Oxford Open Doors is an annual event that offers the public a unique opportunity to explore the historic and modern buildings of Oxford, many of which are normally closed to the public. Organized by Oxford Preservation Trust, the event takes place over a weekend in September and allows visitors to experience the architectural and cultural heritage of the city first hand.

As part of this year’s Oxford Open Doors event, researchers from the Institute for Developmental and Regenerative Medicine (IDRM) invited the public into their labs, offering a rare glimpse into their cutting-edge research. Visitors were guided through research areas and imaging facilities by enthusiastic staff eager to share their work.

The IDRM is a unique flagship institution at the University of Oxford, dedicated to meeting an ambitious challenge: two thirds of all deaths world-wide are due to non-communicable diseases, many of which are cardiovascular, neurological or immune system disorders that have a developmental origin, representing an urgent unmet clinical need.  The mission of the IDRM is the development of new drugs and therapeutic strategies to tackle these chronic illnesses.

At its core is a formal merger of developmental biology and regenerative medicine in the form of over 200 cardiovascular, neuroscience and immunology scientists, integrating their expertise to foster multidisciplinary collaborations. 

Under this research interest, we’ve developed a number of initiatives for Public Engagement that we deployed at this Open Day: the first, Synchrony, is a unique science-art project created in collaboration with computational artist Andy Lomas. Part of the public engagement efforts of the Wellcome funded Human Developmental Biology Initiative, Synchrony brings the complex world of developmental biology to life.

Synchrony allowed attendees to interact with a 3D artificial simulation of growth and pattern formation, changing the rules that govern the formation of some of the fascinating shapes that we can observe during embryonic development. This immersive experience captured visitors’ curiosity and sparked conversations about the intricate biological processes that direct embryo development and organ formation with Matthew Stower who represented the HDBI researchers. The creative process itself involved lengthy discussions between Andy and researchers Matthew Stower, Shifaan Thowfeequ, Claudio Cortes, Tomoko Watanabe and Shankar Srinivas.

In addition to Synchrony, visitors also engaged with scientific discovery through SyGlass, a virtual reality software for exploring, annotating and sharing 3D data. SyGlass allows users to fully immerse themselves in stunning microscopy images, rendered in interactive 3D in a VR environment. A breath-taking image of the embryonic thymus, contributed by David Grainger, enabled attendees to explore the beauty and intricacies of the 3D structure of this organ. Both children and adults delighted in slicing through the tissue sections and navigating the complexities of the thymus in real time.

We would like to thank The Node for their support. Your extremely popular postcards gave us the perfect ‘hook’, to talk about the wonders of development and how it takes place across species. These visuals helped visitors connect the beauty and complexity of developmental biology with the cutting-edge research happening within IDRM, while at the same time sparking very interesting conversations about research and ethics.

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Núria Ros-Rocher¹, Júlia Ramon Mateu², Elizabeth Williams³

1 Institut Pasteur, Université Paris-Cité, CNRS UMR3691, Evolutionary Cell Biology and Evolution of Morphogenesis Unit, 25-28 Rue du Dr. Roux, 75015 Paris (France)

2 Laboratoire de Biologie du Développement de Villefranche-sur-mer (LBDV), UMR7009 CNRS/UPMC, Institut de la mer de Villefranche, Quai de la Darse, 06234 Villefranche-sur-Mer Cedex, France

3 University of Exeter Biosciences, Streatham Campus, Stocker Rd, Exeter EX4 4QD, UK

With some beautifully warm and sunny mid-summer weather, attendees of the EuroEvoDevo2024 gathered on Friday morning 28th June at the University of Helsinki for a symposium on ‘Environmental cues for life cycle transitions’. This was one of twenty-four symposia at this year’s meeting, with a record attendance of over 600 people. It was co-organized by Vincent Laudet (Okinawa Institute of Science and Technology, Japan and Academia Sinica, Taiwan) and Elizabeth Williams (University of Exeter, UK) and involved ten presenters. This symposium aimed to examine life cycle transitions and the influence of the environment on them from a broad perspective across organisms from different kingdoms of life. What aspects of the environment trigger changes in an organism’s form and physiology and how do organisms sense these?

Although starting early on the final morning of the conference, our enthusiastic late-night discussions at the nearby conference pub, the ‘Thirsty Scholar’, were offset by strong Finnish coffee, a selection of fresh vegetarian foods, the promise of the conference dinner and party later that evening, and our shared interest in the evolution of complex life cycles. The symposium brought together diverse researchers and topics, with study organisms ranging from liverwort to diatoms, choanoflagellates, insects, marine invertebrates, and fish. Environmental cues discussed ranged from abiotic factors such as light, temperature and salinity, to biotic cues including microbial biofilms, coralline algae, and food type and availability.    

Overall, five different types of life cycle transition were discussed. Arild Husby, from Uppsala University (Sweden), presented recent work on morphological polyphenism in water striders, which is induced by changes in photoperiod. When exposed to more hours of light in the day, water striders will develop short-wing phenotype and under short day lengths will develop long-winged phenotypes. RNA sequencing followed by RNAi knockdown and ELISA measurements of the insect moulting hormone ecdysone indicated that this polyphenism is regulated by the Fat/Hippo pathway (Gudmunds et al. 2024), with a possible role for ecdysone signalling. The Fat/Hippo pathway controls organ size in animals and relates to a fundamental question in developmental biology: how does an organ know when to stop growing after reaching a particular size? The ecdysone signalling pathway was also the focus of work presented by Giulia Campli, from the University of Lausanne, Switzerland, who investigated the moulting gene repertoire across an impressive 153 proteome dataset, comprehensive of insect, crustacean and chelicerate arthropod species, highlighting changes and novelties in the molecular machinery driving the moulting process, a second type of life cycle transition for exoskeleton renewal (Campli et al. 2024). 

A third common type of life cycle transition among animals is metamorphosis from a larval to a juvenile form. Several speakers focused on this transition across a wide range of animals, from sponges to fish. Océane Blard, PhD student at the University of Queensland (Australia), presented remarkable brightfield timelapses and fluorescent confocal images illustrating the morphological and cellular changes observed during the first 6 hours of metamorphosis in the marine sponge Amphimedon queenslandica. She also tracked the fate of larval sensory-secretory cells, known as flask cells, which are proposed to detect cues from coralline algae to initiate settlement and metamorphosis (Nakanishi et al. 2015). These cells appear to be able to transdifferentiate, and future research will focus on understanding the molecular mechanisms underlying this process using single cell transcriptomics. Elizabeth Williams, research fellow at the University of Exeter (UK), focused on understanding how free-living larvae settle and metamorphose to become seafloor-dwelling juveniles. Elizabeth and colleagues found that microalgae biofilms, notably diatoms, induce a settlement response in the marine worm Platynereis dumerilii (Hird et al. 2024), they described similar sensory-neurosecretory cells across polychaete, cnidarian and mollusc larvae, potentially responsible for timing metamorphosis according to external environmental conditions. In future work, Elizabeth and colleagues will focus on the morphology, function and molecular signalling mechanisms of these cells.

Júlia Ramon Mateu, PhD student at the Laboratoire de Biologie du Développement (LBDV) in Villefranche-sur-mer, presented her PhD work on the cellular and molecular mechanisms of cnidarian settlement. She identified and characterized the cell types forming the aboral end (head) of the planula larva of three cnidarian species belonging to the two main clades of Cnidaria: the jellyfish Clytia hemisphaerica and the corals Astroides calycularis and Pocillopora acuta. To do so, she generated bulk RNA-seq data from the planula aboral end and single-cell RNA-seq data from the same larval stage for each species. Through a comparative analysis of transcriptomic profiles, morphology and distribution of aboral cell types between the larvae of Clytia and the two coral species, she defined the common cellular architecture of the planula aboral end featuring two broad cell-type classes: secretory-like and neural-like. Of particular interest, she identified genes related to taurine uptake and catabolism as a shared feature of specialized cells in the planula aboral end: associated with a neural cell type in Clytia and a specialized epidermal cell type in the two coral species. Using settlement assays developed for both Clytia and Astroides planulae, she could show that exogenous taurine delays or prevents settlement in both species. Júlia concluded her presentation with an updated model of the cells and molecules involved in cnidarian larval settlement, proposing taurine uptake as a key factor in the regulation of the settlement response.

Vincent Laudet (Okinawa Institute of Science & Technology, Japan and Academia Sinica, Taiwan) also investigates the environmental influences of larval metamorphosis and phenotypic disparity in coral reef fish. Like marine invertebrates, the environment can also modulate different aspects of fish metamorphosis from pelagic larvae to reef-associated juveniles. Vincent presented a case of environmental acclimation in fish populations living in the enclosed lagoon of Taiaro Atoll, an extreme environment in French Polynesia, (Reynaud et al. 2023). Compared to their natural habitat (beaches and mangroves), the lagoon displays higher salinity and temperature. Interestingly, fish living in the lagoon and their counterparts living in oceanic reefs were identical at the genetic level, despite differences in their habitat. To understand the molecular mechanisms of fish acclimation to high salinity, Vincent and colleagues compared their transcriptomic profiles and found that each population had particular differences in thyroid hormone regulation. Thyroid hormones are central hormone regulators triggering metamorphosis in vertebrates (Zwahlen et al. 2024). In fish, they regulate a major metabolic shift during metamorphosis from the juvenile to the adult phase, including changes in pigmentation and maturation of the nervous/sensory and digestive systems. Vincent and colleagues proposed a new view of the role of thyroid hormones: rather than functioning independently in metamorphosis, metabolism and seasonality, thyroid hormones perform a 3-in-1 function of timing a biphasic life cycle transition. Thus, local acclimation to the environment is an endocrine-driven process, conferring plasticity on fish from the time they undergo metamorphosis and enabling them to adapt to high salinity conditions.

Núria Ros-Rocher, postdoctoral researcher at Institute Pasteur (Paris, France), presented another case of salinity-dependent life cycle transition in a microeukaryote. Núria showed that Choanoeca flexa, a facultatively multicellular choanoflagellate (Brunet et al. 2019;Reyes-Rivera et al. 2022), develops colonies by an unconventional mixed mode of clonal-aggregative multicellularity (Ros-Rocher and Reyes-Rivera et al. 2024). To further explore the ecological determinants of C. flexa clonal-aggregative multicellularity, Núria, Josean, and colleagues combined cell biology approaches in the laboratory with fieldwork experiments in the Caribbean island of Curaçao, the original site where this species was discovered (see Brunet et al. 2019). In the field, they surveyed 150 seawater-filled splash pools, the natural habitat of C. flexa, and explored the impact of splash pool evaporation and refilling cycles on C. flexa multicellularity. C. flexa colonies could only be observed in a narrow range of salinity and were never observed in higher salinities in the field, yet sometimes appeared in splash pools that got refilled by waves or splash after complete desiccation.

Surprisingly, multicellular C. flexa colonies could be recovered from soil samples collected from dry splash pools upon rehydration with seawater in the lab. Later experiments in the lab mimicking the evaporation of splash pools showed that colonies dissociate into quiescent single cells in high salinities, losing multicellularity. In those conditions, some single cells transitioned into a desiccation-resistant form, losing some of the stereotypical choanoflagellate features (a collar and a flagellum). Repeating similar rehydration experiments, Núria showed that those cells could return to their multicellular form upon rehydration. The authors hypothesized that this mixed clonal-aggregative strategy might represent a versatile adaptation to the ephemeral nature of splash pools where the simultaneous action of both mechanisms could allow for a faster re-establishment of multicellularity. In ongoing work, Núria, Josean and colleagues are exploring the genetic diversity of newly isolated strains from distinct splash pools. They are focusing on kin recognition capacities that may impact choanoflagellate survival in the field and provide insights into the adaptation of this organism in a highly fluctuating environment.

In addition to metamorphoses and multicellularity, three presenters also discussed later-in-life transitions related to reproduction. Mariella Ferrante, researcher at the Stazione Zoologica Anton Dohrn (Italy), presented recent work on centric and pennate diatoms exploring how these different single-celled microalgae switch between sexual and asexual reproduction. While sexual reproduction in centric diatoms can be triggered by environmental cues such as light and salinity, pennate diatoms do not appear to have a specific trigger for initiating sexual reproduction. However, planktonic pennate diatoms need to solve the challenging problem of locating a mate within the vast 3D ocean to undergo sexual reproduction before they drop below a critical size due to several rounds of asexual reproduction which cause them to decrease in size with each asexual division. Mariella and colleagues are using RNA-Seq and single-cell RNA-seq in different diatom species and reproductive stages to explore heterogeneity in diatom populations as well as to identify a panel of molecular markers for detecting diatom sex events at sea (Annunziata et al. 2022).

 Alexandre Alié, researcher at the Laboratoire de Biologie du Développement (LBDV) in Villefranche-sur-Mer, is also using scRNA-Seq to understand asexual reproductive transitions in another organism that inhabits the pelagic oceans – Mediterranean salps, planktonic tunicates that form large chains through asexual reproduction. Alexandre and colleagues hope to use this information to understand more about how the stem cells in the stolons of the salps can bud off dozens of new individuals and how this process is influenced by environmental factors such as temperature and food availability. Finally, David Hoey introduced us to the molecular regulation of transformations for reproduction in the liverwort Marchantia polymorpha, which switches between sexual and asexual reproduction throughout the year depending on light quality, day length and drought conditions (Duckett et al. 2024). David generated knockout lines of a RAS gene and found that it is a regulator of asexual reproductive development in M. polymorpha. Remarkably, this gene is responsive to both light and drought cues.

Overall, a strong theme running through the symposium was the development of molecular lab techniques such as gene knockout and single-cell RNA-Seq for application in a variety of non-model organisms. Ongoing studies on all of these emerging models are advancing our understanding of how environmental cues are detected and lead to important transitions in a life cycle. We also found inspiration in the examples from several researchers of combining their molecular lab studies with field studies of their organism of interest, to place our understanding of organismal and life cycle diversity back into a broader ecological context. We anticipate that this will be a strong theme going forwards, and aligns with the emphasis on eco-evo-devo that was highlighted at this year’s conference by keynote speaker Scott Gilbert, who also drew attention to the importance of microbial ecology in shaping animal life cycles and symbioses. Could inter-species or inter-kingdom interactions be a future focal area to further inform life cycle evolution?

References

• Annunziata et al. 2022. Trade-off between sex and growth in diatoms: Molecular mechanisms and demographic implications. Science Advances 8:3, DOI: 10.1126/sciadv.abj9466
• Brunet et al. 2019. Light-regulated collective contractility in a multicellular choanoflagellate. Science 366: 6463, https://www.science.org/doi/10.1126/science.aay2346
• Campli et al. 2024. The moulting arthropod: a complete genetic toolkit review. Biological Reviews of the Cambridge Philosophical Society, https://doi.org/10.1111/brv.13123
• Duckett et al. 2024 The biology of Marchantia polymorpha subsp. ruderalis Bischl. & Boissel. Dub in nature. Frontiers in Plant Science, 15, https://doi.org/10.3389/fpls.2024.1339832
• Gudmunds et al. 2024. The Fat/Hippo pathway drives photoperiod-induced wing length polymorphism. BioRxiv https://doi.org/10.1101/2024.04.05.588279
• Hird et al. 2024. Microalgal biofilm induces larval settlement in the model marine worm Platynereis dumerilii. Royal Society Open Science, https://doi.org/10.1098/rsos.240274
• Nakanishi et al. 2015. Sensory flask cells in sponge larvae regulate metamorphosis via calcium signaling. Integrative and Comparative Biology 55(6): 1018-1027, DOI: 10.1093/icb/icv014
• Reynaud et al. 2023. The environment: A vector of phenotypic disparity during the settlement phase of coral reef fishes. Journal of Experimental Marine Biology and Ecology, 568: 151937, https://doi.org/10.1016/j.jembe.2023.151937
• Reyes-Rivera et al. 2022. Nitric oxide signaling controls collective contractions in a colonial choanoflagellate. Current Biology, 32(11): P2539-2547, DOI: 10.1016/j.cub.2022.04.017
• Ros-Rocher et al. 2024. Mixed clonal-aggregative multicellularity entrained by extreme salinity fluctuations in a close relative of animals. bioRxiv, https://doi.org/10.1101/2024.03.25.586565
• Zwhahlen et al. 2024. The ecological function of thyroid hormones. Philosophical Transactions of the Royal Society B, Biological Sciences, https://doi.org/10.1098/rstb.2022.0511

Acknowledgements

The Company of Biologists provided financial support for this symposium. We thank the organisers of the EED2024 conference in Helsinki, Finland, and all attendees of this symposium for contributing to discussions of this session. We also thank the symposium speakers for contributing to revision of this report.

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London EvoDevo meeting (LED)

Dear colleagues,

Given all the outstanding evo-devo work going on in the London area, we feel that a regular gathering to share recent discoveries and to network would be a real asset. Not too long ago there was such a meeting, and we are aiming to revive the spirit of it.

Our hope is to arrange a sustainable meeting that would occur once or twice a year. The current plan is to have half-day events that include short presentations/feedback (15 min + 5 min Qs) and socialization. Ideally, the location of these meetings will rotate among the organizer’s campuses. We wish to cast a wide net by defining “evo-devo” broadly and attract researchers working directly in the area but also those working on evolutionary or developmental biology looking to bridge to the other side.

We have arranged the rebooting of London EvoDevo to be November 8^th on Queen Mary’s campus from 13:00 – 17:00. We hope that you or people in your group will join and potentially be willing to either apply for a talk for this meeting or the future ones.

For more information and to register, please follow this LINK.

We hope to see you in November!

London EvoDevo Organizers,

Ferdi Marlétaz (UCL)

Margarida Cardoso Moreira (Crick), James DiFrisco (Crick),

Chema Martin (QMUL), Alex de Mendoza (QMUL), Roman Arguello (QMUL)

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In this SciArt profile, we meet Yoshinari Yoshida, a philosopher of science focusing on the conceptual and historical issues in developmental biology. Yoshinari enjoys drawing animals and plants, and have recently got into drawing primates, such as gibbons and orangutans, because of his research into in vitro models in comparative biology.

Can you tell us about your background and what you work on now?

I was born in Milton Keynes, UK, and grew up in Japan. I did my undergrad and Master’s in Philosophy and History of Science at Kyoto University, Japan, and Ph.D. in Philosophy at the University of Minnesota, USA. I am currently a postdoctoral fellow at the University of Exeter, UK. Throughout this academic career, I have studied philosophy of science, with a special focus on conceptual and historical issues in developmental biology. My central philosophical interest is in how various models and representations (e.g., “living models” such as model organisms and cell culture systems, and visual models such as diagrams and illustrations) play their roles in the production of knowledge about generality and diversity in developmental processes. My current project focuses on the prospects and limitations of in vitro models in comparative biology. I try to clarify how in vitro systems, such as organoids, play their roles as models in the context of cross-species comparison and reconstruction of evolutionary history. I examine experimental and inferential practices as well as interdisciplinary interactions among experts to tackle this philosophical question.

Were you always going to be a philosopher of science?

When I was younger, I was rather going to be a scientist. I was a kid who loved insects and extinct animals, and in my teenage era, I enjoyed learning about evolution and embryonic development. But I also liked the philosophical discussions about conceptual questions. When I was in the second year of my undergrad, I took the courses “philosophy of science” and “history of science.” I enjoyed these courses so much, and started thinking that studying these fields would allow me to fulfill both my interest in science and my love of philosophical discussions. I decided to major in philosophy of science, and since then, I’ve been in this field.

And what about art – have you always enjoyed it?

Almost yes. I was already into art when I was very little. I was a kid who kept drawing during lunch break in elementary school as well as after school. My favorite objects were monsters, robots, and manga characters. There were periods in my life when I was a bit away from doing art, such as during high school. But I’ve maintained my interest in art. Several years ago, when I was in my Ph.D., I bought a tablet and started drawing digitally. Until then, drawing had been a private hobby just for myself. But I began posting my artwork on social media and also drawing more constantly.

What or who are your most important artistic influences?

It’s not easy to name one specific artist. (I still lack systematic knowledge of art history.) But I’m often inspired by realistic oil paintings from the modern and early modern periods. My favorite is those paintings that are so realistic that they look as if they are photographs from distance, but consist of explicit strokes when seen closely. Those paintings make me think about how we see the world.

How do you make your art?

Currently I do everything using a painting app called Procreate in my tablet. When I see an object or scene whose shapes, colors, textures, or patterns I find interesting, I take pictures, which I may use as a reference later. Or, sometimes I become interested in a certain animal or plant, and search for images of the animal or plant and then start drawing it.

Does your research influence your art at all (or vice versa), or are they separate worlds?

Art is my major hobby that is a source of joy distinct from research in my life. But I think there are some connections between the two. For instance, I am interested in the role of images in science. I have some articles (one published, one currently under review) about how the use of specific types of diagrammatic representations contribute to knowledge production in science. I am also working on a side project with my colleagues about the role of speculation in evolutionary thinking, in which we discuss the power of illustrations. I think this philosophical interest of mine in images in science has a common root with my artistic interest in drawing and painting. My art is also sometimes influenced by my academic interests. For example, I’ve recently been into drawing primates, such as gibbons and orangutans. This was triggered by my current research project on in vitro models in comparative biology. I’ve read studies that apply advanced in vitro technologies to primates to elucidate human evolution, which made me interested in primates themselves.

What are you thinking of working on next?

There are a few ideas. I want to keep working on primates and drawing as many primate species as possible. That would become an interesting series! I’m also interested in making art that is related to developmental biology. Even though my primary philosophical focus has always been on developmental biology, I’ve not made art about this field. It would be fun to make drawings inspired by the normal developmental stages of chick embryos, mechanisms of collective cell migration, or tissue movements in gastrulation. Finally, I’m interested in trying oil painting. While I’ve drawn only digitally over the last several years, I love seeing oil paintings at museums. That would be an interesting new step.

Find out more about Yoshinari:

Instagram: @yoshinariysd

X/ Twitter: @YoshinariYsd

Personal website: https://sites.google.com/site/yoshinariyoshida/

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In our recent paper published in Developmental Biology entitled “Localization and origins of juvenile skeletogenic cells in the sea urchin Lytechinus pictus”, we studied the intriguing and unique late larval development of sea urchins with a focus on skeletogenic cells, which deposit the biomineral skeleton. Here, I discuss how this project came about, why we were excited by our findings, some of the challenges faced, and what’s next for the project.

Project beginnings

In 2021, I joined Deirdre Lyons’ lab at Scripps Institution of Oceanography (UC San Diego) to explore my fascination for invertebrate evolution and development. I was excited to develop a project on skeletogenesis and biomineralization in sea urchins, a topic in which Deirdre and I shared interest. As I was new to the field, I dove into the literature and realized that although I was interested in embryonic development, I was most fascinated by the development of the adult body plan, which occurs throughout the late larval stage. This study was also motivated by the development of the sea urchin Lytechinus pictus as a better model species [1-3], which has been an ongoing collaboration between the Lyons lab and the Hamdoun lab, also at UC San Diego. Since L. pictus is a faster developing and more transparent species than other commonly studied sea urchins, we could study and image late larval development in higher detail.

Most sea urchins are indirect developers and undergo a complete switch in body plan during metamorphosis from a bilaterally symmetric swimming larva to a pentaradially symmetric benthic juvenile. Prior to metamorphosis, much of the adult body plan develops on the left side of the larva in a structure called the rudiment, while other smaller adult structures develop in other locations (Fig. 1). I wondered how two drastically different body plans could be made from the same set of genetic information, and how the development of a second body plan could occur outside of a classic embryonic context. I was specifically inspired by a study from 2007 by Mamiko Yajima in which she demonstrated that the juvenile skeletogenic cells arise not from the larval skeletogenic lineage, but from mesodermal cells that had a non-skeletogenic origin in the larva [4], and I found it fascinating that cells with one fate in the larva can be completely reprogrammed to make a new cell type in the adult. I therefore decided to revisit this topic using recent technical advances in the field.

Prior to this study, rudiment development had been described in detail at the tissue level and some immunostaining had been performed [5-9], however mRNA localization via in situ hybridization was challenging due to the thickness and complexity of the late larval tissue. By leveraging HCR RNA-FISH, we were able to conduct some of the first mRNA in situ hybridizations in these late stages as the small probes used in this technique are capable of penetrating into thick late larval tissue. I chose to focus on skeletogenic cells as they have been well-studied during embryonic development of sea urchins for gene regulatory networks and biomineralization [10,11] and had previously been studied in some detail in the late larva [7,8], although not in the newer model species L. pictus. We therefore asked three main questions: where are skeletogenic cells and biomineral located during late larval development in L. pictus within the complex rudiment morphology, what are the most likely origins of the juvenile skeletogenic cells, and are gene expression patterns across skeletogenic cells similar in embryonic and juvenile skeletal development?

What we learned

We first described in detail the localization of presumptive juvenile skeletogenic cells prior to biomineralization of juvenile skeleton, and at later stages, the localization of biomineral and juvenile skeletogenic cells both within and outside of the rudiment in L. pictus (Fig. 2A-B). We were excited to find that prior to the start of juvenile skeletogenesis in the rudiment, we could identify populations of skeletogenic cells using known biomineral marker genes that were not part of larval rods. Additionally, in some of these presumptive juvenile skeletogenic cells, we found co-expression of our skeletogenic cell marker and a marker of blastocoelar cells, which in the larva act as migratory immune cells (Fig. 2C-D). We therefore proposed that since mesodermal cells of non-skeletogenic origin contribute to juvenile skeletogenic cells [4], that blastocoelar cells may at least in part be the cell type that switches to a skeletogenic fate. We then sampled various biomineralization genes from different gene families and found that mRNA expression patterns throughout juvenile skeletogenesis are similar to the patterns during embryonic development (Fig. 2E-F).

Overall, our work gives insight into how indirect developers, like the sea urchin, can reuse cells and developmental mechanisms to make a new body plan from another. Additionally, it is thought that since all echinoderms have adult skeleton but not all have larval skeleton, that the larval skeleton in sea urchins may be a more recent innovation, while the adult skeletogenic cells could arise from a common origin [12]. Since other echinoderms have blastocoelar-like lineages in the larva [13-14], it is possible that this cell type is the ancestral adult skeletogenic cell lineage that expresses an ancestral skeletogenic gene regulatory network, although this hypothesis needs to be clarified with further work.

Unexpected challenges

One of the most challenging aspects of this project was identifying a consistent staging scheme during late larval development, which was an important part of the project to help establish L. pictus as a model sea urchin. Interestingly, the larval body plan and parts of the juvenile body plan, like the rudiment, often develop asynchronously, and so similar looking larvae may not have a rudiment at the same developmental stage [5-9]. Additionally, the extent of rudiment development was often different when metamorphosis occurred, leading to post-metamorphic juveniles in different developmental stages. Although frustrating when trying to stage the animals, this helped me think about development in new ways since in indirect developers, adult body plan development may not be as clear-cut as in embryogenesis.

I was also personally challenged to learn about all aspects of the scientific process, from asking clear scientific questions, thinking hard about the best ways to answer them, and learning to tell a compelling story through publication. As with any scientific project, there were many challenges and failed experiments along the way, particularly as I was learning many techniques for the first time. For instance, learning how to mount, orient, and image these large samples added a lot of time to the project, and I often had to retake very long acquisitions when images did not turn out as expected. This really challenged my ability to persevere when experiments were not working, but I felt especially honored and reinvigorated when one of our images was featured on the cover of Developmental Biology. Overall, I was very proud to have finished this work within the relatively short timeline of my master’s degree, and I am greatly looking forward to applying this newfound knowledge in future work.

What’s next with the sea urchins?

With this paper, we highlighted that sea urchin adult body plan development is fascinating, yet understudied, and how it is important to think of body plan development outside of an embryonic context. However, there are still many remaining questions to be answered. For instance, only skeletogenic cells have been studied in some detail during adult body plan development, and HCR RNA-FISH provides the opportunity to understand whole body patterning by investigating other cell types. However, more complex questions, such as mapping where adult cells are originating, would require more complex techniques. With the development of L. pictus as a better system where mature adults can be raised from egg in 4-6 months and with which transgenic lines can be created [1-3], an abundance of new questions could be studied. I am greatly looking forward to continuing work with this amazing animal in the future to answer innovative questions in adult body plan development and evolution. Finally, I am very thankful for my mentors, particularly Dr. Lyons, for the mentorship throughout this project, as well as other collaborators and colleagues who have supported and guided me through past and present research.

References

1. Nesbit KT, Fleming T, Batzel G, Pouv A, Rosenblatt HD, Pace DA, Hamdoun A, Lyons DC. 2019. The painted sea urchin, Lytechinus pictus, as a genetically-enabled developmental model. Methods in Cell Biology. 150: 105-123.
2. Jackson EW, Romero E, Kling S, Lee Y, Tjeerdema E, Hamdoun A. 2024. Stable germline transgenesis using the Minos Tc1/mariner element in the sea urchin Lytechinus pictus. Development. 151(20).
3. Vyas H, Schrankel CS, Espinoza JA, Mitchell KL, Nesbit KT, Jackson E, Chang N, Lee Y, Warner J, Reitzel A, Lyons DC, Hamdoun A. 2022. Generation of a homozygous mutant drug transporter (ABCB1) knockout line in the sea urchin Lytechinus pictus. Development. 149(11).
4. Yajima M. 2007. A switch in the cellular basis of skeletogenesis in late-stage sea urchin larvae. Developmental Biology. 307(2): 272-281.
5. Cameron RA, Hinegardner RT. 1978. Early events in sea urchin metamorphosis, description, and analysis. Journal of Morphology. 157(1): 21-31.
6. Heyland A, Hodin J. 2014. A detailed staging scheme for late larval development in Strongylocentrotus purpuratus focused on readily-visible juvenile structures within the rudiment. BMC Developmental Biology. 14(22): 1-14.
7. Yajima M, Kiyomoto M. 2006. Study of larval and adult skeletogenic cells in developing sea urchin larvae. The Biological Bulletin. 211(2): 183-192.
8. Smith MM, Smith LC, Cameron RA, Urry LA. 2008. The larval stages of the sea urchin, Strongylocentrotus purpuratus. Journal of Morphology. 269(6): 713-733.
9. Formery L, Wakefield A, Gesson M, Toisoul L, Lhomond G, Gilletta L, Lasbleiz R, Schubert M, Croce JC. 2022. Developmental atlas of the indirect-developing sea urchin Paracentrotus lividus: From fertilization to juvenile stages. Frontiers in Cell and Developmental Biology. 10.
10. Davidson EH, Rast JP, Oliveri P, Ransick A, Calestani C, Yuh C-H, Minokawa T, Amore G, Hinman V, Arenas-Mena C. 2002. A Provisional Regulatory Gene Network for Specification of Endomesoderm in the Sea Urchin Embryo. Developmental Biology. 246(1): 162-190.
11. Wilt FH. 2002. Biomineralization of the Spicules of Sea Urchin Embryos. Zoological Science. 19(3): 253-261.
12. Koga H, Morino Y, Wada H. 2014. The echinoderm larval skeleton as a possible model system for experimental evolutionary biology. Genesis. 52(3): 186-192.
13. Tamboline CR, Burke RD. 1992. Secondary mesenchyme of the sea urchin embryo: Ontogeny of blastocoelar cells. Journal of Experimental Biology. 262(1): 51-60.
14. Metchnikoff E. 1893. Lectures on the Comparative Physiology of Inflammation: Delivered at the Pasteur Institute in 1891.

(2 votes)

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The definition of outreach is “to reach further than”, however, I have found that when it comes to science outreach initiatives, we are only reaching those already in the loop. Although I personally applaud all these events and ideas and have been part of them for many years – either recruiting to my graduate program or showcasing my institution – it always seems like I am speaking to people who are already familiar with what a PhD program is or what academic research entails, and they typically seem to know which career or degree they want to pursue. I believe the key is to approach and interact with those who are not part of the scientific community.

As an international student, I thought that if I wanted to pursue a career in any way related to healthcare, the only road was medical school. It was not until pretty far into high school that I learned of alternate careers and graduate school. Maybe I am the anomaly, but I would bet that many students in similar positions do not know the many opportunities that are available by pursuing a graduate degree in biomedical sciences. Even though I have now moved on from my graduate training, I wish to remain involved in outreach initiatives. However, I want to do so a little differently, and I hope I can encourage others to do the same.

My idea of outreach will now not be limited to going to a local college to speak about my graduate program and such. I want “to reach further than” I have previously and introduce this amazing career path to people even before they reach high school, to truly go out on a limb and pitch this life to those that are not aware of it,rather than simply preaching to the choir. If we were to ask the average 3rd grader to describe a scientist, I’m sure they would all describe the typical Albert Einstein-esque look – messy hair, old and a little unhinged. And (although we all know someone like this in academia) I want to be part of outreach initiatives that challenge this idea, and instead pitch a lifestyle of endless curiosity and non-stop learning to kids that do nothing but wonder and ask questions themselves. If you had told me when I was a kid myself that I would be working with the technology used in Jurassic Park (CRISPR, gene editing, in-vitro cultures), I would have thought that it was the perfect job. I believe I can sell this same idea to kids now: kids that do not know what a PhD is, that do not think being a scientist is something that is feasible for them, or that have not met someone who is as passionate for learning as they are even later in life. There is a reason why younger kids are better at learning new languages; they are just ready to absorb all the knowledge you throw at them. So, this is the time to go and show them a science experiment, hook them in, and get them to ask “why?”.

I’m going to end my rant here, but I just think there is more we could do, there is more I want to do. Institutions around the world are starting initiatives specifically aimed at primary school children such as The Crick’s annual Discovery Day, or the Vanderbilt Institute for Infection, Immunology and Inflammation ‘MEGAMicrobe’ events which target students ages 5 to 14. This is a great starting point, and I hope that more initiatives like these start to take off, and more people are encouraged to get involved with them. Outreach should not be exclusive to the people already in the loop, the point of it all is to go beyond, and “to reach further than”.

What is one outreach event you have been involved in recently? What ideas do you have or have seen that could be used as science outreach? Comment below!

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This article is co-written by Eliza Hirsch and Paul Bump.

Who are we?

Hi, my name is Eliza Hirsch, and I am an undergraduate in the Srivastava Lab at Harvard University. And my name is Paul Bump, and I am a postdoctoral fellow at the Srivastava Lab who works with Eliza. Our lab studies the highly regenerative acoel, Hofstenia miamia. This Hofstenia species originally was collected in Bermuda and now lives in Cambridge, Massachusetts where we study stem cell biology and the evolution of regeneration across animals.

Who is Hofstenia?

Hofstenia miamia is an acoel worm capable of whole-body regeneration, meaning that if you were to bisect one, both halves would regenerate, leaving you with two worms instead of one. Hofstenia maintain a population of adult pluripotent stem cells (aPSCs) that are responsible for regeneration. The regenerative ability of Hofstenia is quite impressive but they are hardly the only systems in which such regenerative ability exists. So why study them? Well, two other key traits of Hofstenia are central to answering this question. 

Hofstenia have experimentally accessible embryos, something that is very central to the work we do on the development of the aPSCs. They lay their embryos when they are zygotes, allowing us to study their development from the single cell stage all the way up through hatching and beyond. Hofstenia embryos are also easily accessible due to their abundance. The worms we care for lay zygotes frequently enough and at a high enough volume that we can collect almost every day of the week and on most days, we get a substantial number of them. Thus, as a system Hofstenia allow us to study the development of a highly regenerative organism as well as its adult regenerative ability. 

How to work with Hofstenia

To understand what it is like to work daily on Hofstenia we thought it would be best to give a perspective from Eliza who is completing her undergraduate thesis research in the Srivastava Lab:

Hofstenia are relatively easy to care for. In the lab, we keep a large portion of our worm colony in plastic Tupperwares of various sizes with up to sixty worms being stored in any given large Tupperware box. When not being cleaned or collected from or used for any other experimental processes, these boxes sit in incubators. The worms themselves live in artificial sea water that is made in the lab, something which perhaps requires the most time and effort in the caretaking of these animals. Adult worms are fed recently hatched brine shrimp as Hofstenia require live food. Hatchlings and sick worms or worms that may be left unattended over a long weekend without cleaning are fed rotifers as they pose a far smaller risk to the pH of the water and by extension the health of the worms in the event of their death. 

To clean the worms and avoid problems with ammonia build up in the water and the like, twice a week we replace the water and wipe down the inside of the container. An important element of worm care is the collection of embryos. The animals lay their embryos on the sides and bottom of the container, and I gently scrape at the walls with a glass pipette to dislodge them and then collect them with the same pipette and deposit them in a dish full of sea water. On top of the wild type worms, the lab also has various lines of transgenic worms that are more sensitive than the wild type worms and much harder to replace and therefore require even more careful care.

Recently the lab has started using a water system for many of the wild type worms that significantly reduces the amount of time spent cleaning the worms as they have circulating water and are no longer in boxes but rather are in tanks. However, since my work depends on very early embryos, my worms are still in boxes so that I have access to those embryos which are more difficult to collect in the water system.

How to crack an egg, starting development in Hofstenia

For my project, I have been collecting embryos every day. Hofstenia embryos are surrounded with a transparent chorion (their eggshell) that must be removed for much of the work that I do with them. Nearly daily I perform a chemical dechorionation on the single-cell zygotes I’ve sorted from the embryos I collected that morning. For this process I use a combination of chemicals and then mouth pipette embryos one by one into twenty-four well plates filled with a combination of artificial sea water and a small amount of antibiotics. Antibiotics are an essential addition as removing the chorion strips the embryos of a key defense against infection. I then raise these one-cell embryos up without a chorion so that I can do experiments at specific developmental stages that would be too sensitive to the dechorionation process.

The future for Hofstenia

Since Hofstenia is still a relatively new research organism, there are many possible questions about them to try to answer in one’s own research. In the future, hopefully many labs will continue to increase our overall understanding of Hofstenia and use that information to evaluate evolutionary and mechanistic questions of how the stem cell lineage develops and allows the regeneration of any missing body part. While one drawback of such a new model system is that many methods for working on the animals have yet to be developed, the future is bright as we will continue developing technologies with which to study Hofstenia development and regeneration. 

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This post is co-authored by Beatrice L. Milnes and Mansi Srivastava, who participated in SciCommConnect organised by the Node, preLights and FocalPlane.

A frequent source of lightheaded acknowledgment at the end of many talks is often not just human collaborators, but a thank you to the model organisms themselves, which are of phenomenal importance for scientific research. To many who work with them, Model Organism Databases (MODs) are indispensable, so much so that imagining one day in a lab without access to them is horrifying. MODs curate information generated from years of experimental research, providing a scaffold for sharing data in a consistent and structured way, thereby enhancing our understanding of both the organisms and the work they are being used for. With modern technologies like nucleic acid sequencing, gene editing, and structural protein biology, the quantum of data is increasing exponentially. MODs categorise information, annotate the quirks of gene naming, and compile references that capture the growth curve of the field. Databases assemble the available informational pieces of the jigsaw puzzle and at the same time highlight the gaps in our understanding — they often serve as handbooks for researchers to delve into new aspects of their system. As biology utilises increasingly ‘omic’ data, integrating big data across species is more important than ever. Modern biology has placed a large emphasis on addressing fundamental questions and assessing the translatability of their findings to human welfare. This often requires relating data not only across scientific disciplines but also between the various model systems employed. Groups like the Alliance of Genome Resources have already begun some of this work by building a cross-species consortium. The utility of integrating data across species has resulted in a call for awareness among scientists to assist in this process by including information in publications that help with indexing in MODs.  

What does it take to build a model organism database? 

As active experimentalists, we rarely get a peek into the process of database consolidation despite near-daily usage of the end product. Many scientists do not interface with the management that puts great effort into intentionally curating these MODs. It is important to acknowledge that the level of data organisation and layering that databases achieve is in itself as important as the experimentation they inform. The web design contributes greatly to the utility of the sites and is responsible for visualising the available tools, which differ notably between species. MODs also facilitate the exchange of information between two researchers from different corners of the globe without the need for direct communication. In this regard, databases are directly responsible for the accelerated pace of research in the last few decades. Experimentalists, as the primary consumers and beneficiaries of MODs, should be aware and supportive of the resources needed to build the databases they rely so heavily on.

Managing and integrating scientific data has become an essential skill that needs to be inculcated in the next generation of researchers. To this end, consortia like the National Institute of Health (NIH), European Molecular Biology Laboratory (EMBL), and DNA Databank of Japan (DDBJ) can be used as benchmark examples of highly structured databases. Whether a new database is created by a lab, within an institute, in a community space, or available through open access, it should have core structural components that accommodate its growth in the future. Reporting detailed data accurately and including experimental commonalities that could benefit the larger community (eg. plasmid names or standardised phenotyping) should lead to a generation of a database seamlessly. Much of this data could already be published and locally available through alternative resources but many larger trends can only be validated once multiple sources have been compiled by curators. Perhaps one of the largest barriers to this work is that as end-users, we do not know much about the generation of MODs.

Funding support for MODs

Generating large-scale databases is a skill and time-intensive task. For example, the National Center for Biotechnology Information – Sequence Read Archive (NCBI) is curating a central repository for genome/ transcriptome sequencing datasets across model organisms. Unfortunately, such endeavours have limited funding and grant opportunities. For example, the NCBI is supported by the US government alone despite MODs being well utilised as a global resource. Several MODs that serve a smaller research community, for example, Axobase, do not have long-term sustenance funding. Model organisms that are evolutionarily closer to humans such as rodents, their disease-causing pathogens, and food crops are heavily researched. On the other hand, some model organisms are used by a smaller research community aiming to crack open a new area of biology. Intuitively, the nature and quantum of information available for each model organism is very diverse. Databases are crucial for both novices entering a field or experts looking for the next big question. For upcoming MODs, the lack of seed funding limits their conception in the first place. Securing the future of MODs must be prioritised by the scientific community, perhaps through diversification of funding sources. 

What’s the future for MODs? 

There is an open question of what measures need to be taken to ensure the sustenance and growth of all common and upcoming MODs. As more emerging model systems come into common use, new areas of comparative biology can be catapulted by the presence of a MOD. Perhaps we can support the generation of databases for new species by building off an existing MOD template to generate another, for example, Echinobase has been built using Xenbase as a reference. The first and foremost step would be to have ample funding opportunities for data curation. The funding bodies of existing MODs can begin by allocating annual funds for database development in addition to maintenance of current databases. To ensure revenue generation, the users of databases may be charged a nominal amount for storage and curation of their data. The databases of model organisms that are of national importance should be supported by multiple government schemes. To ensure its sustainability, database building can become an integral component of scientific training and can be developed as a career opportunity. More and more early career researchers investing their creativity and skill into science communication platforms such as MODs would ensure their growth. In the present scenario, scientific information is a commodity of high value, and its true potential can be harnessed only after compilation into these databases.

The progress of research is directly correlated to the development of databases. MODs are not just scientific repositories; they are now a direct measure of our scientific prowess. On a longer timescale, they are also the knowledge that we will pass on to the next generation of researchers. The responsibility of supporting MODs lies on the scientific community as a whole. All of us as stakeholders should prioritise, strategise, and contribute towards both old and new MODs. 

(4 votes)

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Maps are vital tools for any exploration, guiding us through the unknown and helping us make sense of the world. For some developmental biologists, a map refers to a fate map. During embryonic development, a single fertilized egg divides and differentiates into various cell types. Thus, determining cell lineages over time and space in small embryos is a critical step in understanding how our body is built. One of the earliest experimental embryologists, Vogt’s seminal work with ‘vital dye’ to trace cell lineages and migration paths during amphibian embryo gastrulation remains valuable even today, despite the technological limitations of his time [Vogt, 1929]. With the more sophisticated techniques we have now, what kind of maps might we be able to create?

In our recent paper [Kuroda et al., 2024], we combined modern techniques to elucidate the complex developmental origins of the pectoral girdle in zebrafish, labeling the cells that potentially contribute to the pectoral girdle from the early developmental stage long before the formation of skeletons. By comparing this cell fate map of the fish pectoral girdle with the preexisting map of the amniote shoulder girdle, we were able to infer how the shoulder/pectoral skeletons were rearranged during the transition of vertebrates from water to land. Below, I would like to share the background scene of our research.

How I started this project

After getting my Ph.D. degree in Japan, I was fortunate enough to be hired as a postdoc in the Tetsuya Nakamura lab. Tetsuya has been researching the key morphological evolution involved in the water-to-land transition, mainly from embryological and genomic perspectives. After revealing the hidden and surprising commonality of developmental origins between fins and hands [Nakamura et al., 2016], he established his own lab at Rutgers University in the U.S. The Nakamura lab was also expanding its research scope to the skeletal structure that supports the pectoral appendages, specifically the pectoral girdle, when I moved to his lab.

The ancestry of our shoulder skeleton seems to have already existed in the pectoral girdles of the fish-like body plan before the vertebrate transition from water to land. Phylogenetic evidence for this assumption can be observed in the presence of the scapula and clavicle in extinct lobe-finned fish, such as Eusthenopteron and in extant lobe-finned fish, such as lungfish and coelacanth [Jarvik, 1980]. Their pectoral skeletons had been transformed over generations of bony vertebrates, eventually helping them adapt to terrestrial life by supporting their increased body weight after the transition to land [Romer & Parsons, 1986].

Although the scapula and clavicle were cherished and used over long geological periods, some bones were reduced or completely lost during and/or after the water-to-land transition. Among them, the cleithrum is one of the most mysterious bones. Originally, the cleithrum in aquatic fish was referred to as the clavicle. However, Gegenbaur discovered the true clavicle in fish and thus renamed the bone originally called the clavicle as the cleithrum [Gegenbaur, 1895]. The relative size of the cleithrum gradually decreased after the transition to land and disappeared multiple times independently in tetrapod lineages [Gess & Ahlberg, 2018; Matsuoka et al., 2005].

When I joined the Nakamura lab, some members had already created a mutant zebrafish with  gli3 gene and discovered that this mutant exhibited an interesting phenotype: an enlarged scapulocoracoid (a cartilaginous precursor for the scapula and coracoid) and cleithrum reduction. It was as if these changes mimicked the pectoral girdle morphological changes that occurred during the water-to-land transition in bony vertebrates! They were beginning to uncover that the ratio of scapulocoracoid to cleithrum changes depending on the magnitude of the Hedgehog-Gli signaling and its downstream pathways [Wei et al., 2023]. To better understand what was happening in this mutant fish and, possibly, the actual evolutionary process, I started my project to determine 1) whether the embryonic sources of the fish scapulocoracoid and cleithrum are shared, and 2) whether there are other cell populations involved, including potential contributions from outside the pectoral fin precursors.

How I chose methods for cell lineage tracing

In animals possessing cleithra, the bone forms at the boundary between the head and trunk. So, I narrowed down the candidate cell populations that could contribute to the cleithrum to four groups: two from head mesenchymal populations (cranial neural crest cells and cardiopharyngeal mesoderm (CPM)) and two from trunk mesenchymal populations (anterior somites and fin-field lateral plate mesoderm (LPM)). Once I refined the candidates, all that remained was to investigate whether each of these cell populations contributes to the cleithrum.

There is no perfect method for cell lineage analysis, and the applicability of a particular method is limited to certain occasions or animal species. Thus, several methods should be combined for consistent and reliable conclusions. Fortunately, many tools for cell lineage analysis have been developed in zebrafish, allowing us to test cell lineages of the pectoral girdle with different methodologies. Initially, I considered tracking cell lineages through tissue transplantation, as has been done in zebrafish and medaka in previous studies [Cole et al., 2011; Shimada et al., 2013]. This involves preparing fluorescently labeled zebrafish and unlabeled zebrafish embryos and surgically exchanging tissues between them at the same embryonic stage. The advantage of this method is that, since the fluorescent protein is encoded in the donor’s genome, the label remains stable and doesn’t dilute with cell division in the host embryo, as is the case with vital dyes. This makes it suitable for long-term tracking, such as investigating the developmental origins of the musculoskeletal system. The downside of this method is that a fine surgical skill is required. Imagine precisely cutting out an approximately 100 mm square region from an almost spherical donor embryo, and transplanting it into the target region in a host embryo! Even worse, this procedure has to be performed at the developmental stage before our four target cell populations mingled with each other, i.e., late at night on the day of fertilization. I soon realized that I couldn’t perform such a precise operation with trembling hands coming from sleep deprivation, so I decided to explore other methods. (I have deep respect for the predecessors who managed to pull off such experiments in the past!)

Next, I tried a tracking method using a fluorescent protein called Kaede, which changes color from green to red upon exposure to UV light. I placed zebrafish embryos that express Kaede throughout the body on a confocal microscope stage, determined the target area with the computer monitor, and illuminated the UV laser on the target cells, converting the cell color from green to red. Although working late at night was still necessary, handling the microscope and clicking a computer mouse with trembling hands was much more feasible. This method worked nicely, and I finally succeeded in labeling the cranial neural crest cells, CPM, anterior somites, and fin-field LPM individually!

The downside of manual labeling of the spatially restricted area such as vital dye injection, surgical transplantation, and Kaede-based photoconversion is that the experimental precision depends on the experimenter’s technique. So, we decided to deploy another complementary and more reproducible method, genetic cell lineage analysis. Genetic cell lineage analysis is a technique that utilizes the tissue-specific activity of a cis-regulatory element with an artificial gene induction system, such as Cre/loxp system that permanently labels the target cell population. Of course, the activity of a cis-regulatory element is more or less pleiotropic, so one shouldn’t expect the existence of a perfect ‘marker’ enhancer/promotor. However, genetic cell lineage analysis was still an excellent tool that compensates for the weaknesses of Kaede-based photoconversion. In collaboration with Christian Mosimann and Robert Lalonde at the University of Colorado and Gage Crump and Claire Arata at the University of Southern California, we were able to collect transgenic (Tg) fishes to track progeny cells that experience the following ‘marker’ gene expressions: sox10 and crestin (as markers for neural crest cells), tbx1 (for CPM), draculin (for LPM), and tbx6 (for paraxial mesoderm including anterior somites).

Multiple lineages converge to form a single bone

By combining manual cell labeling by Kaede photoconversion and genetic cell lineage analyses by several transgenic fishes, we eventually found that the cranial neural crest cells, CPM, anterior somites, and fin-field LPM all converge to form the cleithrum. In other words, the cell populations that form the boundary between the head and trunk in vertebrate embryos all converge into the cleithrum.

We have now successfully mapped the four distinct cell populations onto the cleithrum, from somitogenesis embryo to early larva. Let’s take another look at the development and evolution of the pectoral/shoulder girdle with this map in hand. First, the cleithrum indeed shared a developmental origin with the fin-field LPM, which differentiates into the scapulocoracoid. Perhaps the phenotype reported in the gli3 mutant mentioned above is related to the signal responsiveness of the embryonic resources shared by the cleithrum and scapulocoracoid, such as the LPM. If changes in Hedgehog-Gli signaling were also involved in the tetrapod evolution, the cleithrum may have degenerated and eventually disappeared due to losing the “resource competition” with the scapulocoracoid.

Indeed, a hypothesis has been proposed that not just a part of the LPM-derived cleithrum, but the entire cleithrum, was incorporated into the mammalian scapula through evolution [Matsuoka et al., 2005]. However, our results did not perfectly support this hypothesis. This is because recent results of genetic cell lineage analysis using mice could not confirm the contribution of neural crest cells and CPM (collectively referred as head mesenchyme) to the scapula [Adachi et al., 2020; Heude et al., 2018], while these head mesenchyme contributes to the cleithrum in zebrafish. Given this mismatch of embryonic origins of mammalian shoulder and fish pectoral girdle, our view is that the head mesenchyme which would have formed the cleithrum in ancestral tetrapods, no longer participates in the formation of the shoulder girdle skeleton in derived amniotes. This rearrangement of cell populations at the head/trunk interface may be due to the evolution of a specialized LPM region, which should be called the “neck LPM”, required for the formation of the long neck characteristic of the modern amniotes [Hirasawa et al., 2016; Lours & Dietrich, 2005]. This novel LPM population may have spatially separated the cranial mesenchyme that would have formed the cleithrum in ancestral animals from the trunk mesenchyme, leading to a cleithrum loss in multiple lineages.

Remaining questions

We propose that if we assume the complex lineages that make up the cleithrum is mechanistically indispensable for its formation, we can reasonably explain the evolutional acquisition of the functional neck and the concurrent disappearance of the cleithrum in amniotes. However, it is not yet clear what role each cell lineages plays in the formation of the cleithrum. If one or two of the four cell populations that make up the cleithrum are missing, can the other cell populations compensate for the absence of the missing cell populations and still form the cleithrum? If so, are the developmental origins of animals with the cleithra diverse? Furthermore, we emphasize that the evolution of the cleithrum and the neck cannot be discussed separately. So, then, what exactly is the neck LPM in amniotes? When and how did it evolve through changes in the ancestral developmental system? Some of these questions are now being actively pursued in the Nakamura lab, and of course, are open questions for all researchers interested in the evolution of the shoulder and neck as well.

References

Adachi, N., Bilio, M., Baldini, A., & Kelly, R. G. (2020). Cardiopharyngeal mesoderm origins of musculoskeletal and connective tissues in the mammalian pharynx. Development, 147(3), dev185256. https://doi.org/10.1242/dev.185256

Cole, N. J., Hall, T. E., Don, E. K., Berger, S., Boisvert, C. A., Neyt, C., Ericsson, R., Joss, J., Gurevich, D. B., & Currie, P. D. (2011). Development and evolution of the muscles of the pelvic fin. PLoS Biol, 9(10), e1001168. https://doi.org/10.1371/journal.pbio.1001168

Gegenbaur, C. (1895). Clavicula und Cleithrum. Morphologisches Jahrbuch, 23, 1-20.

Gess, R., & Ahlberg, P. E. (2018). A tetrapod fauna from within the Devonian Antarctic Circle. Science, 360(6393), 1120-1124. https://doi.org/10.1126/science.aaq1645

Heude, E., Tesarova, M., Sefton, E. M., Jullian, E., Adachi, N., Grimaldi, A., Zikmund, T., Kaiser, J., Kardon, G., Kelly, R. G., & Tajbakhsh, S. (2018). Unique morphogenetic signatures define mammalian neck muscles and associated connective tissues. Elife, 7, e40179. https://doi.org/10.7554/eLife.40179

Hirasawa, T., Fujimoto, S., & Kuratani, S. (2016). Expansion of the neck reconstituted the shoulder-diaphragm in amniote evolution. Development, Growth & Differentiation, 58(1), 143-153. https://doi.org/10.1111/dgd.12243

Jarvik, E. (1980). Basic structure and evolution of vertebrates (Vol. 1). Academic Press. https://books.google.co.jp/books?id=UMwKAQAAIAAJ

Kuroda, S., Lalonde, R. L., Mansour, T. A., Mosimann, C., & Nakamura, T. (2024). Multiple embryonic sources converge to form the pectoral girdle skeleton in zebrafish. Nature Communications, 15(1), 6313. https://doi.org/10.1038/s41467-024-50734-x

Lours, C., & Dietrich, S. (2005). The dissociation of the Fgf-feedback loop controls the limbless state of the neck. Development, 132(24), 5553-5564. https://doi.org/10.1242/dev.02164

Matsuoka, T., Ahlberg, P. E., Kessaris, N., Iannarelli, P., Dennehy, U., Richardson, W. D., McMahon, A. P., & Koentges, G. (2005). Neural crest origins of the neck and shoulder. Nature, 436(7049), 347-355. https://doi.org/10.1038/nature03837

Nakamura, T., Gehrke, A. R., Lemberg, J., Szymaszek, J., & Shubin, N. H. (2016). Digits and fin rays share common developmental histories. Nature, 537(7619), 225-228. https://doi.org/10.1038/nature19322

Romer, A. S., & Parsons, T. S. (1986). The vertebrate body (6th ed.). Saunders College.

Shimada, A., Kawanishi, T., Kaneko, T., Yoshihara, H., Yano, T., Inohaya, K., Kinoshita, M., Kamei, Y., Tamura, K., & Takeda, H. (2013). Trunk exoskeleton in teleosts is mesodermal in origin. Nature Communications, 4(1), 1639. https://doi.org/10.1038/ncomms2643

Vogt, W. (1929). Gestaltungsanalyse am Amphibienkeim mit örtlicher Vitalfärbung.II. Teil Gestaltungsanalyse am Amphibienkeim mit Örtlicher Vitalfärbung. Wilhelm Roux’ Archiv für Entwicklungsmechanik der Organismen, 120, 384-706. https://doi.org/10.1007/BF02109667

Wei, J., Wood, T. W. P., Flaherty, K., Enny, A., Andrescavage, A., Brazer, D., Navon, D., Stewart, T. A., Cohen, H., Shanabag, A., Kuroda, S., Braasch, I., & Nakamura, T. (2023). Distinct ossification trade-offs illuminate the shoulder girdle reconfiguration at the water-to-land transition. bioRxiv, 2023.2007.2017.547998. https://doi.org/10.1101/2023.07.17.547998

(2 votes)

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Hi everyone! 

My name is Ryan Harrison, and I will be supporting the community sites at The Company of Biologists as part of a three-month internship. 

I am currently pursuing a PhD between the labs of Timothy Saunders and James Briscoe as part of the MRC DTP in Interdisciplinary Biomedical Research, where I am researching organoids that mimic embryonic development of the lower back and spine.  

At university, I am also part of the Ambassadors for a Better Research Culture (ABRC), where we aim to improve the research environment on the medical school campus for postgraduate students and staff. Here, I am part of the ‘LGBTQIA+ Inclusion’ subgroup where we run monthly events to foster a community of LGBTQIA+ researchers at the medical school campus. We have also established a larger series called ‘Pride in STEM’, where we invite external speakers to discuss their experiences of being queer in different STEM career environments. I would like to carry this into my internship and compile some LGBTQIA+ voices for a post in the Honest Conversations blog series here on the Node. If anyone is interested in sharing their experiences of being queer in academia, please feel free to get in touch at ryan.harrison@biologists.com or thenode@biologists.com. 

(1 votes)

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