and allowed to become hypoxic, forcing nemerteans to come out of their shelter and allowing their collection


3.2 Hypoxia-Induced Migration This method, proposed by Kirsteuer [17], is suitable for smaller specimens from the low and mid intertidal region inhabiting withing the encrusting community formed by algae, mussels, barnacles or other creatures adhering to a hard substrate (natural rocks and outcrops, or manmade structures like pilons and jetties). This strategy induces them to migrate out to the open by falling oxygen concentrations.


Rearing


3.3 Specimen Collected specimens can be kept in the laboratory for many months and up to several years with minimal maintenance (see Note 11).


3.4 Feeding Although many nemerteans, including Lineus sanguineus can survive for many months without feeding, keeping well-fed worms will improve overall condition, yield more reliable experimental results and allow increasing population numbers through clonal propagation (see Subheading 3.5). L. sanguineus is a voracious predator, and likely a scavenger too, and will feed readily once it learns the nature of its food. It can be fed a variety of items, including live annelids, processed liver, minced scallops or mussels as well as eggs/oocytes from other invertebrates. In this protocol, we detail preparation of and feeding with liver homogenate, since this is a well-established method used to rear and perform dsRNA- or drugmediated interference experiments on the planarian Schmidtea mediterranea [18, 19].

	- 1. Label one glass dish per worm to be cut (see Note 22).
	- 2. Fill the dishes to about half of their volume with FSW.
	- 3. Add cold (4 C) FSW to a shallow petri dish plate up to about 5 mm.
	- 4. Move the worm to be amputated into the cold petri dish (see Note 23).
	- 5. Wait until the worm starts crawling and extends.
	- 6. Using a #10 scalpel blade (or similar curved edge blade), make a single transverse cut at about one third of the total body length from the anterior end.
	- 7. Move the anterior fragment into a new, labeled culture dish (see Subheading 3.3).

#### 4 Notes

1. Clean sea water is the main requirement for successful maintenance and rearing of Lineus sanguineus and other nemerteans. While most marine research stations are fitted with sea tables and a constant supply of natural sea water, other locations are unlikely to have such facilities. If located near the ocean, sea water can be procured from the shore, brought in tanks or bottles, filter-sterilized and stored. Worm cultures use relatively small amounts of water, so unless there are many specimens being kept, only occasional trips would be needed. If located inland, then procuring natural sea water might not be practical. In such a case, it is possible to use instead one of the many formulations for artificial sea waters sold for aquariums. However, formulations not always yield an artificial sea water equivalent to the one which the worms are habituated and might even prove lethal to them. Specific formulations should be tested on one or two individuals, by passing the worms through a graduated replacement from the natural seawater in which they were placed after collection to the artificial sea water. Even if the worms appear to survive the artificial medium, specifics of the formulation could affect regenerative ability. Thus, if setting up worm cultures for a research project at an inland location, it might be advisable to bring natural seawater from the collection sites and test that regenerative responses are similar in both natural and artificial media.


successfully pass this period, they usually become much more eager to feed on the same substrate in subsequent occasions. However, if worms keep rejecting the food after several attempts, it might be worth trying with a different item.


29. Time to complete regeneration varies, depending on the size of the fragments, temperature, and condition and strain of the original individual, but usually by 2 weeks it should be possible to see ocelli, brains, mouth and a proboscis on the regenerate (Fig. 1c).

#### Acknowledgments

We are grateful to Jon L. Norenburg (National Museum of Natural History, Smithsonian Institution) and Nuria Anado´n (Departamento de Organismos y Sistemas, Universidad de Oviedo) for inducing each of us into the depths of nemertean lore. E.E.Z. was supported through his training by a University of Maryland & Smithsonian Institution Seed Grant, and by the Smithonian Tropical Research Institute. F.A´.F.-A´ . was supported by a JdC-I Postdoctoral Fellowship Grant (ref. IJC2020-043170-I) awarded by CIN/AEI/10.13039/501100011033 and the European Union NextGenerationEU/PRTR. This research was supported by the Spanish government through the 'Severo Ochoa Centre of Excellence' accreditation (CEX2019-000928-S).

#### References


the phylogeny of some cosmopolitan Lineus species (Nemertea). Hydrobiologia 266: 159–168


(eds) Methods in cell biology. Academic Press, New York, pp 243–262


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 13

# Studying Xenacoelomorpha WBR Using Isodiametra pulchra

#### Bernhard Egger

#### Abstract

Xenacoelomorpha are a phylogenetically and biologically interesting, but severely understudied group of worm-like animals. Among them, the acoel Isodiametra pulchra has been shown to be amenable to experimental work, including the study of stem cells and regeneration. The animal is capable of regenerating the posterior part of the body, but not its head. Here, methods such as nucleic acid extractions, in situ hybridisation, RNA interference, antibody and cytochemical stainings, and the general handling of the animals are presented.

Key words Acoela, Isodiametra, Regeneration, Neoblast stem cells, Antibody stainings, Phalloidin, In situ hybridization, RNA and DNA extraction, Anesthesia

#### 1 Introduction

Xenacoelomorpha are one of the few remaining phyla with an unresolved, contested position in the Tree of Life. The group is either recovered as sister group of all other bilaterian animals, or as a member of Deuterostomia [1, 2]. Three groups constitute the Xenacoelomorpha: Xenoturbellida with 6 described species in one genus, Nemertodermatida with 18 described species, and Acoela with more than 300 described species being by far the largest and best known of the three groups. Their simple body plan—lacking a coelom, a circulatory system, a skeleton or respiratory organs other than the epidermis—can either be seen as plesiomorphic, or as a series of reductions [3]. In either case, they are an interesting and still poorly studied group of almost exclusively marine animals.

The regeneration capacity of the few studied xenacoelomorphs varies, where only a few species were shown to be able to completely regenerate their head, including brain and statocyst (a gravity sensing organ), such as Hofstenia miamia [4]. Regeneration capacity is possibly linked to the mode of reproduction, where obligatorily sexually reproducing species are often less capable of regeneration than asexually reproducing species. In different acoels, all modes of

Fig. 1 (a) Squeeze preparation of a live adult specimen of Isodiametra pulchra. (b) Same specimen as in a, nuclei of the epidermis stained blue with DAPI. Anterior is up. dp digestive parenchyma, eg developing eggs, fg female genital opening, mg male genital apparatus, mo mouth, mp male genital opening, st statocyst. Scale bar is 100 μm

asexual reproduction occur: architomy (fission happens before new organs have been built), paratomy (fission happens after new organs have been built), and budding [5].

Regeneration, growth and homeostasis in acoels is powered by neoblast stem cells, the only proliferating cells in the body, located exclusively in the mesenchymal space and thus lacking in the epidermis [4–7].

One of the better studied acoels is Isodiametra pulchra, an animal less than a millimeter in length, transparent, bearing a single statocyst near the anterior end (Fig. 1). It belongs to the speciesrich family Isodiametridae (comprising about 100 species), and can be cultured in large numbers in the laboratory. It is sexually reproducing, and cannot regenerate its head, but posterior body parts [8]. The following protocols are tested with adult and juvenile I. pulchra, or its close (and even smaller) relative, Aphanostoma pisae, or both [6–11].

In particular, RNA and DNA extraction, anesthesia, amputation, fixation, in situ hybridization (Fig. 2), RNA interference, and antibody and cytochemical stainings (Fig. 3) are covered in this

Fig. 2 Wholemount in situ hybridisation of an adult specimen of Isodiametra pulchra against Ipiwi1, a stem-cell gene. (Picture is courtesy of Thomas Zauchner). Anterior is up. Scale bar is 100 μm

chapter. While all methods included here have been published elsewhere, this chapter serves to bring them together in a compact format and to provide tricks and tips and notes on critical steps.

#### 2 Materials

2.1 Nucleic Acid Extractions Use nuclease-free (but not DEPC (diethyl pyrocarbonate)-treated) water. Only use nuclease-free sterile tubes, pestles and pipet tips. Only use molecular biology graded reagents. Work under the fume hood if indicated on the reagent's safety data sheet.


Fig. 3 Confocal laser scanning projection of the body wall and genital musculature of Aphanostoma pisae stained with rhodamine-conjugated phalloidin. (Picture is courtesy of Thomas Zauchner). Anterior is up. mg male genital apparatus, sb seminal bursa. Scale bar is 50 μm


2.2 Antibody and Cytochemical Stainings

There is no requirement to use purified water other than dH2O.


#### 2.3 In Situ Hybridization and RNA Interference

All solutions are to be prepared with either nuclease-free or DEPCtreated water (1 mL DEPC per liter solution; stir over night and autoclave).


#### 3 Methods

Work at RT and use a pipette, if not stated otherwise.

3.1 Anesthesia (Relaxation), Amputation, and Fixation

The soft-bodied animals will contract to unsightly balls when exposed to a fixative without prior anesthesia. In the literature and in the following protocols, anesthesia is referred to as "relaxation." Relaxing animals is not only necessary before fixation but also comes in handy for amputations to stop the animals from bending and turning around.


3.2 In Situ Hybridization In situ hybridization is used to detect mRNA in the tissue where it is expressed, using labeled RNA probes. Probe design and synthesis can be done after standard protocols (e.g., [6]).

If not otherwise specified, procedures are done at RT. Pipet liquids, not the animals, that is, the animals stay in the same container (microcentrifuge tube, petri, or embryo dish if not otherwise specified. Liquids are to be removed before adding new liquids.


3.3 RNA Interference (RNAi) This method is used to knock down expression of targeted genes in vivo with double-stranded RNA (dsRNA). In Isodiametra, RNAi can be simply performed by soaking the animals in a seawater solution with dsRNA. Use 25–40 animals per well of a 24-well plate (see Note 16).


3.4 Antibody Stainings Different to in situ hybridization reagents, there is no requirement for using nuclease-free or DEPC-treated water. While many combinations of antibody stainings are possible, here a double fluorescent wholemount staining using antibodies against BrdU and phosphorylated histone H3 (pH3) is presented.


#### 3.5 Cytochemical Stainings Again, a great variety of cytochemical stainings can be performed. Here, a triple fluorescent wholemount staining using EdU, DAPI and phalloidin is presented.


#### 3.6 RNA Extraction When preparing animals for an RNAseq experiment, antibiotics can be used to remove (or reduce) bacterial RNA. Animals can be starved for several days to avoid contaminating algal RNA. Take care not to breathe in opened tubes to prevent RNases from breaking up RNA.

1. Pipette live animals (typically 10–100) into a microcentrifuge tube (see Note 23).


#### 3.7 DNA Extraction For genome sequencing, antibiotics can be used on live animals to remove or reduce bacterial contamination.


#### 4 Notes


disturbed, it is advisable to wait for the next pipetting step until specimens have sunken to the bottom again.


#### Acknowledgments

I gratefully acknowledge the hard work of former students, working with and painstakingly improving protocols, namely Simona Migliano, Isabel Dittmann, Lucy Nevard, Lucy Neumann, Thomas Zauchner, and Jochen Hilchenbach. I am especially grateful to Peter Ladurner and to Katrien De Mulder, who adapted and established many protocols with Isodiametra.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 14

# Studying Echinodermata Arm Explant Regeneration Using Echinaster sepositus

### Cinzia Ferrario , Yousra Ben Khadra , Michela Sugni , M. Daniela Candia Carnevali , Pedro Martinez , and Francesco Bonasoro

#### Abstract

Echinoderms are marine invertebrate deuterostomes known for their amazing regenerative abilities throughout all life stages. Though some species can undergo whole-body regeneration (WBR), others exhibit more restricted regenerative capabilities. Asteroidea (starfish) comprise one of the few echinoderm taxa capable of undergoing WBR. Indeed, some starfish species can restore all tissues and organs not only during larval stages, but also from arm fragments as adults. Arm explants have been used to study cells, tissues and genes involved in starfish regeneration. Here, we describe methods for obtaining and studying regeneration of arm explants in starfish, in particular animal collection and husbandry, preparation of arm explants, regeneration tests, microscopic anatomy techniques (including transmission electron microscopy, TEM) used to analyze the regenerating explant tissues and cells plus a downstream RNA extraction protocol needed for subsequent molecular investigations.

Key words Echinoderms, Starfish, Regeneration, Arm explants, Transmission electron microscopy, Semithin and ultrathin sectioning, TEM grid staining, RNA extraction

#### 1 Introduction

Echinodermata is a well-known phylum of benthic marine deuterostome invertebrates that includes sea lilies, starfish, brittle stars, sea urchins, and sea cucumbers. Echinoderms and vertebrates belong to the same super phylum named deuterostomes, which includes the chordates and therefore vertebrates, and for this reason share common ancestral traits that were retained during evolution in both lineages. This makes them relevant alternative non-vertebrate models to study potential shared biological mechanisms/processes or to study loss of specific "functions" (e.g., remarkable regenerative abilities, including whole-body regeneration—WBR) along the different evolutionary lineages. Echinoderms have developed phylum-specific morphological features, such as secondary radial symmetry, pentameric organization, a miniaturized and modular hydraulic system, that is, the water vascular system, a calcareous endoskeleton (stereom structure), and mutable connective tissues, namely, collagenous tissues that can rapidly modify their intrinsic mechanical properties under nervous system control [1]. Echinoderms are very common and distributed worldwide at almost all depths and latitudes in all marine environments. Indeed, brittle stars can be the dominant macrobenthic fauna on many muddy seafloors and sea cucumbers sometimes account for up to 90% of biomass in the deep oceans [2]. Successful colonization of such diverse biotopes, despite the presence of different predators, may be explained by echinoderms' incredible adaptive capabilities, including their ability to regenerate lost body parts after predation. Although regeneration is observed in all echinoderm taxa, asexual reproduction followed by wholebody regeneration (WBR) is much less common. Notably, some starfish species, primarily Linckia spp. and Coscinasterias spp., are capable of extensive WBR: the ability to regenerate the entire individual from a single arm [2–4].

Several starfish species (class Asteroidea), such as Leptasterias hexactis, Asterias rubens, Marthasterias glacialis, and, more recently, Echinaster sepositus, represent the most used models for regeneration studies. Molecular analyses have been recently added to the range of well-known morphological approaches, namely, light and electron microscopy, to identify the mechanisms involved in both developmental and regenerative processes [5–10]. While regeneration studies were traditionally performed in animals with distally amputated arms, nowadays we have developed the better controlled model of arm explants. A double-amputated arm explant (an amputated arm reamputated at its distal tip) represents a simplified and easy-to-use model to investigate the regenerative process, including the relevant cells and tissues plus the activity of different regulatory molecules, that is, signaling and transcription factors, involved. Of particular importance is that the regenerative potential of the arm is tested in the absence of any systemic control by the rest of the body, including its supporting metabolic contribution [11–15]. Remarkably, in starfish arm explants, the distal end/tip initiates regeneration following the same stages observed in standard arm-tip regeneration, thus proving the validity of this cultured model. Furthermore, although to a much less extent, both distalization (namely, the regeneration of the distal-most structures (terminal tube foot and ossicle)) and intercalation (the regeneration of new tissues between the terminal differentiated structures and the stump) apparently occur in both the distal tip of the arm explants and the tip of single amputated arms. On the contrary, at the proximal end of the arm explants, only the terminal elements are regenerated (distalization) with no signs of intercalation being detectable [16] (Fig. 1). This can provide information on the ability

Fig. 1 Longitudinal section schemes of E. sepositus arm explants at three selected regenerative stages. For clarity, the tube feet of the nonregenerating arm explant portions have been omitted. Left column: regenerating proximal end. Right column: regenerating distal end. First line: 48 h p.a. Second line: 3 weeks p.a. Third line: 10 weeks p.a. Proximal and distal ends regenerate differently: indeed, while the distal end shows distalization (both terminal ossicle and tube foot) and intercalation (new tube feet), the proximal end shows only distalization (terminal tube foot only) without intercalation. For color coding of tissues, see legend embedded in the figure. Black lines ¼ amputation planes

to "manage" the directionality/polarity of regeneration (i.e., unidirectional vs bidirectional).

Echinaster sepositus, known as the red starfish, is found in the East Atlantic Ocean and in the Mediterranean Sea, where it is one of the most common starfish species. E. sepositus inhabits shallow waters, between 1 and 250 m deep, in a wide range of habitats, including rocky and sandy bottoms and seagrass meadows [17]. Its diurnal habits and evident coloration make it clearly visible on any substrate. They live in habitats that are easily accessible and, hence, their collection does not present logistic problems. Its size is sufficiently large to allow easy experimental manipulation and observation of regenerating stages, but still small enough to allow advanced microscopic analyses (e.g., transmission electron microscopy) of the regenerates. Although, as all echinoderms, it is difficult and time-consuming to achieve a full life cycle in the laboratory, adult specimens are rather robust and can be easily maintained in laboratory conditions for long periods (several months or up to 1 year). Overall, these practical features make it a valid and easy-to-handle research model. Indeed, it has been used for many years as model species to study arm tip regeneration using both morphological [18, 19] and molecular [20] approaches. Now we have extended the potential of the species by introducing the culture of arm explants, allowing a more efficient control over the regenerative process. Critical aspects of arm regeneration, such as the control of polarity, the dependence on positional cues, the origin of cells contributing to the different tissues and the regulatory aspects controlling the patterning of newly grown structures, are here more easily studied. In fact, the WBR potential of a single arm or arm piece can be better investigated using arm explants since culturing them allows both better control and easy reproducibility of growing conditions. A seemingly trivial, but extremely useful, characteristic of our model is that, being pentamerous animals, experimentally manipulated and control arm fragments can always be derived from the same animal.

In this chapter, we report methods for collecting and maintaining E. sepositus in the laboratory, preparing double-amputated arm explants, and studying the histology/ultrastructure of the regenerative processes using light and transmission electron microscopy (TEM), methods that we complement with those for extracting RNA used for intensive molecular analyses, such as transcriptomics.

#### 2 Materials

All reagents should be prepared using autoclaved filtered distilled water (dH2O) and kept at room temperature (RT), unless otherwise specified.



2.3 Semithin and Ultrathin Sectioning and Staining

	- 2. Chloroform.
	- 3. RNA later (e.g., Thermo Fisher)/liquid nitrogen.
	- 4. Isopropanol.
	- 5. RNase-free water.
	- 6. 75% EtOH in dH2O.
	- 7. Handheld homogenizer (Pellet Pestle Motor).
	- 8. Pellet pestles.
	- 9. Refrigerated centrifuge.
	- 10. Microvolume spectrophotometer (e.g., NanoDrop, Thermo Fisher Scientific). Agilent 2100 Bioanalyzer System.

#### 3 Methods

3.1 Starfish Collection and Husbandry

	- 2. Amputate one arm proximally at one third of the arm's length with a razor blade (Fig. 2a).
	- 3. Amputate the isolated arm a second time distally at the level of two third of the arm within the next 5 min (Fig. 2a).
	- 4. Repeat steps 2 and 3 to double amputate all four other arms of the starfish.
	- 5. Collect the five "central" arm segments (the double-amputated arm explants), about 2 cm long.
	- 6. Place the arm explants in properly labeled glass containers filled with ASW (Fig. 2b).

Fig. 2 Starfish arm explant preparation, collection and fixation. (a) Top view (x–z) scheme of E. sepositus (aboral side), in which the disc and five slender arms are visible. Using a blade, the central third of each arm is traumatically amputated proximally (P) and distally (D) to obtain five arm explants approximately 2 cm long. For clarity, the amputation of only a single arm is shown. (b) After amputation, arm explants are placed back in the aquaria and left to regenerate for the prefixed time-points. They are then collected from the aquaria and placed in glass containers filled with ASW until they are completely "relaxed" (neither curled nor twisted). (c) Top view (x–z) scheme of a "relaxed" regenerating arm explant sectioned using a blade in three portions: proximal, central, and distal. The proximal and distal portions (smaller than the central one) of each explant are fixed and will be processed according to TEM protocols, whereas the central portion can be discharged. Abbreviations: D, distal end; P, proximal end. Dashed lines ¼ amputation planes


3.3 Fixation for Transmission Electron Microscopy (TEM)

From now on, always wear gloves and a lab coat and work under a fume hood. All solutions should be carefully and gently transferred using clean disposable glass pipettes. All steps should be performed on an orbital shaker (gentle shaking) to facilitate solution penetration and washes. Prepare glass containers with proper labeling for each sample. Before starting, heat the oven at 65 C. Troubles possibly arising during protocol performance and corresponding troubleshooting are listed in Table 1.

#### Table summarizing common problems arising during microscopy protocol and corresponding troubleshooting


(continued)



(continued)

#### Table 1 (continued)



#### Table 1 (continued)


Fig. 4 Example pictures. Light microscopy images (a–f, i, j) and TEM micrographs (g, h, k, l) showing examples of satisfactory and unsatisfactory results in terms of sample fixation and decalcification, and semithin and ultrathin sectioning and staining. (a) Semithin section showing satisfactory sample fixation. All tissues are well preserved. (b) Semithin section showing unsatisfactory sample fixation. Artifacts are visible as lacunae and spaces in the tissues (arrows) as well as epithelium detachment (asterisk). (c) Semithin section showing satisfactory sample decalcification. The skeletal tissues of a spine, that is, trabeculae are well preserved and no signs of calcium carbonate crystals are visible. (d) Semithin section showing unsatisfactory sample decalcification. Skeletal tissues of the ossicles (asterisks) are not well preserved and tissue integrity is therefore lost. (e) Semithin section showing satisfactory semithin sectioning. All tissues are well preserved. (f) Semithin section showing unsatisfactory semithin sectioning. Vertical lines (arrows) are artifacts due to the sectioning, in particular, to the damaged glass knife edge. (g) TEM micrograph showing satisfactory ultrathin sectioning. All tissues are well preserved. (h) TEM micrograph showing unsatisfactory ultrathin sectioning. Lines (arrows) and holes (asterisks) are artifacts due to the sectioning, in particular, to the damaged glass knife edge. (i) Semithin section showing satisfactory semithin staining. The difference between diversely stained tissues is well visible and different tissue identification is therefore easy and clear. (j) Semithin section showing unsatisfactory semithin staining. Basic fuchsin staining (arrows) is not clearly distinguishable from crystal violet staining. Therefore, tissue identification can be more difficult or even wrong. (k) TEM micrograph showing satisfactory ultrathin staining. Contrast between electron-dense and electron-lucent cellular elements is well defined. (l) TEM micrograph showing unsatisfactory ultrathin staining. Low contrast between electron-dense and electron-lucent portions make it difficult to distinguish among different tissues/cellular elements



drops, prepared using a 1 mL syringe (Fig. 3c).

Fig. 3 Procedures to follow for semithin and ultrathin sectioning and semithin and ultrathin section collection. (a) Frontal view (x–y) scheme of the resin block with the embedded sample (black) tightly fixed in the ultramicrotome sample holder, which is positioned in the ultramicrotome arm. (A') Top view (x–z) image of a resin block with an embedded sample of starfish arm explant. The embedded sample appears black due to postfixation in osmium tetroxide. (b) Lateral view (y–z) scheme of the resin block with the embedded sample (black) positioned in the ultramicrotome arm and of the glass knife (with the plastic knife boat) fixed on the ultramicrotome knife holder. Each time the ultramicrotome arm goes up and down (black arrow on the left), a section (1 μm for semithin sections and 50–90 nm for ultrathin sections) is sectioned by the glass knife and floats onto the liquid within the plastic knife boat, which contains autoclaved filtered dH2O for semithin sections or 20% EtOH in autoclaved filtered dH2O for ultrathin sections. (c) Semithin section collection. Top view (x–z) scheme of the glass knife with a plastic knife boat; two semithin sections are floating onto the autoclaved filtered dH2O inside. Using a disposable glass pipette with a rounded tip, semithin sections can be transferred into autoclaved filtered dH2O drops on a labeled glass slide (top view; x–z). (d) Ultrathin section collection. Top view (x–z) scheme of the glass knife with a plastic knife boat; six ultrathin sections are floating onto the 20% EtOH in autoclaved filtered dH2O inside. Ultrathin sections are much smaller than semithin sections. Using a copper circle mounted on a disposable glass pipette tip, ultrathin sections can be collected from above (maintaining the copper circle parallel to the liquid surface) and positioned on the copper TEM grid (top view; x–z)


From now on, always wear gloves and a lab coat.


3.5 Ultrathin Sectioning


From now on, always wear gloves and a lab coat and work under a fume hood for the ultrathin section staining.


3.6 Ultrathin Section Staining and TEM Grid Carbon-Coating

Fig. 5 Ultrathin section staining procedure. (a) Top view (x–z) scheme of TEM grid staining. TEM grids are gently placed in droplets of staining solution using tweezers. The TEM grid position is accurately labeled. When exposed to 1% uranyl acetate, the glass petri dish must be covered with aluminum foil and must not contain NaOH pellets. The latter are needed only in the lead citrate step. (b) Top view (x–z) scheme of TEM grid-washing steps after staining. TEM grids are gently washed in subsequent autoclaved filtered dH2O droplets, using tweezers (arrows) to completely remove traces of staining solution (1% uranyl acetate or lead citrate). During all steps, ultrathin sections face the liquid. The droplets of various solutions are prepared on a glass covered with Parafilm®, using 1 mL syringes with a 0.2 μm filters. The TEM grid position is accurately labeled


#### 3.7 RNA Extraction The following protocol has been successfully employed to perform RNA extraction from regenerating arm tips of both asteroids and ophiuroids. It has been performed on E. sepositus regenerating arm explants, with the same success (see Note 31). The extraction protocol final aim has been either the cloning of fragments by

conventional PCR [21] or obtaining RNA of high quality for transcriptome analysis [22]. From now on, always wear gloves and a lab coat and work under a fume hood.


Fig. 6 Sample electropherogram of RNA extract from regenerating tissue of E. sepositus. A total RNA sample is analyzed on the Agilent 2100 Bioanalyzer System using the Eukaryote Total RNA Nano assay. RIN (RNA Integrity Number) software algorithm allows for the quality determination of eukaryotic total RNAs, based on a numbering system from 1 to 10, with 1 being the most degraded profile and 10 being the most intact

> 24. Synthesize cDNA following standard procedures for downstream experiments, that is, transcriptome analysis or RNA probe synthesis for in situ hybridization. Samples with highest RIN values should be selected, as they are of the highest quality.

#### 4 Notes

1. ASW should be prepared 2–3 weeks before introducing animals into the aquaria and left to run in aquaria that are completely equipped with all filters. This time is necessary to stabilize the chemical/physical parameters and allow the growth of a sufficient population of nitrifying bacteria. To facilitate this process, the researcher can add commercially available bacteria useful for both initial set up of the aquarium and long-term maintenance and partial water renewal. In the latter case, ASW should be prepared 2–3 days in advance before adding it to the system.


with 70% EtOH should be tightly closed with Parafilm® to prevent EtOH evaporation.


RIN (RNA integrity number) values. RINs are calculated based on the mobility run of an RNA sample through a capillary electrophoresis.

#### Acknowledgments

The authors are grateful to Greta Valoti for providing the image of Echinaster sepositus double-amputated arm explants. The laboratory of Pedro Martinez is supported with a Grant from "Agencia Estatal de Investigacio´n, Spain" (PGC2018-094173-B-I00).

#### References


echinoderm arm regeneration. Biochem Genet 52(3–4):166–180

22. Gabre JL, Martinez P, Sko¨ld HN, Ortega-Martinez O, Abril JF (2015) The coelomic epithelium transcriptome from a clonal sea star, Coscinasterias muricata. Mar Genomics 3:245–248

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 15

# Studying Hemichordata WBR Using Ptychodera flava

#### Asuka Arimoto and Kuni Tagawa

#### Abstract

Hemichordates are benthic marine invertebrates closely related to chordates. Several species, including Ptychodera flava in the phylum Hemichordates, can undergo whole body regeneration from a small fragment. P. flava is widely distributed in the warm Indo-Pacific region and is easily collected in the lower tidal zone of a shallow beach with a coral reef. Here, we describe the methods for animal collection and preparation of regenerating tissues. The prepared tissues can be used for various molecular and/or histological experiments. We also demonstrate how to examine gene expression patterns in the tissues using whole mount in situ hybridization.

Key words Hemichordates, Gene expression, Whole mount in situ hybridization, DIG-labeled RNA probes, Preabsorbed antibodies, NBT/BCIP staining

#### 1 Introduction

Hemichordates, which are commonly known as acorn worms or pterobranchs, are benthic marine invertebrates. These animals belong to deuterostome and show morphological similarities to chordates such as branchial gills. Moreover, the orthology of some of these features is supported by gene expression and whole-genome analyses [1, 2]. In contrast to the limited capability of regeneration in solitary chordates, hemichordates can undergo whole body regeneration from a fragment of their body [3]. The regeneration of acorn worms tends to occur in anterior-posterior direction rather than other body axes, especially, two complete individuals are formed through regeneration if the body is split into two pieces behind the hepatic region. Although regeneration is observed in many hemichordates, asexual reproduction through the regenerative process or regeneration from a small piece occurs only in a few species [4, 5]. In addition, although acorn worms are found in the sea floor of various environments, only a few species can be easily collected. By combining these two advantages,

Fig. 1 External morphology of Ptychodera flava. This figure shows the dorsal view of the animal and left is the anterior tip. The body consists of three parts, proboscis, collar and trunk. The posterior end of trunk is terminated at the anus. The genital wings swell up in the reproductive season. The hepatic sacs are small projections in dark brown

Ptychodera flava is thus a suitable species for the study of regeneration (Fig. 1).

Here, we demonstrate how to collect the animal and prepare regenerating tissues in the laboratory. P. flava is widely distributed in the Indo-Pacific region, and some populations have been reported in the temperate zone [6]. This species generally lives just under a sand flat in the intertidal zone of the coral reef with a relatively high population density [7]. These ecological characteristics allow easy access to the habitats and efficient collection of the worms. We also describe the techniques to handle the fragile worms. Although the body is very fragile, the small body size of P. flava reduces the difficulties of avoiding animal damage during collection on the beach. P. flava can be kept without any special aquarium equipment during regeneration. Physiological tolerance allows a high-density system for preparing regenerating tissues at a low cost.

Several hemichordates, including P. flava, have been used for studies using molecular techniques [8–11]. Extensive studies of gene expression patterns using embryonic and/or larval specimens have helped elucidate the mechanisms of animal evolution and development. However, visualization of gene expression in adult tissues is still challenging. In this chapter, we also describe the method of whole mount in situ hybridization of regenerating tissues. This method displays gene expression patterns without ambiguous staining.


#### 2.4 Whole Mount In Situ Hybridization


#### 3 Methods


Fig. 2 Comparison of the habitat and the sampling methods of Ptychodera flava. (a, b) Animal collection on a tidal flat in Okinawa, Japan. A trowel is used to dig the sand. The animals in the dug sand are exposed by addition of seawater. (c, d) The animals were collected by snorkeling in shallow water in Hawaii, USA. In this case, the sand was dug using a vigorous wave of the hand to find the animals


3.2 Preparation of Regenerating Tissues 1. Select undamaged individuals (see Note 9). 2. Remove the filmy mucus attached on the surface of the animals using tweezers (see Note 10). 3. Replace seawater in the dish with 30 mL of anesthetic seawater (see Note 11). 4. Incubate at 25 C for 10 min (see Note 12). 5. Cut the body into two pieces using iris scissors (see Fig. 4a and Note 13). 6. Transfer each piece immediately to a new 9 cm diameter plastic petri dish filled with 30 mL of FSW (see Note 14).

Fig. 3 Animal treatment and maintenance at the laboratory. (a, b) The tangled animals in the plastic bag after transportation and a dissociated individual in a pipette. (c) The sand with filmy mucus attached on the surface of the animal was removed using tweezers and a toothpick. (d) A knot made in the animal body was loosened by toothpicks

Fig. 4 Preparation of regenerating tissues of Ptychodera flava. (a) An undamaged animal was cut into two pieces using iris scissors behind the hepatic sacs. The sand which are indicated arrowheads remaining in the intestine can be confirmed from the outside of the body as in Panel (b). (c) Collection of a regenerating tissue using iris scissors. The animal was not treated with anesthetic seawater. The arrowhead indicates the regenerating proboscis


Fig. 5 Anterior regeneration process of Ptychodera flava. (a) An undamaged individual before amputation. (b) A posterior piece just after amputation. (c) The wound is healing at 2 days postamputation (dpa). (d) The completion of wound healing at 3 dpa. (e) The regenerating tissue called blastema becomes visible at 4 dpa. (f) Two rudiments of collar are swelled on both lateral sides of the blastema at 5 dpa. (g) The collar rudiment surrounds the most prominent mass of the blastema at 7 dpa. The mouth opens at the ventral region between the regenerating proboscis and collar. (h) Complete function of proboscis and collar are recovered at 12 dpa. (i) At 17 dpa, the missing branchial region becomes visible. The process of gill regeneration continues for approximately 2 months. (Reprinted by permission from the Zoological Society of Japan: Zoological Science, Regeneration in the Hemichordate Ptychodera flava, Humphreys et al., 2010)


3.3 Preparation of RNA Probes The method for preparation of digoxigenin-labeled RNA probes is modified from [12].


3.4 Preparation of Preabsorbed Antibodies

	- 1. Chill a pestle and mortar using liquid nitrogen.
	- 2. Put a whole body of adult P. flava in the chilled mortar (see Note 23).
	- 3. Immediately add liquid nitrogen to the mortar.
	- 4. Grind the frozen sample to a fine powder using the pestle and mortar.
	- 5. Transfer the powder to a clean, prechilled 50 mL plastic tube.
	- 6. Add four volumes of the powder of prechilled acetone.
	- 7. Incubate on ice for 30 min.
	- 8. Centrifuge the sample at 10,000 rcf at 4 C for 5 min.
	- 9. Discard the supernatant.

3.5 Whole Mount In Situ Hybridization


Fig. 6 Examples of whole mount in situ hybridization using regenerating tissues of Ptychodera flava. Gene expressions of soxb1 at 7 days postamputation were examined using an antisense probe (a) and a sense probe (b), respectively. The upper side is dorsal. The arrowheads show regenerating proboscis in each panel. The gene expressions of soxb1 were identified at regenerating rudiments of proboscis and collars located at the tip of anterior and surrounding region of them


#### 4 Notes

1. The animals are commonly found in the low tide zone of a shallow beach with a coral reef (Fig. 2). Small individuals that are suitable for preparing regenerating tissues prefer a place where fine coral sand accumulates. These animals are abundant in sand less than 10 cm in depth and are rarely found in an anaerobic bottom layer. We generally collect small individuals with a 2–4 mm diameter of the trunk. Prominent fecal castings are accompanied by burrow systems of some large-sized acorn worms; however, P. flava does not form such a structure. The animals can also be collected in high tide conditions by snorkeling (Fig. 2c, d and [13]). Key points of species identification are shapes and colors of proboscis, genital wings and hepatic sacs. Any closely related species has not been reported in habitats of P. flava [6].


#### Acknowledgement

We wish to express our sincere thanks to the Zoological Society of Japan for allowing us to reproduce their data as Fig. 5 in this chapter. Special thanks to Prof. Nori Satoh for his continued support of our research.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 16

# Studying Tunicata WBR Using Botrylloides anceps

# Arzu Karahan , Esra O¨ ztu¨rk, Berivan Temiz, and Simon Blanchoud

#### Abstract

Tunicates are marine filter-feeding invertebrates that can be found worldwide and which are the closest phylogenetic group to the vertebrates (Craniata). Of particular interest, colonial tunicates are the only known chordates that can undergo Whole-Body Regeneration (WBR) via vascular budding. In Botrylloides anceps, a fully functional adult regenerates from a fragment of the vascular system in around 2 weeks after amputation. In this chapter, we present protocols to collect B. anceps colonies, confirm their species, breed them in the lab, monitor WBR and perform histological staining on cryosections.

Key words Whole-body regeneration, Botrylloides anceps, Vascular budding, DNA barcoding, Chordate, Histological section

#### 1 Introduction

Tunicates are filter-feeding invertebrates that have colonized virtually all marine habitats. Although they were classified in the Mollusca phylum during the early twentieth century, the Tunicata subphylum belong to the Chordata and is the closest phylogenetic group to the vertebrates (Craniata) [1]. Consequently, and despite their apparently simpler body morphologies, tunicates display all chordate features (notochord, post-anal tail, endostyle, neural tube and gill slits) as well as a relatively high tissue complexity (heart, neural ganglion, tunic, circulatory system) [2]. Tunicates is a very diverse group of animals that displays quite different reproductive features, repair abilities, development strategies, and life cycles [3– 5]. The majority of tunicates are sessile hermaphrodites that reproduce through a motile tadpole larval stage. After a short freeswimming life stage [2], the tadpole settles on a substrate using the adhesive papillae located at tip of its head. It undergoes a rapid metamorphosis during which its tail and notochord are resorbed, its organs mature and filter-feeding starts. Water enters through the oral siphon, is filtered by the pharyngeal basket and is evacuated through the atrial siphons [6, 7].

In addition to sexual reproduction, a number of tunicates can reproduce asexually by budding in a process termed blastogenesis (reviewed in [8]). The adult animal, called zooid, starts the development of its daughter, called bud, by the thickening of its epithelium together with that of the underlying layer of tissue. The location of the bud and thus the origin of the underlying tissue can vary depending on the species [8]. These tissue invaginate until forming a double vesicle stage common to all types of asexual reproduction in tunicates. The inner layer will then further invaginate to produce the various organs and the whole bud will mature until it becomes a filter-feeding zooid. In some species of budding tunicates, buds remain connected to its zooid, typically through an interzooidic vascular system, thus forming colonies. In some colonial tunicates, in particular among members of the Botrylloides and Botryllus sister genera, blastogenesis is a highly synchronized process where the new generation of buds matures simultaneously throughout the colony to replace the old zooids that get resorbed during the so-called takeover stage.

In botryllid tunicates, researchers have identified a second nonembryonic development that can lead to the formation of zooids. Botryllids can undergo whole-body regeneration (WBR) from a fragment of their interzooidic vascular system [4, 9]. Most notably, this is the only know occurrence of WBR in the Chordata phylum. WBR is a type of vascular budding, which is initiate by an injury that leads to the loss of all zooids and buds from the colony [9–11]. Many of the up-regulated metabolic pathways during the WBR play a crucial role in stem cell maintenance, proliferation, differentiation, and tissue organization [12–15]. Pluripotent stem cells (potentially undifferentiated hemoblasts) are assumed to be the precursor cells for WBR [4, 9, 16, 17]. It has been reported that regeneration in Botrylloides leachii is initiated by the activation of population of dormant stem-like cells that line the surface of the vascular epithelium [14, 16, 18]. In Botrylloides diegensis, a population of Integrin alpha 6 positive circulating stem cells have been shown to be the source of the WBR capacity [19]. In both cases, upon activation, these cells migrate to the vessel lumen where regeneration begins, and these precursor cells differentiate and eventually transform into a single adult individual within the regeneration lumen [9, 16, 17, 20]. The epithelial layer close to the wound area is generally thought to be the origin of the activation source for regeneration [13, 21].

Vascular budding has also been reported in Botrylloides gascoi and Botrylloides leachii under field conditions when colonies recover from their aestivation during which all zooids are lost [22, 23]. Interestingly, vascular budding is a part of the life history of Botryllus primigenus where it happens spontaneously throughout its adult life cycle [24–26]. Altogether, botryllid ascidians display the rare feature of using three distinct developmental pathways to produce the same final organism. This property is of particular interest for comparative developmental studies. Moreover, botryllid ascidians are used as model organisms in a wide range of studies including apoptosis, immunobiology, allorecognition and angiogenesis [4, 18, 20, 27–32]. These animals are thus highly suitable as research specimens, their usage will be widely popularized in the near future.

To promote these exciting organisms, we here present a number of protocols for the study of colonial tunicates that we developed for Botrylloides anceps. The species originates from the Pacific Ocean. It was recorded for the first time in the Mediterranean Sea along the coastline of Israel in 2009, most likely after an opportunistic migration through the Suez channel [7]. More recently, we logged this species on Turkish coasts in 2018. In this chapter, we present protocols to collect, identify, induce WBR and study the regenerative process using histological staining of cryosections. These protocols should be readily applicable to other botryllids, and likely to other sessile colonial tunicates as well.

#### 2 Materials

2.1 Animals Collection, Handling, Feeding, and DNA Barcoding

All solutions should be prepared with ultrapure water and stored at room temperature unless otherwise state.


1. Cryostat microtome sectioning machine.


2.2 Histological Cryosectioning and Staining


#### 3 Methods

#### 3.1 Sampling and Adaptation Botrylloides anceps colonies can be found in stony areas of the intertidal zone, less than 1 m deep. So far, we collected this species from three different stations (Konacık-Iskenderun, Mezitli-Mersin, and Alanya-Antalya) but we have found other suitable colonial ascidians in different sites of the North-eastern Mediterranean coastlines (Fig. 1a, see Note 3).


Fig. 1 Collecting wild botryllids. (a) The map shows the sampling locations from the southern part of Turkey. Botrylloides were observed in all four location, but B. anceps could not be found in Kızkalesi-Mersin. (b) A picture from the Kızkalesi station. (c) A picture of a complete setup with an inhabited rock, a fragment transferred on a glass slide and a tube with a sample for DNA barcoding. (d) Attaching a sample on a slide. (e) Magnification of a B. anceps colony secured on a slide by using fine cotton thread


#### 3.2 DNA Barcoding Total DNA isolation uses handmade buffers adapted from a previously published protocol [33].


Fig. 2 Colony maintenance. (a) A B. anceps husbandry setup in our aquaculture room. (b) A side-view of a staining rack with botryllid colonies in it. (c) A top view of a Botrylloides anceps colony. Scale bar is 1 cm. (d) Magnification of a part of a system during the "takeover" (main zooids are being resorbed while the primary buds replace them). a ampulla, bv blood vessel, pb primary bud, sb secondary bud, Z zooid



the blastogenic stages (Fig. 2d, see Note 28).

Fig. 3 Whole-body regeneration. (a) A B. anceps colony before removing zooids and buds. Scale bar is 1 mm. (b) The same colony after removal all the zooids and buds. (c) Fourth day post ablation. (d) Magnification of a portion of the regenerating colony where a regeneration niche is developing (arrow). Scale bar is 200 μm. (e) Fully completed WBR (day 13th). (f) The frozen mold (blue arrow) attached to the cryostat for sectioning. (g) A section of B. anceps colony at 1.2 magnification


#### 3.5 Histological Sectioning Although the most common protocols for histology use paraffin embedding, we prefer to use cryosectioning for its quick turnaround time.


#### 3.6 Hematoxylin-Eosin Stain


#### 4 Notes


other invertebrates (mostly sponge) and other botryllids (e.g., Botryllus schlosseri, Botrylloides aff. leachii, Didemnum sp.). All of them can be collected and kept in the aquaculture room using the same protocol.



#### Table 1 Staging method of the blastogenic cycle

and submit it in a museum with a unique voucher ID. Load this ID to NCBI or the BOLD system when you upload your barcodes.



#### Table 2 The whole-body regeneration stages of B. anceps

(m minute, h hours)


sure that the blade is secured and the sample is blocked before you start moving it.


#### Acknowledgments

The study was supported by the YO¨ P-701-2018-2666 project (Middle East Technical University support program) and by the MARISTEM European COST Action (CA16203). The experiments were done in the IMS-METU, DEKOSIM laboratory (BAP-08-11-DPT2012K120880-Turkey) and the Blanchoud Group laboratory, University of Fribourg (Switzerland). We are grateful to Prof. Rinkevich (Israel Oceanographic and Limnologic research center) and his research group; Dr. Amalia Rosner, Dr. Jacob Douek, Dr. Ziva Lapidot, and Mr. Guy Paz for the original protocols adapted in this chapter. We appreciate the support of the Erdemli Municipality in protecting the Kızkalesi sampling area.

#### References


https://doi.org/10.1080/24701394.2017. 1404047


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part III

Cellular Approaches

# Chapter 17

# In Situ Hybridization to Identify Stem Cells in the Freshwater Sponge Ephydatia fluviatilis

### Chiaki Kojima and Noriko Funayama

#### Abstract

Sponges (Porifera) are a large phylum that includes an enormous number of species. They are classified into four classes. Among these four classes, class Demospongiae is the largest and contains more than 90% of sponge species. In the last decade, methodologies for molecular studies and sequencing resources in sponge biology have dramatically advanced and made it possible to clearly define particular types of cells based on the genes they are expressing. Here we describe in detail the method of high-resolution WISH (whole mount in situ hybridization) and dual color fluorescent detection of in situ hybridization (dual color FISH) that we have established to detect particular types of cells, especially their stem cells known as archeocytes, in juveniles of freshwater demosponge, E. fluviatilis.

Key words Stem cells, Piwi, Musashi, Porifera, WISH, FISH, Archeocyte

#### 1 Introduction

Molecular studies of sponges are important for both evolutionary developmental biology (since sponges are one of the earliest branching metazoan phyla) and ecological developmental biology (since sponges are sessile organisms). Furthermore, many sponges have high regenerative ability and thus potentially have totipotent/ pluripotent stem cells. Uncovering the cellular and molecular bases of sponge stem cells will not only be crucial for understanding the ancestral gene repertoire of animal stem cells, but will also give us clues for understanding the evolution of molecular mechanisms for maintaining multipotency (pluripotency) and for elucidating the regulatory mechanisms of their differentiation.

Molecular and cellular studies in juveniles of the freshwater Ephydatia fluviatilis suggested that demosponges, which contain more than 90% of all sponge species, have two types of stem cells: mesenchymal Archaeocytes/Archeocytes and food-entrapping Choanocytes [1–4]. Recent studies of sponges in other classes, suggest that this model could be generalized at least in three classes

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_17, © The Author(s) 2022

of sponges, demosponges, calcareous sponges, and homoscleromorpha [4]. However, the type of cells (archaeocytes or choanocytes) that acts as stem cells seems variable, depending on the cellular organization of each class of sponges. Traditionally, both "archeocytes" and "archaeocytes" are used as terms meaning amoeboid cells that contain a large nucleus with a large nucleolus, and are capable of phagocytosis [5, 6]. These cells are suggested to be totipotent somatic stem cells based on microscopic analysis. They are suggested to produce both somatic differentiated cells and gametes [5–7], just like the multipotent stem cells "interstitial stem cells" in hydra, and "neoblasts" in planarians.

In situ hybridization enables the detection of mRNA and thus it is a powerful tool to characterize cells expressing a particular gene, or to identify specific types of cells that express a particular gene. Actually, by the establishment of the methods of WISH and FISH with high resolution ([8, 9] respectively), together with the identification of cell-type specific genes [8–13], cells with morphological features of Archaeocytes/Archeocytes were defined as at least multipotent stem cells that can undergo self-renewal and directly differentiate into multiple types of cells [1]. Thus, EflPiwiA-, EflPiwiB-, EflMusashiA-expressing cells have been defined as at least multipotent stem cells in demosponges [1–4, 11, 12]. It has also been suggested on the basis of their gene expression and microscopic analysis that these cells are in fact totipotent stem cells.

Here we describe in detail the method of WISH and dual color fluorescent detection of in situ hybridization (dual color FISH) that we have established to detect particular types of cells, especially stem cells, in juveniles of freshwater demosponge, E. fluviatilis.

#### 2 Materials

Prepare all solutions using ultrapure water (deionized water) or RNase-free deionized water (when necessary) and analytical grade reagents.


#### 3 Methods

3.1 Preparation of Sponge Samples

	- 2. Fix the animal overnight at 4 C.
	- 3. Replace the fixative with 1/4 HS.
	- 4. Gently shake plate at 4 C for 30 min.
	- 5. Replace the solution with ice-cold 50% MetOH.
	- 6. Gently shake plate at 4 C for 30 min.
	- 7. Replace the solution with ice-cold 100% MetOH.
	- 8. Store the sample at 30 C.



Day 3


3.4 Dual-Color Fluorescent Whole Mount In Situ Hybridization


#### 4 Notes


PCR DNA fragments should be purified by agarose gel electrophoresis followed by extraction from the gel to eliminate the template circular DNA that was used for PCR. Alternatively, plasmid DNA (GmATC methylated DNA) in the reaction mixture of PCR can be specifically digested by DpnI (e.g., Takara Bio), and then PCR fragments can be column purified using a Gene gel/PCR Extraction kit (e.g., Nippon Genetics). Do not use the PCR reaction solution without such purification, because contamination by the RNA that was synthesized using the PCR template could cause nonspecific signals in WISH or FISH.


Fig. 1 Evaluation of the detection efficiency of the combinations of nucleotide analogue and peroxidase-conjugated antigen, or streptavidin. After RNA synthesis, the reaction mixture was dot blotted on a nitrocellulose membrane at dilutions from 1 to 10<sup>5</sup> , and then heated at 80 C for 2 h, and rinsed with Buffer I. After blocking using 1% blocking reagent in Buffer I for 30 min at RT, RNA probes were fluorescently detected using TSA. Note that the combination of biotin-streptavidin gave a stronger detection signal than the combination of FITC-RNA probe and anti-FITC-HRP

Fig. 2 Fluorescent detection of biotin-RNA probe by streptavidin-HRP in FISH. Our previous studies suggested that the expression of EflSlicateinM1 in sclerocytes is much higher than that of EflMusashiA in archaeocytes (probably more than 10 times). Thus, EflSlicateinM1was used as a positive control for FISH. As shown in the dot blot analysis shown in Fig. 1, biotin-RNA probe detected with streptavidin-HRP did not have high background signals, and the sensitivity of detecting specific signals was as high as the sensitivity using a DIG biotin-RNA probe with anti-DIG-HRP for FISH

> and streptavidin-HRP combination can give high background signals in several freshwater organisms such as planarians (personal communication), that was not the case in juveniles of E. fluviatilis, and EflMusashiA-expressing archeocytes and Efl-SilicateinM1-expressing sclerocytes could be specifically detected (Fig. 2). Thus, recently we are using the combination of biotin-labeled RNA probe and streptavidin-HRP [3].

#### Acknowledgments

I thank Drs. Simon Blanchoud and Brigitte Galliot for giving me the opportunity to contribute to this book. I also thank Dr. Elizabeth Nakajima for her proofreading of the manuscript. This work was supported by MEXT/JSPS KAKENHI grants 20H05942, 19H00994, and 17KT0019 to N. F.

#### References


the morphogenesis of siliceous spicules in freshwater sponge: differential mRNA expression of spicule-type-specific silicatein genes in Ephydatia fluviatilis. Dev Dyn 237:3024– 3039


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 18

# Isolation and Maintenance of In Vitro Cell Cultures from the Ctenophore Mnemiopsis leidyi

### Abigail C. Dieter, Lauren E. Vandepas, and William E. Browne

#### Abstract

The ability to isolate, monitor, and examine specific cells of interest enables targeted experimental manipulations that would otherwise be difficult to perform and interpret in the context of the whole organism. In vitro primary cell cultures derived from ctenophores thus serve as an important tool for understanding complex cellular and molecular interactions that take place both within and between various ctenophore cell types. Here we describe methods for reliably generating and maintaining primary cell cultures derived from the lobate ctenophore Mnemiopsis leidyi that can be used for a wide variety of experimental applications.

Key words Ctenophore, Mnemiopsis leidyi, Suspension culture, Serum, Nonbilaterian

#### 1 Introduction

Ctenophora, also known as comb jellies, are gelatinous invertebrates that inhabit marine ecosystems and represent one of the earliest diverging branches of metazoans [1–3]. The unique rotationally symmetric body plan of ctenophores is composed of two germ layers—an outer ectodermal layer and an inner endodermal layer—separated by a thick collagenous mesoglea populated with a variety of cell types, including muscle and motile stellate cells (Fig. 1) [4]. Recent studies in ctenophores have begun to characterize the range of cell types identifiable by both morphological and functional criteria [5–8] as well as gene expression criteria [9].

Across the metazoan tree of life, the extent to which organisms can heal and regenerate varies dramatically. All animals retain some capacity to repair and replace damaged cells, and the ability to restore injured tissues and organs is widespread among metazoan lineages [10]. Ctenophores have remarkable wound healing and regenerative capabilities (Fig. 2) [11–13]. Among ctenophores, Mnemiopsis leidyi has become a model system for understanding a variety of cellular, molecular, and developmental phenomena. Mnemiopsis can regenerate wounded tissues, restore entire organ

Fig. 1 Mnemiopsis leidyi. (a) Adult M. leidyi, axes labelled on the right, oral oriented up. (b) Representative field from primary cell culture 96 h postisolation. (c) Isolated proliferating ectodermal cells. (d) Isolated proliferating endodermal cells. (e) Isolated motile stellate cells. (f) Isolated giant smooth muscle cells

Fig. 2 Mnemiopsis leidyi ectoderm epithelium stained with neutral red vital dye. (a) Area of contiguous ectodermal epithelium prewounding. (b) 2.5 min postwounding with scalpel blade. (c) 12 min postwounding. Arrows indicate sites of cell aggregations along edges of the healing wound

systems, and recover large scale deletions of their body plan via whole-body regeneration (WBR) [10, 14]. WBR encompasses a complex set of context dependent cellular activities that includes wound healing, immune response, signaling, proliferation, and differentiation that ultimately result in tissue growth and reorganization of the affected region [14–18]. The phylogenetic position of the ctenophore lineage suggests that an improved understanding of WBR in ctenophores will offer unique insight into the evolution of metazoan regeneration [13, 14, 19].

Primary cell cultures provide a useful tool for the study of ctenophore cell biology. Reliable methods for generating and maintaining primary cell cultures from the model lobate ctenophore Mnemiopsis leidyi opens the door for assaying cell biological attributes in specific cell types of interest. In this chapter, we detail cell culture techniques for the selection and preparation of cell sources, the preparation of tissue explants, the dissociation of cells for small and large scale preparations, and cell culture maintenance. These protocols provide simple robust techniques to generate in vitro cell cultures (e.g., Fig. 1b) that can be used for a wide variety of downstream applications including live cell imaging, gene expression profiling, pharmacological assays, flow cytometry, and nextgeneration sequencing applications.

#### 2 Materials

Store all solutions at 4 C unless indicated otherwise.


#### 3 Methods


3.4 Mechanical Cell Dissociation for Smallto-Medium Scale Cultures


3.5 Mechanical Cell Dissociation for Large-Scale Cultures

Fig. 3 Filter stack used for cell suspension size selection. (a) Filter stack components. The adapter includes a plug for syringe attachment. Legend at lower right indicates filter screen sizes. (b) Assembled filter stack with descending 300 μm, 100 μm and terminal 70 μm screens for cell size selection. Mounted syringe allows for the application of light suction below the filter stack to "pull" the initial viscous cell homogenate through the filter screens

	- 2. Excise tissue fragments using a razor blade.
	- 3. Transfer the excised tissue to a glass Dounce.
	- 4. Add 500 μL of FSW P/S.
	- 5. Gently homogenize with 10–15 strokes of a loosely fitted pestle.
	- 6. Centrifuge the resulting homogenate for 10 min at 350 rcf at RT to pellet cells.
	- 7. Remove and discard the supernatant.
	- 8. Add 10 mL of dissociation media.
	- 9. Pipet up and down gently to break up cell pellet.
	- 10. Transfer homogenate to a 15 mL tube.

3.7 Primary Cell Culture Maintenance

#### 4 Notes


#### Acknowledgments

This material is based upon work supported by the National Science Foundation under Grant No. 2013692 [W.E.B]; National Research Council Postdoctoral Fellowship [L.E.V.]. University of Miami's College of Arts and Sciences [A.C.D. & W.E.B.].

#### References


Biol 17:16. https://doi.org/10.1186/ s12915-019-0633-9

18. Kassmer SH, Nourizadeh S, De Tomaso AW (2019) Cellular and molecular mechanisms of regeneration in colonial and solitary ascidians. Dev Biol 448(2):271–278. https://doi.org/ 10.1016/j.ydbio.2018.11.021

19. Sánchez Alvarado A, Tsonis PA (2006) Bridging the regeneration gap: genetic insights from diverse animal models. Nat Rev Genet 7(11): 873–884. https://doi.org/10.1038/nrg1923

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Analysis of Spatial Gene Expression at the Cellular Level in Stony Corals

# Nikki Traylor-Knowles and Madison Emery

#### Abstract

Scleractinians, or stony corals, are colonial animals that possess a high regenerative capacity and a highly diverse innate immune system. As such they present the opportunity to investigate the interconnection between regeneration and immunity in a colonial animal. Understanding the relationship between regeneration and immunity in stony corals is of further interest as it has major implications for coral reef health. One method for understanding the role of innate immunity in scleractinian regeneration is in situ hybridization using RNA probes. Here we describe a protocol for in situ hybridization in adult stony corals using a digoxigenin (DIG)-labeled RNA antisense probe which can be utilized to investigate the spatial expression of immune factors during regeneration.

Key words Coral reefs, Coral, Cnidaria, Regeneration, Innate immunity, Wound healing, In situ hybridization

#### 1 Introduction

Scleractinian corals are stony corals that build coral reefs. They are part of Cnidaria, a diverse phylum that possesses over >10,000 known species and is the sister group to Bilateria. It is estimated that these groups split approximately 604–748 million years ago (Fig. 1) [1]. Stony corals are primarily colonial consisting of many clonal polyps that are interconnected through a web of gastrovascular canals [2]. Stony corals are known to possess a high capacity for tissue regeneration which is hypothesized to be driven by stem cell differentiation and proliferation. However, the mechanisms of this regeneration are still not well understood (Fig. 2) [3]. They also possess a highly diversified innate immune system [4, 5]. From the available coral genomes, we understand that many corals possess a high diversity of immune factors which originated both from neo- and subfunctionalization events. This indicates that a complex interaction of immune factors and regenerative factors may be involved in whole body regeneration [6–10].

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_19, © The Author(s) 2022

Fig. 1 Cnidarian phylogenetic tree. Scleractinians, or stony corals, are part of the phylum Cnidaria. This phylum is a diverse primarily marine phylum and is important for evolutionary study due to its placement as the sister group to Bilateria. The split between Bilateria and Cnidaria is estimated to have occurred 604–748 million years ago

Fig. 2 Schematic of whole-body regeneration in a colonial coral. During whole body regeneration of a colonial coral polyp, immune and stem cell factors are upregulated in response to the injury, and initiate regeneration of the body. Communication between the adjacent polyps and the regenerating polyp are critical for whole body regeneration. Stem and immune cells from adjacent polyps are presumed to migrate into the regenerating polyp area. Depending on the coral species and environmental conditions whole polyp regeneration can take 7–30+ days [3]

The underlying functional mechanisms of whole-body regeneration in stony corals is not as well understood as in other cnidarian models such as Nematostella and Hydra [3]. This is due to a historical emphasis within coral rsearch to focus on the heat stress response, challenges of manipulating the porous stony skeleton, and the high amounts of obligate microbial symbioses [11– 13]. But with recent advances in sequencing technologies and cell biological techniques many of these challenges are starting to be addressed and tools to study whole-body regeneration are being developed [14–16]. Many genes and proteins have now been identified as involved in whole-body regeneration and immunity in corals, however the functional mechanisms of many of these genes are not known (Table 1) [3, 6].

One of the ways to investigate the interplay of innate immunity and regeneration is to use the method in situ hybridization (ISH) of RNA probes to assess the spatial gene expression of specific genes of interest. ISH is a very versatile technique because the RNA probes can be designed for any gene that is expressed. This method was first developed for the study of embryogenesis and has been further developed to understand the spatial gene expression across different live stages and stress response of many different organisms [17]. This method can be done by using many different types of probes labeled with nonradioactive nucleotides such as digoxigenin (DIG), fluorophores, or radioactive nucleotides [17–19]. The power of this technique is that it can detect and visualize small amounts of RNA at a cellular level. This is particularly useful for nonmodel organisms where the link between cell types and specific gene expression may not be understood.

To use this technique to study immunity during whole body regeneration, immune stimulation can be performed using synthetic elicitors such as lipopolysaccharides, peptidoglycans or exposure to known pathogens such as Vibrio to target immune and stem cell related genes that may be expressed during regeneration (Table 1) [20–24]. Additionally, no immune stimulator may be necessary if investigating the early process of regeneration, as the early signals of regeneration are early wound healing gene related to innate immunity [25].

In preparation for this method, DIG-labeled RNA antisense probe, and its accompanying sense control probe should already be designed and ready for use. Additionally, serial sections of paraffin embedded tissue should be prepared for testing both the sense and antisense probes. The sense probe is used as a control for nonspecific binding. If the sense probe has positive staining, then it will indicate that your antisense probe is not targeting the intended RNA. DIG-labeled probes are highly sensitive and can be developed from expressed RNAs for many different stressors [26–30]. It can also be applied to a wide range of tissues and organisms. This technique is not new, however, the application of it on adult stony coral tissues is an emerging technique that has promise for understanding the spatial expression of genes associated with whole-body regeneration. While this technique has primarily been used to assess the expression of developmental genes in cnidarians embryos and larvae, it has recently been modified to be used for adult cnidarians including stony corals [29]. In this book chapter we will outline the steps for performing in situ hybridization on stony coral tissue slices


#### Table 1 Summary of immune factors implicated in regeneration assays in corals

to yield cellular level resolution. This procedure could be easily modified for other cnidarians such as Nematostella and Aiptasia.

#### 2 Materials

All solutions and dilutions should be made using molecular grade, RNase-free reagents, equipment and consumables. This procedure is highly sensitive to RNase contamination which can degrade the RNA probe.

#### 2.1 Removal of Paraffin 1. Thin-sectioned paraffin-embedded slides. 2. 100% xylene. 3. Glass Coplin jars (see Note 1).

	- 2. RNA Probe (see Note 4).
	- 3. Hybridization-probe solution: 0.5 μL probe, 24.5 μL hybridization buffer. Prepare just prior to probe hybridization.
	- 4. Heat block set to 86–90 C.
	- 5. PAP pen.
	- 6. Plastic coverslips.
	- 7. Slide moisture chamber.

#### 2.2 Hybridization of RNA Probe


#### 2.3 Visualization of RNA Probe


#### 3 Methods

3.1 Removal of

Paraffin

All manipulations should be done using sterilized equipment and at room temperature, unless otherwise stated. 1. Under a well-ventilated fume hood, pour 50 mL of 100% xylene into a sterile glass Coplin jar.

	- 1. Set your incubator to 37 C and turn on the hot water bath to 100 C.
	- 2. Prepare the prehybe buffer by placing it in a boiling water bath for 15 min.
	- 3. Place the prehybe buffer in an ice bath for 5 min.
	- 4. Turn on your hybridization oven to the hybridization temperature (see Note 6).
	- 5. Add 18 mL of prehybe buffer to a new sterile slide mailer.
	- 6. Warm the slide mailer in the hybridization oven.
	- 7. Remove slides from the 60% (v/v) ethanol incubation using sterile tweezers.
	- 8. Place slides in a sterile slide mailer filled with 18 mL of 1 PBS.
	- 9. Wash for 5 min on an orbital shaker set to 100–150 rpm.

#### 3.3 RNA Probe Hybridization


3.2 Slide Pretreatment and Prehybe Preparation


#### 1. Incubate slides for 1 min in 18 mL of AP-buffer without MgCl2 at room temperature.

2. Block the slides overnight at 4 C on an orbital shaker according to the DIG Nucleic Acid Detection Kit manufacturer's instructions (see Note 11).

3.4 RNA Probe Visualization


#### 4 Notes


Fig. 3 Representative example of in situ hybridization results in the pacific stony coral, Acropora hyacinthus to demonstrate general outcomes of specific and nonspecific binding. (a) This panel shows the staining of Chordin, a marker expressed during cnidarian regeneration [20] in Acropora hyacinthus tissue which has been exposed to a heat stress. The expression the antisense () probe for Chordin was found throughout the gastrodermis, and in gastrodermal cells. The sense (+) control probe showed some staining within the cnidocytes indicating nonspecific binding of cnidocytes. Cnidocytes are indicated by black arrows. (b) This panel shows the staining of Fructose Bisphosphate Aldolase again in Acropora hyacinthus tissue exposed to heat stress. The antisense () probe had primary staining within the cnidocyte cells in the epidermis. Cnidocytes are indicated by black arrows. The sense (+) control had no nonspecific staining indicating that the cnidocyte staining in the antisense () probe was specific


#### Acknowledgments

This work is supported by startup funds provided by University of Miami, Rosenstiel School of Marine and Atmospheric Sciences and by NSF-1951826. The authors would like to thank Traylor-Knowles Lab, as well as thank Bradford Dimos and Emily Buckley for discussions concerning the chapter development.

#### References


https://doi.org/10.1007/s00338-020- 01952-4


heat stress studies. Ecol Evol 9:10055–10066. https://doi.org/10.1002/ece3.5576


Pocillopora damicornis to bacterial stress from Vibrio coralliilyticus. J Exp Biol 214:1533– 1545. https://doi.org/10.1242/jeb.053165


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Studying Stem Cell Biology in Intact and Whole-Body Regenerating Hydra by Flow Cytometry

# Wanda Buzgariu , Jean-Pierre Aubry-Lachainaye, and Brigitte Galliot

#### Abstract

The freshwater Hydra polyp is a versatile model to study whole-body regeneration from a developmental as well as a cellular point of view. The outstanding regenerative capacities of Hydra are based on its three populations of adult stem cells located in the central body column of the animal. There, these three populations, gastrodermal epithelial, epidermal epithelial, and interstitial, continuously cycle in homeostatic conditions, and their activity is locally regulated after mid-gastric bisection. Moreover, they present an unusual cycling behavior with a short G1 phase and a pausing in G2. This particular cell cycle has been studied for a long time with classical microscopic methods. We describe here two flow cytometry methods that provide accurate and reproducible quantitative data to monitor cell cycle regulation in homeostatic and regenerative contexts. We also present a cell sorting procedure based on flow cytometry, whereby stem cells expressing a fluorescent reporter protein in transgenic lines can be enriched for use in applications such as transcriptomic, proteomic, or cell cycle analysis.

Key words Hydra, Cell cycle, Adult stem cells, Epithelial stem cells, Interstitial stem cells, Flow cytometry, GFP cell sorting, Click-iT EdU labeling of regenerating animals, Cell cycle analysis

#### 1 Introduction

1.1 Hydra and the Unusual Properties of Its Stem Cells

The hydrozoan Hydra polyp, which belongs to Cnidaria, is well known for its robust regenerative abilities, thus providing an attractive model system for regenerative biology [1–5]. This freshwater polyp, which is about 1-cm long, reconstructs any missing part of its body, such as the basal (foot) or apical (head) regions, within a few days after amputation. The Hydra body exhibits a radial symmetry (Fig. 1a), with a cylindrical shape terminated at the apical pole by the head region centered on a unique opening called mouth, and at the basal pole by the basal disc, which helps the animal to adhere to various environmental substrates. The anatomy of Hydra is formed of two tissue layers, one epidermal, the other gastrodermal, held together by the mesoglea, a complex extracellular matrix that maintains their cohesion. This double-body wall

Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_20, © The Author(s) 2022

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols,

Fig. 1 Hydra anatomy and cycling properties of Hydra stem cells. (a) The Hydra polyp exhibits a radial symmetry centered on an oral–aboral axis. At the apical pole also called the head, a ring of tentacles surrounds a dome called hypostome that is centered on the mouth opening, while at the basal end called foot, the basal disc that produces mucus allows the animal to attach to substrates. The animal consists of two epithelial layers, the epidermis on the outside consisting of epidermal epithelial stem cells (eESCs), and the gastrodermis on the inside consisting of gastrodermal epithelial stem cells (gESCs). All epithelial cells along the gastric region are ESCs that terminally differentiate when they reach the extremities. Interstitial stem cells (ISCs) are abundant in the central region of the animal, interspersed with eESCs. (b) The cell cycle of ESCs lasts 3–4 days, while multipotent ISC cycle every 24–30 h. In each cycle, ISCs provide asymmetrically divided interstitial progenitors (IPs) that cycle faster than ISCs (less than 24 h). IPs are migratory cells that differentiate in G0 phase along the body axis and at the extremities. As for somatic derivatives, nematocytes are strictly located in the epidermis, nerve cells are found in both layers, and gland cells in the gastrodermis. Fast cycling cells such as ISCs and IPs are predominantly killed by hydroxyurea (HU) pulse treatment. In contrast, ESCs paused in G2 are resistant to such treatments. Passively moving toward the apical and basal poles, ESCs differentiate in G2 phase into head- and foot-specific epithelial cells. Scheme adapted after [6]

houses three populations of non-interchangeable adult stem cells: two populations of unipotent epithelial stem cells (ESCs), either epidermal or gastrodermal, and the multipotent interstitial stem cells (ISCs), which are found predominantly in the central body column [7–10] (Fig. 1a). By contrast, the tissue at both the ends of the animal consists mainly of differentiated cells, either epithelial or derived from ISCs, such as neurons and mechano-sensory cells called nematocytes [11, 12]. In addition, ISCs can also differentiate into glandular cells, distributed among the gastrodermal epithelial cells, and germ cells when the animal becomes sexual and differentiates gonads [9, 13].

Hydra tissue is characterized by a dynamic homeostasis, as stem cells from the central body region from all three lineages are continuously self-renewing, replacing every 20 days the differentiated cells that progressively get sloughed off at the extremities [14]. The length of the cell cycle of these adult stem cells differs between ESCs that divide every 3–4 days and ISCs that progress faster through the cell cycle, dividing every 24–30 h (Fig. 1b) [15, 16]. Intriguingly, all three stem cell populations share quite unusual features, i.e., a very short G1 phase that lasts 1 h, and an extended G2 phase that ranges from 24 up to 72 h for ESCs [6, 15, 17] and from 6 up to 22 h for ISCs [15, 16]. As a consequence, pulse treatments with drugs that block DNA synthesis such as hydroxyurea (HU) preferentially target the fast cycling cells, the ISCs, and their progenitors (Fig. 1b). Typically, three 24-h periods of HU exposure, each separated by a 12-h drug-free period, suffice to selectively deplete the interstitial cell line and, after a few weeks, produce animals that are purely epithelial. These animals, which have lost their nervous system, cannot catch preys yet can survive for months and years if manually force-fed. In such conditions, the epithelial tissues get properly renewed and the animals retain their developmental potential, i.e., are able to bud and regenerate [18, 19].

The continuous self-renewal of Hydra stem cells in the central part of the body column indicates that this region can be considered as a pro-blastema where stem cells paused in G2-phase are ready to differentiate, divide, and proliferate immediately after amputation [20, 21]. Similar to other cnidarians [22–24], proliferating cells play an important role in the regeneration of apical structures, and a synchronous cell division event is actually rapidly induced upon amputation [20, 25]. Seventeen evolutionarily conserved cell cycle genes are then synchronously upregulated (Shox1, E2F7, TFDP1, POLQ, CCNF, PLK4, CCNE1, CCND2, CDC7, SIPA1L3, MCM5, DLEC1, MCM9, CDC6, CCNA2, CCNB3, PLK1) [20], and a local wave of cell proliferation follows at an early-late stage (around 24 h post-amputation) [26]. When the S-phase progression is blocked with HU prior to amputation, it alters apical regeneration, although only partially, as the stock of epithelial cells stopped in G2 can differentiate without entering a final mitosis [21]. In summary, the regenerative capacity of Hydra relies on large stocks of continuously cycling stem cells, whose unusual properties explain their immediate contribution to the regenerative process, which is achieved in few days.

1.2 Stem Cell Sorting and Methods to Monitor the Cell Cycle Activity in Hydra

The cell cycling behavior of Hydra stem cells was intensively studied by classical microscopical methods that either analyze the incorporation of thymidine analogs (e.g., <sup>3</sup> H-thymidine, BrdU 50 -bromo-deoxyuridine) into replicating DNA, or evaluate the DNA content microfluorimetrically, or allow the counting of mitotic figures [16, 17, 27]. Among these methods, the counting of BrdU-labeled nuclei detected with anti-BrdU antibody allows the establishment of a precise BrdU-labeling index [26, 28]. However, these methods are time-consuming, and the quantitative results are obtained on a rather small number of analyzed cells. To overcome these limitations, we have applied to Hydra flow cytometric methods that are commonly used in model organisms such as algae, sea anemones, planarians, flies, as well as in mammalian cells, to address a variety of biological questions [29–33].

Fig. 2 Schematic view of the flow cytometry protocols presented in this chapter. In this study, two distinct protocols were used to dissociate the Hydra tissues, the first one is based on pronase activity and the second on trypsin–EDTA activity. Pronase dissociation is suitable because it provides living, viable Hydra cells that can be sorted by flow cytometry (see Subheading 3.2). The trypsin–EDTA dissociation is fast and convenient to use when rapid dissociation is needed as required to analyze the cell cycle activity (see Subheadings 3.3 and 3.4)

As a general definition, flow cytometry is a laser-based technology that allows the characterization of properties of cells in suspension, i.e., their size, volume, morphological complexity, and fluorescence-labeled components. This technology ensures that a large number of cells are analyzed quantitatively in a very short time. Advances of the flow cytometry-based cell sorting methodology (FACS, fluorescent-activated cell sorting) opened large possibilities for downstream applications such as transcriptomic [34, 35], proteomic [36], biochemical, or cellular [33] analyses. In this context, the production of transgenic lines in Hydra that constitutively express green fluorescent protein (GFP) in one or the other stem cell population, e.g., ecto-GFP [37], endo-GFP [38], or Cnnos1-GFP [39], opened the possibility to study the molecular signatures of each population after cell sorting [39–41]. The two first sections in this chapter describe how to sort Hydra stem cells based on their selective GFP expression (Figs. 2 and 3).

The flow cytometry methodology is also applied in Hydra to measure the modulations of the cell cycle behavior in homeostatic, regenerative, or ecotoxicological contexts [6, 20, 42–44]. A simple and easy method to characterize the distribution of cell populations in the different phases of the cell cycle is to measure the DNA

Fig. 3 Fluorescence-activated cell sorting (FACS) of epidermal ESCs constitutively expressing GFP. (a) Scheme depicting the FACS procedure to obtain live GFP-positive epidermal ESCs (eESCs) from ecto-GFP transgenic polyps [37] (see Subheadings 3.1 and 3.2). (b) Gating strategy: The cells are first gated based on the forward scatter (FSC-H) and side scatter (SSC) properties (gate A). Next, the intact, viable cell population is selected based on the intensity of Draq7 fluorescence, detected on the FL4-Area channel (gate B). Subsequently, the GFP-positive cells are identified considering the GFP fluorescence collected on the FL1 channel (gate C) and sorted after exclusion of cell doublets, based on the area and height of the FSC signal (gate D). (c) Analysis of the enrichment score of the GFP+ eESCs after re-running the sorted cells on the flow cytometer (FACS-2). Note that 96.6% of sorted cells are viable (gate B) and 99.5% are GFP-positive (gate C). The sorting was done with a Biorad cell sorter S3. (d) Microscopical control of the enrichment of the GFP+ eESCs after sorting. Green fluorescence (left panel) and bright field (right panel) images were acquired with a Leica DM5500 fluorescence microscope 3 h after sorting. Arrows point to cells with low GFP fluorescence. Scale bars: 75 μm

content that doubles during S-phase and is divided by twofold at the end of the G2-phase upon mitosis. The DNA content can be assessed quantitatively from staining with propidium iodide (PI), a DNA intercalating dye whose fluorescence is enhanced upon insertion between the bases [45]. This method is widely used to estimate the relative distribution of the cells between the G0/G1, S, or G2/M phases of the cell cycle, as shown in mammalian [46] or invertebrate cells from Drosophila [47], planarians [48], or Daphnia [49]. The DNA staining protocol presented here integrates a fast dissociation step of the tissue with trypsin–EDTA enzymatic digestion, followed by PI DNA labeling in a hypertonic staining solution in the presence of detergents [50]. This method allows the successful analysis of very small tissue fragments and the processing of a large number of samples in parallel while obtaining good quality DNA histograms as shown by the good coefficient of variations (CV) measured across the G0/G1 peak [6] (Fig. 4).

As an alternative, the monitoring of cell proliferation with the click-iT EdU detection assay was developed by Invitrogen. Similar to BrdU, EdU (5-ethynyl-2<sup>0</sup> -deoxyuridine) is a thymidine analog that gets incorporated into the newly synthesized DNA chain during the replication phase. In contrast to the antibody-based BrdU detection, the click-iT reaction consists of the chemical detection of EdU after a 30-min reaction between EdU and an Alexa photostable dye catalyzed by copper [51, 52]. This method, which can provide a dynamic view of the progression of DNA replication during the S-phase when several time-points are compared, is faster and easier than BrdU immunodetection because neither DNA denaturation nor immunodetection are needed. However, in Hydra, the cellular absorption of EdU in intact animals is low when compared to BrdU, and the number of S-phase cells detected with the click-iT EdU proliferation kit is not reliable. During regeneration, the tissues of amputated animals absorb well the thymidine analogue, and the EdU labeling procedure is well suited for monitoring cell proliferation (Fig. 5).

In this chapter, we report two distinct flow cytometry procedures to detect the cell cycle distribution in homeostatic or regenerative contexts. The first procedure analyses nuclei whose DNA is stained with PI after tissue dissociation while the second procedure relies on the pulse labeling of S-phase nuclei with EdU, a DNA-labeling process that takes advantage of the physiological replication process occurring in live animals. The protocols presented here rely on two distinct procedures for tissue dissociation that precedes flow cytometry. The relatively slow pronase dissociation is well suited to sort fluorescent cells by FACS, whereas the fast trypsin–EDTA dissociation is well suited for cell cycle analysis such as PI labeling or EdU click-iT labeling (Fig. 2). As this latter method discriminates between the cells in G1 or G2 from those in early or late S-phase, it provides a dynamic assessment of the

Fig. 4 DNA content measurement in Hydra cells after PI staining. (a) Workflow of the flow cytometry method used to analyze the cell cycle distribution after PI staining of DNA (see Subheading 3.3). (b) Typical successive gating procedure where the events are acquired based on the FSC/SSC parameters, then gated first on the channel that detects the PI fluorescence (gate A), followed by doublets and clumps exclusion after setting up a singlet discrimination PI-Area/PI-Width window (gate B). The cell cycle distribution (right panel) is deduced from the DNA content reflected by the intensity of the PI-Area signals of PI-labeled cells in gate B. (c) Cell cycle distribution in samples obtained after dissociation of whole animals (intact) or from different Hydra regions

number of cells progressing through the S-phase, when samples corresponding to different labeling periods are compared. In contrast, the PI labeling method, which is cheaper and faster, only provides a static view of the DNA content, allowing to deduce the cellular distribution between the different phases of the cell cycle at the time tissues were dissociated. Depending on the context and the experimental objective, these two flow cytometry methods are valuable tools for quantifying the rate of proliferating cells and the cell cycle progression during whole-body Hydra regeneration.

#### 2 Materials

All stocks and working solutions are prepared with ultrapure, deionized MilliQ water in screw-capped bottles, sterilized either by autoclaving or filtration using a 0.22-μm filter. The bottles are stored at 4 -C or room temperature (RT) as indicated below. Check stock solutions regularly for any sign of contamination, discard them, and replace them with fresh ones when needed. To prepare the working solutions (1), dilute any concentrated stock (10 or 20) with ultrapure MilliQ water.

	- 2. Wild-type AEP Hydra strain.
	- 3. 500 Hydra Medium Stock solution A: 60.57 g Tris–base– HCl in 900 mL H2O, pH 7.7, bring to 1000 mL with H2O. Sterilize by autoclaving, store at 4 -C for several weeks.

Fig. 4 (continued) (apical, gastric column, and basal). Note that the broader G0/G1 peak detected in samples from intact animals or apical region corresponds to a double G0/G1 peak formed by two different cell populations. (d) To identify these two distinct G0/G1 cell populations, a specific gating procedure was applied to the samples obtained after dissociation of tissues obtained from the apical region or from the whole animal. An additional gating based on the FSC-Area and PI-Area parameters allows the identification of these two populations as gate R1 and gate R2. Sorting of the nuclei from gate R1 revealed that this population is mainly composed of terminally differentiated nematocytes arrested in the G0 phase (data not shown), which are mainly found in the apical region. (e) Scheme depicting the flow cytometric procedure applied to headregenerating tips (see Subheading 3.3). Animals bisected at the mid-gastric level were allowed to regenerate. At indicated time-points after bisection, the head regenerating tips were excised and prepared for analysis as shown in (a). (f) Cell cycle profiles measured in H. magnipapillata animals undergoing head regeneration. Five regenerating tips about 500 μm long were processed and analyzed as above. Note the increase in S-phase cells between 4 and 6 h post-amputation (hpa), followed by an increase in the G2 population at 8 hpa as previously reported [20]. The samples were run on a Gallios flow cytometer (Beckman Coulter) (b–d) or a BD FACSCalibur (f), and the data were analyzed with the FlowJo software and subjected to Watson's mathematical model to calculate the proportion of cells in each phase of the cell cycle

Fig. 5 Flow cytometry analysis of DNA replication in EdU-labeled Hydra cells. (a, c) Scheme illustrating the workflow of the flow cytometry procedure performed on samples obtained after dissociation of intact (a) or head-regenerating polyps (c) taken 4 h after bisection and incubated in EdU (5 mM) for 3 h. (b, d) To analyze cell proliferation, the cells were labeled with the click-iT EdU-Alexa 647 kit, the samples were run on a Fortessa flow cytometer, and the data collected based on the FSC and SSC signals (gate A). Next, the debris were removed from gate A by applying a second gating based on the PI-Area (PI-A) signal. The singlet cells were separated in a PI-Width (PI-W) and PI-A window, which helps to exclude the doublets (gate C). Finally, the


Fig. 5 (continued) cells selected from gate C were analyzed in a PI/ EdU-Alexa Fluor 647 dot-plot area, where the negative and positive EdU gates are defined by the fluorescence intensity in the EdU-Alexa 647 channel. Thus, cells in S-phase, which have incorporated EdU, form the EdU(+) population, while cells in G0/G1 or G2/M correspond to the EdU() population. Note the lower percentage of EdU(+) cells and the lower fluorescence intensity on the Alexa 647 channel when intact animals (b) were incubated with the EdU solution compared to the regenerating ones (d). This result indicates that EdU incorporation is lower in gastric cells when intact Hydra are exposed to EdU compared to the regenerating ones. Beside a regulation linked to regeneration, this difference might actually be artefactual, reflecting the barrier effect of the Hydra cuticle to EdU in intact animals when compared to the wounded ones. This hypothesis is supported by the fact that the observed proportion of EdU-labeled cells (11%) after a 3-h labeling is lower than that measured by microscopic analysis after BrdU incorporation or by flow cytometry after PI staining method [6, 17, 20, 25, 26]. Indeed, in starved conditions, about 20% of the cells are cycling in gastric tissues from intact animals (see Fig. 4c). (e, f) Comparison between two experimental conditions: one where intact animals were exposed to EdU for 4 h before the central gastric tissue is dissected (e), another where the extremities of the animal are first amputated and then the severed gastric pieces are incubated in EdU for 4 h (f). After washing, the samples from both experiments were treated as described in Subheading 3.4. Analysis of the EdU-Alexa 647/PI-A data graph (a and b, lower panels) shows that the percentage of EdU-Alexa 647-positive cells is 50% higher when the central gastric pieces are amputated before EdU incubation compared to that obtained in gastric pieces amputated after EdU incubation. Note also that prolonging EdU incubation improves the fraction of EdU (+) cells: 17.5% after 4 h vs. 11% after 3 h


2.3 Click-iT EdU-Detection of Proliferating Hydra Cells


#### 2.4 Equipment 1. Stereomicroscope (for example Olympus SZX10 with a 1.25 objective).


#### 3 Methods

Suspension

3.1 Live Hydra Cell

#### 1. Collect 250 GFP-expressing transgenic animals with the help of a glass Pasteur pipette (see Note 13) in a Pyrex dish.


#### Basic terminology used in this chapter related to flow cytometry and cell cycle analysis


#### 3.2 Flow Cytometry Sorting of Hydra GFP-Expressing Stem Cell (FACS Protocol)


3.3 Cell Cycle Measurement: PI Staining of Trypsin-Dissociated Cells


3.4 Cell Cycle Measurement: Click-iT EdU Labeling of Hydra Cells Replicating Their Genomic DNA (S-Phase)


39. Analyze the data with available software and estimate the percentage of EdU-positive cells that correspond to the proliferating population based on the EdU-Alexa 647 fluorescence intensity (Fig. 5b, d–F).

#### 4 Notes


ISCs are much smaller than ESCs, and they show a higher nuclear to cytoplasmic ratio and a better resistance to centrifugation [56]. If ISCs are considered for sorting, an additional centrifugation step at 300 rcf is requested.


tubes or similar tubes available for your sorter. For RNA extraction, a high RNA yield (1 mg) was obtained with the minikit RNAeasy Plus (Qiagen) from 3 <sup>10</sup><sup>5</sup> sorted cells.


peak that corresponds to two different populations of nuclei. Therefore, to avoid misestimation of the S phase, we apply a different gating procedure to analyze the cell cycle profile of samples taken from the apical part of the animal (Fig. 4d), where the CV value did not exceed 3.0.


#### Acknowledgments

This work was supported by the Swiss National Science Foundation (SNF grants 31003A\_149630, 31003\_169930, and 310030\_189122), the Claraz donation, and the Canton of Geneva.

#### References


Dev 77(10):837–855. https://doi.org/10. 1002/mrd.21206


attenuata. I. Homeostatic control of interstitial cell population size. J Cell Sci 20(1):29–46. https://doi.org/10.1007/BF00848421


cytometry for monitoring microbial cells. J Immunol Methods 243(1–2):191–210. https://doi.org/10.1016/s0022-1759(00) 00234-9


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Noninvasive Intravascular Microtransfusion in Colonial Tunicates

## Lluı`s Albert Matas Serrato, Alessandro Bilella, and Simon Blanchoud

#### Abstract

Tunicates are a diverse group of worldwide marine filter-feeders that are vertebrates' closest invertebrate relatives. Colonial tunicates are the only know chordates that have been shown to undergo whole-body regeneration (WBR). Botrylloides in particular can regenerate one fully functional adult from a minute fragment of their vascular system in as little as 10 days. This regenerative process relies on the proliferation of circulating stem cells, likely supported by the activity of some of the 11 identified types of hemocytes. To study and challenge WBR, it is thus important to have the capacity to isolate, analyze, and manipulate hemolymph in regenerating colonies. Here we present a microtransfusion technique that permits the collection of pure hemocytes, the quantification of their purity, their labeling, and reinjection into colonial tunicates. To exemplify our approach, we present in addition a protocol to analyze the isolated hemocytes using flow cytometry. Our approach is minimally invasive, does not induce lethality, and therefore allows repeated transfusion into exactly the same colony with minimal disruption to the process being studied.

Key words Hemolymph, Transfusion, Colonial ascidians, Botrylloides, Tunicates

#### 1 Introduction

Tunicates are a group of worldwide highly diverse marine invertebrates separated into three classes that include over 3000 known extant species [1, 2]. Most tunicates are benthic sessile animals that reproduce sexually through a motile tadpole larval stage [3, 4]. In addition, a number of tunicate species are able to reproduce asexually through budding [5, 6]. Budding in tunicates typically leads to the formation of colonies where animals, called zooids, are interconnected by an external vascular system (Fig. 1a). Furthermore, some species of colonial tunicates are able to undergo whole-body regeneration (WBR) [7–9]. This is the only known occurrence of WBR among chordates [10].

In the Botrylloides genus, WBR is completed in as little as 10 days following the isolation of a fragment of their external vascular system [11]. WBR starts with the healing of the injury

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_21, © The Author(s) 2022

Fig. 1 Collecting hemocytes in the vascular system of Botrylloides diegensis. (a) A top-view of a colony of Botrylloides diegensis (left) and Botryllus schlosseri (right) growing on a microscopy glass slide. (b) Microinjection setup with a colony holder, a stereoscope, a microinjector, and the micromanipulator. (c) Close-up of the microtransfusion setup, with a colony placed on the colony holder and the marked micropipette close to the insertion point. (d) The external vasculature of the colony. (e) Magnification of the area delimited by the white rectangle in (d). Good candidate points for transfusion are indicated using black arrows, subendostylar points using gray arrows, and bad transfusion points using white arrows. (f) Microcollection of hemocytes. Micropipette is inserted inside the vessel lumen through the tunic and hemocytes (arrows) are flowing in. (g) Average collection rate over time (n ¼ 5). Changes in collection point are indicated using circles, while changes in micropipette using triangles. Display are the individual collections (thin), the average collection (thick), and the corresponding standard deviation (gray)

sites followed by a major remodeling of the isolated vascular system [12]. Circulating stem cells are then mobilized to regeneration niches, i.e., a discrete regeneration locus within the vascular system. These niches compete through a yet undetermined process that consistently leads to the development of a single adult zooid, while all other niches are resorbed by the vascular system [12].

In addition to this role during regeneration [12], hemocytes are involved in numerous biological processes in colonial tunicates, including immune response [13–15], allorecognition [16–18], and asexual reproduction [19–21]. Colonial tunicates' hemolymph is typically composed of less than a dozen 4- to 25-μm-wide cell types classified into four functional classes: undifferentiated, phagocytic, cytotoxic, and storage cells [22–25]. For instance, in Botrylloides leachii, 11 different cell types ranging from 5 to 20 μm have been described [12], as well as the two additional mast-like and transport functional classes. Hemolymph extraction, manipulation, and alteration are thus essential tools for analyzing as well as challenging the functions of hemocytes. A large palette of approaches have been established for this purpose in colonial tunicates. Extracted hemolymph has been used for starting primary cultures of hemocytes [26], its cell composition studied through histological staining [12] as well as flow cytometry [27], and specific hemocyte populations sorted by fluorescently activated cell sorting [28]. Hemocytes have been labeled and injected in a recipient colony [29, 30], a variety of staining solutions delivered into the vasculature [31–33] and functional characterization undertaken by injection of small interfering RNA probes [33, 34] as well as chemical compounds [35, 36].

The most common approach for performing hemolymph collection is by mechanical injury of the vessel of an anti-clottingtreated colony. Hemolymph is then collected with a syringe or a glass micropipette as it bleeds out from the colony [12, 14, 22]. When larger amounts of hemolymph are required, mechanical dissociation of the entire colony is often used [21, 28–30, 37]. In both the approaches, the colony thus bears severe injuries and material exogenous to the hemolymph can contaminate the samples.

Here we describe a microtransfusion technique that allows to collect hemolymph intravascularly with high purity. We also present methods to characterize cytologically the hemolymph, label its hemocytes, and reinject them using the same setup used for the collection. This process can be routinely performed in the same colony with minimal damage, making it useful for long-term in vivo experiments aimed to study the role of the hemolymph in colonial tunicates.

#### 2 Materials

All reagents are prepared with deionized water and stored at room temperature unless otherwise stated.


#### 3 Methods


Fig. 2 Manipulating hemocytes. (a) Counting cells on a hemocytometer using Trypan blue to detect dead cells (none visible) with (b) a magnification of the area delimited in (a) where five cells are visible. (c) Measuring the purity of a collection using a linear regression of the absorbance of a purity scale. Absorbance of the reference samples are depicted using gray dots, purity scales as gray lines, the linear regression as a bold black line, and its 95% confidence interval by a light gray surface. Samples used to measure medium (n ¼ 23) and high (n ¼ 18) purity are depicted in blue and red, respectively. Their corresponding average values and standard deviation are shown as a bold line overlapping a light area. R<sup>2</sup> is the regression coefficient and Δ the slope of the regression. (d) Labeled hemocytes with (e) a magnification of the area delimited in (d) and (f) the corresponding fluorescence of the lipophyllic dye (MemGlow 640) with (g) the corresponding magnification. All scale bars are 100 μm


3.2 Characterization of Hemolymph Collection Depending on the downstream application, careful characterization of the sample needs to be undertaken. Here we measure the purity of the collection, the amount of collected cells, the viability of these hemocytes and calculate the in vivo cell concentration and hemolymph volume.


As described in the introduction, hemolymph has been used for a variety of applications in a number of publications. Here we present our protocol to label hemocytes, analyze them using flow cytometry, and fix them for morphological purposes.


3.3 Flow Cytometry Analysis of Hemolymph


#### 3.4 Injection of Compounds and Cells

Given that the vasculature is a closed system, there is a limitation in the volume and rate of injection that can be achieved, which depends on the size of the colony. This protocol typically yields to the injection of 40 μL of solution in 15 min in colonies composed of more than five adults (Fig. 3c, see Notes 33 and 34).

Fig. 3 Microinjections in Botrylloides diegensis. (a) Vascular system with a micropipette inserted in the vessel lumen loaded with PAS. (b) Properly inserted micropipette starting to inject a dyed solution. (c) Average injection rate over time, separated between intravascular (red, n ¼ 5) and subendostylar (blue, n ¼ 5), overlaid with the global average (black). Display are the individual collections (thin), the average collection (thick), and the corresponding standard deviation (light area). (d) Systemic distribution of the injected medium 5 min after the end of the microinjection. (e) Injection of labeled hemocytes (arrows) in the recipient colony. All scale bars are 500 μm


#### 4 Notes


process freeze-dried samples for metabolomic and proteomic analysis.


tips is pretty fast too, but these tips are relatively expensive. Alternatively, mounting the empty micropipette on the microinjection setup and loading it through its tip by applying negative pressure are slow but inexpensive.


#### Acknowledgments

Funding support was provided by the Swiss National Science Foundation (SNSF) grant number PZ00P3\_173981 as well as by the European COST Action MARSITEM CA16203 short-term scientific mission grants (STSM) 40760 and 43747. We would like to thank Francesca Cima and Loriano Ballarin for the numerous discussions and advice; Sefano Tiozzo for hosting LAMS during his STSMs and for the help setting up microtransfusions; Aude Blanchoud for proofreading the manuscript.

#### References


generations. FASEB J 21:1335–1344. https:// doi.org/10.1096/fj.06-7337com


555–564. https://doi.org/10.1007/s00441- 007-0513-4


regeneration in the invertebrate chordate Botrylloides diegensis. Nat Commun 11:4435. https://doi.org/10.1038/s41467-020- 18288-w


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part IV

Genetic Approaches

# Chapter 22

# Gene Manipulation in Hydractinia

### Eleni Chrysostomou, Febrimarsa, Timothy DuBuc, and Uri Frank

#### Abstract

The ability to regenerate lost body parts is irregularly distributed among animals, with substantial differences in regenerative potential between and within metazoan phyla. It is widely believed that regenerative animal clades inherited some aspects of their capacity to regenerate from their common ancestors but have also evolved new mechanisms that are not shared with other regenerative animals. Therefore, to gain a broad understanding of animal regenerative mechanisms and evolution, a broad sampling approach is necessary. Unfortunately, only few regenerative animals have been established as laboratory models with protocols for functional gene studies. Here, we describe the methods to establish transgenic individuals of the marine cnidarian Hydractinia. We also provide methods for transient gene expression manipulation without modifying the genome of the animals.

Key words Hydractinia, Transgenesis, CRISPR-Cas9

#### 1 Introduction

The phylum Cnidaria comprises some 11,000 species of rather diverse animals [1] that share a unique cell type—the cnidocyte, also known as nematocyte or stinging cell. These cells, which belong to the neural lineage, are used for prey capture, defense, and adhesion [2, 3]. Phylogenetically, members of the Cnidaria are divided between two main clades, Anthozoa (e.g., corals, sea anemones, and sea pens) and Medusozoa (e.g., hydrozoans, scyphozoans, and cubozoans), plus a group of parasitic cnidarians, the Myxozoa, that are a sister taxon to Medusozoa [4]. Anthozoans are being characterized by the lack of a medusa stage, which has been lost and gained multiple times in the Medusozoa clade. As a result, many modern medusozoans lack the medusa stage in their life cycle.

Cnidarians possess a remarkable regenerative ability, being able to regrow whole bodies from small tissue fragments, and in some cases also from reaggregated cell suspensions [5–7]. However, the mechanisms used for regeneration are different not only between

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_22, © The Author(s) 2022

species but also within different body parts of single species [8, 9]. Hence, to gain full understanding of the mechanisms driving regeneration in such a diverse group, it is necessary to study regeneration in as many species and contexts as possible. The main challenge in doing so at the molecular level is the enormous effort required to develop transgenic technologies for each studied species. Contrary to common sense, it appears that substantially different protocols are required to obtain transgenic animals even within one phylum [10–13]. Here, we present the current protocols used to manipulate gene expression in the colonial hydrozoan Hydractinia.

The genus Hydractinia is represented in the literature primarily by two North Atlantic sibling species, H. echinata and H. symbiolongicarpus. Despite the similarity between the two, they do not readily hybridize [14] and differ in genome size (774 and 514 Mb, respectively) as well as in some aspects of postmetamorphosis growth form. Both the species can be grown in artificial seawater tanks, but selected laboratory strains are only available in H. symbiolongicarpus [15, 16].

Hydractinia is one of only four cnidarian genera for which transgenesis technologies have been well established (Fig. 1) [10, 11, 17, 18]; the other three being Hydra, Nematostella, and Clytia. First steps to generate transgenic Aiptasia have been reported recently [19]. Transient gene knockdown protocols using short hairpin RNA, RNAi, and morpholino oligonucleotides have been established in Hydractinia as well [20–22]. Transient ectopic/overexpression experiments can be achieved with mRNA injection into embryos [15].

Hydractinia has been primarily used to study stem/germ cells [15, 20], regeneration [19], neurogenesis [23], and allorecognition [24]. However, recent developments by the Hydractinia research community call for expansion of the usage of this tractable animal model to other disciplines as well.

#### 2 Materials

2.1 Animal Maintenance, Spawning, and Metamorphosis


Fig. 1 A selection of transgenic reporter animals. (a) Tfap2::GFP. This male sexual polyp expresses GFP in early germ cells. (b) Piwi1::GFP. This animal expresses GFP in i-cells and germ cells. The image shows a feeding polyp with no germ cells. (c) A double transgenic female obtained by crossing a Tfap2::GFP animal with a Piwi1::mScarlet partner. (d) A CRISPR-Cas9 knockin animal in which GFP has been inserted in-frame in to the Ef1a endogenous coding sequence. (e) A Rfamide::GFP reporter animal. A subset of neurons express GFP. (f) A G0, Actin1:: GFP mosaic reporter animal. This transgene is epithelial specific. All images were taken from live animals

#### 2.2 Embryo Microinjection


Fig. 2 Hydractinia culture. (a) Overview of a commercial tank system used in our laboratory. (b) A close-up on the slides-in-a-staining-rack system. (c) A close up on a colony growing on a microscope slide

2.3 shRNA Prepare all solutions using ultrapure nuclease-free water (prepared by purifying deionized water, to attain a resistivity of 18 MΩ/cm at 25 C) and analytical grade reagents. Diligently follow all waste disposal regulations when disposing waste materials.


2.4 CRISPR-Cas9 Editing and Genotyping CRISPR RNAs (crRNA) for gene targeting experiments are synthesized commercially (e.g., Integrated DNA Technologies, IDT). crRNAs need to be hybridized to tracer RNA (tracr RNA) prior to use and can be stored at 20 C after hybridization. Cas9 enzyme aliquots are stored at 80 C and are mixed with crRNAs:tracrRNA prior to use.


#### 3 Methods

3.1 Animal Maintenance, Spawning and Metamorphosis


#### 3.2 Capillaries' Preparation and Injection

Fig. 3 Constructs structure. (a) Reporter construct. (b) Ectopic expression construct. (c) mRNA synthesis template. (d) shRNA synthesis template. RS restriction site, P2A self-cleavage P2A peptide coding sequence 5<sup>0</sup> GGTTCAGGT GCTACAAATTTTTCATTATTAAAACAAGCTGGTGATGTTGAAGAAAATCCAGGTCCA 30 , LP: linker peptide coding sequence 50 TGGCCAGGAGGCTCCGGCTCC30 , UTR untranslated region 5<sup>0</sup> TGCAGCCCCGGTAGAAAAA3<sup>0</sup> . URS: upstream regulatory sequence; DRS: downstream regulatory sequence


Constructs


3.4 Strategy for Ectopic Expression Construct Cloning Vectors for transgenic reporter lines can be used to ectopically express or overexpress your gene of interest. URS and DRS regions of the reporter line do not change and a cassette containing UTR + CDS + P2A peptide is inserted upstream of the fluorescence protein (Fig. 3B).

	- 2. Use target gene protein sequence from other animals as query to search the homologous transcripts from Hydractinia symbiolongicarpus.
	- 3. Perform TBLASTN search at https://blast.ncbi.nlm.nih.gov/ against transcriptome shotgun assembly (TSA) database (do not use the default nr database) of Hydractinia symbiolongicarpus. Limit the search by TSA project and type in "GAWH: TSA: Hydractinia symbiolongicarpus, transcriptome shotgun assembly" into the box.
	- 4. From the TBLASTN results page, retrieve the top hit Hydractinia transcript sequence. This is the coding sequence of gene of interest (CDS-GOI).
	- 5. Retrieve the coding sequence of a fluorescence of protein (CDS-FP, e.g., eGFP or mScarlet) from a public database or use the available fluorescence protein sequence that you have already in an existing plasmid in your lab.
	- 6. Codon optimize FP sequences for Hydractinia symbiolongicarpus using http://genomes.urv.es/ OPTIMIZER /. Codon usage table can be found at http://www.kazusa.or.jp/codon/ cgi-bin/showcodon.cgi?species¼13093.
	- 7. Arrange these two CDS according to the scheme shown in Fig. 3C (see Note 15).


3.6.1 Generating Plasmid for Template Construct


3.6.2 Synthesizing Gibson Assembled dsDNA Fragment for Template Construct


3.7 T7 In Vitro Transcription mRNA Synthesis, Microinjection, and Evaluation

	- 2. Perform TBLASTN search at https://blast.ncbi.nlm.nih.gov/ against transcriptome shotgun assembly (TSA) database (do not use the default nr database) of Hydractinia symbiolongicarpus. Limit the search set by TSA project and type in "GAWH: TSA: Hydractinia symbiolongicarpus, transcriptome shotgun assembly" to the filling box.
	- 3. From the tblastn results page, retrieve the top hit Hydractinia's transcript sequence (only the sequence and not in fasta format).
	- 4. Use the top hit sequence to find siRNA motif at http://www. invivogen.com/sirnawizard/design.php. Choose the desired motif size (21 nt is the default). Leave blanks the option of mRNA database and miRNA SEED database.
	- 5. Choose several sequences from the motif outputs with higher GC content, 5<sup>0</sup> GG, and 3<sup>0</sup> AT-rich stretches (see Note 19).
	- 6. Perform BLASTN on several motif output sequences against the Hydractinia symbiolongicarpus TSA with loose parameter (e.g., expected value 1) to confirm specificity (see Note 20).
	- 7. Create the shRNA design by converting the selected 21-nt motif (passenger sequence) into RNA by replacing any T with U.
	- 8. Add the reverse-complemented sequence (guide sequence) to the 3<sup>0</sup> end and separate them by loop sequences [5- 0 -AUUUACU-30 ].
	- 9. Add "UU" to the 3<sup>0</sup> of the sequence to create overhang.
	- 10. Select sequences that are not predicted to form secondary structures using http://rna.tbi.univie.ac.at/cgi-bin/ RNAWebSuite/RNAfold.cgi. Add one or two mismatches in the middle of the passenger sequence (but not to the guide sequence).


#### Table 1 Injection solution recipe for shRNA/mRNA injection

	- 1. Design crRNAs using Geneious or other software.
	- 2. BLAST crRNAs against the Hydractinia genome to confirm their specificity and exclude those with multiple matches in the genome.
	- 3. Hybridize the crRNA with tracrRNA before use to create a viable short guide RNA (sgRNA) that can work with Cas9 for editing. Alternatively, obtain sgRNAs from a commercial supplier.
	- 4. Incubate crRNA:tracrRNA with Cas9 enzyme for 15 min on ice prior to use.
	- 5. Dilute Cas9 to 1 μg/μL and crRNA:tracrRNA to 500 ng/μL.

3.9 CRISPR/Cas9 Design, Microinjection, and Genotyping


#### 4 Notes


#### Acknowledgments

Research in the Frank lab is supported by the SFI-HRB-Wellcome Research Partnership (grant No. 210722/Z/18/Z) and by National Science Foundation (grant No. 1827635).

#### References

1. Zapata F, Goetz FE, Smith SA, Howison M, Siebert S, Church SH, Sanders SM, Ames CL, McFadden CS, France SC, Daly M, Collins AG, Haddock SHD, Dunn CW, Cartwright P (2015) Phylogenomic analyses support traditional relationships within Cnidaria. PLoS One 10(10):e0139068. https://doi.org/10.1371/ journal.pone.0139068


GFP-transgenic animals and chimeras. Dev Biol 348:120–129


Baxevanis AD, Frank U (2017) An evolutionarily conserved SoxB-Hdac2 crosstalk regulates neurogenesis in a cnidarian. Cell Rep 18: 1395–1409. https://doi.org/10.1016/j.cel rep.2017.01.019


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Manipulation of Gene Activity in the Regenerative Model Sea Anemone, Nematostella vectensis

### Eric M. Hill, Cheng-Yi Chen, Florencia del Viso, Lacey R. Ellington, Shuonan He, Ahmet Karabulut, Ariel Paulson, and Matthew C. Gibson

#### Abstract

With a surprisingly complex genome and an ever-expanding genetic toolkit, the sea anemone Nematostella vectensis has become a powerful model system for the study of both development and whole-body regeneration. Here we provide the most current protocols for short-hairpin RNA (shRNA)-mediated gene knockdown and CRISPR/Cas9-targeted mutagenesis in this system. We further show that a simple Klenow reaction followed by in vitro transcription allows for the production of gene-specific shRNAs and single guide RNAs (sgRNAs) in a fast, affordable, and readily scalable manner. Together, shRNA knockdown and CRISPR/Cas9-targeted mutagenesis allow for rapid screens of gene function as well as the production of stable mutant lines that enable functional genetic analysis throughout the Nematostella life cycle.

Key words Nematostella vectensis, Cnidaria, CRISPR, shRNA, Mutagenesis, Electroporation, Knockdown, Cas9, Microinjection, Genome editing

#### 1 Introduction

As a member of phylum Cnidaria, the sister group to Bilateria, the starlet sea anemone Nematostella vectensis is an important model for the study of animal development and evolution (Fig. 1) [1– 4]. More recently, Nematostella has also become a popular system for the study of whole-body regeneration [5–7]. Nematostella polyps are highly regenerative, capable of replacing all missing parts through cellular proliferation in approximately 1 week (Fig. 2) [8, 9]. The presence of similar capabilities throughout most cnidarian species [10] makes whole-body regeneration a potentially shared characteristic of basal metazoans and uniquely positions Nematostella as a model to study the molecular and

Cheng-Yi Chen, Florencia del Viso, Lacey R. Ellington, Shuonan He, and Ahmet Karabulut contributed equally to this work.

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_23, © The Author(s) 2022

Fig. 1 Life cycle of Nematostella vectensis. The lifecycle of Nematostella vectensis is an example of indirect development. Eggs divide rapidly following fertilization and gastrulate to set up the germ layers of the animals and form a transient stage called a gastrula. The planula stage comes next. The planula is a motile larva that moves due to a ciliary organ called the apical tuft. After a few days, the planula undergoes metamorphosis to become a primary polyp. Growth from feeding causes the primary polyp to grow to a sexually mature adult polyp. Both primary and adult polyps are capable of whole-body regeneration. Currently, the egg and blastomeres formed by early mitotic cleavages are the most readily available stages for genetic manipulation

Fig. 2 Regeneration in Nematostella vectensis. (a) An uninjured Nematostella vectensis polyp. (b) Time course of oral regeneration after amputation aboral to the pharynx. All missing tissue is restored by 6 days of regeneration (6 dR). Tentacle growth continues for approximately an additional week (12 dR). Scale bar – 400 μm

genetic basis for ancient and conserved mechanisms of regeneration.

Gene-specific knockdown by shRNA was only recently reported in Nematostella [11]. The Cnidarian microRNA pathway most commonly regulates target mRNAs by cleavage, similar to the mechanism of small interfering RNAs and allows for highly specific knockdown of individual genes [11–13]. For shRNA design, a free web-based interface or downloadable programs support algorithms that can be used to design gene-specific 19-mer targeting motifs. Optimal targeting motifs are then incorporated into a standardized oligo design that allows for cost-effective DNA template production and in vitro transcription of shRNAs by T7 RNA polymerase (Fig. 3a, a<sup>0</sup> ). Additionally, DNA templates for shRNA production can be produced from standard commercially produced oligonucleotides making it cost effective. Once produced, shRNA can be delivered either by microinjection or electroporation of eggs [11, 14]. The method we describe produces micrograms of shRNA per in vitro transcription (IVT), making each reaction suitable for hundreds of microinjections or multiple electroporation experiments.

Initial genome editing efforts in Nematostella used TALENs (Transcription Activator-Like Effector Nucleases) or multistep, cloning-based methods for CRISPR sgRNA production [15]. We have recently adopted a method for sgRNA production leveraging sgRNA design using CRISPRscan [16, 17] followed by T7-mediated transcription from a DNA template produced by a one-step Klenow extension reaction (Fig. 3b, b<sup>0</sup> ). As with shRNA, this method produces micrograms of sgRNA molecules that can be used for hundreds of microinjection experiments. While the current preferred method for targeted mutagenesis is the injection of preloaded ribonucleoproteins (RNPs) of sgRNA and Cas9 protein, similar mutation rates can be achieved using a Nematostella codon-optimized Cas9 mRNA and slightly lower mutation rates can be obtained with a zebrafish codon-optimized Cas9 mRNA [18].

Although current methods for the introduction of functional RNA molecules are limited to early embryogenesis (Fig. 1), the future is bright for functional genetic analysis restricted only to polyp stage animals. A functional heat shock promoter [16] as well as the increased efficacy of genetic insertions by homologydirected repair [16, 19] will likely soon lead to the establishment of transgenic lines containing spatially and temporally controlled genetic tools, such as inducible site-specific DNA recombinase systems (e.g., Cre:Lox and FLP:FRT) or other synthetic conditional alleles found in traditional model organisms [20–24]. Continued development of the Nematostella toolkit will not only allow the functional interrogation of the genetic requirements of whole-body

Fig. 3 Design and production of shRNAs and sgRNAs. (a) Schematic of universal shRNA and gene-specific shRNA oligonucleotides design. (a<sup>0</sup> ) Graphic outline of shRNA production protocol. (b) Schematic of universal sgRNA and target-specific sgRNA oligonucleotides designed using CRISPRscan [14]. (b<sup>0</sup> ) Graphic outline of sgRNA production protocol

regeneration but will also enable the direct comparison of embryogenesis and regeneration in the same animal.

Here, we discuss current strategies for functional genetic analysis by shRNA-mediated knockdown and CRISPR/Cas9-targeted mutagenesis in Nematostella. Both the shRNA knockdown and CRISPR/Cas9-targeted mutagenesis protocols detailed here have four main steps: (1) Design; (2) Production; (3) Delivery; and (4) Screening and Care. While the general methods for these two protocols are similar, their purposes are complementary and allow for genetic analysis throughout the life cycle of the animal.


#### 2.2 CRISPR/ Cas9 Targeted Mutagenesis


50 -TAATACGACTCACTATA-[target-specific region, 20 bp]- GTTTTAGAGCTAGAA-30 (Fig. 3b).


#### 2.3 Microinjection 1. De-jellied Nematostella eggs (see Notes 4–6).



#### Table 1 CRISPR/Cas9 mutagenesis injection conditions.


#### 2.4 Electroporation of shRNA 1. Polysucrose 400 (e.g., Ficoll PM 400, Sigma Aldrich).

	- 3. 4 mm electroporation cuvettes.
	- 4. Electroporation Solution: 15% polysucrose 400 in 12 ppt ASW (see Note 11).

#### 3 Methods

3.1 shRNA Production

#### 1. Identify the coding sequence (CDS) region for each gene of interest (see Note 12).


[reverse complement of shRNA candidate sequence, 19 bp]- TATAGTGAGT-3<sup>0</sup> (Fig. 3a, see Notes 16–18).


3.2 shRNA Microinjection An alternative protocol for shRNA delivery using electroporation is provided in Subheading 3.3. When choosing which shRNA delivery method, it can be important to consider your experiment. The main benefit of microinjection is consistent shRNA delivery due to calibrated injection volumes and the use of long-term labeling injection dyes. Therefore, ideal experiments for shRNA microinjection may include functional screening of a single or a small number of genes of interest or optimization of a consistent shRNA phenotype for production of a consistent samples for further analysis.


#### 3.3 shRNA Electroporation

An alternative to this shRNA delivery method using microinjection is provided in Subheading 3.2.

When choosing which shRNA delivery method, it can be important to consider your experiment. The main benefits of electroporation are ease of shRNA delivery, rapid delivery to a large number of eggs, higher throughput genetic screening, low equipment cost, and potential applicability for use with other Cnidarian species where shRNA injection is very difficult or impossible [25]. Therefore, ideal experiments for shRNA electroporation may include genetic screening of a large number of genes as well as the opportunity for evolutionary comparison of gene function across different Cnidarian species.

	- 2. Discard unhealthy, unfertilized, or lysed animals as well as debris.
	- 3. Repeat steps 1 and 2 daily.
	- 4. As development proceeds, morphological, molecular or cellular phenotypes may begin to appear. Always compare experimental shRNA-treated groups to both mock treatments as well as to untreated wild type controls from the same spawning group (see Notes 37–41).
	- 1. Identify the genomic region of interest (see Note 42).
	- 2. Copy and paste DNA sequence into the CRISPRscan Sequence Submission web interface (https://www.crisprscan.org/? page¼sequence), select "Cas9 – NGG" and "In vitro T7 promoter" from dropdown menus and click "Get sgRNAs" button (see Notes 43–45).
	- 3. The program will suggest many different sgRNA sequences (in upper case) and will automatically design necessary oligo sequences for production by T7 in vitro transcription for each (Fig. 3b).
	- 4. Select 3–5 candidate sgRNA sequences for each DNA sequence (see Note 46).
	- 5. BLAST search each sgRNA sequence (upper case only from CRISPRscan-designed oligos) in the Nematostella genome using an expectation value of 1.0E-2 to compensate for shorter sequence inputs. Each candidate sequence should only have one match in the genome. Additionally, it is important to confirm that the sgRNAs avoid intron–exon junctions (see Notes 15 and 47).

3.4 Screening and Care of shRNA-Treated Animals

3.5 sgRNA Production for CRISPR/ Cas9-Targeted Mutagenesis


1. Incubate RNP Injection Mixture at 37 C for 10 min.


3.7 Genotypic Screening and Care of CRISPR/Cas9 Injected Animals

3.6 sgRNA/Cas9 Delivery by Microinjection

> Genotypic screening by standard PCR and molecular biology techniques is required to confirm DNA cutting and genomic mutagenesis.


#### 4 Notes


Dextran Texas Red™ is a red, non-toxic, long-term tracer dye that will last for at least 7 days within injected animals. However, since it is much harder to visualize injection in the red channel of a fluorescent dissecting scope, it is generally recommended to add FITC to the mixture even when using Dextran as a long-term tracer. Alternatively, Dextran AlexaFluor488 (Thermo Fisher, D34682; Stock Concentration ¼ 8.3 μg/μ L) can be used as single long-term dye that works well for injection. However, it should be noted that primary polyps frequently exhibit endogenous green autofluorescence surrounding the pharynx which may make long-term labeling with this green dextran conjugates less ideal for many experiments.


more formal annotations. As improved genomic resources for Nematostella vectensis emerge, this protocol can and should be used with the most up-to-date and trusted assemblies.


fact, incomplete removal of the TRI reagent can affect the quality of shRNA (an abnormally low 260/280 reading) and result in injection toxicity. If the user decides to use TRI reagent®, simply expand the reaction to 100 μL and add 300 μL of TRI reagent®. Mix well and add another 400 μL of 100% ethanol before proceeding according to the instruction from the kit. If no TRI reagent® is being used, the user can safely skip the first two washes with RNA prewash buffer as detailed in this protocol. Empirical testing in our laboratory of RNA purification of IVT reactions described here have shown no significant reduction in product yield when omitting the TRI reagent®; however, individual users may want to confirm this on their own.


were successfully electroporated. Eggs should recover and return to normal morphology within 30–60 min.


#### shAnthox1a\_R:

50 -AAGGTCTGACGACGAATGTGATCTCTTGAATCA CATTCGTCGTCAGACCTATAGTGAGT-30 ;

Working injection concentration: 500 ng/μL;

Working electroporation concentration: 900 ng/μL;

Observable phenotype: loss of endodermal segment s5, tentacle fusion at polyp stage.

#### shβ-catenin\_R:

50 - AAGTGGCACCAAACGTATCATTCTCTTGAAAT GATACGTTTGGTGCCACTATAGTGAGT-30 ;

Working injection concentration: 100 ng/μL;

Working electroporation concentration: 300 ng/μL;

Observable phenotype: gastrulation failure, disruption of epithelial tissue and lethal at 4 dpf.

#### sheGFP\_R:

50 - AAGACGTAAACGGCCACAAGTTCTCTT GAAACTTGTGGCCGTTTACGTCTATAGTGAGT-3<sup>0</sup> ;

Working injection concentration: 100–1000 ng/μL;

Working electroporation concentration: 200 ng/μL;

Observable phenotype: reduction of GFP in transgenic animals. Knockdown effect lasts to 7 dpf when used at 1000 ng/μL.

#### Scrambled\_R:

50 - AAGCAACACGCAGAGTCGTAATCTCTTGAAT TACGACTCTGCGTGTTGCTATAGTGAGT-3<sup>0</sup> ;

Working concentration: negative control, always match your test shRNA;

Observable phenotype: no observable phenotype when used under 1500 ng/μL. Slight developmental delay when used at 2000 ng/μL.


>1000 sgRNAs injected in zebrafish [14]. sgRNAs are given scores to predict their ability to cut genomic DNA. Scores above 0.55 are termed "efficient for cutting" and above 0.70 are called "highly efficient for cutting (17)." In general, the direct correlation between CRISPRscan score and cutting efficiency holds true in Nematostella. However, it is worth noting that we have seen productive mutagenesis using sgRNAs with scores as low as 40 and when considering which sgRNA to use, the genomic location of the target with regard to ideal site for the experiment should be weighted more heavily than a CRISPRscan score. Both canonical ("GG18") and non-canonical ("Gg18," "GG17," etc.) will work for cutting [14].


all individuals harboring mutant alleles will show a phenotype as F0 animals.


[11, 29]. It should be noted that different genomic loci very regularly require individually optimized injection mixtures.

#### eGFP sgRNA (negative control for wild type animals):

50 -taatacgactcactataGGTCAGGGTGGTCACGAGGGgttt tagagctagaa-30 ;

Working sgRNA injection concentration: 250–500 ng/μL;

Working Cas9 protein injection concentration: 500 ng/μL;

Observable F0 phenotype: No observable phenotype in wild-type animals; reduction or loss of fluorescent protein expression in GFP transgenic lines [11].

eGFP genotyping oligo Forward: 5<sup>0</sup> - AAGGCGTTATGGTC GGTATG-3<sup>0</sup>

eGFP genotyping oligo Reverse: 5<sup>0</sup> -TGCTTGTCGGCCA TGATATAG-3<sup>0</sup> ,

APC (positive control for wild-type animals):

50 -taatacgactcactataGGGGGGCCCTAGTCAGCAGGgttt tagagctagaa-3<sup>0</sup> ;

Working sgRNA injection concentration: 500 ng/μL;

Working Cas9 protein concentration: 500 ng/μL;

Observable F0 phenotype: Formation of ectopic oral structures (tentacles and pharynx) at primary polyp stage, 7–10 dpf [27].

APC genotyping oligo Forward: 5<sup>0</sup> - AGAATCCTGCAG AAGATGAACA-3<sup>0</sup>

APC genotyping oligo Reverse: 5<sup>0</sup> - CCTGGCATACAA AGGTGACA-3'.


zebrafish codon-optimized Cas9 mRNA and Nematostella codon-optimized Cas9 mRNA are functional in Nematostella embryos. Nematostella codon-optimized Cas9 mRNA exhibits mutagenesis rates similar to Cas9 protein and higher than the zebrafish codon-optimized version. However, again, it must be stressed that toxicity is often high with any Cas9 mRNA. See Table 1 for additional information.


#### Acknowledgments

We thank all Gibson Lab members past and present for their help in the development and optimization of the protocols detailed here. We thank Mark Miller for illustrations. We also thank the Stowers Institute Molecular Biology Core, especially Kym Delventhal, MaryEllen Kirkman, and Kyle Weaver, for help with screening design as well as the Stowers Institute Reptile and Aquatics Core facility for animal husbandry help. This study was supported by NIH F32 GM131522 (EMH) and the Stowers Institute for Medical Research (MCG).

#### References


Nematostella vectensis, in vitro fertilization of gametes, and dejellying of zygotes. Cold Spring Harb Protoc 2009(9). https://doi. org/10.1101/pdb.prot5281


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Monitoring Telomere Maintenance During Regeneration of Annelids

# Nithila A. Joseph, Chi-Fan Chen, Jiun-Hong Chen, and Liuh-Yow Chen

#### Abstract

Telomere shortening is a hallmark of aging and eventually constrains the proliferative capacity of cells. The protocols discussed here are used for monitoring telomeres comprehensively in Aeolosoma viride, a model system for regeneration studies. We present methods for analyzing the activity of telomerase enzyme in regenerating tissue by telomeric repeat amplification protocol (TRAP) assay, for comparing telomere length between existing tissue and newly regenerated tissue by telomere restriction fragment (TRF) assay, as well as for visualizing telomeres by fluorescence in situ hybridization (FISH).

Key words Aeolosoma viride, Telomere maintenance, Regeneration, TRAP assay, TRF assay, Telomere FISH

#### 1 Introduction

Regeneration aids in repairing or rebuilding damaged tissue or entire body parts in animals. The ability to regenerate is not uniform among animals. For instance, metazoans like planarians can regenerate entirely with symmetry and proportion from tiny fragments of their existing body using adult somatic stem cells called neoblasts [1]. Advanced mammals like humans can only regenerate certain body parts like liver after partial resection [2]. Salamanders like axolotls and newts can regenerate several body parts including limbs or tail by activating progenitor cells [3]. While axolotls lose the ability to regenerate eye lens around 2 weeks after birth, newts can regenerate lens without being limited by age [4]. Observations akin to this implicate aging as one of the factors that affect regeneration potential. Although the real mechanism of how age affects wound healing and regeneration is not clear, most people accept that wound healing is affected by aging.

This raises questions about the aging situation of the regenerated tissues. Are the regenerated tissues similar to the existing tissue or younger than them? Does regeneration evoke rejuvenation? Aging is a complicated life phenomenon, and it is quite difficult to evaluate the age of animals especially that of invertebrates. Length of telomere, one of characteristics to realize aging in animals, was chosen as the parameter to figure out the relationship between aging and regeneration. The inability of DNA replication mechanism to fully replicate the lagging strand of chromosomes causes progressive shortening of telomeres [5].

Telomerase is a ribonucleoprotein composed of TERC (RNA component) and TERT (reverse transcriptase) to lengthen telomeres. Expression of telomerase is varied among organisms. Remarkably, certain species of animals with good regenerative ability have upregulated telomerase expression [6, 7]. Also, regenerative ability waning with age supports the notion that shorter telomeres contribute to diminished proliferation.

Established model animals for regeneration studies, such as amphibians and reptiles, usually have long life spans, which disallows the exploration of age as a factor restricting regeneration. To circumvent this problem, we used Aeolosoma viride, a freshwater annelid as a new model for regeneration and aging-related research. Annelids are well known for their ability to restore anterior and posterior ends from few of their body segments. Lifespan of A. viride is approximately 2 months, making it an appealing model for aging-related study.

A. viride of varying age groups when observed for regeneration showed that regeneration declines as age of the worm increases [8]. Telomere length was maintained at the regeneration sites after amputation of the head [8].

Here, we describe protocols to monitor the telomere maintenance in A. viride during tissue regeneration. We discuss three main strategies: (1) PCR-based telomeric repeat amplification protocol (TRAP) assay for analyzing the activity of telomerase in regenerating sites (2). Telomere restriction fragment (TRF) assay (3). Telomere fluorescence in situ hybridization (FISH).

#### 2 Materials

All solutions are prepared using filtered and deionized ultrapure water at room temperature (RT) and analytical grade reagents (unless otherwise indicated).

2.1 Telomeric Repeat Amplification Protocol (TRAP) Assay


1. Nuclei lysis buffer (e.g., Wizard Genomic DNA Purification Kit Promega).


2.2 Terminal Restriction Fragment (TRF) Assay

	- 1. 0.02% (w/v) colchicine in ASW.
	- 2. Carnoy's fixative: 750 mL methanol, 250 mL glacial acetic acid.

2.3 Telomere Fluorescence In Situ Hybridization (Telomere FISH)


#### 3 Methods


Fig. 1 TRAP assay of A. viride extract. Crude and heat-inactivated (85 C) A. viride extracts were used for TRAP reaction. Lysate from human 293 T cells was used for positive control reaction and lysis buffer for negative control reaction. A ladder pattern indicates telomerase-extended products. Asterisk indicates internal control products. (Adapted from [8]).


#### 3.3 Genomic DNA Extraction from Regenerating Tissues


3.4 Terminal Restriction Fragment (TRF) Assay

Fig. 2 TRF assay of A. viride genomic DNA. Intact () and restriction digested (+) genomic DNA were resolved by agarose gel electrophoresis for southern blotting. Restriction fragments of telomeric DNA shows a span of 0.2–5 kb in length. Asterisks indicate internal repetitive sequences. Sizes of selected bands of DNA ladder (L) are indicated on the left. (Adapted from [8]).


3.5 Telomere Fluorescence in Situ Hybridization (Telomere FISH)

Fig. 3 FISH images of A. viride interphase nuclei (left) and metaphase spread (right) stained with telomeric PNA probes (red) and DAPI (blue). Scale bars: 10 μm. (Adapted from [8]).


#### 4 Notes


Additionally, each reaction benefits from the addition of internal controls. This is done by adding a forward primer (e.g., K1in TRAPeze Telomerase Detection Kit) and an internal control template (e.g., TSK1 in TRAPeze Telomerase Detection Kit). Together with the TS primer, K1 and TSK1 amplify an internal control standard (a 36 bp amplicon in TRAPeze Telomerase Detection Kit). The internal control band is an indicator of PCR amplification efficiency.

3. TRAP reaction in its classical form uses a wax barrier to prevent CX primer from mixing with the other reaction components. This was later simplified by the use of "hot start" Taq DNA-polymerase such as Platinum™ Taq DNA polymerase. The "hot start" Taq DNA-polymerase enzyme is activated when the sample is heated to 95 C for 2 min before PCR cycling.


#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Analysis of DNA Double-Stranded Breaks Using the Comet Assay in Planarians

### Paul G. Barghouth, Salvador Rojas, Lacey R. O'Dell, Andrew M. Betancourt, and Ne´ stor J. Oviedo

#### Abstract

Comet assay provides the opportunity to detect and characterize DNA strand breaks. Cellular lysing followed by embedding in agarose slide is used to visualize under an electrical current migration patterns corresponding to DNA fragments of different sizes. Here we describe the process of detecting and characterizing DNA damage by Comet assay on planarians, which is a model organism commonly used to understand the process of whole-body regeneration, stem cell regulation, and adult tissue maintenance.

Key words Alkaline Comet Assay, Planarian, Double-stranded breaks, DNA damage

#### 1 Introduction

Single-cell gel electrophoresis or Comet assay is an attractive tool to assess the integrity of DNA molecules [1–4]. DNA maintains its structure through its negatively charged supercoils around the histone core. DNA strand breaks may result from endogenous and exogenous sources (e.g., ROS and ionizing radiation), leading to the disruption of DNA integrity [5]. The Comet assay takes advantage of the relaxation of DNA supercoils to assess the levels of DNA strand breaks. Briefly, cell suspensions are embedded in agarose-coated slides and are lysed with detergent (i.e., Triton-X 100) to remove nuclear membranes and DNA histone structures, resulting in gel-embedded nucleoid bodies [4]. Increases in DNA breaks and subsequent relaxation in its loops can be exposed when an electric field is applied, and a comet tail-like structure is formed in relation to the amount of damaged DNA. Higher amounts of DNA strand breaks yield more prominent comet tail-like structures [3, 4, 6–8]. After the initial comet protocols were established in 1980s [1, 2], many variations to the protocols have been established (e.g., Comet-FISH, Comet-BrdU) [6–10]. However, the

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_25, © The Author(s) 2022

Comet assay under alkaline conditions (i.e., pH >13) has remained the most widely used method by converting all types of DNA damage (i.e., crosslinks, strand breaks, adducts, etc.) to doublestranded DNA breaks [2–4, 11].

Detection of DNA damage and its repair can be studied in a variety of ways in planarian flatworms. These include immunohistochemistry and Western blot techniques to assess the expression of markers associated with the DNA damage response (i.e., RAD51, H2AX, and PARP) [12–15]. Planarian stem cells known as neoblasts are the only cells with replicative capacity in planarians. Techniques to assess neoblast chromosomal stability and telomeric maintenance have been established [12, 16, 17]. Recent research implemented the use of Comet assay in planarians to characterize the extent of DNA strand breaks [12, 14–16]. The procedure can be guided toward specific cell types by using flow cytometry to sort cells (e.g., neoblasts) or may involve evaluation of different cell types obtained by the dissociation of whole animals. Future adaptations of the Comet assay may also involve double labeling with immunostaining, gene expression probes, and BrdU, which altogether may facilitate characterization of DNA damage and repair on specific cell types.

The Comet assay not only provides a qualitative representation of the extent of DNA strand breaks but can also be used to obtain a precise quantification between different degrees of damage and repair. Here we demonstrate that Comet assay can be used to efficiently detect the extent of DNA strand breaks in a variety of conditions using the highly regenerative planarian model. This includes exposure to gamma irradiation, knockdown of genes, and pharmacological treatments with genotoxic compounds.

#### 2 Materials

#### 2.1 Handling Equipment


#### 2.2 Comet Assay Solutions All the solutions are prepared with Nanopure water unless otherwise stated.


#### 3 Methods


Fig. 1 Comet protocol preparation and visual protocol. (a) Comet workflow and timeline. (b) Representative images of unfrosted and 1% NMPA-coated slides. A good slide to conduct the Comet protocol with alongside coated slides that are not usable due to voids in agarose (i.e., red circles). (c) Setup for planarian dissociation. Needed a dissecting microscope, Peltier cooler, petri dish, and tweezer with a razor blade. (d) Representative images of steps in worm dissociation. Note that by the end of the dissociation process, the end product should look homogenous void of remaining tissue structures. (e) Image showing the "bottom-to-top" method to


#### 3.3 Slide Preparation 1. Microwave and equilibrate 30 mL 0.5% LMPA to 37 C for 30–60 min.

	- 2. Place 40 mL of cooled transparent final lysing solution in a Coplin jar.
	- 3. Gently insert slides in the filled Coplin jar.
	- 4. Protect slides from light by wrapping Coplin jar with aluminum foil.
	- 5. Place Coplin jars in the 4 C refrigerator overnight (see Note 17).

#### 3.5 Comet Slide Electrophoresis 1. Refrigerate 500 mL of 1 electrophoresis buffer at 4 C for 30 min.


Fig. 1 (continued) prepare cell embedded slides without the generation of bubbles. (f) Electrophoresis setup within the 4 C fridge connected to a voltage power source. The slides are aligned tightly side-by-side on the cathode side of the box


3.6 Comet Slide Neutralization and Fixation

3.7 Comet Slide Staining and Visualization

Fig. 2 The effect of various treatments on DNA damage using the Comet assay. (a) Visual representation of single nuclei post alkaline Comet electrophoresis and staining. Nuclei are stained with DNA dye (i.e., SYBR green) and imaged using florescent microscopy, revealing the severity of DNA damage per cell. Tail length can be ranked, categorizing cells as undamaged, moderate, and severe damage (e.g., yellow (0), orange [1], and red [2], respectively). (b) Quantification of three independent Comet assays using the ranking score method on 7-day starved animals. Approximately 40% of planarian cells contain undamaged DNA (i.e., score of 0) and is consistent with other experimental models [18]. (c) Comet-tail length after exposure to 1 K rad gamma irradiation (sub-lethal) in a 7-day time course post treatment. Planarian stem cells are lost by 1–2 days post sub-lethal irradiation due to increased DNA damage. However, by days 4–5 post irradiation, planarian stem cell begin to repopulate, and this is accompanied by an increase in DNA damage and DNA repair proteins as shown previously [12]. By day 7 post radiation, DNA integrity begins to reestablish. (d) Increase in DNA damage can be achieved through RNA interference (RNAi). Graph represents comet-tail length of 30-day

repeat each experiment in triplicated form for each independent biological replicate.

7. Measure the comet-tail length from the edge of the comethead to the edge of the comet-tail using imaging software (e.g., ImageJ). Rank comet-tail lengths from 0 to 2, where a score of 0 shows little DNA damage, 1 moderate, and 2 is a dispersed tail with no nucleus visible (i.e., 0–10 μm, 11–39 μm, and 40–60 μm, respectively) (Fig. 2).

#### 3.8 Comet Slide Storage 1. Remove coverslip.


#### 4 Notes


Fig. 2 (continued) starved animals for both the control and experimental group Rad51(RNAi). Rad51(RNAi) animals contain cells that harbor increased DNA damage and chromosomal abnormalities [13]. RAD51 is a key component in DSB repair within the planarian during homeostasis and pore-radiation stem cell repopulation. Chromosomal abnormalities. (e) Treatment with pharmacological agents such as Aphidicolin can result in increased DNA damage within the planarian model system. Aphidicolin (APH) is an inhibitor of DNA replication, blocking DNA polymerase Alpha and Delta during S-phase of the cell cycle. Comet assay was performed on animals exposed to DMSO and APH [0.5 mM] for 6 hours (i.e., control and experimental group, respectively). It is evident that APH treatment increases DNA DSBs within the planarian. (c–e) Each dot represents an individual planarian cell's comet-tail length. (b–e) All graphs represent mean s.e.m. Statistics were obtained by two-way ANOVA; \* <0.05 and \*\*\*\* < 0.0001

a volume of 40 mL or an adequate volume to fill one Coplin jar. Increase volume depending on the number of Coplin jars required to hold all slides. Furthermore, this solution will turn opaque and requires cooling at 4 C to turn clear. Solution must be clear prior to use.


#### Acknowledgments

This work was supported by the University of California Cancer Research Coordinating Committee (Award# CRR-18-525108) and the National Institutes of Health (NIH) National Institute of General Medical Sciences (NIGMS) award R01GM132753 to N.J. O.

#### References


electrophoresis. Genes Cells 22(1):84–93. https://doi.org/10.1111/gtc.12457


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 26

# Random Integration Transgenesis in a Free-Living Regenerative Flatworm Macrostomum lignano

Jakub Wudarski , Kirill Ustyantsev , Filipa Reinoite , and Eugene Berezikov

#### Abstract

Regeneration-capable flatworms are highly informative research models to study the mechanisms of stem cell regulation, regeneration, and tissue patterning. Transgenesis is a powerful research tool for investigating gene function, but until recently, a transgenesis method was missing in flatworms, hampering their wider adoption in biomedical research. Here we describe a detailed protocol to create stable transgenic lines of the flatworm M. lignano using random integration of DNA constructs through microinjection into single-cell stage embryos.

Key words Macrostomum lignano, Flatworms, Regeneration, Transgenesis, Microinjection, Random integration, Irradiation

#### 1 Introduction

Macrostomum lignano is a free-living marine flatworm capable of regeneration anterior to the brain and posterior to the pharynx [1]. During the last decade, the interest in this research model steadily increased [2]. Similar to other flatworms, regeneration in M. lignano is fueled by stem cells called neoblasts [3]. It is a small and transparent animal that is easy to culture in laboratory conditions. M. lignano is a non-self-fertilizing hermaphrodite with a short generation time of 2–3 weeks [4, 5]. When cultured in standard laboratory conditions, animals lay approximately one egg per day. Embryonic development takes 5 days, and hatchlings reach adulthood in about two weeks. The laid eggs are fertilized, relatively large (100 μm) and follow the archoophoran mode of development [4, 5], i.e., they have a large, and yolk-rich oocyte instead of a small oocyte supplied by dedicated yolk cells. These features, together with the recently reported genome and transcriptome assemblies [6–8], make M. lignano a versatile model organism for

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_26, © The Author(s) 2022

research on stem cells and regeneration [2, 9]. In addition, the availability of transgenic techniques renders this flatworm a unique research model among Platyhelminthes [8]. Here we present a method for transgenesis in M. lignano using microinjection of different components into single-cell stage embryos. The method includes preparation and maintenance of animal cultures, design of transgenic constructs, microinjection procedures, and selection of transgenic animals.

#### 2 Materials

A typical M. lignano transgenesis work space is similar to configurations used for transgenesis in other animals, where DNA is delivered by microinjection into cells. It includes instruments for preparation of microinjection needles (a micropipette puller and a microforge), a stereomicroscope and an inverted microscope equipped with micromanipulators and a microinjector (Fig. 1).


Fig. 1 Typical microinjection working station equipment. (a) On the right: a stereomicroscope for worm transferring and egg picking. On the left: a microforge for fine preparation of pulled microcapillaries into holders and/or needle opening. (b) A micropipette puller. (c) An inverted microscope equipped with micromanipulators and a microinjector. (d) A fluorescence stereomicroscope for the selection of eggs and worms positive for a transgene expression


#### 3 Methods



3.3 Preparation of Plastic Pickers for Egg Collection M. lignano eggs are covered with a sticky mucus, which helps to fix the eggs on a surface. Most of the time, eggs are laid closely to each other and form clumps. Plastic pickers are used to separate eggs in the clumps. Additionally, the mucus around the eggs adheres to the tip of the picker, which helps to transfer the eggs from a petri dish to a microinjection slide and then attach them to the slide surface. A second picker is usually necessary to assist the release of the egg from the first picker. After that, the eggs can be easily manipulated to the desired location on the microinjection slide by gentle touching with the tip of the picker.


3.4 Preparation of

the Holders

	- 2. Break the pulled glass capillary using a microforge to create a tip of approximately 140 μm outer diameter and 50 μm inner diameter.
	- 3. Heat-polish the pipette tip to create smooth edges using the glass bead on the microforge filament.
	- 4. Using the microforge, bend the tip to a ~20 angle. To do so, rotate the tip vertically and apply heat close to the point where the bend is needed. Do not touch the heat source to prevent

Fig. 2 Typical microinstruments used to manipulate and inject M. lignano eggs. (a) A plastic picker for egg collection made from a microloader tip. (b) A close up on the plastic picker tip. (c) A close up on the tip of a holder. (d) A close up on the taper and the tip of a microinjection needle. Note the filament inside

the glass from melting into the microforge. See Fig. 2c for a typical holder example.


Fig. 3 Selection of the promoter for a gene of interest using M. lignano genome browser. A genomic region encompassing Mlig005144.g2 gene (APOB homolog) shows the structure of the gene, and RNA-seq and CEL-seq tracks. Region selected for the promoter cloning is annotated as a black rectangular block upstream of the ATG

genome browser search field or by searching its sequence using the BLAT tool (see Note 8).


3.7 Microinjection Mix: DNA for Random Integration The chosen DNA can be used in three different forms: as a circular plasmid, as a linearized plasmid, or as a PCR product. In the first case, a plasmid suspension in DNase/RNase-free water or TE-buffer with concentration of 150–300 ng/μL is recommended. To prepare the cut plasmid:


To prepare the PCR product:

	- 2. Keep the transferred worms overnight at 20 C to slightly starve them.
	- 3. In the morning on the following day, transfer the worms again to fresh ASW and put them in the dark at 20 C for approximately 2–3 h (can be in a shelf or a drawer at room temperature).
	- 4. Move the worms into light (they can be returned to the incubator) and keep them there for 30 min.
	- 5. Once the first eggs are laid, the egg collecting step can be started using a stereomicroscope.
	- 6. Put a drop (150–200 μL) of ASW on a 30-mm non-treated round glass cover slide or any other glass slide that fits into a well of a six-well plate.
	- 7. Use the plastic pickers to collect the laid eggs and transfer them to the drop of ASW (see Notes 16–18).
	- 8. Put the slide with the eggs on the microinjection stage and focus the inverted microscope on the first egg using low magnification (5 or 10 objective) (Fig. 4a).
	- 9. Mount the holder capillary on the micromanipulator and position it near the egg (Fig. 4a).
	- 10. Load the microinjection needle with 1 μL of your choice of material-to-be-injected. There are no differences in the microinjection procedure in regard to the material used for injections (see Note 19). The needle can be loaded using a microloader tip or by capillary force by applying material to its back.
	- 11. Mount the loaded needle on the micromanipulator and connect the pressure tube to the pressure supply unit of the microinjector.

Fig. 4 Highlights of a typical microinjection procedure into the M. lignano single-cell eggs. (a) Positioning of the eggs, a loaded needle, and a holder under 5 objective of the inverted microscope. (b) The holder touching the edge of the egg, and the needle in the position to "clean" before the injection (40 objective). (c) The needle touching the egg shell before puncturing. (d) The needle puncturing through the egg shell. (e) The moment of injection is seen as a burst inside the egg. (f) The needle is removed from the successfully injected egg. Scale reference: M. lignano egg size ~100 μm


3.10 Transgenic Eggs Maintenance and Selection of Homozygous Lines


Fig. 5 An overview of an APOB::GFP::Ef1a\_30 UTR transgene expression in M. lignano eggs and in the whole worm. (a) Comparison of positive and negative eggs under fluorescent stereomicroscope 1 day after the injection with the PCR DNA fragment encoding the transgene. From top to bottom: FITC channel, bright-field, and merged. Scale bars: 100 μm. (b) Promoter of the M. lignano APOB homolog (Mlig005144.g2) exhibits gut-specific expression pattern in the worm. From left to right: FITC channel, bright-field, and merged. Scale bars: 100 μm


#### 4 Notes


probability of generating mosaic animals increases. Thus, inject only an egg when it is in a single-cell stage.


#### Acknowledgments

The work on the design of transgenic constructs and microinjection procedure was done at the Institute of cytology and Genetics SB RAS by KU and financially supported by Russian Science Foundation grant No. 19-74-00029. FR was supported by the UMCG Graduate School of Medical Sciences fellowship.

#### References


Falciatori I, Visozo DB, Smith AD, Ladurner P, Sch€arer L, McCombie WR, Hannon GJ, Schatz M (2015) Genome and transcriptome of the regeneration-competent flatworm, Macrostomum lignano. Proc Natl Acad Sci U S A 112:12462–12467. https:// doi.org/10.1073/pnas.1516718112


lignano. Sci Rep 8:1–10. https://doi.org/10. 1038/s41598-018-21107-4


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 27

# RNAi Screening to Assess Tissue Regeneration in Planarians

### Rachel H. Roberts-Galbraith

#### Abstract

Over the past several decades, planarians have emerged as a powerful model system with which to study the cellular and molecular basis of whole-body regeneration. The best studied planarians belong to freshwater flatworm species that maintain their remarkable regenerative capacity partly through the deployment of a population of adult pluripotent stem cells. Assessment of gene function in planarian regeneration has primarily been achieved through RNA interference (RNAi), either through the feeding or injection of double-stranded RNA (dsRNA). RNAi treatment of planarians has several advantages, including ease of use, which allows for medium-throughput screens of hundreds of genes over the course of a single project. Here, I present methods for dsRNA synthesis and RNAi feeding, as well as strategies for follow-up assessment of both structural and functional regeneration of organ systems of planarians, with a special emphasis on neural regeneration.

Key words Planarian, Schmidtea mediterranea, Dugesia japonica, RNAi, dsRNA synthesis, Screening, Regeneration, Functional genomics

#### 1 Introduction

Planarian flatworms have grown popular as a study system for regeneration because they can regrow all cell types after nearly any injury. Over one hundred years ago, scientists determined that planarians can achieve whole-body regeneration starting with a small fragment of an adult animal. More recent work, mostly using the species Schmidtea mediterranea (Fig. 1a, b) and Dugesia japonica (see Note 1), revealed many important cellular and molecular contributors to planarians' regenerative capacity, including a population of adult, pluripotent stem cells, and constitutive bodywide axial polarity signaling (Fig. 1c, d; [7] and for reviews, see [2, 3, 8]). Planarians also possess diverse tissue types, allowing dissection of the molecular mechanisms that power structural and functional regeneration at the level of organ systems. The planarian body consists of: a tri-lobed intestine and a tube-shaped feeding

Fig. 1 Introduction to planarians. (a) A live planarian is pictured with its anterior (head) toward the top of the page. Eyespots are visible. (b) A planarian is diagrammed as pictured in a, with eyespots labeled. The pharynx (feeding organ) is tucked inside the body of the planarian when an animal is not feeding, but the outline of the pharynx can be faintly visible from the dorsal side. During feeding, the tube-shaped pharynx emerges from its pouch to extend through an opening on the ventral surface of the planarian body (not shown). (c) Planarian stem cells are a heterogeneous population, containing both pluripotent (dark blue) and specialized (light blue, teal) cells (reviewed in [1]). Stem cells are present throughout the planarian body, with two main exceptions. The pharynx has no resident stem cells, and few stem cells exist in the tip of the head (anterior to the eyespots). (d) A suite of polarity determinants regulates body patterning in the planarian (for review, see [2, 3]). The anterior/posterior axis of polarity signaling is depicted here. Wnt ligands (e.g., Wnt1, Wnt11-1, and Wnt11-2) are produced in the tail of the planarian. Wnt inhibitors (e.g., Notum, sFRP-1) are produced in the head to oppose Wnt signaling. Additional signaling molecules pattern the trunk of the planarian and pattern additional axes (e.g., dorsoventral) (for review, see [2, 3]). (e) The digestive system of the planarian is diagrammed, with the pharynx connecting to the intestine (green), which has one anterior primary branch and two posterior secondary branches. The intestine is composed of multiple cell types and is surrounded by enteric muscle [4, 5]. (f) The central nervous system of the planarian is diagrammed (for review, see [6]). Two ventral nerve cords connect with horseshoe-shaped cephalic ganglia which are also referred to as the planarian brain. Brain branches project outward from the cephalic ganglia to the edge of the planarian head

organ called a pharynx (Fig. 1e); muscle cells in many orientations throughout the body, which function to facilitate animal movement and produce signals for body patterning; a cephalized nervous system (Fig. 1f); osmoregulatory protonephridia; an epidermis, much of which is ciliated and promotes movement by gliding; secretory organs that produce mucus; connective tissue called the parenchyma, within which stem cells are embedded; ovaries, testes, and other reproductive tissues; and other novel cell and tissue types still to be explored [4]. Even within these organ systems, an amazing degree of complexity is present. For example, the planarian nervous system is composed of dozens of neural cell types and glia, all arranged spatially within horseshoe-shaped cephalic ganglia that connect to two ventral nerve cords (Fig. 1f, [6]). Separate peripheral and pharyngeal nerve networks are also present.

As we have learned more about planarian regeneration and physiology, RNA interference has emerged as the most common tool with which to query gene function. Planarian biologists typically produce double-stranded RNA (dsRNA) either in bacteria or in vitro. dsRNA is administered to planarians by feeding, soaking, or injection to trigger RNAi [9–14]. This approach, often repeated several times, depending on the RNAi paradigm, causes a reduction in levels of the target mRNA that can range from nearly a 50% decrease to more than a 95% decrease [15]. The RNAi effect is even stronger in regenerated planarian tissues, which often experience a more penetrant reduction in mRNA [16]. The ease of performing RNAi in planarians allowed several screens of hundreds of genes within a single project (e.g., [15, 17]). Thus, this method is a powerful tool for medium-throughput analysis of gene function during whole-body regeneration and in the context of replacement of specific tissues or cell types after injury. In this chapter, I will outline typical methods for RNAi treatment by feeding of synthetic dsRNA in Schmidtea mediterranea. I will also present a range of possible approaches for assessment of regeneration downstream of RNAi.

Taken together, the following approaches can determine the extent to which regeneration occurs normally after gene perturbation by RNAi. dsRNA synthesis and feeding to achieve RNAi in planarians will be an accessible strategy for studies of regeneration, particularly those focused on whole-body regeneration, regeneration of complex organ systems or tissues, and brain regeneration in particular.

#### 2 Materials

All solutions should be prepared with sterile, ultra-pure water, and stored at room temperature (RT) unless otherwise stated.

#### 2.1 Template Preparation


Fig. 2 Molecular strategy for dsRNA synthesis. (a) The pJC53.2 plasmid used for cloning upstream of dsRNA synthesis is pictured here [18]. This plasmid is digested with Eam1105I for TA cloning of PCR products generated from cDNA. (b) The resulting plasmids contain fragments of each gene of interest. These plasmids are subjected to PCR using a primer that recognizes the T7 promoter sequence to create an amplified product for each target and flanking promoters (c). The PCR products are used as a template for in vitro synthesis reactions using T7 RNA polymerase. Each in vitro synthesis reaction generates gene-specific dsRNA (d)

	- 2. 1 M spermidine. Filter sterilize, aliquot 500 μL per tube and store at 20 C.
	- 3. 1 M dithiothreitol (DTT). Filter sterilize, store at 20 C.
	- 4. 10 high yield transcription buffer: 4 mL 1 M Tris pH 8.0, 2 mL 1 M MgCl2, 200 μL 1 M spermidine, 1 mL 1 M DTT, in 2.8 mL RNase-free water. Sterile filter, aliquot 200 μL per tube and store at 20 C.
	- 5. rNTP mix: 25 mM rATP, 25 mM rUTP, 25 mM rCTP, 25 mM rGTP. Aliquot 100 μL per tube and store at 20 C.
	- 6. T7 RNA polymerase.
	- 7. Thermostable inorganic pyrophosphatase (TIPP) enzyme.
	- 8. Ribonuclease (RNase) inhibitor (e.g., RNasin®).
	- 9. Formaldehyde loading dye.
	- 10. RNase-free DNase.
	- 11. 5 M ammonium acetate. Prepare with RNase-free water. Sterile filter.
	- 12. 100% ethanol.
	- 13. 70% (v/v) ethanol. Prepare with RNase-free water.

#### 2.3 RNAi Treatment and Amputation 1. 1 μg/μL purified dsRNA. 2. Planarians—Schmidtea mediterranea. 10 animals of 3–5 mm length per gene of interest, plus 10 animals of similar size for negative control. 3. 60–100 mm petri dishes. 4. Cafeteria trays. 5. 1x Montjuı¨c salts: 1.6 mM NaCl, 1 mM CaCl2, 1 mM MgSO4, 0.1 mM MgCl2, 0.1 mM KCl, 1.2 mM NaHCO3, pH 7.5 with HCl or NaOH. 6. Large bulb, wide mouth transfer pipettes (e.g., 8.6 mL). 7. Disposable pellet pestles. 8. Planarian food (e.g., liver puree prepared as per [19]). 9. Green food coloring.


#### 3 Methods

#### 3.1 Template Preparation by PCR The starting material for template preparation is ~750 bp of each target gene cloned from cDNA into pJC53.2 vector (Fig. 2a, b, see Note 3). Positive and negative controls should also be included (see Note 4). 1. Prepare one PCR per template to be amplified (see Note 5). 2. Combine 5 μL of the hot start mix solution and 5 μL of 35 mM MgCl2 in each tube. 3. Incubate for 15 min at RT to precipitate the MgCl2.


#### 3.2 In Vitro dsRNA Synthesis The starting material for dsRNA synthesis is template DNA with T7 promoter sequences on each side generated in Subheading 3.1 (see Note 7). For all steps in this section of the protocol, use RNase-free materials, including RNase-free filter tips and RNase-free tubes.


Determine the design for the RNAi experiment. The time course frequently used in my laboratory is to complete three feedings every 5 days with 3–5 μg dsRNA per feeding for 10 planarians (Fig. 3a). We wait for 7 days after the last feeding and amputate animals pre-pharyngeally. After amputation, we wait for 7 days until observing or fixing animals for assessment of regeneration as detailed below. Other feeding paradigms, injection paradigms, dsRNA doses, and amputation strategies may be used (Fig. 3b–e, see Note 9).


#### 3.3 dsRNA Feeding for RNA Interference (RNAi)

Fig. 3 Paradigms for RNAi. (a) The typical strategy used in my laboratory for dsRNA feedings and amputation is shown here. Planarians receive three feedings of dsRNA over the course of ~11 days. Pre-pharyngeal amputation occurs approximately 1 week after the final feeding and animals are observed for head or brain regeneration 1 week after amputation. (b) Additional amputation strategies are presented. Amputation is indicated with a dashed line on the left image of each pair and regeneration is diagrammed in the right image of each pair, with blastema tissue shown in a lighter color. (1) Animals can be amputated post-pharyngeally to observe tail regeneration. (2) Sagittal amputation can be used to observe regeneration of lateral structures and reestablishment of mediolateral patterning. (3) Chemical amputation can be used to remove the pharynx to observe pharyngeal regeneration [20]. (4) Other excisions can be made to determine local wound responses after minor injuries. (c) A long-term RNAi strategy with weekly dsRNA feedings can be used to determine


Fig. 3 (continued) whether genes are required for growth or tissue maintenance under homeostatic (non-injury) conditions. (d) Frequent feeding strategies can be used to increase the efficiency of gene knockdown by RNAi and to improve phenotype penetrance. (e) Injection strategies can be used instead of or in addition to dsRNA feeding [13]. Though this strategy is more time-consuming, it can be especially valuable when gene knockdown interferes with proper feeding of the planarians


3.4 Strategies for Assessing Regeneration After RNAi

Performing RNAi for 10 or more animals per sample is usually sufficient to assess whether significant differences exist between control and experimental RNAi-treated animals. After RNAi treatment and amputation, one or more of the following approaches may be used to assess and quantify regeneration: blastema


Fig. 4 Assessing regeneration phenotypes. (a) Blastema size can be measured by outlining the blastema (yellow dashed line) and measuring blastema area in ImageJ [30]. The body size can similarly be measured (red line). By dividing blastema size by body size to normalize for variable animal size, the resulting value can be compared across populations and RNAi treatments. (b) Similarly, I use in situ hybridization with a choline acetyltransferase (ChAT) riboprobe [23] to stain the central nervous system for measurement of brain size after regeneration (yellow dashed line). Brain size can also be normalized to body size (red line) for comparison of brain regeneration across RNAi treatments. (c) Some cell types are present in numbers low enough that they can be accurately counted, like cells expressing glutamic acid decarboxylase (GAD) [31]. Counting these cell types for regenerated animals following RNAi treatment would allow determination of genes that affect regeneration of GAD<sup>+</sup> cells. In the image shown, 34 cells are labeled (arrowheads). (d) This table lists some available antibodies and examples of riboprobes that can be used for staining diverse cell types or organ measurement (see Note 12); in situ hybridization to examine specific organs or cell types ([21, 22], see Note 13); immunofluorescence to detect cell types or tissue regeneration ([23–29], see Note 14); reverse transcription and quantitative PCR (RT-qPCR) to examine gene expression (see Notes 15 and 16); functional assessment including behavioral assays (see Note 17). Potential data from these types of experiments are shown in Fig. 4.

#### 4 Notes


Fig. 4 (continued) systems [23–29, 32–51]. These approaches may be used to determine whether regeneration proceeds normally after RNAi, including the shape of organs and the renewed expression of key markers. (e–j) In situ hybridization examples are presented. These expression patterns could be used to explore regeneration of the following cell types, structures, and organs: stem cells (smedwi-1 [32]); neurons (nAChR, dd\_Smed\_v6\_8058\_0\_1); subsets of neurons (CNG1 [15], ppp-2 [18]) including brain branches (gpas [34]); the intestine (dd\_Smed\_v6\_2841\_0\_1); muscle (mhc [39]); and protonephridia (smedinx-10 [42]). Note some neural staining in the pharynx (arrows in f and g). smedinx-10 also stains cells in the pharynx (arrowhead in j) and pigment cups of the eyespots (small arrow in j)

negative control. In parallel, for a positive control, I recommend that dsRNA be generated from a fragment of Smedwi-2 or another gene for which RNAi produces a known phenotype. Smedwi-2(RNAi) causes lysis and death of the planarians in a short period of time [32]. Observation of this phenotype can help ensure that synthesis and dosage are appropriate and consistent.


quickly or when the phenotype prevents feeding (e.g., paralysis, loss of pharynx).


modifications. For example, histone H3 phosphorylated at Serine 10 is a marker of mitotic stem cells [29].


#### Acknowledgments

I would like to thank Bidushi (Tulip) Chandra and Jennifer Jenkins for constructive feedback on this chapter. I would like to thank our funding sources, including the Alfred P. Sloan Foundation and the McKnight Foundation, for financial support.

#### References


32:37–46. https://doi.org/10.1016/j.gde. 2015.01.009


Science 332(6031):811–816. https://doi. org/10.1126/science.1203983


cell function, and tissue homeostasis by systematic gene perturbation in planaria. Dev Cell 8(5):635–649. https://doi.org/10.1016/j. devcel.2005.02.014


optimization of sample processing for immunolabeling. BMC Dev Biol 14:45. https://doi. org/10.1186/s12861-014-0045-6


An RNAi screen reveals intestinal regulators of branching morphogenesis, differentiation, and stem cell proliferation in planarians. Dev Cell 23(4):691–704. https://doi.org/10.1016/j. devcel.2012.09.008


epidermal differentiation. eLife 4:e10501. https://doi.org/10.7554/eLife.10501


Nature 500(7460):81–84. https://doi.org/ 10.1038/nature12414


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Monitoring Chromatin Regulation in Planarians Using Chromatin Immunoprecipitation Followed by Sequencing (ChIP-seq)

### Divya Sridhar and Aziz Aboobaker

#### Abstract

Planarians are an accessible model system to study animal regeneration and stem cells. Over the last two decades, new molecular techniques have provided us with powerful tools to understand whole-body regeneration and pluripotent adult stem cells specifically. We describe a method for performing Chromatin Immunoprecipitation followed by sequencing (ChIP-seq) on planarian cells that relies on FACS to isolate different cell populations followed by immunoprecipitation and library preparation for next-generation sequencing. Whole-genome profiling of histone modifications enables a greater understanding of epigenetic mechanisms in development, pluripotency, and differentiation. This protocol adds to the growing list of functional genomic approaches to study whole-body regeneration in animals.

Key words Planarian, Chromatin, Immunoprecipitation, Epigenetic regulation, Pluripotency, FACS, Histone modifications

#### 1 Introduction

Planarians are best known for their ability to regenerate their whole bodies and owe this remarkable ability to neoblasts, their sole population of pluripotent stem cells. Being the only dividing cell in planarians, neoblasts replace cells lost due to normal physiological turnover as well as injury [1–3]. Neoblasts are thus a promising model system to investigate the epigenetic regulation of pluripotency, stem cell function and differentiation, and tissue patterning during regeneration. Various studies have established conservation of key features of stem cell biology with other animals. Planarians also allow the study of stem cell heterogeneity and lineage progression from undifferentiated stem cells due to the availability of molecular markers for stem cells and their progeny [4, 5]. An advantage of using Schmidtea mediterranea as a model organism for studying epigenetics is the availability of an excellent array of genomic resources and tools to make these studies possible. These include an excellent genome assembly [6], annotations [7], genome database [8], and a transcriptome repository [9]. Nevertheless, while planarians are a promising model system for in vivo stem cell biology, we are only beginning to understand the molecular principles governing the associated regulatory mechanisms. Further research into stem cells in planarians and other model organisms will help us understand fundamental stem cell properties, including disentangling pluripotency and self-renewal [10].

Our understanding of the epigenetic control of regeneration in planarian is in its relative infancy compared to other model organisms. As the epigenetic regulation of gene expression depends on DNA methylation, histone modifications, and overall chromatin organization, understanding these in pluripotent planarian stem cells is of interest to the community [10]. With respect to DNA methylation, a number of strong lines of evidence suggest that 5-methyl cytosine (the major form of DNA methylation in animals) is not part of epigenetic regulation in planarians [10, 11]. S. mediterranea was found to have only the conserved DNA methyltransferase 2 (DNMT2) that despite its name is only thought to methylate RNA [10]. DNA methylation is read by methyl binding domain (MBD) proteins that form key components of histone modifying and chromatin remodeling complexes. In planarians, a single MBD protein, called MBD2/3, has been described. This protein actually lacks the conserved residues known to contact methylated DNA and thus is unlikely to bind 5-methyl cytosine [11]. The absence of the 5-methyl cytosine modification in the S. mediterranea genome was also confirmed in various ways, including the lack of antibody staining against 5-methyl cytosine, and undetectable levels of 5-methyl cytosine in high-performance liquid chromatography mass spectrometry [11]. These different lines of evidence suggest that the function of planarian MBD2/3 is likely independent of DNA methylation, and that DNA methylation is not involved in the epigenetic control of planarian neoblasts. The MBD2/3 protein is known to function in the Nucleosome remodeling and Deacetylase (NuRD) complex in animals. In planarians MBD2/3 and the functions of four other NuRD components have been investigated by RNAi-mediated knockdown in planarians: Smed-HDAC1 [12–14] Smed-CHD [15], RbAp48 [16, 17], and GATAD2 (or p66) [18]. Knockdown of each of these genes affects stem cell differentiation.

The phenotypic effects of the loss of epigenetic regulators that control gene expression can be effectively assessed during planarian regeneration using RNAi. Stem cell survival and differentiation defects can be monitored with in situ hybridization using a growing list of markers. The phenotypes observed are caused by the mis-regulation of gene expression across the genome and often, the mis-regulation of a few key genes have a large effect with respect to the observed phenotype. With the advent of Chromatin Immunoprecipitation followed by sequencing (ChIP-seq) on planarian cells, we can now correlate the phenotypic effects with epigenetic changes at loci across the genome by measuring changes in histone marks in populations of cells as a result of RNAi. By measuring changes in histone marks that induced by RNAi experiments and correlating these changes with gene expression, we can begin to identify epigenetic targets involved in normal stem cell regulation and regeneration. So far studies have confirmed that relationships between gene expression and the enrichment of particular histone marks on nucleosomes proximal to gene promoters present in other animals are conserved [19–21]. For example, as in other animals, higher levels of H3K4me3 are associated with the promoters of actively transcribed genes in planarian stem cells, while H3K27me3 is associated with the promoters of silenced genes [19–21].

ChIP-seq was first used on whole dissociated planarians to show that the histone methyl-transferase enzymes Set1 and MLL1/2, the main mediators of H3K4me3 in animals, target markedly different genomic loci in vivo, respectively [19]. Set1 targets were shown to be associated broadly with the maintenance of basic cell function and survival, while MLL1/2 targets were specifically enriched for genes involved in ciliogenesis. These observations correlate with loss of stem cells in set1(RNAi) animals and the specific loss of cilia and associated locomotion in mll1/2(RNAi) animals. Mihaylova et al. investigated the role of planarian orthologs of a third H3K4 methyltransferase enzymes MLL3/4 [20]. In mammals, loss of MLL3/4 function has been implicated in tumorigenesis [22–24]. RNAi of MLL3/4 in planarians led to the formation of tumor-like outgrowths, suggesting that this histone methyltransferase has tumor suppressor activity in planarians [20]. RNA-seq and ChIP-seq analyses performed on G2/M planarian stem cells from MLL3/4 knockdown animals indicate that genes downstream of MLL3/4 limit or promote stem cell proliferation during regeneration. The MLL3/4 protein plays a role in transcriptional regulation via mono- and/or tri-methylating H3K4 at promoters and enhancers [20] RNA-seq on the same cells revealed that a number of genes involved in cell proliferation and differentiation, including potential oncogenes, were significantly upregulated. The transcriptional changes of some genes following knockdown of planarian MLL3/4 correlate with differences in H3K4me1 peaks at the promoter region, suggestive of direct effect of MLL3/4.

A study by Dattani et al. applied an improved ChIP-seq protocol for neoblasts in S. mediterranea to generate genome-wide profiles for the active marks H3K4me3 and H3K36me3, and suppressive marks H3K4me1 and H3K27me3 [21] in order to look at epigenetic regulation of gene expression in neoblasts. As predicted from work in vertebrates and other protostomes, these marks showed conserved patterns of association with active and suppressed gene expression in planarian neoblasts. Significantly, loci that have little or no transcriptional activity in the neoblast compartment and are known to activate transcriptionally in the postmitotic progeny during differentiation show bivalent histone modifications, with both H3K4me3 and H3K27me3 marks at promoter regions. ChIP-seq also revealed high levels of paused RNA Polymerase II at the promoter-proximal region as further evidence that these genes are bivalent in neoblasts, becoming actively transcribed upon differentiation. These findings suggest that epigenetic regulation of potency through bivalency at promoter regions is conserved across bilaterians, rather than a special feature of vertebrates [21]. Overall, these studies have established that ChIP-seq can be efficiently used in neoblasts to investigate epigenetic regulation of stem cell fate.

In this chapter, we provide step-by-step robust protocols for cell dissociation and isolation of planarian cells, chromatin extraction and sonication, immunoprecipitation, and preparation of ChIP libraries (Fig. 1). We also outline a range of quality control steps that could be used at various stages of the protocol.

#### 2 Materials

All solutions should be prepared using ultrapure water and analytical grade reagents. Reagents should be prepared and stored at the temperatures indicated. Local waste disposal regulations should be adhered to when disposing of chemical and plastic waste.

#### 2.1 Cell Dissociation and Isolation of Stem Cells


Fig. 1 An overview of the ChIP-seq workflow

Make up the volume to 50 mL using H2O. Make fresh for each use (see Note 2).


2.2 Chromatin Extraction and Sonication

	- 1. Protein-A covered beads.
	- 2. Magnetic separation rack for 1.5-mL tubes.
	- 3. Blocking solution: 0.5% BSA in 1 PBS.
	- 4. ChIP-grade antibodies: 7 μg antibody per sample (see Note 4).
	- 5. Commercial Drosophila S2 chromatin as internal immunoprecipitation control (store-bought, e.g., Active Motif).
	- 6. ChIP wash buffer: 50 mM HEPES–KOH, pH 8, 0.5 M LiCl, 1 mM EDTA, 1% NP-40, 1% sodium deoxycholate, 1 protease inhibitors tablet. Make fresh for each use.
	- 7. Tris–EDTA buffer (TE): 1 TE buffer (store-bought).
	- 8. 0.1 TE: 10 μL TE, 90 μL water.
	- 9. TE-SDS: 2% (v/v) SDS in 1 TE.
	- 2. Agencourt AMPure XP beads (Beckman Coulter).

2.3 Immunoprecipitation and Reverse Crosslinking

2.4 Preparation of ChIP Libraries for Sequencing


#### 3 Methods

3.1 Dissociation and Isolation of Stem Cells


#### 1. Transfer cells from FACS tubes to protein low-binding 1.5-mL tubes.


#### 3.2 Chromatin Extraction and Sonication


Fig. 2 Example fragment size distribution analyzed on TapeStation (a) after chromatin shearing, (b) after a test de-crosslink. The optimal chromatin size distribution for ChIP-seq is between 200 and 800 bp


#### 3.3 Chromatin Immunoprecipitation


3.4 Preparation of ChIP Libraries for Sequencing


#### 3.5 PCR Enrichment and Purification of DNA


Initial denaturation at 98 C for 30 s/N [denaturation at 98 C for 10 s, annealing/extension at 65 C for 75 s] followed by final extension at 65 C for 5 min. N ¼ 3–15 (see Note 18).


Fig. 3 Size distribution of (a) input along with 3 ChIP DNA libraries, (b) ChIP DNA library

#### 4 Notes


Scaling of PCR amplification cycles based on input DNA. The ideal number of PCR cycles to amplify libraries for sequencing depends on the amount of DNA that goes into the end repair reaction (Subheading 3.4). In our experience, this can vary depending on the antibody used and the amount of chromatin used in immunoprecipitation


Table 1 and serve as a starting point to determine the number of PCR cycles best for standard library preparation.

19. Qubit, agarose gel, TapeStation or bioanalyzer can be used to test the suitability of the libraries for sequencing.

#### References


Schmidtea mediterranea and the evolution of core cellular mechanisms. Nature 554(7690): 56–61. h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature25473


A (2013) Planarian MBD2/3 is required for adult stem cell pluripotency independently of DNA methylation. Dev Biol 384:141–153. https://doi.org/10.1016/j.ydbio.2013. 09.020


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Assessing Chromatin Accessibility During WBR in Acoels

### Andrew R. Gehrke and Mansi Srivastava

#### Abstract

Dynamic gene expression seen during whole-body regeneration is likely controlled by genomic regulatory elements that dictate the spatiotemporal activity of the regeneration transcriptome. Identifying and characterizing these non-coding regulatory sequences are key to understanding how genes are connected into networks to deploy the process of whole-body regeneration. Here, we describe the application of the Assay for Transposase Accessible Chromatin (ATAC-seq) in the acoel Hofstenia miamia to identify regions of open chromatin that represent putative regulatory elements. Notably, when paired with gene knockdown techniques such as RNAi, ATAC-seq can be implemented in a functional genomics approach to validate putative regulatory elements. ATAC-seq requires no species-specific reagents, is amenable to small input cell numbers, and can be completed in a single day, making it an ideal assay to identify dynamic chromatin at high resolution during whole-body regeneration in virtually any species with a quality genome assembly.

Key words ATAC-seq, Chromatin, Gene regulation, Acoel, Whole-body regeneration

#### 1 Introduction

The crucial role of regulatory elements that comprise the non-coding genome has been demonstrated in development, disease, and evolution [1, 2]. Advances in genomics (e.g., the ability to sequence and assemble myriads of animal genomes) and techniques in molecular biology have now made it possible to explore the role of the regulatory genome in the process of whole-body regeneration. Previous techniques to characterize regulatory elements have relied either on species-specific reagents or a large number of input cells, hindering the genome-wide identification of putative enhancers in emerging model systems. The Assay for Transposase Accessible Chromatin (ATAC-seq) [3], which is relatively wet-lab simple and requires a small amount of input material, has the potential to revolutionize the fields of functional genomics and evolutionarydevelopmental biology by providing a method to identify putative enhancers at high resolution in emerging systems of study (Fig. 1).

ATAC-seq works by treating a small number of permeabilized cells or exposed nuclei to a transposase enzyme that preferentially

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_29, © The Author(s) 2022

Fig. 1 Overview of an ATAC-seq seq experiment to assay regeneration-responsive chromatin. ATAC-seq involved applying a transposase (left panel) capable of cutting open chromatin and simultaneously ligating in sequencing primers ("tagmentation"). The transposase enzyme will integrate less in closed chromatin (WT) and will preferentially insert into open chromatin, e.g., a region that harbors an enhancer (blue) that opens during regeneration ("regen," bottom left). The final library consists of small regions of open chromatin that are ready to be sequenced. After alignment of these sequences to the genome (green lines, right panel), "peaks" of open chromatin (green) can be called and compared across regenerating samples (differential accessibility). Transcription factor (TF) binding can be inferred by viewing the number of transposase cutting events (# cuts) around TF binding sites. When a TF is bound, it occludes the transposase from inserting into that region and leaves a "footprint," which can be compared across samples ("differential TF footprinting")

accesses regions of open chromatin, simultaneously cutting DNA and inserting primers for sequencing ("tagmentation") (Fig. 1). Following sequencing, reads mapped to the genome provide information on open chromatin, nucleosome position, and transcription factor binding. The main benefits of the assay are (1) no speciesspecific reagents, (2) low input required, from 50,000 cells down to a few thousand, (3) reproducibility, in that replicates are highly concordant, and (4) speed, one can go from intact tissue to a sequencing-ready library in a single day.

Due to the experimental ease and high resolution of ATAC-seq, a number of methods papers have been published that describe the assay in detail. These include step-by-step instructions for cell lines [4], zebrafish [5, 6], echinoderms [7], xenopus [8], and plants [9]. Recent advances to the protocol ("Omni-ATAC") have improved the sensitivity of the assay and made it possible to perform in frozen tissues [10]. In addition to the wet-lab protocols for ATAC-seq, there are a number of methods papers that describe the bioinformatic data analysis portion of ATAC-seq [11–13]. The majority of the wet and dry lab portions of ATAC-seq are quite similar across organisms and do not deviate much from the original methods paper describing the assay [4]. The critical factor when performing ATAC-seq in a "new" species is attaining the correct number of cells for proper transposition. Keeping this as a focus, here we describe step-by-step instructions for ATAC-seq in the acoel worm Hofstenia miamia. A defining step of this protocol is direct disruption of tissue in lysis buffer (as opposed to traditional dissociation and cell counting), followed immediately by transposition. This rapid processing of samples likely reduces background noise and better captures transcription factor binding as inferred by footprinting. We envision that this protocol will work robustly for all invertebrate animals that are generally easy to lyse or dissociate into single cells.

#### 2 Materials


Primer sequences. Primer sequences used for PCR, table reproduced from Supplementary Table 1 of [3]


(continued)



#### 3 Methods

Care should be taken to move as quickly as possible from tissue extraction to dissociation to retain chromatin state at the appropriate timepoint. This protocol is based on the original ATAC-seq protocol [3]. Modifications that improve the assay have been described ("Omni-ATAC") [10], but use a detergent mixture that may be harmful to more sensitive cells. Thus, we suggest attempting the original protocol first and subsequently exploring the Omni-ATAC modifications to potentially improve the experiment.



3.2 PCR Amplification of Library and Sequencing

Fig. 2 Tapestation examples of different quality ATAC libraries. ATAC libraries were run on an Agilent Tapestation 2200 using an HD5000 tape. "Ideal" trace shows extensive nucleosomal "laddering," indicating a high-quality library. The "acceptable" trace also shows nucleosomal laddering, but with an extended sub-nucleosomal peak that may indicate slight "over-tagmentation" (over-cutting of the enzyme, likely due to too few cells in the reaction). If no "ideal" libraries are present this library is acceptable to sequence. "Overtagmented" shows no laddering and only a single peak, likely due to too few cells being added to the reaction. This library should not be sequenced, and the experiment should be repeated with more input cells. The "undertagmented" (insufficient cutting by the enzyme) trace shows no nucleosomal laddering but also no clear sub-nucleosomal peak, which could be the result of too many cells in the reaction. This library should not be sequenced, and the experiment should be repeated with fewer cells


Fig. 3 Standard bioinformatic workflow for processing and analyzing ATAC-seq data. Raw reads are removed of adapters, then aligned to the genome of interest with no additional quality trimming. Duplicate reads are removed, as well as reads that map to the mitochondrial genome, and reads that are not properly paired. We use Genrich to call peaks on each biological replicate, and then use IDR to call reproducible peaks between replicates. Finally, we use bedtools merge on all peaksets from all samples to create a non-overlapping set of peaks that represents a consensus peakset. For analysis, we use Diffbind to call differentially accessible peaks between samples, ChIPseeker to make peak-to-gene connections, and TOBIAS for footprinting and bound site calling. Results are best visualized first using IGV, and then pyGenomeTracks to create publicationquality figures

> changed to the designation of the mitochondrial genome in the input assembly (e.g., scaffoldX).


#### 4 Notes


Fig. 4 Expected results. (a) ~15 kb region of the Hofstenia genome that encompasses a gene, runt, showing ATAC data for 0 h and 6 h, along with with the consensus peakset, Diffbind-called regeneration peaks, and sites that are bound by the TF egr at 6 h by TOBIAS. There are two major regeneration-responsive peaks (shown in red), the first being the promoter, and the second in an intron. Both the regeneration peaks have "bound" sites for egr. (b) A second region of the Hofstenia genome, this time showing an intergenic regeneration-responsive peak upstream of a putative target gene


calls for all predicted sites of a particular TF in the genome, as well as create aggregate plots that show overall binding differences between timepoints. See Fig. 3 for a genome browser example of these types of data combined.

12. Depending on downstream analysis, removing multi-mapped reads and retaining only properly paired reads may be unnecessary or detrimental (too few reads retained). For instance, peak-calling with Genrich can utilize both multi-mapped and unpaired alignments in generating peak calls. We recommend checking the documentation of software for the desired downstream application to determine how best to handle these reads.

#### Acknowledgments

We thank Jose Luis Gomez Skarmeta (Centro Andaluz de Biologı´a del Desarrollo, Sevilla, Spain) and Elisa de la Calle-Mustienes (Centro Andaluz de Biologı´a del Desarrollo, Sevilla, Spain) for assistance with initial Hofstenia ATAC-seq experiments, performed at the Marine Biological Laboratories. We thank Kyle McCulloch for critical reading of the manuscript. A.R.G. is supported by the Simeon J. Fortin Charitable Foundation, Bank of America, N.A. Trustee, and the Helen Hay Whitney Foundation, M.S. is supported by the Milton Foundation of Harvard University and the National Science Foundation (award no.1652104).

#### References


transposase-accessible chromatin and circularized chromosome conformation capture, two methods to explore the regulatory landscapes of genes in zebrafish. Methods Cell Biol 135: 413–430. https://doi.org/10.1016/bs.mcb. 2016.02.008


genomes using ATAC-Seq. Methods Mol Biol 1675:183–201


(2019) Assaying chromatin accessibility using ATAC-Seq in invertebrate chordate embryos. Front Cell Dev Biol 7:372


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part V

OMICS Approaches

# Single-Cell Transcriptomic Analysis in the Regenerating Cnidarian Nematostella vectensis

# Flora Plessier, Sandrine Schmutz, Sophie Novault, and Heather Marlow

#### Abstract

Cnidarians have historically served as excellent laboratory models for regenerative development given their capacity to regrow large portions of the adult organism. This capacity is notably absent or poorly developed in the powerful genetic laboratory models Drosophila, C. elegans, and mouse. Increasingly, development of genetic and genomic resources and the application of next-generation sequencing-based techniques in cnidarian systems has further expanded the potential of cnidarian regenerative models. Here, we present a workflow for the characterization of the regenerative response in the sea anemone Nematostella vectensis utilizing fluorescence-activated cell sorting and a plate-based single-cell RNA-sequencing pipeline. This approach can characterize the transcriptional response during regeneration in distinct populations of cells, thus providing a quantitative view of a whole organism process at cellular resolution.

Key words Single-cell RNA-seq, Cnidaria, Nematostella vectensis, Regeneration, Flow cytometry

#### 1 Introduction

The ability to regenerate is widespread among marine invertebrates. Understanding the molecular and developmental basis of regeneration in a diversity of taxa has the potential to provide insight into shared principles of the regenerative response. Cnidarians (Hydra, jellyfish, sea anemones, corals) belong to one of the earliest-branching metazoan phyla and are well known for their extensive regenerative abilities [1–3]. The sea anemone Nematostella vectensis is an experimentally tractable Cnidarian species for studies into mechanisms of regeneration. Significant advances in genetic and genomic resources, ease of culture, and conservation of gene content with Bilaterian taxa contribute to the utility of this sea anemone as a developmental and regenerative model system [4–6].

An experimentally bisected Nematostella vectensis polyp can regenerate two whole animals in about a week [1]. How the sea anemone regenerates lost tissue, maintains axial coordinates, and generates the appropriate number and distribution of constituent

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_30, © The Author(s) 2022

cell types during regeneration or asexual reproduction is not well understood. Studies focusing on the cellular dynamics following oral pole regeneration in juvenile polyps have highlighted the involvement of cellular proliferation and apoptosis [1, 7]. Wholeanimal transcriptomic time-courses have been employed to uncover genes modulated during the regeneration process [8–10]. However, the behaviors of individual cell populations during regeneration have not been molecularly or functionally characterized in Nematostella vectensis thus far [11]. In order to better characterize the process of regeneration, additional tools that allow for the characterization of behaviors of single cells are needed. As regeneration is dynamic, gaining an understanding of the molecular events happening in individual cells throughout this process is critical.

Over the past few years, newly developed single-cell RNA-Sequencing (scRNA-Seq) techniques have enabled investigators to sample the transcriptome of single cells from complex tissues and whole organisms, to assess their developmental trajectories and reconstruct gene regulatory networks governing cell identity and function [12]. Different single-cell sequencing technologies have been developed ranging from plate-based, droplet or microfluidic platforms, each with distinct advantages and limitations [13]. Recently, single-cell RNA-Seq of whole Nematostella vectensis polyps and planula larvae via a plate-based approach, MARS-seq, has enabled the generation of a molecular atlas of the cell types present in the sea anemone [14]. In combination with targeted genetic labeling and trajectory inference methods, scRNA-Seq data from regenerating animals could enable the reconstruction of key cellular trajectories, as well as the relative contributions of different cell populations to the newly regenerated tissues.

From cell suspension, most scRNA-seq protocols rely on a similar workflow, where single cells are physically isolated and lysed, in wells or droplets, captured mRNA molecules are reverse transcribed and amplified, and multiplexed dsDNA libraries are generated and sequenced [13]. Complete, high-viability dissociation of starting tissue is a critical pre-requisite of scRNA-seq. Single cells can then be isolated through Fluorescence-activated cell sorting (FACS), which enables a single cell to be separated from a cell suspension with a high degree of purity. Following single-cell isolation, mature mRNAs are often captured through their polyadenylated tails using barcoded oligos bearing polyT stretches, then reverse transcribed into cDNAs before being amplified via PCR or in vitro transcription [15]. During capture or in subsequent steps, cell- and/or pool-specific barcodes are added, to enable attribution of the sequenced reads to a specific cell and sample of origin. To limit the impact of amplification artifacts, unique molecular identifiers (UMIs), short stretches of random DNA bases, are attached to the captured mRNA molecules [15, 16]. Duplicate reads from the same mRNA molecule can be merged by UMI. For data processing and analysis, several quality control and clustering analysis packages have been developed [17–19].

In this protocol, we focus on a single-cell RNA-sequencing experiment in a regenerating Nematostella vectensis polyp including the dissociation of samples of interest into live single-cell suspensions, the flow cytometer parameters and gating strategy to sort live single cells into separate wells of a 384-well plate, as well as an overview of quality control steps after library sequencing, read demultiplexing and mapping, QCs, and general notes and considerations specific to single-cell RNA-Seq Nematostella vectensis data (Fig. 1). Briefly, polyps along a regeneration time-course following oral pole amputation are dissociated into single-cell suspensions at timepoints of interest (Fig. 1a), then live single cells are sorted on a flow cytometer into single wells of a 384-well plate (Fig. 1b). Following cell lysis and mRNA capture with barcoded oligonucleotides in individual wells, multiplexed cDNA libraries are generated using the MARS-Seq single-cell RNA-Seq method [15, 20] (Fig. 1c). The scRNA-seq library preparation method has been adapted from an established workflow and an extensive protocol detailing the experimental steps, and an associated computational pipeline has recently been published [20]. After library sequencing, reads are filtered, demultiplexed, and mapped using the MARS-Seq computational pipeline onto the Nematostella genome to generate count tables, listing the number of recovered molecules from each gene in each sequenced cell [20, 21] (Fig. 1d). The count tables can then be filtered and clustered using various clustering methods for biological analyses [22] (Fig. 1d). Here, we generate an example clustering using the MetaCell clustering package, a published pipeline for the analysis of scRNA-seq data [17].

#### 2 Materials

All solutions should be prepared using ultrapure water and stored at room temperature (RT) unless otherwise stated.


Fig. 1 Overview of single-cell RNA Sequencing in the regenerating sea anemone Nematostella vectensis. (a) DIC images of the oral region of uncut control, 2 and KCl, 27.6 mM NaHCO3, 50 mM Tris–HCl pH 8. Filter sterilize.



Fig. 1 (continued) 6 day post-amputation (2 dpa and 6 dpa) juvenile Nematostella vectensis polyps (~5 weeks) at room temperature. Amputation site is indicated with a red line. (b) Enzymatic dissociation of sample into a singlecell suspension and single-cell FACS sorting into 384-well plates. (c) Simplified overview of the labeling strategy for single-cell-derived mRNA which includes barcoding for each cell in each well using capture oligonucleotides with a polyTtract, a random unique molecular identifier (UMI) and a cell barcode. (d) Multiplexed single-cell RNA-seq short-read libraries are sequenced, reads are filtered, mapped, and demultiplexed to attribute recovered molecules, through their unique UMIs tags, to their cell of origin. Cells are then clustered using unsupervised clustering based on gene expression for downstream analyses


#### 3 Methods


3.2 Live Single-Cell Sorting Using Flow Cytometry for Single-Cell RNA-Sequencing


Fig. 2 FACS gating strategy for live cell-sized singlets with live/dead double-staining. (a) Gating strategy to select live single cells from sample stained with calcein AM and propidium iodide (PI). (b) Overall events SSC-A vs. FSC-A profile. Note that we are using bi-exponential scaling on both FSC-A and SSC-A because of the size heterogeneities in our samples. (c) Singlet gate to exclude multiplets based on FSC-H vs. FSC-W profile. Singlet proportion is usually in the 90–95% range. (d) From the singlet population, the live cell gating strategy is based on calcein (live) signal inclusion and propidium iodide (permeabilized dead cells) signal exclusion. (e) From the live singlet population, setup of the cell-sized particle gate, to keep only live single cells for sorting. For setting up this cell-sized gate boundaries, see Fig. 3a, g. The proportion of cells that fall within the represented gate is indicated in red

marker and dead cell marker (gated population in purple in Fig. 2d).

Fig. 3 FACS gating strategy setup for live cell-sized singlets with absolute size comparison. (a) Gating strategy to select live single cells. (b) All events gate from a whole animal cell suspension (SSC-A vs. FSC-A). Note that we are using bi-exponential scaling on both FSC-A (except in panel g) and SSC-A given the inherent size heterogeneities of Nematostella vectensis whole organism cell suspensions. (c) Overlay of three sizing beads FCS-A profiles (1.34 μm, 3.1 μm, and 6 μm beads) to set the FSC-A boundary of the cell-sized gate presented


1. Process plates through the MARS-Seq library preparation pipeline with N ¼ 17 cycle of PCR amplification at the last step.


Fig. 3 (continued) in panel g. (d) Singlet gate to exclude multiplets based on their FSC-H vs. FSC-W profile. (e) Live cell gating strategy based on calcein Violet signal to keep only cells with calcein (live) signal. (f) and (g) Cell-sized particle gate. Gate boundary is based on beads sizing gate, including all particles ~3 μm and above to maximize cells and minimize debris. Representation in a biexponential scale enables clearer demarcation of cell-sized live particles. The proportion of cells that fall within the represented gate is indicated in red.

3.3 Single-Cell RNA-Seq Library Generation, QC, and Initial Characterization

Fig. 4 Library quality metrics of a sequenced single-cell RNA-seq experiment. (a) Example of a typical library tapestation profile of a single MARS-Seq half-plate library (red arrow). Expected fragment size is usually in the 410–440 bp range. Black arrowheads indicate the upper and lower lane markers on a D1000 high sensitivity tape. (b) Sequencing depth statistics across sequenced libraries, by half-plate (192 cells) library. Median read depth per library is indicated in dark red at the top. Median sequencing depth target is 40,000 reads/cell when sequencing 32 MARS-Seq libraries on a NextSeq 500 high output kit. Note the two libraries flagged in orange

depth, cell, and sample type. UMI recovery per cell correlates with the overall cell size, with bigger cells yielding higher UMI counts.


#### 4 Notes


Fig. 4 (continued) whose median sequencing depth is much lower, indicating potentially either a sample issue, or library undersequencing or library quality issues. As their matched other half-plates show the expected sequencing depth, it seems likely to be a technical issue arising during processing or library pooling. The other libraries pass this check. (c) Unique Molecular Identifiers (UMI) recovery statistics across half-plate libraries. Same libraries as shown in b. Note that flagged libraries (in orange in b and c) also display a lower UMI distribution compared to their other half-plate. The two empty wells kept as negative controls (red Xs) per half-plate display very low UMI recovery (noise) compared to wells that held one cell (blue dots)

Fig. 5 UMI distribution in cells passing QC and clustering example from a N. vectensis regeneration experiment. (a) UMI distribution across all cells. Here, cells from which between 120 and 3000 (red lines) UMIs were recovered were kept for downstream analyses. (b) 2D-projection of 1300 cells clustered here using the MetaCell analysis package, colors are used to demarcate individual MetaCell clusters. (c) Gene expression (UMIs per 1000 UMIs in the cluster) for known marker genes from [14] are used to annotate the clusters. (d) 2D-projection of control and regenerated oral regions at 4/6 days post-amputation. Overall 2D clustering is

visualization, by selecting mOrange signal-positive cells that are not also GFP-like positive.


Fig. 5 (continued) split by sample origin (control or regenerated), and colored according to a preliminary MetaCell annotation based on known markers published in the N. vectensis cell atlas [14]. Most cells are split according to their prospective cell state, with newly regenerated oral side samples falling within all broad cell types


#### References


development and regeneration. Wiley Interdiscip Rev Dev Biol 5:408–428


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Characterization of Soluble Cell-Free Coelomic Fluid Proteome from the Starfish Marthasterias glacialis

# Laidson Paes Gomes , Catarina Gouveia e Silva, Jean-Charles Gaillard, Jean Armengaud , and Ana Varela Coelho

#### Abstract

Proteomics combined to advanced bioinformatics tools is acquiring a pivotal role in the comprehensive understanding of living organism's biology, in particular for non-model organisms, which includes most marine and aquatic invertebrates. Depicting of protein composition in a whole organ/organism followed by their assembling in functional protein association networks promotes the understanding of key biological processes. Here, we provide a detailed description of the extraction procedure of cell-free coelomic fluid soluble proteins and the characterization of the proteome of the starfish Marthasterias glacialis. Due to coelomic fluid richness in glycoproteins, which complicates protein identification, extracts of soluble proteins are deglycosylated prior to tandem mass spectrometry. This experimental approach is useful at improving knowledge on the coelomic fluid physiological role and deciphering its involvement in regeneration of starfish body parts when comparing different regeneration conditions.

Key words Starfish, Marthasterias glacialis, Coelomic fluid, Proteomics, Tandem mass spectrometry

#### 1 Introduction

Proteomics is the study of multiple protein systems with focus on the interplay of multiple, distinct proteins, and their roles as part of a larger system or network [1]. Mass spectrometry (MS), coupled to liquid chromatography (LC–MS), is the most used wellestablished methodology for proteome investigations [2]. A mass spectrometer measures the mass/charge ratio (m/z) of generated peptide molecular ions and their relative abundances allowing to characterize, identify, and quantify peptides and proteins. Protein extracts rich in glycoproteins can be enzymatically deglycosylated prior to LC-MS analysis to extend the number of identified proteins [3]. To make sense of the extensive sets of identified proteins by proteomics, a functional analysis is required. It tends to organize the identified proteins in biochemical, cellular, biological, and disease-related categories according to the process that they are

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_31, © The Author(s) 2022

associated with. These different steps are hampered in non-model organisms due to the lack of biological knowledge and complete genome information.

Regenerative potential is expressed to a maximum extent in echinoderms. Starfish are capable of reconstructing external appendages and internal organs often subjected to amputation, selfinduced or traumatic, rapidly followed by complete successful re-growth of lost parts. Marthasterias glacialis (Linne´ 1758) is a fairly common asteroid echinoderm widely distributed throughout the northern Europe; it is often found on the Atlantic coast of central/northern Portugal due to its preference for glacial waters. This spiny starfish, which surface is covered with thorns, can be found in waters up to 200 m in a variety of habitats from muddy, protected locations to rock exposed waves [4]. M. glacialis normally reaches 25–30 cm in diameter, has five arms, each having three rows of thorns, and differs in color from dark brown to greenish gray. It is a voracious predator feeding on various animals, dead or alive, such as mollusks, fishes, crustaceans, or other echinoderms [5]. Starting from only one fifth of the central disc, M. glacialis can survive and regenerate a new individual [4]. However, its regeneration process is slow and complex. Its arm tip regeneration, or even of an entire arm lost by autotomy, can take from a couple of weeks to several months [4]. This species can be seen as an important model for the study of regeneration due to 70% genome similarity with humans [4], its extraordinary regeneration ability [6], and its easy collection from the wild and maintenance in aquaria.

The coelomic fluid circulates the water vascular system, an internal network composed of channels that contacts their internal organs. The liquid part of the coelomic fluid consists of seawater and is extremely rich in secreted molecules. This fluid is responsible for the transport of circulating cells (coelomocytes), nutrients, and metabolites bathing the internal organs [7]. Major coelomocyte morphotypes have the primary function of mediating immune responses, being able to recognize and neutralize foreign material [8, 9]. Humoral responses on echinoderms are represented by a great variety of molecules, like lectins, perforins, and cytokines, secreted by coelomocytes or surrounding tissues, and promoting cell migration, agglutination, and healing [9]. Recently, the Asterias rubens coelomocyte-free coelomic fluid proteome was characterized by LC-MALDI tandem mass spectrometry, identifying 91 proteins [10]. The most represented functional categories were pattern recognition receptors and peptidase inhibitors. Proteins known to be involved in the process of sea cucumber intestinal regeneration, such as ependymin, β-microseminoprotein, serum amyloid A and avidin-like proteins, have also been identified. Proteome characterization of cell-free coelomic fluid soluble proteins suggests that this fluid plays an important role in cell signaling, transport, and responses to injury in starfish, constituting a relevant tissue to be studied to deeply elucidate the molecular processes associated with starfish organ regeneration. The protocol presented here below can be easily applied to other invertebrate fluids and to quantitative differential proteomics studies involving regeneration or other physiological challenges.

Figure 1 summarizes the shotgun proteomics experimental workflow for the characterization of the soluble proteome of cellfree coelomic fluid (CFF) from M. glacialis. In order to extend the number of identified proteins, half of the precipitated protein extract is deglycosylated with Peptide-N-Glycosidase F (PNGase F). PNGase F enzymatic treatment removes the N-linked oligosaccharides from glycoproteins since it cleaves between the innermost N-acetylglucosamine (GlcNAc) and asparagine residues of high mannose, hybrid, and complex oligosaccharides [3]. Since this step can reduce the sensitivity of the analytical procedure, the non-deglycosylated extract is also assayed. Proteins are in-gel digested with trypsin previous to identification by LC-MSMS analysis. Functional analysis of the identified proteins, including prediction of N-glycosylation sites [11], is used to allow a comprehensive description of the metabolic and biological processes occurring in this tissue.

#### 2 Materials

Use bidistilled water and room temperature unless otherwise stated.


Fig. 1 Experimental workflow for the characterization of CFF soluble proteome from M. glacialis. Each step of this protocol is described in text boxes and the workflow follows the sequence of events indicated by the arrows

	- 2. Enhancer of trypsin enzymatic performance: 0.01% (v/v) ProteaseMax™ in AB.
	- 3. 50% (v/v) methanol in AB.
	- 4. Acetonitrile pro-analysis grade (ACN).
	- 5. 50% (v/v) ACN in AB.
	- 6. 25 mM DTT in AB.
	- 7. 55 mM iodoacetamide in AB.
	- 8. 5% (v/v) trifluoroacetic acid.

2.4 Untargeted Proteomics and Data Analysis


#### 3 Methods

3.1 Cell-Free Coelomic Fluid Collection and Protein Precipitation

	- 3. Hold the animal with the punctured arm hanging.
	- 4. Place a 15-mL tube below the arm with the Venipuncture capillary inside it.
	- 5. Collect 15 mL of the coelomic fluid by gravity into the tube.
	- 6. Centrifugate the sample at 1000 rcf for 5 min at 4 C (see Note 5).

1. Transfer 40 μg of the CFF protein extract to a 1.5-mL tube.


3.2 Peptide-N-Glycosidase F Enzymatic Treatment and SDS-PAGE Sample Preparation


3.3 Denaturing Polyacrylamide Gel Electrophoresis (SDS-PAGE): Gel Casting and Sample Running

Fig. 2 Polyacrylamide gel electrophoresis: gel casting and sample running. (a, b) Assembling the gel cassette using two glass plates and two spacers. (c) The comb is introduced between the two glass plates after pouring the acrylamide stacking gel solution. (d) The gel cassette with the polymerized acrylamide (polyacrylamide) is included in the electrode assembly before introducing it inside the tank of the electrophoresis system. (e) Protein bands stained with Coomassie Blue after 2 cm migration of the non-deglycosylated and deglycosylated CFF total protein extract. The rectangle delimits the region of the gel to be excised and divided in four strips, here defined by the square brackets, before in-gel trypsinization

#### 3.4 Coomassie Blue Gel Staining

	- 2. Transfer the bands to separate 1.5-mL tubes.
	- 3. Add 200 μL of 50% (v/v) methanol in AB.
	- 4. Vortex for 1 min at 500 rpm.
	- 5. Repeat steps 3 and 4 to destain the gel bands.
	- 6. Replace the solution with 200 μL 50% (v/v) ACN in AB.
	- 7. Vortex for 5 min at 500 rpm.
	- 8. Replace the solution with 100% acetonitrile.
	- 9. Vortex for 1 min at 500 rpm.
	- 10. Remove the supernatant.
	- 11. Dry the gel bands in a vacuum centrifuge.
	- 12. Add 100 μL of 25 mM DTT in 50 mM AB.
	- 13. Incubate for 10 min at 56 C under 500 rpm agitation in a ThermoMixer® to rehydrate the gel and reduce the proteins persulfate bonds.
	- 14. Add 100 μL of 55 mM iodoacetamide in 50 mM AB.
	- 15. Incubate for 10 min in the dark to alkylate the proteins thiol groups.
	- 16. Wash twice the gel bands with water as in steps 9 and 10.
	- 17. Dehydrate the gel bands as described in step 11.
	- 18. Incubate the dried in-gel digests with 40 μL 0.01 μg/μL trypsin in 0.01% (v/v) trifluoroacetic acid for 15 min on ice.
	- 19. Discard the supernatant.
	- 20. Add 70 μL of the enhancer of trypsin enzymatic performance solution.
	- 21. Incubate for 1 h at 50 C.
	- 22. Stop the proteolysis by adding 5 μL of 5% (v/v) trifluoroacetic acid (see Note 13).
	- 1. Load 3 μL of each tryptic peptide mixture to be online desalted on the RP C18 trapping column.
	- 2. Resolve the tryptic peptides on the RP C18 analytical column at a flow rate of 200 nL/min with a 90 min gradient of 4–25% of solvent A in 75 min and 25–40% of solvent B in 15 min (see Notes 14 and 15).
	- 3. The mass spectrometer is operated in data-dependent method consisting in a scan cycle initiated with a full scan of peptide ions, followed by selection of the precursor molecular ion, high energy collisional dissociation, and MS/MS scans on the

3.6 Liquid Chromatography– Mass Spectrometry Untargeted Proteomics Analysis

3.5 Protein In-Gel Trypsination

Fig. 3 Pie chart illustrating chosen relevant KEGG pathways from STRING. A total of 51proteins were classified in seven biological processes other than carbon metabolism

20 most abundant precursor ions. Full scan mass spectra are acquired from m/z 350–1500 with a resolution of 60,000. Each MS/MS scan is acquired with a threshold intensity of 83,000, on potential charge states of 2+ and 3+ after ion selection performed with a dynamic exclusion of 10 s, maximum IT of 60 ms and an m/z isolation window of 2.

4. MS/MS spectra at a resolution of 15,000 are search against the established echinoderm assembled protein database using MASCOT 2.5.1 software. The peptide matches with a MAS-COT peptide score below a p-value of 0.05 were filtered and assigned to proteins.


#### 4 Notes


identification was performed using the protein sequences available from other echinoderm specimens. This database was constructed compiling protein data from several databases, such as NCBI (https://www.ncbi.nlm.nih.gov/protein/), UniProt (https://www.uniprot.org/), and Echinobase (https://www.echinobase.org/entry/).


interaction, mTOR, AGE-RAGE, and Wnt signaling pathways).

20. PNGase F removes the internal N-linked oligosaccharides from glycoproteins. This enzyme catalyzes the cleavage between N-acetyl-D-glucosamine and asparagine residues. The asparagine residue from which the oligosaccharide is removed is deaminated to aspartic acid. For M. glacialis coelomic fluid 43 proteins were only detected after PNGase F treatment and with an asparagine residue to aspartic acid modification. Thirty-seven out of these proteins had glycosylation site predictions, and 31 had more than one glycosylation site predictions.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Using RNA-Seq for Transcriptome Profiling of Botrylloides sp. Regeneration

### Michael Meier and Megan J. Wilson

#### Abstract

The decrease in sequencing costs and technology improvements has led to the adoption of RNA-sequencing to profile transcriptomes from further non-traditional regeneration model organisms such as the colonial ascidian Botrylloides leachii. The relatively unbiased way in which transcripts are identified and quantified makes this technique suitable to detect large-scale changes in expression, and the identification of novel transcripts and isoforms. Of particular interest to many researchers is the discovery of differentially expressed transcripts across different treatment conditions or stages of regeneration. This protocol describes a workflow starting from processing raw sequencing reads, mapping reads, assembly of transcripts, and measuring their abundance, creating lists of differentially expressed genes and their biological interpretation using gene ontologies. All programs used in this protocol are open-source software tools and freely available.

Key words Regeneration, Sequencing, RNA-seq, Ascidian

#### 1 Introduction

Ascidians, also known as tunicates or sea squirts, are marine invertebrate filter-feeding animals. They belong to the Tunicata subphylum, an extant sister clade of the vertebrate clade in the chordate phylum. The close evolutionary relationship is most evident during embryo development where ascidians share vertebrate morphological features such as a notochord and a neural tube. Broadly, ascidians can be classified as solitary or colonial and whether they are pelagic or sessile. Botrylloid colonial ascidians have become an important model organism to study whole-body regeneration, chordate evolution, immunobiology, and allorecognition [1– 7]. Fueled by recent advances in next-generation sequencing technology, RNA-sequencing has become a standard tool to characterize entire transcriptomes of cells, tissues, and whole organisms. This protocol uses publicly available RNA-seq data from a regeneration time course experiment in Botrylloides leachii [1]. Additionally, with the availability of a sequenced genome [8], a wider range of software tools (for sequenced organisms) are now also available to analyze this RNA-seq data from B. leachii.

Despite the wide use of RNA-seq, data analysis workflows are equally numerous and not yet standardized [9]. RNA-seq can provide a snapshot of all transcripts present in a cell or tissue of interest. Often though, the research questions are centered around quantifying changes in gene expression across time points or treatment conditions. To accurately compare different samples to each other important post-processing steps of RNA-seq data must be done before meaningful conclusions can be drawn. The focus of this chapter is to provide the user with a workflow to produce a table of differentially expressed genes (DEGs) from an RNA-seq experiment. The dataset used in this protocol has been previously published by our lab and consists of eight samples, each consisting of pooled RNA from multiple regenerating fragments isolated during a regeneration time-course [1]. Although no replicates were included in this earlier study, limiting differential expression analysis of lowly expressed genes, this protocol includes a "simulated" workflow for DESeq2 [10] to obtain a list of differentially expressed genes across stages of regeneration. We also provide an overview of library preparation strategies and overall experimental design.

Figure 1 presents a general workflow for species with a sequenced and annotated genome.

In principle, this protocol can also be applied to RNA-seq datasets generated from other organisms. Although a genome reference is needed when following the protocol described here, we also suggest software to quantify gene expression without such a reference.

#### 2 Materials

One of the challenges in conducting RNA-seq analysis or any other bioinformatic task is the installation and maintenance of various software packages and programs which can represent a major hurdle for users new to this field. Commands written in shell will be indicated by the "\$" prefix, commands in R will be proceeded by ">" and outputs by "#".


Fig. 1 Overview over the protocol. The first step in a typical RNA-seq workflow is removal of low quality and adapter sequences (step 1). Reads are mapped to the genome either using STAR where it is optional to include a transcriptome reference annotation (step 2). Star maps, assembles and quantifies transcripts simultaneously producing count tables (step 3). Count tables are then used as input for differential gene expression analysis with DESeq2 which produces tables of differentially expressed (DE) genes (step 4). Gene ontology analysis of DE genes is performed with GOATOOLS


#### 3 Methods

This protocol will be exemplified using our paired RNA-seq data of regeneration in B. leachii [1]. This dataset consists of eight samples which are listed in Table 1 and can be retrieved from the SRA archive SRP064769 (see Note 16).




Note 18).

#### 2. Create the genome index (see Note 19):

\$ STAR --runMode genomeGenerate --runThreadN 4 --genomeDir genome\_Boleac --genomeFastaFiles Boleac\_SBv3\_genome.fasta - sjdbGTFfile Boleac\_transcripts.gtf

#### 3. Align the reads (see Notes 20 and 21):

\$ STAR --runMode alignReads --genomeDir genome --outFileNamePrefix BL\_regeneration/reg\_0 --sjdbGTFfile Boleac\_transcripts\_v5.gtf --quantMode GeneCounts --runThreadN 4 - outSAMtype BAM SortedByCoordinate --readFilesIn reg\_0\_1\_val\_1.fq reg\_0\_2\_val\_2.fq

4. Repeat for other samples (see Note 22).

3.3 Normalizing Count Data with DESeq2


```
> setwd("path to your local folder")
```

4. Concatenate count files by typing (see Notes 23 and 24):

```
> ff <- list.files( path = "./", pattern = "*ReadsPerGene.out.
tab$", full.names = TRUE )
 > counts.files <- lapply( ff, read.table, skip = 4 )
> counts <- as.data.frame(sapply( counts.files, function(x) x
[ , 4 ]))
 > ff <- gsub( "[.]ReadsPerGene[.]out[.]tab", "", ff )
 > ff <- gsub( "[.]/counts/", "", ff )
 > colnames(counts) <- c("embr","wcol","reg_0","reg_1","re-
g_2","reg_3","reg_4","reg_5")
 > row.names(counts) <- counts.files[[1]]$V1
```
5. DESeq2 requires a sample information sheet (see Note 25) that can be created using a spreadsheet software such as MS Excel and imported into R by typing:

coldata <- read.csv(file="coldata.csv", row.names=1)

6. Generate a dds object which normalizes all count data across samples to library size by typing:

Fig. 2 PCA plot created from the top 500 variable genes among samples

```
> dds <- DESeqDataSetFromMatrix(countData = counts,colData =
coldata,design = ~ stage)
  > dds <- DESeq(dds)
```
7. Create a sample distance matrix type (see Note 26):

```
> res <- results(dds)
> vsd <- vst(dds, blind=FALSE)
> sampleDists <- dist(t(assay(vsd)))
> data <- plotPCA(vsd, returnData=TRUE)
> percentVar <- round(100 * attr(data, "percentVar"))
```
8. Load ggplot2 by typing:

```
> library(ggplot2)
```
9. Plot the principal component analysis (PCA) of your data (Fig. 2) with:

```
> p<-ggplot(data, aes(PC1, PC2)) + geom_text(label=rownames
(data), nudge_x=0.25, nudge_y=0.25,check_overlap=T) + xlab
(paste0("PC1: ",percentVar[1],"% variance")) + ylab(paste0
("PC2: ",percentVar[2],"% variance")) + theme_light()
> p
```
#### 3.4 Differential Gene Expression Analysis Using DESeq2

To extract differentially expressed genes (DEGs) between conditions (stages 1 and 2 and 0, arbitrarily chosen), we define the contrasts we are interested in with the results() function from DESeq2 (see Note 27).

1. To get differentially expressed genes between stages 1 and 0, we type:

```
> res1_0 <- results(dds, contrast=c("stage","1","0"))
```
2. We then using the subset function to generate result tables of differentially up- and down-regulated at an adjusted p-value also described as false discovery rate (FDR) of 5%.

```
> res_1_0_sig<-subset(res1_0, padj < 0.05)
> res_1_0_sig_up<-subset(res_1_0_sig, log2FoldChange > 0.5)
 > res_1_0_sig_up_order<-res_1_0_sig_up[order(res_1_0_sig_up
$padj),]
 > write.csv(res_1_0_sig_up_order, file="res_1_0_sig_up_or-
der.csv")
> res_1_0_sig_down<-subset(res_1_0_sig, log2FoldChange < 0.5)
 > res_1_0_sig_down_order<-res_1_0_sig_down[order(re-
s_1_0_sig_down$padj),]
 > write.csv(res_1_0_sig_down_order, file="res_1_0_sig_dow-
n_order.csv")
```

der.csv")

4. Perform the same process to generate results for comparing stages 2 and 0.

> res2\_0 <- results(dds, contrast=c("stage","2","0")) > res\_2\_0\_sig<-subset(res2\_0, padj < 0.05) > res\_2\_0\_sig\_up<-subset(res\_2\_0\_sig, log2FoldChange > 0.5) > res\_2\_0\_sig\_up\_order<-res\_2\_0\_sig\_up[order(res\_2\_0\_sig\_up

\$padj),] > write.csv(res\_2\_0\_sig\_up\_order, file="res\_2\_0\_sig\_up\_or-

> res\_2\_0\_sig\_down<-subset(res\_2\_0\_sig, log2FoldChange < 0.5)

> res\_2\_0\_sig\_down\_order<-res\_2\_0\_sig\_down[order(res\_2\_0\_sig\_down\$padj),]

> write.csv(res\_2\_0\_sig\_down\_order, file="res\_2\_0\_sig\_down\_order.csv")


#### Table 2 List of DEGs ordered by increasing significance

#### 3.5 Gene Ontology Enrichment Analysis

To get more insight into the biological significance of DE genes we obtained in the above step, we will assign gene ontologies and perform Gene Ontology (GO) enrichment analysis using goatools [13].

1. In R create a list of gene IDs from the result file:

```
> gene_ids_up<-rownames(res_1_0_sig_up_order)
 > write.table(gene_ids_up,file="gene_ids_up.csv", row.name-
s=FALSE, quote = FALSE, col.names=FALSE)
```

```
> res1_0_universe<-subset(res1_0, baseMean > 10)
> gene_ids_univ<-rownames(res1_0_universe)
 > write.table(gene_ids_univ,file="gene_ids_univ.txt", row.
names=FALSE, quote = FALSE, col.names=FALSE)
```
4. Modify the GAF file so the identifiers match the gene IDs in our lists:

```
> gaf<-read.table(file="Boleac_slimTunicate.gaf",skip=5,
sep="\t")
> gaf_red<-gaf_new[ ,c("V6","V5")]
 > gaf_red_col<-aggregate(V5 ~V6, gaf_red, paste, col-
```
Fig. 3 Example of a GO plot for molecular function terms overrepresented using the list of stage 1 upregulated genes. The dotplot generated using ggplot shows the 15 most significant (adjusted p-value < 0.05) enriched GO terms within the molecular function (MF) category

```
lapse=";")
 > write.table(gaf_red_col,file="ids2go_BL.txt", row.name-
s=FALSE, quote = FALSE, col.names=FALSE)
```
5. Find enrichments with a python script run in terminal (see Note 28).

\$ find\_enrichment.py gene\_ids\_up.txt gene\_ids\_univ.txt ids2 go\_BL.txt --pval=0.05 --method=fdr\_bh --pval\_field=fdr\_bh - outfile=results\_id2gos\_1\_0\_up.xlsx

6. Plot top 15 enriched terms of each category (see Notes 29 and 30, Fig. 3).

```
> library("readxl")
> go<-readxl::read_excel("results_id2gos_1_0_up.xlsx")
> go_MF<-subset(go, NS=="MF")
> go_MF_15<-go_MF[1:15, ]
 > b<-ggplot(go_MF_15, aes(x=reorder(name, -p_fdr_bh),
y=study_count,color=p_fdr_bh,size=study_count))+geom_point()
+coord_flip()
 > b + theme_minimal() + labs(x="Molecular Function",y="Gene
count",color="p.adjust", size="Gene count")
```
#### 4 Notes


price difference in stranded vs. non-stranded kits almost warrants a stranded kit.


\$ wget https://www.aniseed.cnrs.fr/aniseed/download/?file= data%2Fboleac%2FBoleac\_SBv3\_genome\_gff3\_fasta.zip

\$ wget https://www.aniseed.cnrs.fr/aniseed/download/?file= data%2Fboleac%2FBoleac.gaf.gz

#### 7. Download the gene annotation and gene ontology files in terminal by typing:

\$ wget https://www.aniseed.cnrs.fr/aniseed/download/?file= data%2Fboleac%2FBoleac\_slimTunicate.gaf.gz -O Boleac\_slimTunicate.gaf.gz

\$ gunzip Boleac\_slimTunicate.gaf.gz

\$ wget http://purl.obolibrary.org/obo/go/go-basic.obo -O go- basic.obo


organism. This step could be also performed on a high-capacity server more suitable for this task.


```
$ curl -O https://repo.anaconda.com/miniconda/Miniconda3-
latest-${ARCH}.sh
 $ sh Miniconda3-latest-${ARCH}.sh
```
where the environment variable ARCH should be set to the type of your local operating system (e.g., Linux-ppc64le, Linux-x86\_64, MacOSX-x86\_64, Windows-x86). This installation includes software tools used in this protocol such as STAR [11], statistical environment R [12], and DESeq2 [10]. For a full list of installed packages run:

```
$ conda list
$ conda install -c bioconda bioconductor-deseq2
```
#### 13. To install SRA-Tools run:

```
$ conda install -c bioconda sra-tool
```
Detailed instructions and information on setting up downloads from SRA archives can be found on the website https:// ncbi.github.io/sra-tools/

14. Install goatools [13] by typing the code below in terminal.

\$ conda install -c bioconda goatools


\$ fastq-dump --split-files SRR2729873

This will create two FASTQ files for each sample, rename all files with their short identifier according to Table 3, e.g., SRR2729873 to embr before proceeding.


\$ mkdir genome\_Boleac



#### Table 3 An example of a text file showing the first few lines of gene raw count data

percentage of unmapped reads is higher than 25% of total reads, the sample is problematic and should be reconsidered.


\$ grep -v "N\_" reg\_0ReadsPerGene.out.tab | awk '{unst+=\$2; forw+=\$3;rev+=\$4}END{print\ unst,forw,rev}'

which results in "\$ 8842238 508841 8718480." This is interpreted as 8,718,480 reads map to the reverse or second strand and only 508,841 reads map to the first strand, which indicates a stranded library was made and the reverse strand was sequenced.

24. Depending on the library preparation method, it is crucial to select the right column in this step. In our case, second-strand synthesis was used, so column 4 was selected for further analysis.

> counts <- as.data.frame( sapply( counts.files, function(x) x [ , 4 ))

25. To define which condition or time-points samples are associated, we create a coldata object which can be made in a text-editor or Microsoft Excel and saved in a CSV file format in the working directory specified earlier. In this example, the coldata object has a unique identifier for each sample (column 1) and one column specifying conditional information (Table 4). The unique identifier of samples must be identical in the coldata object as well as in the counts object. To check if that is the case type:

```
> all(rownames(coldata) == colnames(counts))
```
In this case, the counts object column names can be replaced with the sample identifiers in coldata. Please note that the condition in this chosen arbitrary is for demonstration purposes only.


Table 4 Coldata object with sample information

26. A principal component plot (PCA) is a useful tool to assess if and how individual samples cluster and if the variation in the expression of the most heterogeneously expressed genes can be explained by the nature of the sample (e.g., samples clustering along developmental time).

In this case, we can see (Fig. 2) a strong separation of the embryonic stage (explaining most of the variation) and separation of regeneration stages and the whole colony as the secondlargest component of variation.


> install.packages("readxl")

30. To plot the other categories or more terms, you can change the subset and name the output accordingly.

#### Acknowledgments

This work is supported by a Royal Society of New Zealand grant (UOO1713) to M.J.W. We would like to thank Berivan Temiz for critical reading of the manuscript.

#### References


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part VI

Integrative Approaches

# Chapter 33

# Studying Mechanical Oscillations During Whole-Body Regeneration in Hydra

### Jaroslav Ferenc and Charisios D. Tsiairis

#### Abstract

Cells of the freshwater cnidarian Hydra possess an exceptional regeneration ability. In small groups of these cells, organizer centers emerge spontaneously and instruct the patterning of the surrounding population into a new animal. This property makes them an excellent model system to study the general rules of selforganization. A small tissue fragment or a clump of randomly aggregated cells can form a hollow spheroid that is able to establish a body axis de novo. Interestingly, mechanical oscillations (inflation/deflation cycles of the spheroid) driven by osmosis accompany the successful establishment of axial polarity. Here we describe different approaches for generating Hydra tissue spheroids, along with imaging and image analysis techniques to investigate their mechanical behavior.

Key words Tissue spheroids, Mechanical oscillations, De novo axis formation, Symmetry breaking, Self-organization

#### 1 Introduction

Hydra is a simple freshwater animal composed of two epithelial layers, gastrodermis and epidermis, and organized along a single oral/aboral axis. Its regenerative capacities and amenability to experimental manipulation have made it a rich source of insights about regenerating missing body parts already at the dawn of modern experimental biology [1]. Experiments where regeneration has been challenged, as well as transplantation experiments, have substantially shaped the theories of biological pattern formation [2, 3]. Importantly, Hydra does not only offer a platform for manipulating existing patterns but also for observing their emergence de novo. This was shown when cells from dissociated body columns were reaggregated, and they managed to recreate functional animals in a few days [4, 5]. Astonishingly, unlike organoid systems, they are able to do so without the external addition of signaling factors (see ref. 1 for further comparison with organoids).

Initially, the cells in the aggregates sort to re-establish the epidermal and gastrodermal layers, thus creating a symmetric hollow epithelial spheroid composed of cells whose positional identity along the main body axis is to be specified [6]. After approximately 24 h, symmetry is broken and Wnt-expressing organizing centers start to emerge. These centers guide the appearance of head structures at the oral end of the axis [7]. The role of Wnt signaling as a key driver of oral identity is well established both in the homeostatic conditions and in regenerating Hydra [8, 9]. Depending on the size of the aggregates and the axial origin of dissociated tissue, one or more organizing centers can appear [10]. A similar fate awaits spheroids created from small tissue fragments [11]. When a small piece of the body column tissue is excised, it will fold into a hollow spheroid, visually indistinguishable from the one created from reaggregated cells. Since all the cells have a shared identity, body poles need to be defined in this case as well. Wnt signaling centers will emerge and eventually develop into new animal heads.

Interestingly, regenerating spheroids of any origin experience cycles of inflations and deflations on the way to symmetry breaking [12]. These mechanical oscillations are osmotically driven and appear to be important for proper regeneration [13]. Water from the hypotonic medium is entering the cells, which pump it inside the spheroid cavity to maintain their osmotic balance [14]. As a result, the whole spheroid inflates until reaching a threshold of tissue rupture. The accumulated liquid is thus released, the spheroid deflates, and the cycle is repeated. During the spheroid development, the profile of oscillations changes. Initial high-amplitude and low-frequency oscillations (termed Phase I oscillations) eventually transition to faster cycles with lower amplitude (Phase II oscillations). This is a hallmark of symmetry breaking and reflects the emergence of a stable mouth opening that releases the accumulated liquid under lower pressures [15].

Spheroids prepared from small tissue fragments or by singlecell re-aggregation (Fig. 1) are useful to study these mechanical events during regeneration. However, one method might suit specific experimental demands better than the other. Making reaggregates is more laborious, yet offers better control of the spheroid size by using a fixed number of cells. In addition, different cell populations (e.g., expressing different fluorescent markers) can be mixed in one aggregate. Cut tissue pieces, on the other hand, preserve tissue integrity, close faster, and allow better selection of original tissue axial position. Importantly, such spheroids also retain supracellular actin myofibers, which have been recently implicated in mechanically guiding the axis emergence [16]. These structures dissolve upon tissue dissociation and begin to reappear with random orientation in the aggregates. Moreover, they only seem to align and reorient after the symmetry has been broken [17]. This

Fig. 1 Overview of spheroid preparation and development. (a) Using cut pieces as starting material, (b) using cells from dissociated body columns, (c) both methods generate spheroids that will break symmetry and regenerate into full animals

difference thus offers an opportunity to dissect the impact of these actin structures on the tissue mechanical and biological properties.

Importantly, Hydra spheroids as an experimental model system do not only offer versatile starting conditions. Additional advantages include short regeneration time, simple culture conditions, amenability to imaging, and experimental manipulations. Perturbing the osmolarity by adding solutes, such as sucrose or sorbitol, to the medium allows slowing down the oscillations in a concentration-dependent manner [13]. Different small molecules (e.g., cytoskeleton-affecting drugs) have also been used to perturb the oscillation dynamics. For example, treatment with the Rho-kinase inhibitor Y-27632 results in a sigmoidal rather than linear inflation behavior [18]. Furthermore, since similar oscillations occur in many other cyst-like structures [19], Hydra spheroids offer a unique system to tackle the biological significance of such phenomena. In the following protocols, we detail techniques for making Hydra spheroids from both cells originating from dissociated animals and cut tissue pieces (Fig. 1). Instructions and tools for live imaging and computational extraction of basic oscillation characteristics from the acquired image data are also provided.

#### 2 Materials

Use distilled water to prepare all the solutions. If not indicated otherwise, solutions can be stored at room temperature.

#### 2.1 Culture Media and Animal Handling

	- 2. Stereomicroscope.
	- 3. Dissociation medium: 3.6 mM KCl, 6 mM CaCl2, 1.2 mM MgSO4, 6 mM sodium citrate, 6 mM sodium pyruvate, 4 mM glucose, 12.5 mM TES-HCl, pH 6.9. Add antibiotics (0.05 g/ L kanamycin, 0.1 g/L streptomycin) and filter sterilize. This solution can be stored at 4 C for a month.
	- 4. Reaggregate medium 1: A 1:3 mixture of Hydra medium and dissociation medium.
	- 5. Reaggregate medium 2: A 1:1 mixture of Hydra medium and dissociation medium.
	- 6. Reaggregate medium 3: A 3:1 mixture of Hydra medium and dissociation medium.
	- 7. Dissociation pipettes: Flame Pasteur pipettes to obtain a narrow opening smaller than 1 mm in diameter. This requires some practice.
	- 8. 0.4-mL microcentrifuge tubes (e.g., APEX Scientific mini).

#### 3 Methods

2.2 Cutting and

3.1 Spheroid Preparation from Cut Tissue Pieces


Fig. 2 Critical steps in spheroid preparation protocols. (a) foot half of a bisected animal (note the slight tissue swelling next to the cut side), red line indicates the position of the next cut, (b) ring of tissue, red lines indicate the positions of cuts to prepare fragments of equal size that will give rise to spheres, (c) properly closed spheroids just after closing (left), and after inflation begun (right), (d) improperly closed spheroid releasing cells (arrowheads), (e) tubes with cell suspension positioned in a 50-mL tube and ready for centrifugation, (f) tubes standing in a dish of dissociation medium before the release of aggregates, (g) spheroids mounted in the agarose wells in imaging chamber. All scale bars correspond to 500 μm

	- 1. Fill the lid of a 90-mm Petri dish with Hydra medium.
	- 2. Transfer ~30 Hydra individuals into the dish using a handling pipette.
	- 3. Orient the animals with the help of the pipette so that they are lying flat and wait until they relax (see Note 4).
	- 4. Cut away the heads of the animals (cut below the tentacles).
	- 5. Cut away the feet (cut above the less pigmented zone) of the animals (see Note 11).
	- 6. Transfer the resulting body columns into a 15-mL tube with 3 mL of dissociation medium.
	- 7. Vortex briefly and wait until the body columns settle at the bottom of the tube.
	- 8. Remove as much of the medium as possible.

3.2 Spheroid Preparation from Dissociated Body Tissue


#### 3.3 Imaging 1. Boil the 1% agarose gel (see Note 19).


3.4 Image Segmentation and Quantification of Oscillation Parameters

Fig. 3 Imaging setup and image analysis. (a) Proper imaging setup, (b) the image from (a) segmented using the described strategy, (c) example of a relative radius trace, shaded area indicates Phase I oscillations, deflation points detected by the "findcollapse" function with a threshold of 1.15 are shown as red dots, (d) plot generated using the "sphereslope" function for the Phase I data in c, black dots indicate the corrected radius (note the absence of deflation), the fitted line is shown in red

Image segmentation macro that can be used for batch processing in ImageJ. Adjust the parameters to fit the pixel size of your images as described in Note 29

```
run("8-bit");
run("Median...", "radius¼60 stack");
run("Auto Local Threshold", "method¼Phansalkar radius¼50 parameter_1¼0
 parameter_2¼0 white stack");
run("Fill Holes", "stack");
run("Median...", "radius¼20 stack");
```

#### 4 Notes

1. The stock solutions should not be mixed before adding them to the water. This will cause the salts to precipitate.

The "findcollapse" function. This function identifies deflations and outputs for each instance the time (as frame of the time course) and the amplitude (as the difference of relative radius). Inputs: rad relative radius data as a column vector, threshold—threshold for detecting the collapse. We recommend a threshold of 1.15 for Phase I oscillations

```
function [frame,amplitude]¼findcollapse(rad,threshold)
if ~iscolumn(rad)
error('expect rad to be a column vector');
end
amplitude ¼ -diff(rad);
b ¼ rad(1:end-1)./rad(2:end);
frame ¼ (2:(length(rad)))';
frame(b <¼ threshold) ¼ [];
amplitude(b <¼ threshold) ¼ [];
end
```

The "sphereslope" function. This function extracts the slope of inflation for a specified period of the time course and plots the fit if requested. Inputs: rad—relative radius data as a column vector, threshold—threshold for detecting the collapse, tinterval—time step of imaging in minutes, frstart defines the beginning (frame in the time course) of the measured interval, frstop—defines the end of the measured interval, varargin—use either "plot" or "noplot" depending on your preferences. If nothing is specified for varargin, the default option is no plot

```
function slope ¼ sphereslope(rad,threshold,tinterval,frstart,frstop,
 varargin)
if ~isempty(varargin)
switch varargin{1}
case 'plot'
gen_plot ¼ true;
case 'noplot'
gen_plot ¼ false;
otherwise
error('plot options are plot and noplot')
end
else
gen_plot ¼ false; % default plot option
end
if ~iscolumn(rad)
error('expect rad to be a column vector');
end
% detecting collapses and correcting for them
a ¼ -diff(rad);
b ¼ rad(1:end-1)./(rad(2:end));
corrad ¼ rad + [0;cumsum(a.*(b > threshold))];
% linear fit
time ¼ ((0:(length(rad)-1))*(tinterval/60))';
P ¼ polyfit(time(frstart:frstop),corrad(frstart:frstop),1);
slope ¼ P(1);
% plot result
if gen_plot
scatter(time(frstart:frstop),corrad(frstart:
 frstop),15,'k','o','filled')
yfit ¼ P(1)*time + P(2);
title(['slope¼',num2str(slope),'h-1'])
ylabel('cumulative radius')
xlabel('time [h]')
hold on;
plot(time(frstart:frstop),yfit(frstart:
 frstop),'r','LineWidth',2);
hold off;
end
end
```
Similarly, rings with big diameter can be divided into three or more pieces.

9. Repeat steps 4–6 in Subheading 3.1 if you want to obtain more rings/spheres from the same animal.


For better reproducibility among experiments, cell concentration should be determined using a counting chamber and aggregates prepared with the same number of cells. In our experience, a few thousands of epithelial cells should be used to prepare an aggregate of a final size comparable to a cut spheroid.


best determined empirically. Ultimately, the epidermal layer should be continuous, flat, and cells should not be released from spots in the surface.


105 strain. The usual causes of unsuccessful closing include cutting with a blunt blade, damage while handling the cut pieces, and poor health of the animals.


the shifts created by spheroid deflations. A straight line is then fitted to the corrected data. This allows measuring the overall slope for long periods, such as the whole Phase I duration (Fig. 3d). To enable slope measurements for different time windows (e.g., one oscillation), the function requires the user to specify an interval for this measurement.

#### Acknowledgments

We thank Jacqueline Ferralli for useful additions to the protocols and Melinda Liu Perkins (UC Berkeley) for helpful suggestions on the Matlab functions. Our research is supported by the Novartis Research Foundation and by the Schweizerischer Nationalfonds zur Fo¨rderung der Wissenschaftlichen Forschung (grant 31003A\_182674).

#### References


pivotal element of de-novo symmetry breaking in hydra spheroids. bioRxiv


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 34

# Combining RNAi-Mediated β-Catenin Inhibition and Reaggregation to Study Hydra Whole-Body Regeneration

### Matthias Christian Vogg and Brigitte Galliot

#### Abstract

In addition to its ability to regenerate any amputated body part, the Hydra freshwater polyp shows the amazing ability to regenerate as a full polyp after a complete dissociation of its tissues. The developmental processes at work in reaggregates undergoing whole-body regeneration can be investigated at the molecular level by RNA interference (RNAi). Here we provide a protocol that combines β-catenin RNAi with reaggregation. This protocol serves as a basis to generate "RNAi-reaggregates," followed by the extraction of high-quality RNA for the precise quantification of gene expression by real-time PCR. This protocol is efficient, providing both a molecular signature, with the significant downregulation of β-catenin and Wnt3, as well as a robust phenotype, the lack of axis formation, which is observed in all reaggregates.

Key words Hydra, Reaggregation, Whole-body regeneration, siRNA electroporation, Gene knockdown, β-catenin, Wnt3, qPCR, Patterning

#### 1 Introduction

Hydra is a small freshwater organism that belongs to the phylum Cnidaria (Fig. 1a). The animals exhibit a tube shape with a head at the apical pole and a foot (basal disk) at the basal one. The head region is composed of an apical dome-shaped structure centered around the mouth opening, named hypostome, and at its base, a ring of tentacles (Fig. 1b). When a Hydra is cut into two halves, each half will regenerate within 3–4 days a new complete body, including a fully functional head from the lower half and a foot from the upper half [2]. This is achieved through the rapid formation of an organizer (a group of cells that can induce and pattern adjacent cells) at the regenerating tip as reviewed in [3]. Over the last two decades, it has been demonstrated that Wnt/β-catenin signaling is a component of the head organizer, with the growth factor Wnt3 acting as a head activator [4–8]. In brief, Wnt3 is mainly expressed in the hypostome, is the earliest upregulated

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols,

Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_34, © The Author(s) 2022

Fig. 1 Phylogenetic position and Hydra anatomy. (a) Hydra is a member of the phylum Cnidaria and the class Hydrozoa. Phylogenetic tree, after Collins et al. [1]. (b) The Hydra head is composed of a hypostome (dome-shaped structure surrounding the mouth opening) with a mouth opening at the apex and a tentacle ring at the basis. The body column separates the head from the basal region with the basal disk also named foot. Scale bar: 500 μm

Wnt gene during head regeneration, and acts in an auto-regulatory loop to maintain Wnt/β-catenin activity in the apical region [6, 7]. Noteworthy, Wnt3 expression and thus head formation outside the head region are suppressed by the transcription factor Sp5 [8]. At the basal pole, BMP signaling seems to play a key role in basal regeneration [9]. Overall, two regulatory networks organize each pole of a regenerating Hydra.

The extreme regenerative capacity of Hydra culminates in the regeneration from reaggregates. The phenomenon of Hydra cells to self-organize into a new animal has fascinated researchers since its discovery in the 1970s [10, 11]. Once Hydra tissues are dissociated into a single-cell suspension, reaggregates either form spontaneously by keeping the cells at a high density or can be induced by mechanically compacting the cells in capillary tubes or by gentle centrifugation [10]. In the first immediate phase of the reaggregation process, epidermal and gastrodermal cells sort out to re-establish the original cell layers [10]. Over the next days, new polyps emerge from the mass of cells, equipped with tentacles, and a hypostome at the apical pole that become fully functional (i.e., able to feed) around day 6 [10]. At a later stage, a basal disk develops on each polyp, which will eventually detach 1–4 weeks later. The number of polyps that develop from a given reaggregate depends on the initial number of cells that form the reaggregate (classically several polyps emerge from a 70,000-cell reaggregate).

Hydra reaggregates can be seen as the forefather of organoids as they share common features, i.e., their formation relies on self-organization and requires Wnt/β-catenin signaling for symmetry breaking [2]. In Hydra reaggregates, the transition from a ball shape to an elongated shape of the reaggregate is a critical first axisdefining step, characterized by the emergence of Wnt3 expressing clusters that will develop into apical poles [12]. To functionally assess the involvement of genes that act in the patterning of a reaggregate, RNA interference (RNAi) serves as a powerful tool to silence gene expression.

RNAi can be induced in intact Hydra by electroporating small interfering RNAs (siRNAs) [8, 13]. In short, siRNAs are loaded onto an RNA-induced silencing complex (RISC), whereas the "passenger" strand is removed by Argonaute-2 (Ago-2). This leads to an activation of the RISC complex with a single-stranded "guide" RNA molecule that targets mRNAs in a sequence-specific manner. Due to the action of the RNase-H like activity of Ago-2, mRNAs are degraded, which results in gene silencing [14, 15]. We recently demonstrated that gene silencing persists over several days even when RNAi animals are dissociated to a single-cell level, which has opened up new perspectives to study the developmental processes at work in reaggregates [8].

In this chapter, we provide a detailed protocol that combines RNAi-mediated gene silencing with reaggregation (Fig. 2). The effectiveness of this protocol is illustrated by the case of β-catenin, leading to a failure of axis formation. Indeed, the quantification of β-catenin transcripts, in reaggregates exposed to β-catenin siRNAs, by real-time PCR (qPCR) shows a significant downregulation of βcatenin. In addition, these β-catenin RNAi reaggregates show a reduced Wnt3 expression and do not develop axes, which is consistent with a function of Wnt/β-catenin signaling in the formation of the Hydra body axis.

#### 2 Materials

2.1 Electroporation and Reaggregation

Prepare all solutions with ultrapure water (Milli-Q system 18.2 MΩ-cm at 25 C) at room temperature except noted otherwise.


Fig. 2 Method overview. See text for details


2.2 RNA Extraction, Small Interfering RNAs, and Real-Time PCR


2.3 Kits and Equipment


#### 3 Methods


4. Wait 5 min to let tissue pieces sediment.


#### 2. Add 350 μL of the RNA extract kit's lysis buffer.


3.3 RNA Extraction and Real-Time PCR (qPCR)


#### 4 Notes


Fig. 3 Regeneration of Hydra body axes from reaggregated cells after silencing β-catenin. The reaggregation experiment was performed with animals exposed twice (RNAi1, RNAi2) to scramble (top) and β-catenin (bottom) siRNAs. Shown are four representative reaggregates (agg 1–4). Reaggregates were imaged at indicated time points. The red arrow indicates the time point of animal dissociation, taken as t0 for reaggregation. Note that axes are clearly visible in control RNAi reaggregates on day 4 (white arrowheads), while β-catenin RNAi reaggregates fail to develop axes while forming a few tentacles (white arrows). Scale bars: 200 μm

Fig. 4 Real-time PCR of RNAi reaggregates. RNAi was performed as depicted in the scheme. The red arrow indicates the time point of animal dissociation, taken as t0 for reaggregation. RNA was extracted on day 4 of the reaggregation process, followed by real-time PCR (qPCR) to measure the expression of β-catenin (blue symbols) and Wnt3 (purple symbols) in control and β-catenin RNAi reaggregates. Each data point represents one biologically independent experiment. Statistical p-values: \* 0.05, \*\* 0.01 (unpaired t-test)

#### Acknowledgments

The authors thank Charisios Tsiairis for helpful comments and discussions. The research in the Galliot laboratory is supported by the Swiss National Science Foundation (SNF 310030\_189122), the Canton of Geneva, and the Claraz donation.

#### References


Curr Top Dev Biol 116:391–414. https://doi. org/10.1016/bs.ctdb.2015.11.002


https://doi.org/10.1016/j.devcel.2009. 07.014


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Creating a User-Friendly and Open-Access Gene Expression Database for Comparing Embryonic Development and Regeneration in Nematostella vectensis

Olivier Croce and Eric Ro¨ttinger

#### Abstract

The sea anemone Nematostella vectensis has emerged as a powerful research model to understand at the gene regulatory network level, to what extend regeneration recapitulates embryonic development. Such comparison involves massive transcriptomic analysis, a routine approach for identifying differential gene expression. Here we present a workflow to build a user-friendly, mineable, and open-access database providing access to the scientific community to various RNAseq datasets.

Key words Database, Data mining, Open-access, RNAseq, Cnidarian, Nematostella vectensis, Regeneration, Embryonic development

#### 1 Introduction

The anthozoan cnidarian Nematostella vectensis (Fig. 1) has initially been developed as a research model organism to gain insights into the evolution of developmental mechanisms and novel cell types/ biological features as it is easily cultivable under laboratory conditions [4]. The first cnidarian genome to be sequenced was the one from Nematostella that has revealed astonishing similarities with the ones from mammalians [5, 6]. Since then, a wealth of resources and tools have been developed ranging from embryonic RNAseq datasets [1, 2] to meganuclease-induced transgenesis [7], as well as functional approaches for gene know-downs [8–11] and CRISPR/CAS9 mediated knock-outs and knock-ins [11–13].

More recently, Nematostella is emerging as a powerful and complementary whole-body regeneration model, as it is able to regrow missing body parts within days after amputation [14– 18]. In combination with its historical use as an embryogenesis research model, Nematostella is thus very well suited to compare embryonic development and whole-body regeneration within the

Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1\_35, © The Author(s) 2022

Fig. 1 Nematostella vectensis, a research model to assess the relationship between embryonic development and regeneration. (a) General anatomy of the sea anemone Nematostella. The body of this small anthozoan cnidarian (<5 cm) is organized along an oral/aboral axis. The oral region is formed by tentacles (ten) that surround the mouth (\*) and a pharynx (pha). The body column (bco) contains internal structures called mesenteries (mes) that end at the aboral most region with the so-called physa (phy). (b) Schematic representation of the phylogenetic position of cnidarians (including Nematostella, indicated in orange) within the metazoan tree of life. (c) Confocal images (phalloidin/actin filaments in black, DAPI/nuclei in red) of representative stages of Nematostella embryonic development and regeneration. Depicted below are the time points for which published RNAseq samples [1–3] are present in the NvERTx database

same organism [3, 17, 19]. This is particularly convenient not only to determine to what extent regeneration recapitulates the gene regulatory program deployed during embryonic development addressing one of the long-lasting questions in regeneration biology [20], but importantly to highlight if regeneration is controlled by specific toolkits only present in differentiated tissues.

Several studies have performed global gene expression analysis during embryonic development [1–3, 6, 21] or regeneration [3, 22]. In the present chapter, we describe the methods we used to create NvERTx, a gene expression database for comparing embryonic development and regeneration gene expression data in Nematostella vectensis [3, 23]. This database features transcriptomic data combining 22 time points during embryonic development and 16 time points during regeneration (Fig. 1c). In addition, we have developed a web interface to facilitate the manipulation of the data contained in this database. This website simplifies data mining and can generate some figures such as graphs to compare groups of sequences. This interface also integrates various common online tools like Blast to retrieve all transcripts homologous for a given sequence entered by the user. Building such in silico tool is a mean to make RNAseq data accessible to the larger scientific community and enable additional usage of the vast amount of data that were initially produced for a given scientific question.

The workflow presented in this protocol is only a general framework for developing an open-access scientific database. More in-depth knowledge of various aspects of programming and of computer science theory in general might be necessary to implement such tool. Although collaborations will bring together knowledge about the biology to be incorporated and the technology to deploy it online, skills to develop such database can be acquired through a wealth of documentation and literature.

#### 2 Materials

The protocol and material presented here is the one that was used to develop NvERTx, a website to access and mine quantitative transcriptomic data from the sea anemone Nematostella vectensis [3]. The protocol will need to be adapted to develop websites presenting other types of data as well as to take into account the unavoidable evolution of software and data format. Yet, this protocol provides a generic and flexible framework to make data accessible and usable to the larger scientific community. In addition to programming skills, we here detail the basic material/data/tools required for the development of a user-friendly transcriptome data mining website. Data implemented in the developed database and website is assumed existent. Refer to [23] for a recent protocol describing a standard workflow for quantifying and performing initial analysis of next-generation sequencing (NGS) reads from an RNA-seq analysis.

# 2.1 Data Resource Requirements



2.2 Technical Requirements

#### Genomic and transcriptomic resources for Nematostella vectensis


(using AUGUST, Prodigal, or common ORFing prediction tools) if no annotation is available in public databases.

	- 2. Database management system: Database management should follow a relational model. The model organizes data into many tables with a unique key identifying each row. Queries can be performed using the SQL that is the reference language since many decades. We encourage to use MySQL (see Note 4).
	- 3. Web framework: A modern web development requires to use a robust and reusable syntax. We used Django which is a highlevel Python web framework that encourages rapid development and clean, pragmatic design (https://www. djangoproject.com/). Moreover, Django facilitates the communication with the database directly or through additional


Fig. 2 Screenshot of the NvERTx homepage. (a) Users can search for genes using the gene name, Nemve1 accession number, NCBI GenBank accession number. (b) NvERTx IDs can also be used to directly obtain the temporal expression profiles for the genes of interest. Multiple, up to five, transcripts can be queried simultaneously. (c) Users can also directly explore co-expression clusters from embryonic development and regeneration to identify groups of co-expressed genes. (d) The transcriptome can also be searched using BLASTn or tBLASTn

> libraries such as SQL-Alchemy (https://www.sqlalchemy.org) (see Note 5). For the design of the site, we have used "Bootstrap" (https://getbootstrap.com) which is an additional framework. It facilitates the creation of web interfaces (HTML + JavaScript) and avoids display problems that can occur between web browsers.

4. Data mining software suite: The web interface of NvERTx includes online software to perform basic manipulations of the RNA-seq sequences (Fig. 2). We added the BLAST suite to facilitate the search of all transcripts of the NvERTx database homologous to a given sequence entered by the user. The aligner MUSCLE was also added to visually compare a group of similar sequences (SNPs, indels, etc.). The integration of these tools in the Django code is facilitated by some specific API such as PyBlast (https://pypi.org/project/pyblast/) or django-blastplus (https://github.com/michalstuglik/djangoblastplus).

#### 3 Methods

#### 3.1 Defining the Scope and Application of the Tool

An important aspect for developing a database for internal use or for the scientific community is to define the scope and intended use of the tool. Some preliminaries are therefore required to be clearly defined such as:


#### 3.2 Setup the Relational Database

3.2.1 Database Design

Prior to starting the building of the database and the website per se, the probably most important aspect of the process is to take time to conceptualize the database regarding its intended usage. Taken this into account will make this online tool user-friendly and as useful as possible for the scientific community.

To do so, we encourage you to make an exhaustive inventory of all information and meta-information that will have to be integrated into the database (see Note 7). This information consists of the biological resources acquired from experiments such as RNA-seq results, but also include every data or metadata that could be related to the numerical values: annotations (genes, go-terms, pathways, etc.), information of the experiments protocol itself (samples sources, extraction method, library preparation, sequencer device used), general or user comments, image descriptions. The data inventory should also include the external sources of information that could be useful to associate: GSEA, ontologies, PubMed, websites of collaborators, etc.

It is also necessary to consider other information that are not directly related to the biological data, but which will be useful for the management of the database and the website: e.g., a table grouping the users with their respective access permissions (guest, administrator, permissions to read or modify part of the data in the database, etc.) (see Note 8).

3.2.2 Build the Database Structure A popular method to build databases is based on a relational model that uses linked tables. Depending on the size of the expected database, it is advisable to create a model before directly building the Tables. A database model is a way to set a representation of the data and its relations understandable by a non-expert and independent from any further technical choice of development.

There are several tools or methods to make this model. For example, the Merise method is a general-purpose modeling methodology in the field of information systems development. Merise proceeds to separate treatment of data and processes, where the data-oriented view is modeled in three stages, from conceptual, logical through to physical. The UML (Unified Modeling Language) is also a general-purpose, developmental, modeling language in the field of software engineering that is intended to provide a standard way to visualize the design of a system. A model built using UML proposes a diagram structure representation that can be easily interpreted.

An intermediate solution is to use a graphical table construction and visualization. This type of software allows the biologist to visually verify that all the information is present in the database and in the right place. Also, the relationships between tables can be represented with this type of tools. Once the database schema is finalized, the tool can automatically generate the tables in SQL format for import into a MySQL system for instance. Among these tools, we can mention "MySQL Workbench" which is a free tool and both simple and powerful (https://www.mysql.com/ products/workbench/).

Finally, the design of the database must prioritize the independence from the technical solutions that will be used to make the GUI. It must be sufficiently modular to not constrain the use of a specific coding language. It should be possible to interact with the database via an external website, a scripting API, or any interfacing solution that can be developed later (see Note 9).

3.2.3 Populate the Database There are several possibilities to populate the database. The most direct is to fill the data directly in SQL command lines or through an external graphical tool like "PHP MyAdmin" (https://www. phpmyadmin.net) which facilitates the access and the management of a database.

> Some recurring operations will be easier by using some dedicated scripts that will have to be developed. For example, importing data from RNA-Seq to populate the database is almost impossible to do without using a script that retrieves the values from a raw file and converts them into SQL entries. It will then be necessary to develop a series of scripts, in Python or in a language familiar to the database manager.

> If the database is extended, it will be necessary to develop several scripts and to check that the logic of filling the database is respected: some tables must be filled before others, some records must be mandatory, others optional (see Note 10).

#### 3.3 Build the Website

3.3.1 Design and Create the Main Webpages

In addition to the classical sections such as a Home page describing the project and an About page displaying the various contact details, the important parts of the website will have to be decided in close collaboration with the main end-users. In our example, NvERTx presents a dedicated page for displaying the data of co-expression clusters of embryogenesis versus regeneration and a dedicated page displaying RNA-Seq differential expression by sample type. Each of these webpages are linked to selection menus allowing to display specific information.

Forms to search information must be clearly visible. The entry boxes to search using terms like gene names or by IDs will be often used. They must be accessible directly, e.g., from the left or top main menu. The development of the frontend will be done via the chosen framework (see Note 11).

It is necessary to establish the list of features that will be required to achieve the project (e.g., count tables, expression plots, expression clusters). These functionalities can be classified into two sets: tools to visualize the data and tools to mine the data.


3.3.2 Implementing Data Visualization and Mining Tools

Fig. 3 Screenshot of the expression plots. Shown are plots for regeneration (top) and embryonic development (bottom) for Nematostella runx (NvERTx.4.92297), tcf/lef (NvERTx.4.46364), c-ets1B (NvERTx.4.68511). Selecting the tabs in (a) enables the users to obtain (1) count data from each of the data sets, (2) transcript annotations, (3) sequences in FASTA format, (4) bibliographical resources including PubMed links and PaperBlast queries (if available), and (5) MUSCLE alignment to compare similar transcripts

> which allows to mix different sets of value to generate online graphics (3D histograms, heatmaps, or more sophisticated representations).

If the website is hosted on a local web server, unless for a strict internal use, you will need external access permissions for the site to be visible to the rest of the world. The database itself does not necessarily need to have an open access to the outside because it will go through the website. It is recommended to have a real domain name associated with the website to improve its visibility.

The NvERTx website is freely and openly accessible to the community (http://nvertx.kahikai.org). The source code for the website can be found at https://github.com/IRCAN/NvER\_ plotter\_django. Data sets from the database can be found at http:// nvertx.ircan.org/ER/ER\_plotter/about.

#### 3.4 Data and Services Availability

3.4.1 Accessibility

Fig. 4 Screenshot of the differential gene expression (DE Genes) feature from NvERTx. A scroll down menu indicates all possible differential gene expression analysis, whose results are represented by volcano plots. Selecting a given dot will provide additional information such as the NvERTx ID, nr\_hits, and fold change values

3.4.2 Maintenance It is highly recommended to use a Docker container system (https://www.docker.com) or similar. This system allows to encapsulate the database, the site, and all the technical requirements in a single file. This makes the project independent of the native server environment and avoids compatibility problems between different versions or other websites hosted on the same server. Moreover, a container is easily exportable and installable on another server without complicated technical modifications. NvERTx uses a Docker container (https://hub.docker.com/repository/docker/ ircan/nvertx).

We recommend depositing all the developments of the project (database structure, website code, API, documents) on a source code development (SCM) platform such as GitHub. This has the advantage of making the code open-source and usable by anyone. Moreover, this kind of sharing platform facilitates collaborative development of IT projects: people who want to improve the project can contribute by modifying the code. The latest version of the project can be made available from this platform. Note that this does not concern the data of the database itself, but only the main source code related to the project.

3.4.3 Documentation Several documents must be provided. These documents can be text files included at the root level of the GitHub project for example. There will be a README file giving the main lines of the project, an INSTALL detailing the technical procedure to install the database and the associated code of this project, a CHANGELOG specifying the modifications made since the last update of the code. Moreover, a document specific to the practical use of the database and the website is strongly recommended. It could be a pdf that can be downloaded directly from the site detailing the data available and all the items/tools available on the site or an "how to" tutorial.

3.4.4 Evaluating the Impact of the Database User feedback is the best way to evaluate the usage and user friendliness of a given database. Those feedbacks may not only help to improve the database usefulness itself but also for the development of any potential upgrade or future in silico database. Several tools exist to gain information on the statistics such as number, origin, session durations, etc. of the database users (e.g., GOOGLE analytics). However, certainly the most valuable evaluation factor for scientific databases is its referenced usage by the larger scientific community, i.e., the resulting citations or collaborations. Thus, providing information on how to cite the database is crucial and should be implemented in the webpage (e.g., the FAQ section).

#### 4 Notes


with some differences like a fully object-oriented language with a more powerful frontend part. Ruby is probably slightly more difficult to handle for beginner developers.


#### Acknowledgments

The authors thank Jacob Warner and Vincent Guerlais for the development of NvERTx as well as Boris Meyer for technical support after the launch of the online database. This work was supported by the French Government (National Research Agency, ANR) through the grant RENEW (ANR-19-PRC) to E.R.

#### References


course analysis of oral vs. aboral whole-body regeneration in the sea anemone Nematostella vectensis. BMC Genomics 17(1):718


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 36

# Formalizing Phenotypes of Regeneration

#### Daniel Lobo

#### Abstract

Regeneration experiments can produce complex phenotypes including morphological outcomes and gene expression patterns that are crucial for the understanding of the mechanisms of regeneration. However, due to their inherent complexity, variability between individuals, and heterogeneous data spreading across the literature, extracting mechanistic knowledge from them is a current challenge. Toward this goal, here we present protocols to unambiguously formalize the phenotypes of regeneration and their experimental procedures using precise mathematical morphological descriptions and standardized gene expression patterns. We illustrate the application of the methodology with step-by-step protocols for planaria and limb regeneration phenotypes. The curated datasets with these methods are not only helpful for human scientists, but they represent a key formalized resource that can be easily integrated into downstream reverse engineering methodologies for the automatic extraction of mechanistic knowledge. This approach can pave the way for discovering comprehensive systems-level models of regeneration.

Key words Formalization, Modeling, Inference, Regeneration

#### 1 Introduction

The resultant phenotypes of regeneration experiments can include complex morphologies, spatial patterns, anatomical manipulations, and temporal dynamics—an extraordinary rich dataset to aid in the understanding of the mechanisms of regeneration [1–3]. Organisms such as planarian worms can regenerate any body part after almost any amputation, including new heads, eyes, brain, etc., and produce aberrant morphologies after genetic perturbations [4–7]. Amphibians and insects can regenerate their amputated limbs and appendages by re-growing them from the rest of their bodies, and perturbations can result in ectopic segments [8–10]. In addition, grafting experiments surgically transplanting tissue or appendages to different locations can highlight the system response to ectopic signals and result in the regeneration of a completely new anterior– posterior axis, such as with an inverted graft in planaria [11], or the regeneration of supernumerary limbs, as in the case of graft rotations in axolotl [12] and cockroach [13]. These experiments are essential to discern the spatial localizations of the signals dictating the tissue organization during regeneration.

However, the experimental perturbations and the anatomical features of the regenerated morphologies are usually characterized in the literature with microscopy images containing much variability between similar phenotypes [14] and described in natural language, which can be ambiguous and miss crucial details [15, 16]. As a result, these rich and large datasets cannot be readily analyzed by neither human scientists nor computational methods for the extraction of mechanistic knowledge. Instead, a first step formalizing the experimental procedures and the phenotypes of regeneration is needed to encode both their qualitative and quantitative features in a standardized way.

For the unambiguous formalization of regeneration phenotypes, we have developed mathematical formalisms that can precisely describe anatomical structures, organ locations, and overall morphological shapes. We have developed such formalisms for encoding the regeneration phenotypes of planaria whole-body morphology [17], and salamander, frog, crustacean, insect, and arachnid limb and appendages [18]. These formalisms are based on mathematical graphs, where the nodes of the graph represent morphological regions or organs, such as the head or tail of a planarian worm, and the edges represent the connections and topological information between these regions, such as the segments forming a limb. Importantly, the topological information of the graphs can be extended with quantitative data regarding the overall shapes and sizes of the morphological structures, their angles of interconnection, and the localization vectors and rotations of the organs, a process that removes the unimportant differences between individual organisms. These formalizations can also include unambiguous descriptions of the experiments that produce the resultant phenotypes. A mathematical tree can define how different amputations, graftings, and irradiations are performed in a particular experiment. In this way, these formal descriptions based on mathematical structures can encode unambiguously both the experimental procedures and the resultant phenotypes of regeneration.

To facilitate working with these formalizations, we have developed user-friendly tools that allows any user to easily encode the phenotypes of regeneration. Planform [19] is a software tool for the formalization of regeneration experiments and phenotypes of planarian worms. Any user without special training can easily create the mathematical formalisms with the provided drag-and-drop interface for the specification of the different types of worm regions, sizes, and overall shapes. Similarly, the interface allows the definition of precise surgical manipulations, the grafting of pieces between worms, and the total or partial irradiation of regions of the worm body. For the formalization of limb regeneration experiments, we have developed Limbform [20], a similar software tool with a user-friendly interface. Limbform can be used to define experiments and phenotypes for salamander, frog, crustacean, insect, and arachnid. The number and position of each anatomical structure, such as digits and ectopic limbs, can be easily defined. Importantly, these tools automatically produce from the graph encodings the cartoon diagrams of the created reference morphologies, which can guide the user during the formalization of complex morphologies. The morphologies, experiments, and expression patterns formalized with these tools are stored in a centralized local database file. Using this protocol, we have curated a database of planarian regeneration with more than 1500 different experiments from the literature (see Note 1). In addition, we have curated a database of limb regeneration experiments with more than 800 different experiments (see Note 2). Both databases are freely available resources for the community, and the tools include a search functionality to find particular regeneration phenotypes resulting from any manipulation, gene, and drug of interest included in these large databases.

Gene expression patterns represent another fundamental phenotype in regeneration experiments. In situ hybridization [21, 22] and immunohistochemistry [23–25] assays can reveal the spatial gene expression pattern of a specific gene at the level of the whole organism. Gene expression patterns through space and time can reveal the genetic regulation of the key elements controlling regeneration. We have developed PlanGexQ [26] as a user-friendly software tool to formalize planarian gene expression patterns in standard reference morphologies. The spatial localization of gene expression patterns can be easily input into a standard 2D worm morphology defined with a similar mathematical graph formalism as in Planform. Predefined expression patterns for organs are also available for the user to select, which transfers directly to the current standard morphology. In addition, the tool automatically assigns gene ontology terms from the Planarian Anatomy Ontology [27] by scanning the spatial locations of the gene expression in the reference morphology as well as searching for keywords in the entered captions and descriptions.

Crucially, the mathematical nature of these formalized datasets can be integrated in a reverse-engineering methodology for the inference of mechanistic models or regeneration directly from experimental phenotypes [28] (see Note 3). Figure 1 illustrates the main steps of this iterative methodology and its integration with the formalization of phenotypes. First, functional experiments are performed at the bench to obtain morphological phenotypes from surgical, pharmacological, and genetic perturbations. These experiments and phenotypes are then formalized with the methodology presented in this chapter. Mathematical models based on ordinary or partial differential equations are built to include the

Fig. 1 The formalization protocol presented in this chapter can be integrated as a key component in an iterative methodology for the reverse engineering of the phenotypes of regeneration. Functional experiments at the bench provide the input datasets, which need to be formalized before knowledge can be computationally extracted. Mathematical models based on dynamical systems are designed to include the relevant regulatory networks and biophysical interactions. The model parameters, regulatory interactions, and particular equations can be discovered by machine learning to infer a mechanistic hypothesis explaining the input experimental dataset. Predictions are made with the model, which are then tested with new experiments, closing the methodology loop

main components of the system, their regulations, and the relevant cell and tissue biophysical forces [29–31]. Features of the model such as the parameters and the particular interactions in the regulatory network can be automatically inferred from the experimental datasets with a machine learning approach [32–34]. Finally, the inferred mechanistic model can be employed to formulate testable predictions in terms of novel phenotypes, genes, and perturbations that can be validated with further experiments at the bench [35–37], closing the reverse engineering cycle. In this way, the phenotypes of regeneration formalized with the presented protocols can be computationally analyzed toward the inference of mechanistic knowledge directly extracted from them.

This chapter presents detailed protocols for the formalization of regeneration experiments and phenotypes using user-friendly interactive tools (see Note 4). The first protocol aims to formalize planarian regeneration. The curation process includes the definition of experimental details, detailed spatial manipulation procedures, and morphological outcomes—body configurations, organ positions, and geometric properties. The second protocol can be used to formalize limb regeneration experiments for a variety of model organisms. Complex amputation and grafting procedures can be unambiguously encoded together with the precise limb and appendage configurations during regeneration. Finally, the last protocol details a procedure to encode planarian gene expression patterns using reference morphologies. This methodology facilitates the curation of a formal dataset of standardized and ontologically annotated gene patterns that can streamline their comparative analysis and downstream mechanistic inference. Practical notes are included for clarifying key steps and implementation details.

#### 2 Materials


#### 3 Methods

This protocol includes detailed instructions to formalize a planarian experimental phenotype with the Planform software tool. Start by executing the program and continue with the following steps.

3.1 Formalizing

Planarian Regeneration Phenotypes with Planform

Fig. 2 Screenshots of the user-friendly software tool Planform for the curation and formalization of experimental procedures and resultant phenotypes of planarian regeneration. (a) An experiment formalization includes procedure details, manipulations, and the resultant morphologies. (b) Manipulations are unambiguously defined as a hierarchy of simpler actions, such as amputations and graftings. (c) Worm morphologies are defined with a mathematical graph, where the nodes represent worm regions and organs

> 1. Create a new database by clicking File and then New database. Input a new file name in any computer folder when prompted (see Note 5). Alternatively, open any existing database already created or downloaded with Open database (see Note 6).


polygons with an arbitrary number of points (vertices), which can be edited with the right-click menu. The position and orientation of grafting manipulations can be defined as well with the drag-and-drop cartoon interface. Clicking and dragging a grafted piece translates it with respect the host morphology, while clicking and dragging on empty space rotates the graft.


3.2 Formalizing Limb Regeneration Phenotypes with Limbform Limbform follows the same general design than Planform, but for the formalization of regeneration phenotypes of salamander, frog, crustacean, insect, and arachnid limb and appendages. To formalize a new phenotype, follow the same procedure as with Planform except for the following steps.


Fig. 3 Screenshots of the software tool Limbform for the curation and formalization of salamander, frog, crustacean, insect, and arachnid limb and appendage regeneration experiments and morphologies. (a) Example of a formalized grafting experiment with salamander resulting in multiple ectopic forearms. (b) User-friendly interface for the formalization of a grafting experiment from the left to the right arm. (c) Limb phenotype with ectopic forearms, including details such as the number of bones in each limb segment and digit


3.3 Formalizing Planarian Gene Expression Patterns with PlanGexQ

PlanGexQ is a software tool for the formalization of planarian gene expression patterns into standard morphologies. Start by executing the program and continue with the following steps.

Fig. 4 Screenshots of the software tool PlanGexQ for the registration and ontological annotation of gene expression patterns in planarian worms. (a) Interface for the curation of a gene expression pattern in a double head regeneration phenotype, including the original microscopy image and ontology terms. (b) Formalized standard morphology together with curated gene expression patterns. Registering expression patterns in standard morphologies facilitates their downstream analysis with machine learning tools


starting from v.3.0 of the tool. If the expression pattern includes specific organs, click on their diagram to directly include their expression pattern. Optionally, select a different expression color by clicking Change Expression Color.


#### 4 Notes


native tools that are multiplatform, as the same code can be compiled for Windows, Mac OS X, and Linux operating systems. The Qt libraries are especially useful for implementing the user interface—including forms, widgets, menus, and any other graphical element—and abstracting the access to the database file. Importantly, Qt also provides a Graphics View Framework, which was used for the visualization and interaction with the morphology and manipulation graphs as well as for the worm and limb cartoon diagrams. This framework provides a surface for displaying interactive custom-made 2D graphical items and support zooming and rotation. In this way, the nodes and lines of a morphological graph can be shown as interactive objects for the user to easily formalize a worm, limb, or expression pattern. Furthermore, the framework supports the creation of custom shapes using Be´zier curves, which was used to implement the dynamic real-time generation of worm and limb cartoon diagrams directly from the corresponding graph formalization.


that differ, rather than starting completely from a new empty manipulation.


anatomical ontology terms according to the worm regions that include gene expression as currently input in the standard morphology. This function also updates the relative gene expression levels for each worm region of the current expression pattern.

26. The check marks in the ontology tree terms, selected terms, and suggested terms are always synchronized so it is clear which terms are assigned to the current expression pattern. To facilitate the annotation process, hovering the mouse over a term shows the description of the ontology term.

#### Acknowledgments

I thank the members of the Lobo Lab for their dedicated work and discussions. This work was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award number R35GM137953. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

#### References


175:327–345. https://doi.org/10.1016/j. cell.2018.09.021


patterns from 2D RNA in situ hybridization images. Bioinformatics 26:761–769. https:// doi.org/10.1093/bioinformatics/btp658


and protocols. Humana Press, New York, pp 353–366. https://doi.org/10.1007/978-1- 4939-7802-1\_10


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# INDEX

#### A


#### B


#### C

Cas9 ...........................................423, 430, 439, 442–444, 448–450, 459–462


Simon Blanchoud and Brigitte Galliot (eds.), Whole-Body Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2450, https://doi.org/10.1007/978-1-0716-2172-1, © The Editor(s) (if applicable) and The Author(s) 2022


#### E


#### F


#### G


#### H



#### I


#### K


#### L


#### M



WHOLE-BODY REGENERATION: METHODS AND PROTOCOLS

#### N


#### O


#### P


#### 684 WHOLE-BODY REGENERATION: METHODS AND PROTOCOLS Index


#### Q


#### R


#### S


#### T


#### WHOLE-BODY REGENERATION: METHODS AND PROTOCOLS Index 685


#### Ultrastructure.............................................. 123, 168, 266

#### V


#### W


#### X

