# Developmental transitions involve common biological processes across living beings

Evan Force, Dominique Lamy, Stéphane Debernard, Arnould Savouré,

Matthieu Dacher

► To cite this version:

Evan Force, Dominique Lamy, Stéphane Debernard, Arnould Savouré, Matthieu Dacher. Developmental transitions involve common biological processes across living beings. Heliyon, 2025, 11, pp.e42995. 10.1016/j.heliyon.2025.e42995. hal-04983546

HAL Id: hal-04983546

https://hal.science/hal-04983546v1

Submitted on 9 Mar 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
Review article

Developmental transitions involve common biological processes
across living beings

Evan Force*, Dominique Lamy, Stéphane Debernard, Arnould Savouré, Matthieu Dacher

Sorbonne Université, Université Paris-Est Créteil, INRAE, CNRS, IRD, Institute for Ecology and Environmental Sciences of Paris, iEES Paris, F-75005, Paris, France

ARTICLE INFO

Keywords:

Integrative biology

Unicellular organisms

Multicellular organisms

Morphological changes

Functional modifications

Cellular processes

Molecular signaling pathways

Environmental factors

Adaptations

A B S T R A C T

Pivotal biological processes unfold within a limited time during the life cycles of living beings. They are characterized by changes in morpho-anatomy, physiology, and/or in behavior, such as sexual maturation in many animals, flowering in plants, or sporulation in some unicellular organisms. Such events collectively represent transitions between distinct stages of the life cycle of organisms and are often termed "transitions of development" or "developmental transitions". Although these terms are mentioned in biological literature, no definition has yet been established. We thus propose to define developmental transitions as adaptative events marked by modifications occurring at different biological scales ranging from biochemistry to ecology during a brief period in the life cycle of living beings, namely the passage from one developmental stage to another. From this definition, this review covers the similar mechanisms and functions of developmental transitions, drawing insights from extensively studied models as well as lesser-known examples, from unicellular to multicellular organisms of diverse taxonomic groups. Cross-checking works in various fields highlight that developmental transitions are underlain by common processes. Indeed, they require fundamental cellular mechanisms (e.g., proliferation, differentiation, apoptosis) that are tightly coordinated in time and space through molecular signaling pathways and genetic networks, which in turn are regulated by biotic and abiotic factors, thus reflecting adaptations to the environment. This broad and transverse perspective aims to highlight commonalities that transcend taxonomic boundaries, providing insights into the fundamental principles governing dynamic transformations of living organisms.

1. Introduction

Living beings or organisms are defined as any system able to maintain its integrity by exchanging matter and energy with the
environment (nutrition sensu lato), and to ensure its perpetuation through reproduction. During their life, they undergo a succession of
developmental stages differing in size, shape, organization, physiology, the ability to reproduce or even passage from unicellular to
pluricellular forms (or the opposite), etc ... Cross-checking pioneer biological studies reveal remarkable parallels in the changes
leading from one stage to the other in both unicellular and multicellular organisms. Since the discovery of spore formation in *Bacillus
subtilis* by Ferdinand Cohn [1], floral induction in angiosperms by the pioneering work of Mikhail Chailakhyan [2], and tail regression

<footnote>* Corresponding author. iEES Paris, Sorbonne Université, 4, Place de Jussieu, 75005, Paris, France.
E-mail address: evan.force@sorbonne-universite.fr (E. Force).</footnote>
of ascidian tadpoles during metamorphosis by Henri Lacaze-Duthiers [3,4], plethora of works has brought in evidence the existence of common cellular and molecular mechanisms involved in developmental processes. Thus, transitions between distinct stages in the life cycle of organisms are ubiquitous, and are often termed “transitions of development” or “developmental transitions” [5,6].

Although these terms are commonly used in biological literature, no general definition has been established to date. We therefore propose to define developmental transitions as changes in organisms occurring at different levels of biological organization: morphology (ultrastructure for microorganisms), physiology, biochemistry, and/or ecology. These changes are associated with the passage from one developmental stage to another (i.e., growth or embryogenesis are not developmental transitions). They are finely coordinated and last for a short period of the life cycle relatively to one or both the developmental stages they separate. Developmental transitions typically involve cellular mechanisms including migration, proliferation, differentiation, and death. All of these events are tightly controlled in time and space by complex combinations of molecular signaling pathways and gene networks, and this fine orchestration is also directly and indirectly influenced by endogenous and exogenous factors.

In biology, developmental transitions are universal phenomena. Yet, they have never been studied in an integrated and transverse way. Thus, this review focuses on transition between developmental stages rather than on the stages themselves. The general definition we propose covers a wide variety of physiological contexts in different types of organisms. In spite of this heterogeneity, it is justified to use a unique term to cover all of them as it can be shown they involve similar mechanisms and fulfil related functions. Thus, the goal of this review is to reveal common characteristics of developmental transitions in diverse taxonomic groups: bacteria, amoebozoans, and pluricellular organisms among which fungi, angiosperms, and animals (including cnidarians, tunicates, and several vertebrates). Whereas specific cell populations inside multicellular organisms may also accomplish transitions (for instance maturation of xylem in angiosperms or macrophage maturation in mammals), our focus will remain on developmental transitions at the level of the individual, while sometimes considering populations of microorganisms. This broad and transverse perspective aims to highlight commonalities that transcend taxonomic boundaries, providing insights into the fundamental principles governing dynamic transformations of living organisms.

## 2. Developmental transitions result in morphological and physiological changes

Life cycles of organisms often involve changes in living environments and/or lifestyles. Such modifications lead to morphological and physiological alterations through developmental transitions at the origin of the modulation of the nutrition function (sensu lato).

<figure><img src="image_2.png" /><figcaption>**Fig. 1.** Global and drastic changes related to nutrition function during a developmental transition. (A) Significant shifts during the transition from heterotrophy to autotrophy, a pivotal stage in the life cycle of a plant characterized by the shift from relying on internal nutrient sources to self-sustained photosynthesis. (B) Brain maturation and onset of the foraging behavior in younger worker bees is regulated by the density of older foragers inside the hive. (C) *E. coli* undergoes morphological modifications depending on their nutritional conditions: flagellar biosynthesis and bacterial motility are affected by the nutrient resource of short-chain fatty acids (SCFAs) in the animal intestine. Figure created in BioRender.com.</figcaption></figure>
## 2.1. Modulation of a physiological function associated with morphological modifications: example of nutrition

Nutrition encompasses all the physiological functions that ensure the supply of matter and energy to an organism as well as its maintenance such as during embryogenesis. In plants, the development of embryo primarily relies on nutrients stored within the seed. The embryo grows and differentiates, forming various structures such as the embryonic root, hypocotyl, and cotyledons to form a seed. At this stage, the seedling is still heterotrophic, relying on the stored nutrients in the seed to ensure its growth and development.

With further development, the seedling undergoes a crucial transition as it establishes its capacity for photosynthesis. The precise timing of the heterotrophic-to-autotrophic (also named de-etiolation) transition plays a crucial role in plant survival, which is tightly orchestrated by the interplay of developmental and environmental cues [7]. Upon perception of light signals through the phytochrome, cotyledons and subsequently leaves undergo structural and physiological changes to develop chlorophyll-containing cells and other components essential for photosynthesis (Fig. 1A) [8]. Photosynthesis allows the conversion of carbon dioxide into organic compounds and generates the energy required for growth and development, thus marking the transition from heterotrophy to autotrophy (Fig. 1A). This reversible transition is a pivotal stage in the life cycle of a plant and is characterized by the shift from relying on internal nutrient sources to self-sustained photosynthesis [9].

Global and drastic changes related to nutrition can also concern animals. The honey bee Apis mellifera is arguably one of the most studied insects. Being eusocial, its biology can be assessed using the concept of superorganism [10,11]. Just like cell society or cells of a pluricellular organism, the individuals making a superorganism achieve labor division, i.e., specialize in different tasks [12,13]. The establishment of this labor division often rests on developmental transitions related to the nutrition function of individuals.

While after mating a queen spends all its life laying eggs, workers perform a succession of tasks [12]. One of the most salient developmental transitions in their behavior is the onset of foraging, which is also the first time they fly, leave the hive, defecate, and are exposed to light. As this is the first time they start collecting food, this is an important behavioral transition. This transition is regulated by the presence of older foragers so that as long as these keep on foraging, younger workers' flight onset is delayed (Fig. 1B) [12,14]. Interestingly, the onset of foraging is correlated with the development of the mushroom bodies, paired brain regions involved in integrating sensory information as well as in learning. Such brain development is consistent with the demanding cognitive tasks performed by foragers (learning flower characteristics, orientating around the hive) (Fig. 1B).

Up to now, we have presented data indicating nutrition-linked developmental transitions in multicellular organisms, but they also exist in unicellular organisms. Some bacteria also undergo morphological shifts depending on their nutritional conditions. *Escherichia coli* (serotype O157:H7) can survive and persist in various environments such as soil, water, or the digestive tract of an animal [15], requiring a considerable capacity to adapt to variations of the environmental conditions. In this regard, motility confers an important advantage to bacteria by allowing them to move towards environments with favorable conditions or away from harmful environments;

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

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

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Irreversible and reversible morphological changes of part of organisms involved in the reproduction of living beings. (A) Formation of aerial hyphae, septation and differentiation of germinal spores during the asexual reproduction in *Streptomyces*. (B) Transition from vegetative meristem to a floral meristem at the origin of the sexual reproduction in flowering plants. (C) Ascidian metamorphosis and sexual maturation in imago, and color changes in male stickleback before the search of sexual partner and mating. Figure created in BioRender.com.</figcaption></figure>
in particular, it is known to be involved in the search for nutrients within niches conducive to their survival. Generally, and in the case of *E. coli*, motility is ensured by the bacterial flagellum resembling a macromolecular machine composed of a basal body (rotating motor), a hook (universal joint), and a filament (propeller) [16].

Interestingly, motility as well as flagellar biosynthesis by *E. coli* are notably affected by the nutrient resource of short-chain fatty acids (SCFAs) (Fig. 1C). For example, in the digestive tract of animals, SCFAs are end products resulting from the fermentation of dietary fiber by many intestinal microorganisms [17] and are composed of acetic, propionic, and butyric acids [18]. However, the composition of SCFAs as well as their concentration in the intestine are uneven [19]. Depending on the presence and quantity of SCFAs, certain genes regulating flagellum biosynthesis (notably *flhC* and *flhD*) are expressed or inhibited [20]. It has been reported that exposure to a high concentration of butyrate, similar to that of SCFAs in the large intestine, causes the expression of flagellar genes and leads to motility of the bacteria (Fig. 1C) [20]. Therefore, precise regulation of flagellar biosynthesis in response to SCFAs gives *E. coli* a considerable advantage in finding a trophic niche appropriate to its survival within the digestive tract of the animal host [16,20].

To sum up, the transition from heterotrophy to autotrophy in seedlings, triggered by light signals, highlights a crucial shift in plant nutrition through photosynthesis. Similar nutrition-linked shifts are associated with morphological changes in animals, such as honey bees, where workers begin foraging after development of specific cerebral areas according to the density of older foragers inside the hive. Nutrition-driven morphological changes are also evident in unicellular organisms like *Escherichia coli*. These bacteria adjust their motility and flagellar biosynthesis in response to nutrient conditions to optimize their survival. This underscores developmental transitions involving morphological adaptations related to nutrition across diverse life forms. Alongside nutrition, developmental transitions also concern reproduction-linked morphological modifications.

## 2.2. Acquisition of a physiological function associated with morphological modifications: example of reproduction

Reproduction is a hallmark of living beings, and involves either combining genomes from different individuals (sexual reproduction) or only the genome of one organism (asexual reproduction). Sexual reproduction occurs thanks to differentiated cells (gametes). While asexual reproduction is often based on the multiplication of an undifferentiated cell, it sometimes involves cellular differentiation (e.g., oocyte in parthenogenesis). Here, we will discuss how the acquisition of reproduction function occurs through developmental transitions.

Asexual reproduction is illustrated by the soil-dwelling bacterium *Streptomyces* producing germinative spores [21,22]. These spores can resist the lack of nutrients and an aggressive environment allowing the bacteria to survive harsh conditions and to efficiently disperse in new habitats.

The vegetative mycelium of *Streptomyces* includes hyphae that grow by tip extension and by developing new branches, spreading across and deeply into the substrate (Fig. 2A). This hyphal growth is governed by a cytoplasmic protein complex called the polarisome [21], that determines the direction of the growth and ultimately initiates the development of a new branching hyphal tip. This results in a ramifying hyphal network, consisting of long and branching filamentous cells, with an exponential growth (Fig. 2A) [22].

Sporulation requires dramatic shifts in the morphology and physiology of the cells, governed by a global cascade of internal factors [23]. The hyphal branches first specialize into aerial hyphae that form a hair-like layer on the surface colonies. The extension and growth of the aerial hyphae requires the sporogenic structures to be covered by a hydrophobic sheath, to escape the aqueous environment of the vegetative mycelium [24–26]. Following the erection of the aerial hyphae, septation occurs and each multigenomic aerial hypha synchronously differentiates into a long chain of unigenomic pre-spores. Finally, pre-spores mature before being released into the surrounding environment as thick-walled dormant spores. Once favorable conditions are encountered, the spores germinate and produce a new vegetative mycelium (Fig. 2A).

This example of *Streptomyces* illustrates a developmental transition consisting of physiological and morphological changes characterized by the differentiation of several cell types inside a cell society in which some members are involved in reproduction [27].

In multicellular organisms such as angiosperm plants, the floral transition involving tissue differentiation represents a pivotal developmental switch orchestrated by the shoot apical meristem (SAM), which also influences the production of leaves, flowers, and fruits. Flowering is controlled by a complex genetic network that integrates a multitude of external factors along with internal signals [28,29].

The latter initiates a cascade of molecular events that trigger the profound morphological changes which will alter the identity of the meristem fate to initiate its transformation from vegetative to floral identity [30]. During this transition, the vegetative meristem gives rise to an inflorescence meristem, which serves as a precursor to the floral meristems or directly transforms into floral meristems, depending on the specific plant species (Fig. 2B).

These floral meristems, in turn, undergo a series of morphological changes in size and shape, culminating in the formation of four fundamental differentiated structures within the flower: sepals and petals, which constitute the sterile organs, and stamens and carpels, representing the fertile organs (Fig. 2B) [31]. Each of these structures arises sequentially from distinct whorls that originate from the floral meristem.

In spite of the phylogenetic distance, reproduction through differentiation can be found in both bacteria and plants, but also in animals such as ascidians. Ascidians are important members of marine benthic communities and represent the largest group of the subphylum Urochordata, which is the sister group of the Vertebrata within the phylum Chordata [32,33]. Almost all ascidians have a biphasic life cycle that is characterized by the irreversible and dramatic transformation of an asexual larva into a hermaphrodite imago (final form of the adult). After adhesion on an appropriate substratum, the larva enters a rapid metamorphosis marked by numerous morphogenetic and physiological events including resorption of the long and muscularized tail, and also the formation of adult-specific organs such as differentiated gonads involved in reproduction (Fig. 2C, animal on the left) [34].
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Internal factors elicit and mediate cellular processes involved in developmental transitions. Developmental transitions are the result of various cellular processes such as migration, division, differentiation, and death (A). Their mechanisms are elicited by molecules that act in a targeted and specific manner (B). Developmental transitions are mediated through the activation of molecular signaling pathways either acting at the level of effector genes or on other molecules; these signaling pathways are highly conserved during evolution (C). Figure created in **BioRender.com**.</figcaption></figure>
The sexual maturity of imago is reached in a few weeks after metamorphosis and is marked by the differentiation of a her-
maphrodite reproductive apparatus (Fig. 2C, animal on the left) [35]. The female and male reproductive organs develop on the lateral
body wall with the gonoducts opening in the cloacal cavity. Concomitantly, the primordial germ cells embedded in the endoderm of
the tail migrate towards the trunk during the tail regression to differentiate into gametes: spermatozoa and oocytes. After the formation
of the testicular wall, the testis becomes a compact mass in which the meiotic spermatogonia is initiated. The wall separates the testis
from the developing female gonad and divides it into testicular follicles. It follows a gradual growth of the testis with the production of
mature sperm in the adult (Fig. 2C, animal on the left) [36]. In addition to sexual reproduction, most colonial ascidians are also able to
reproduce asexually by budding and strobilation [37].

So far, the developmental transitions described above share the common point of being irreversible. Morphological changes initiated are permanent, although they can be reabsorbed through programmed degeneration. Nevertheless, some developmental transitions involved in reproduction are reversible such as coloration change in sticklebacks, small teleost fishes.

Along with other authors, Tinbergen [38] has described morphological changes linked to their reproduction, particularly in males *Gasterosteus aculeatus* known as the three-spined sticklebacks. During the reproduction period, males change their coloration by exhibiting a bright and red color on the throat and ventral regions, as well as blue eyes (Fig. 2C, animal on the right). This red coloration is due to the filling of chromatophores cells with carotenoid pigments obtained through food [39].

While much work has been carried out to understand the morphological changes in male sticklebacks during reproduction, fewer studies have focused on females. However, females also exhibit a developmental transition that is concomitant with those of males. The eyes of the females become more sensitive to red wavelength [40] and, in some populations, they display a dark mottled or bar-like pattern along their back and upper flanks during reproduction, and this mottled pattern stimulates the male [41]. Similarly, egg production induces swelling of the abdomen and this morphological change also triggers the courtship behavior of males [38]. Thus, more than a simple consequence of egg production, this morphological switching acts as a courtship-releasing signal for males.

Overall, one of the interesting points about this developmental transition is its reversibility and cyclicity associated with that of the reproductive activity. As a result, it can occur several times in the stickleback's life [42].

In short, asexual reproduction in the soil bacterium *Streptomyces* involves producing resilient spores to survive harsh conditions. These spores emerge from a complex hyphal network that undergoes morphological changes, demonstrating a modulation of reproduction linked to morphological transformations. Similarly, in angiosperm plants, the floral transition involves tissue differentiation and profound morphological changes, transforming the shoot apical meristem to flowers. Sticklebacks exhibit cyclic morphological changes during reproduction, with males developing bright colors and females altering their eye sensitivity and body patterns. These examples across bacteria, plants, and animals highlight the commonalities of reproduction-linked morphological modifications through developmental transitions.

Thus, during a short and timely period of the life, organisms undergo developmental transitions resulting in morphological shifts related to the acquisition of a physiological function such as reproduction or modulation of a function as in nutrition. Although developmental transitions can be considered irreversible at first approximation, some are distinguished by their reversibility. A reversible process is a process in which the system and its environment can return to exactly the same state they were. Indeed, etiolation (transition from autotrophy to heterotrophy) and de-etiolation (transition from heterotrophy to autotrophy) of plants, the formation or the loss of flagellum in *E. coli*, or the change in body pattern in sticklebacks, are all examples of reversible developmental transitions which occur several times during the life cycle depending on endogenous and exogenous factors. This raises the question of how such transitions are induced and regulated; the next two sections aim to cover these aspects.

**3. Common internal factors elicit, mediate, and regulate cellular processes involved in developmental transitions**

Developmental transitions imply various cellular processes including migration, proliferation, differentiation, and death. These cellular mechanisms are initiated by molecules that act in a targeted and specific manner.

3.1. *Developmental transitions are the result of cellular processes: migration, proliferation, differentiation and death*

The development of multicellular organisms is essential to the growth or even the regression of tissues and organs. In ascidians, the
orchestration of metamorphosis is rapid in comparison with other animals and the cell lineages are well-described; thus, they provide a
convenient experimental model to decipher the cellular mechanisms underlying this developmental process [43]. As discussed above,
the free-swimming larva transforms into a sessile adult through a series of synchronized morphogenetic modifications that are
enhanced by the anchoring of the larva to its substrate [43,44].

Over the last decades, there has been an increasing body of evidence that three types of changes take place during this ascidian larva-to-adult transition. These are the remodeling and degeneration of larval structures coupled with the *novo* development of adult tissues, which appear to involve various cellular processes including cell migration, proliferation, and differentiation as well as apoptosis and phagocytosis [45]. For instance, the initiation of metamorphosis is accompanied by an intense mitotic activity leading to the growth of adult organs [46], which occurs in parallel with the loss of adhesive papillae and the resorption of the larval tail that is the most spectacular metamorphic event in ascidians. This latter involves apoptotic cell death that begins at the end of the tail and progresses until its base (Fig. 3A) [47].

Cell proliferation and differentiation are also cellular processes involved in development of angiosperm plants. In the vegetative phase of plant development, meristem growth is characteristically indeterminate (i.e., it can continue as long as appropriate conditions are maintained), exhibiting a continuous and unrestricted pattern. By contrast, as the plant transitions into the reproductive phase,
meristem growth assumes a predominantly determinate nature [48,49].

The transition from the vegetative to the reproductive phase, known as the floral transition, involves significant changes in the shape and identity of the shoot apical meristem (SAM). The SAM transforms into an inflorescence meristem, which gives rise to floral meristems responsible for flower formation [50] by an increase in cell division and size within the SAM (called doming). These cellular events are earliest indicators of the impending floral transition [48,49].

The cell proliferation status of cells in the SAM undergoes modifications during the floral transition, a phenomenon first reported in the 1960s through cytological approaches [51]. More recently, Jacqmard et al. [48] have shed further light on this process in *Arabidopsis thaliana*. The study revealed that there is a notable increase in mitotic activity, marking one of the earliest events in the SAM during the transition from vegetative to reproductive growth. This increase in mitotic activity precedes other critical events such as the enlargement and doming of the SAM, bolting, and the initiation of the first floral meristem (Fig. 3A) [48]. Recently, RNA-seq analysis of an apical meristem time series revealed important changes in cell cycle gene expression during floral initiation in *Arabidopsis* (Fig. 3A) [52].

In angiosperms, a pattern emerges, where the vast majority of flowers exhibit four distinct types of floral organs: sepals, petals, stamens, and carpels. The widespread prevalence of this pattern strongly suggests the existence of an underlying robust mechanism governing floral development which exhibit a dynamic shift in the balance between the rates of cell differentiation and proliferation. Each angiosperm species seems to follow this pattern, albeit with a certain degree of variation.

As discussed above, in bacteria, cellular differentiation also exists both in isolated individuals (e.g. *Bacillus subtilis*) and in multi-cellular formations as in certain actinomycetes. Indeed, studies showed that cell differentiation is involved in cooperation between the different bacterial phenotypes, in particular during the formation of aerial hyphae in *Streptomyces*. The vegetative mycelium secretes proteases that participate in substrate decomposition, thereby providing nutrients for sporulation [53,54], in parallel of a releasing nutrient by apoptosis of vegetative mycelium [53,55-57]. The nutrient intakes conduct to the differentiation of other bacterial types involved in the formation of aerial hyphae during sporulation (Fig. 3A) [53,55].

Furthermore, other work performed on mobile prokaryotes, *Bacillus subtilis*, has demonstrated the importance of adhesion to a substrate and the formation of a specialized bacterial communities under the form of aggregates embedded in a newly synthesized matrix composed of extracellular polymeric substances (EPS) including polysaccharides, lipids, proteins, and/or DNA. These clumped microbial communities are referred to biofilm [58] that represents a prevalent bacterial life form [59] in natural environments, industrial processes, and disease contexts [58]. Bacterial biofilms reflect a reversible developmental transition from a planktonic solitary phase to a multicellular sessile life stage, triggered by specific environmental cues, such as nutrient depletion, temperature, and/or pH changes [60-62]. The multicellular, even multi-species, composition of biofilms is marked by functional and phenotypic specializations of various bacterial types, microbial populations exhibit different genetic expression and metabolic profiles exhibiting different metabolic and genetic expression profiles.

For example, bimodal distributions in *B. subtilis* gene expression levels have been associated with the division of labor of different bacterial populations during biofilm formation such as motility, surfactant production, matrix production, proteases synthesis, and sporulation [63-65]. At the periphery of the biofilm and in contact with the surrounding environment, the motile bacteria positively express the set of genes necessary for the production of flagella [66]. However, other genes inhibit the rotation of flagella, thus stopping the motility of bacteria encased in the biofilm and enhancing the production of the extracellular matrix [67]. More internal to biofilm, the surfactin-producing bacteria synthesize this surfactant [68-70] acting as a signal molecule to promote the production of the matrix in the periphery [71,72]. Then, at the center of the biofilm where the absence of nutrients induces intense cellular stress, there are dormant and resistant bacteria after the achievement of the sporulation process [73-76].

This capacity for cell differentiation in the biofilm formation is attributed to a certain number of factors influencing the expression of microbial genes: external stress factors, the absence or presence of nutrients and wastes, but also the differential diffusion of intercellular signaling molecules [77-79].

To sum up, developmental transitions in multicellular organisms result from intricate cellular processes. In ascidians, metamorphosis from a larva to a sessile adult involves synchronized morphogenetic changes like cell migration, division, differentiation, and apoptosis. For example, mitotic activity leads to adult organ growth, while larval structures degenerate *via* apoptosis. Similarly, in angiosperm plants, the transition from vegetative to reproductive phases involves changes in the SAM sustained by increased cell division and differentiation. In bacteria, cellular differentiation also plays a crucial role. *Streptomyces*, for instance, forms aerial hyphae during sporulation through apoptosis. Furthermore, *Bacillus subtilis* demonstrates the importance of substrate adhesion and community formation in differentiation, leading to biofilm formation. These transitions underscore the universal role of cellular processes such as proliferation, differentiation, and apoptosis in the developmental transitions across various life forms. All these events are controlled by internal factors. These will now be discussed by covering their implications in developmental transitions.

## 3.2. Hormones and autoinducers are molecules that elicit developmental transitions

Several molecules produced by living beings are known to orchestrate developmental processes by governing the cellular events described above. They exert their influence in a targeted and specific way, acting both locally and over extended distances. They contribute to the proper functioning of organisms by controlling a myriad of physiological functions, and specifically in reproduction. Indeed, bird reproduction is mediated by multiple hormones, which also act on the plumage. The plumage of certain species can change during sexual maturation and according to seasons [80,81].

Plumage colorations depend on differentiation and distribution of specific pigment cells named melanocytes (Fig. 3B). As in mammals, the plumage coloration of birds is mainly due to endogenous pigments such as melanin [82,83]. Plumage changes are
largely controlled by hormones synthesized by the pituitary and thyroid glands, which are also involved in reproduction (Fig. 3B) [84,85]. Injection of pituitary hormones such as α-melanocyte stimulating hormone, follicle-stimulating hormone, and luteinizing hormone was shown to induce unusual black pigmentation of feathers in *Lagopus leucura* during the winter [86], thus revealing the hormonal control of feather pigmentation. Besides the actions of pituitary hormones, thyroid hormones play also an important role in regulating plumage change: a thyroid ablation prevents feather color change [87]. Hence, in the context of sexual maturation and more broadly in reproduction, hormones are crucial molecular actors in the color shift of plumage, by acting in particular on the differentiation of melanocytes [88].

As described above, in male stickleback display a bright red coloration on the throat and ventral regions [39]. This change in coloration is also under the influence of hormones. Pharmacological treatments combined with castration provided evidence for a predominant role of the androgenic hormone 11-ketotestosterone in the control of male developmental transition. Indeed, the red coloration appears in castrated males when they are treated with 11-ketotestosterone [89].

The examples of birds and sticklebacks highlight the critical roles of hormones in the initiation of developmental transitions, particularly steroid hormones. The latter also exist in plants. Brassinosteroids (BRs) are a group of polyhydroxylated steroid hormones that play a crucial role in plant development [90,91]. Moreover, besides their role in controlling plant development, BRs have a multifaceted role in integrating environmental signals that govern the floral transition. A notable aspect of their function is their ability to modify chromatin methylation levels, which in turn controls the expression of the key floral repressor, Flowering Locus C (FLC) [92]. As a result, BRs play a dual role in the control of flowering. On the one hand, they can delay flowering by triggering FLC expression, while on the other hand, they integrate environmental cues to tightly control flowering time.

BRs also have a positive impact on male fertility by directly controlling key genes that govern the development of anthers and pollen (Fig. 3B) [93]. It is noteworthy that mature pollen is rich in BRs, indicating their essential role in enhancing fertilization efficiency, particularly under adverse conditions.

Similar to hormonal communication in plants and animals, most bacteria use a density-dependent cell-cell communication system known as *quorum sensing* (QS) [94,95], which relies on the secretion of diffusible signaling molecules called autoinducers, that diffuse through the bacterial membrane or are exported outside the cell. These autoinducers enable bacteria to adjust their phenotypic responses according to cell density and to synchronize the regulation of gene expression involved in social behaviors, such as virulence, antibiotic production, and biofilm formation and maturation [96]. The biofilm provides a stable structure for bacteria, wherein they can perform multiple tasks such as nutrient acquisition, protection against predation or host antimicrobial compounds, and stress tolerance as in *B. subtilis* described above [61,97–101].

Several phenotypes involved in biofilm formation are often QS-regulated [62,102], such as adhesin production required for cell attachment, production of extracellular polysaccharide that form the biofilm matrix, or motility and chemotaxis [95,96,101,103]. The most common and well-described QS molecules are acylhomoserine lactones (acyl-HSL), mediating QS signals coordinating biofilm formation in many bacteria (Fig. 3B) [62,104,105].

In short, hormones and autoinducers are critical molecules that elicit developmental transitions in living organisms. In birds, hormones from the pituitary and thyroid glands regulate plumage changes and reproduction, influencing melanocyte differentiation and migration. In male sticklebacks, steroid hormones are essential for the development of red coloration during reproduction, as in plants where brassinosteroid hormones are crucial for flowering and development of male fertility. Furthermore, in bacteria, *quorum sensing* uses autoinducers to coordinate gene expression related to social behaviors like biofilm formation. These signaling molecules enable bacteria to adjust their responses based on cell density, synchronizing activities essential for survival and adaptation. Thus, hormones in animals and plants, and autoinducers in bacteria, are pivotal in regulating developmental transitions by acting on cellular processes previously described. Hormone and autoinducer synthesis are also under the control of environmental factors which, by extension, modulate their actions through molecular signaling pathways.

### 3.3. Molecular signaling pathways mediate the execution of cellular processes involved in developmental transitions

A signaling pathway is a sequence of steps involving a cascade of molecules located on the plasma membrane and/or in the cell cytoplasm. As part of developmental transitions, signaling pathways are important in mediating cellular processes such as cell migration, proliferation, differentiation, and death by activating or inhibiting the expression of effector genes.

Some molecular signaling pathways are highly conserved during evolution. One example is nitric oxide (NO) signaling pathway; NO is a gaseous molecule controlling many physiological responses in eukaryotes and prokaryotes [106,107].

For instance, at low (nanomolar) concentrations, NO is involved in signaling pathways in bacteria during *quorum sensing*-elicited biofilm formation [108]. It has been recently shown that endogenously produced NO by NO synthase (NOS) could control one of the developmental transitions (aerial mycelium formation) in the *Streptomyces* life cycle as described above (Fig. 3C) [109]. Among key developmental factors in this transition, *bld* genes control the formation of the hair-like reproductive aerial hyphae in a complex extracellular signaling cascade [23], with a central regulatory role of the nucleotide second messenger cyclic di-GMP (c-di-GMP) [110]. Indeed, c-di-GMP signaling controls the initiation of the morphogenesis, by forming a complex with BldD, a master repressor of sporulation genes, thereby controlling the hypha-to-spore transition [22]. At high and low levels of c-di-GMP, BldD induces vegetative growth and aerial mycelium formation, respectively [111]. Honma et al. [109] reported that the intracellular c-di-GMP production and the activity of BldD depend on endogenously produced NO.

In multicellular eukaryotes, the NO pathway is also crucial in plant development and signaling, with a wide range of impacts on plant reproduction. Instead of NOS [112], in land plants, the enzyme Nitrate Reductase (NR) plays a crucial role in NO synthesis [113]. In *Arabidopsis*, high levels of NO were linked to delayed flowering by suppressing *LEAFY* (*LFY*) and *CONSTANS* (*CO*) genes, which
control floral induction, and increasing the expression of the *FLC* floral repressor gene [114,115]. Overall, NO has diverse effects on plant biology, influencing flower development and pollen production. Is this signaling pathway also fundamental in the modulation of developmental transitions in animals?

NO is known to act as a regulator of metamorphosis [116] and apoptosis [117] in several animal groups. As described above, the ascidian larvae exhibit a tail regression step during its metamorphosis through the induction of a polarized apoptotic wave leading to the removal of most tail cells [47,118]. A high level of expression of *NOS* is present at the end of larval life, just before tail regression, and conjointly with a wide diffusion of NO through the whole body. It has been demonstrated in various species of ascidians that NO functions in a dose-dependent fashion as an inhibitory or inducer of settlement and metamorphosis of tadpole larvae [119–121]. In addition, there is a correlation between NO levels and the activation of apoptosis during the last steps of tail regression, thus placing the NO signaling pathway as another potential modulator of apoptosis in ascidian metamorphosis [120].

Therefore, NO plays a conserved role in mediating developmental transitions, although its ubiquitous nature grants it many other roles (e.g., see Dacher & Gauthier, 2008 [122]). Similarly, multicellular eukaryotes share another ubiquitous signaling pathway in developmental processes, the mitogen-activating-protein kinase (MAPK) pathway.

Recent studies highlighted in the solitary ascidian *Ciona intestinalis* that the intracellular activation of caspases (a family of cysteine proteases well known for their central roles during apoptosis) is required for mediating the proteolytic events indispensable for the initiation and execution of tail degeneration [123,124]. This apoptotic process was also found to be controlled by the activation of various intracellular signaling pathways, including the MAPK pathway [125–128]. It has been reported in *C. intestinalis* that some components of the MAPK signaling cascade are highly expressed in tail tissues before the triggering of apoptosis [125]. Moreover, their inhibition was shown to prevent the development of metamorphosed larvae as well as tail regression through a blockage of apoptosis; this was associated with a decline in the number of dying cells at the tip of the tail [125].

These molecular signaling pathways are known to act on effector genes. It is well-known that *Hox* genes are effector genes of the MAPK pathway and encode an evolutionarily conserved set of transcription factors. These comprise a homeodomain DNA-binding domain which play a key role in forming the anterior-posterior body axis pattern during the early developmental stages of animals. Some ascidian *Hox* genes especially *Hox1*, *Hox10*, and *Hox12* are highly expressed during embryonic and larval development but also in juveniles [129–131]. The loss of function of the *Hox1*, *Hox10*, and *Hox12* provokes severe body malformations and even death in juveniles whereas larvae displayed a normal tadpole morphology. For instance, *Hox10*-disrupted juveniles had lost their intestine, whereas *Hox* genes are not critical for the construction of tadpole shape of larva; however, they are expressed from the larval stage and fulfill their functions during the larval-juvenile transition by controlling the morphogenesis of adult organs including digestive tract (Fig. 3C) [130,132]. *Hox10* was shown to be involved in this softening of the extracellular matrix by repressing the expression of *collagen type IX* gene, thus reflecting a relevant role of *Hox10* in the cell migration and the formation of adult digestive apparatus during the ascidian metamorphosis [45]. Recently, several studies in ascidians have revealed regulatory interactions of the *Hox* genes with the MAPK signaling pathway [133,134].

Over the past decade, research on MAPK has notably unveiled their roles in plant growth and development. Expression pattern analyses have shown that MAPKs are widely expressed in various plant tissues and organs, including inflorescences and anthers (Fig. 3C) [135]. MAPK signaling has been found to control floral development, size, and abscission. Notably, MAPK phosphatases have been linked to petal growth rate and size during flowering [136].

In summary, molecular signaling pathways, such as nitric oxide (NO) and mitogen-activated protein kinase (MAPK), play crucial roles in modulating cellular processes involved in developmental transitions across diverse living beings. In bacteria, NO signaling influences *quorum sensing* and biofilm formation, while in *Streptomyces*, it regulates morphogenesis. In plants, NO modulates flower development and pollen growth, delaying flowering by influencing gene expression. Moreover, in animals, NO regulates cellular processes such as apoptosis, exemplified by its role in ascidian tail regression during metamorphosis. Similarly, the MAPK pathway is integral to developmental transitions. For instance, in ascidians, MAPK signaling is involved in apoptosis during tail regression by regulating *Hox* gene expression. In plants, MAPK pathway mediates flower transition through impacts on floral organ size and growth. These conserved pathways underscore the fundamental mechanisms by which molecular signals drive developmental transitions across various life forms.

Thus, in this section, we highlighted remarkable similarities in developmental transitions across living beings (Fig. 3). These developmental transitions involve cellular processes (migration, proliferation, differentiation, and death) initiated by molecular elicitors (hormones and autoinducers in *quorum sensing*); they act on effector genes through molecular signaling pathways highly conserved during evolution (NO and MAPK pathways). Interestingly, elicitor synthesis, activation of previously described molecular signaling pathways, and gene expression are regulated by external factors; the following sections will explore this aspect.

# 4. Common external factors mediate developmental transitions

The environment features predictable changes (such as seasons) or predictable components (such as other living beings). Thus, these could play a crucial role in initiating developmental transitions in all organisms. Indeed, abiotic factors such as temperature and photoperiod are developmental synchronizers.

## 4.1. Abiotic conditions influence developmental transitions

Shifts in day length led to physiological modifications in photoperiod-sensitive living beings, allowing them to adapt to predictable annual climate changes. Light and photoperiod act on the biosynthesis of molecules, the expression of effector genes, and the cellular
processes. The myxomycete (Amoebozoa) *Physarum polycephalum*, a unicellular slime mold known under the name “Blob”, undergoes different morphological features from the vegetative to the sporulation stages in its life cycle [137]. This developmental transition is notably mediated by light [138–143]. As the plasmodium differentiates toward the sporulation-competent state, sensitivity to light develops with cessation of growth, condensation of cellular material, division by mitosis, and migration of the multinucleated cell.

It is known that light interferes with the NO signaling pathway discussed above; light-pulse-triggered sporulation is processed by a phytochrome-type photoreceptor [143]. Stimulation of this photoreceptor induces the expression of several genes of cell cycle a few hours after photoactivation of phytochrome (Fig. 4A) [139].

In addition, some proteins associated with light-induced pathways have been characterized [144,145]. The actin cytoskeleton of *P. polycephalum* is essential for its locomotion, division, and other biological processes [146]; the assembly and disassembly of actin filaments are controlled by a group of actin-related proteins, whose activities are regulated by specific signaling pathways. Barrantes et al. [147] showed that some actin-binding genes were specifically involved before and after photoinduction in the starved *P. polycephalum* plasmodium. Genes encoding actin-binding proteins are downregulated after photoinduction [147]. Thus, during sporulation, the cessation of production of such proteins is probably involved in the reorganization of subcellular compartments through remodeling of the actin cytoskeleton and prepares for the entrance in sporulation (Fig. 4A).

The case of *P. polycephalum* exemplifies the connection between environmental stimuli and the internal process previously discussed, as most of the plant developmental processes. Environmental cues enable plants to develop and bloom during favorable season for reproductive success and seed production. Light and especially day length or photoperiod provide one of the environmental cues used to achieve this. Through the photoperiod pathway, day lengths trigger the production of a systemic flowering signal FLOWERING LOCUS T (FT) termed florigen, widely conserved in angiosperms [148], to trigger the floral transition [149]. In long-day plants, FT expression is rhythmically activated by the output of the photoperiod pathway CONSTANS (CO), specifically at the end of long days. CO plays a central role in the photoperiod response pathway by mediating between the circadian clock, the floral regulators FT, and the integrator SOC1 [150]. The photoperiodic cue interacts with the repression pathway, which regulates the competence of the SAM (Fig. 4B).

These intricate responses of plants to photoperiod, highlight how quantitative changes in signals can finely modulate developmental transitions in plants. They also offer insights into the evolutionary adaptations that underlie the observed variability in transitions among different plant species: rather than automatically occur, they integrate internal constraints and environmental situations, ultimately providing adaptive response to external conditions. This is also the case in anuran amphibians which undergo complex and extensive morphological changes as tadpoles progress through metamorphosis to become juveniles. These changes include the coordinated development and remodeling of organs, generation of limbs, tail regression, and the acquisition of new

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Light and photoperiod act on the biosynthesis of molecules, expression of effector genes, and the cellular processes responsible for developmental transitions. (A) Light effects on gene regulations involved in modifications of cell cycle and actin filament polymerization at the origin of the sporulation of *Physarum polycephalum*. (B) Photoperiod acts on the repression pathway of flowering plants which inhibits the competence of the shoot apical meristem (SAM). CO: CONSTANS, FT: florigen, SOC1: integrator encoding a MADS box transcription factor. (C) Photoperiod impacting the control of anuran metamorphosis. Figure created in BioRender.com.</figcaption></figure>
behaviors (e.g., shift from herbivory to carnivory and emergence of predatory behavior), related to changes in diet and the transition from an aquatic to a terrestrial environment for most [151].

During this metamorphosis, the molecular, cellular and physiological events are initiated and mediated by thyroid hormones (TH) [152] under the influence of environmental factors such as photoperiod. Indeed, the amphibian endocrine system depends on the activity of a circadian clock: TH biosynthesis follows a rhythmic 24-h cycle [153]. Studies have shown that an increasing in the duration of the light phase of a 24-h light/dark (LD) cycle, or even a decrease in the duration of the LD cycle, promotes metamorphosis after TH stimulation [154–156]. The mechanisms by which photoperiod impacts metamorphosis are still poorly understood. Nevertheless, it has been demonstrated that changes in the photoperiod lead to variations in the biosynthesis of THs, through the action of another hormone: the melatonin [153]. Melatonin is produced by the pineal gland and the retina, and is a photoperiod-sensitive hormone providing organisms with temporal information [157]. Moreover, this hormone exerts a direct inhibitory effect on the thyroid gland at all stages of the anuran amphibian life cycle [158]. Melatonin acts by stopping the development of the thyroid gland and also by limiting cell proliferation as described in *Rana pipiens* [159]. In larvae, the melatonin blood level is high and its decline initiates metamorphosis. Therefore, photoperiod is an environmental factor impacting metamorphosis in anuran amphibians, through its indirect action on the thyroid pathway activity via melatonin, considered as the time-keeping hormone (Fig. 4C).

To sum up, abiotic factors, such as light and photoperiod, significantly influence developmental transitions in living beings through molecular signaling pathways. In *Physarum polycephalum*, light triggers the transition from vegetative to sporulation stages, mediated by a phytochrome-type photoreceptor that influences gene expression and cellular processes. Similarly, in plants, photoperiod regulates flowering through the production of crucial actors, aligning reproductive success with favorable seasons. In amphibians, photoperiod also affects metamorphosis by modulating thyroid hormone biosynthesis through circadian rhythms. These examples demonstrate how environmental cues like light and photoperiod finely tune developmental transitions across diverse species, enabling adaptation to seasonal changes. In addition to all these abiotic parameters, biotic conditions can also modulate developmental transitions.

## 4.2. Living beings can impact the developmental transitions of other living beings

The dynamic networks within ecosystems involve intricate interactions among all living organisms. These connections vary in intensity and can manifest at different stages of an organism's life. Whether between individuals of the same species or between individuals of different species, certain biotic interactions are essential to the development of living beings and in particular during the sex change in some vertebrates. Sex is generally determined by either endogenous genetic component, external environmental component, or a combination of both. The primordial gonad differentiates either into an ovary or a testis and thus determines the animal's sex. Thereafter, the differentiated gonads undergo active maintenance by repressing the opposite sex throughout adult life [160–162].

While most species are gonochoric (i.e., sex is determined at the end of sexual differentiation and will remain unchanged in the adult state), it is common for some animals to possess both male and female gonads at the same time: these are simultaneous hermaphroditic organisms. This sexual state is present in several clades of animals as another type of hermaphrodism involving sex changes during the adult life: successive hermaphroditism. Teleost fishes show both simultaneous [163–165] and successive hermaphroditism [166].

For example, in gobies (e.g., *Lythrypnus dalli*), subordinate individuals perceive the dominant animal through visual, chemical, and tactile cues, or a combination of these senses, depending on species-specific sensory modalities. Within a harem comprising protogynous fishes (i.e., mature females), some females will become males, the latter often having better reproductive success [167]. This developmental transition occurs when a male withdraws and enough females make up the harem. Since then, the larger female changes sex to male [168–172]. Several researches have shown that sex change may be a plastic response to the local social environment [173,174] characterized by the inhibition of behavior and/or reproductive function in subordinates by dominant animals, and could considered as a reversible developmental transition. More generally, beyond fishes, sex change occurs in animals as diverse as annelids, echinoderms, crustaceans, and mollusks, but also in plants [175].

Besides interactions between multicellular organisms, the presence of unicellular organisms can induce developmental transitions of multicellular organisms, such as between cnidarian larvae and bacteria of the *Pseudoalteromonas* genus which are responsible for metamorphosis induction in these animals.

Cnidarians are mainly benthic marine animals. Their life cycle is mostly characterized by a planula larva settling on a substrate and then metamorphosing into a sessile benthic organism called a polyp. In addition, the search for and recognition of a conductive substrate for its development is restricted to the larval stage. However, the larvae do not possess sensory structures allowing them to remotely locate a suitable substrate. As a result, they distinguish their habitat by a hierarchy of specific key stimuli. This is the case for certain anthozoans (*Nematostella* sp., *Acropora* sp.), scyphozoans (*Cassiopea* sp.), and hydrozoans (*Coryne* sp., *Proboscidactyla* sp., *Hydractinia* sp.) [176]. The last stimulus initiating the installation of the larva and the metamorphosis comes from bacteria located on the substrate.

Chemical signals in marine biofilms induce larval settlement and metamorphosis are well known [177–179]. However, the identification of the signaling molecules is difficult and only a few key bacterial signals have been characterized [180–182], such as bromopyrrole produced by a bacterium of the *Pseudoalteromonas* genus [183,184]. Interestingly, stimulation by bromopyrrole alone is not sufficient to attach the larvae to the substrate and induce metamorphosis. Recent research on bacteria-induced metamorphosis of *Hydroides elegans* demonstrated the involvement of contractile injection systems (CISs, phage tail-like machineries found in a wide range of bacteria) in *Pseudoalteromonas*. These CISs are responsible for the colonization of the substrate by the cnidarian as well as its
metamorphosis through the transfer of an effector protein stimulating the signaling pathways of mitogen-activated protein kinase (MAPK) [185-187].

As in cnidarians, the biotic environment induces developmental transitions involved in the Rhizobium-legume symbiosis. Indeed, in this case a joint developmental transition occurs in both organisms: their intricate leads to the development of a plant-derived structure efficient for biological nitrogen fixation (Fig. 5). Symbiosis begins with a complex exchange of signals, enabling the *Rhizobium* to penetrate roots down towards the nodule primordial cells. Indeed, the initial stage of symbiosis begins with the activation of transcription regulators known as NodD proteins by plant root exudates, which contain a mix of flavonoid compounds. The recognition between flavonoids and NodD proteins is specific and triggers the expression of nodulation (*nod*) genes by binding to conserved sequences called *nod-box* located upstream of bacterial *nod* operons (Fig. 5) [188]. This activation leads the rhizobia to produce Nod-factors signal molecules recognized by specific plant receptors [189] and thus are key inductors of nodule development. Nod-Factors elicit, when appropriate, diverse transitional responses in the plant, such as root hair curling, cortical cell division, and the formation of an infection thread that guides the bacteria toward the emerging primordium [190-192]. Once there, the rhizobia are endocytosed into these cells and enveloped within a specialized symbiotic membrane [193].

Subsequently, the rhizobia transform into bacteroids, essentially functioning as a plant structure that converts atmospheric nitrogen into ammonia (Fig. 5). This ammonia becomes a valuable resource for the host legume. Recent scientific interest has focused on a subset of plant-produced anti-microbial peptides (AMPs) known as nodule-specific cysteine-rich (NCR) peptides and their role in the establishment of this symbiotic relationship [194]. Beyond their antimicrobial properties, there is emerging evidence to suggest that certain AMPs may also play a crucial role in signaling [195]. Remarkably, in specific legume species, bacteroids within plant nodules ultimately undergo terminal differentiation, rendering the nodules unable to resume growth. Interestingly, these legumes are known to produce NCRs anti-microbial peptides [196]. In other legumes, the fate of the bacteroids is not irreversible and endosymbionts can return to free-living life: in this case, there is no change in morphology, size, DNA content, and membrane properties of the bacteria, and these legumes lack genes coding for the mentioned symbiotic peptide families in their genome. Moreover, a mutation in *dnf-1 M. truncatula* gene prevents the proper targeting of NCRs to the bacteroid [197] and impairs its terminal differentiation. This compelling evidence strongly suggests that NCRs are directly involved in the intricate process of terminal differentiation in symbiotic bacteria. This symbiosis, which can be either reversible or irreversible, is an adaptive process: both species get nutritional advantages from it.

In summary, biotic factors significantly influence developmental transitions in living organisms through intricate interactions within ecosystems. These interactions are crucial for biological process such as sex change in certain vertebrates, where social cues and dominance hierarchies can trigger sex reversal in gobies. Furthermore, unicellular organisms can induce developmental changes in multicellular organisms. For example, specific bacteria in marine biofilms release chemical signals that trigger metamorphosis in

<figure><img src="image_8.png" /><figcaption>**Fig. 5.** Interaction between a unicellular and a multicellular organism can cause a joint developmental transition in both organisms: example of the Rhizobium-legume symbiosis. ER: Endoplasmic Reticulum, NCRs: Nodule-specific Cysteine-Rich peptides, NFRs: Nod Factor Receptors, Nod: Nodulation gene or factors. Figure created in BioRender.com.</figcaption></figure>
cnidarian larvae. Similarly, the Rhizobium-legume symbiosis exemplifies biotic-induced developmental transitions.

Overall, the examples covered in this section illustrated that developmental transitions are adaptation to abiotic and biotic components of the environment, promoting both reproduction and nutrition, thus enhancing fitness.

## 5. Evolutionary perspectives and conclusions

Developmental transitions are defined as global physiological and/or structural changes in organisms, leading it from one developmental stage to the other. They are observed at different biological scales (morphology, physiology, biochemistry, and ecol- ogy), in all groups of living beings. During transitions of development, biological processes are highly coordinated and restricted to a short period of the organisms' life cycle. Furthermore, developmental transitions are characterized by changes in the living beings' morphology and concern all or only a specific part of an organism. Morphological changes modulate certain physiological functions (e. g., nutrition), or involve the acquisition of new physiological functions crucial to the organism's survival, and more broadly to the species' survival (e.g., reproduction).

Developmental transitions are the result of common cell processes (such as migration, proliferation, differentiation, and death) and are induced by molecules, acting in targeted and specific ways at both short and long ranges, such as hormones and bacterial auto- inducers. Hormones and autoinducers can activate similar molecular signaling pathways (NO and MAPK) that activate or inhibit the expression of effector genes involved in cellular processes driving developmental transitions. Then environmental factors such as light and photoperiod are synchronizers of developmental transitions, which can also be triggered by interspecific interactions (e.g., symbiosis).

Up to now, the material covered in this review highlights the universal presence of developmental transition among living beings. While these developmental transitions are clearly adaptive processes, their evolutionary history can be challenging to reconstruct as it is difficult to determine how selection pressures operated in the past. The field of evolutionary developmental biology (evo-devo) explores the mechanistic relationships between individual developmental processes and phenotypic change over evolutionary time [198,199]. In a general framework, evo-devo aims to explain the origin of the morphological complexity of organisms by comparing the genes that regulate their development. This field also traces the evolution of the development of organisms, enabling comparisons of developmental processes across species while accounting for their phylogenetic relationships [198,199]. Specifically, evo-devo has addressed the conservation of developmental processes, including the transitions examined in this review. For example, recent studies have promoted sticklebacks as valuable evo-devo models [200,201]. These fish exist in the marine, brackish, or freshwater envi- ronments with varying morphologies, despite all being derived from marine ancestors. Phylogenies built on reproductive behaviors and morphologic characters [202] align closely, and the phylogeny of the reproduction-linked developmental transition in Gaster- osteridae is well-documented [203]. Therefore, it would be interesting to integrate this knowledge in sticklebacks into evo-devo studies on their morphological diversification. This connection is supported by Boughman's [204] or Scott's [205] results, who re- ported that diversification of sticklebacks can result from sexual selection facilitated by their developmental transition. This suggests that, beyond resulting from sexual selection, developmental transition also drives evolution. Similarly, Cronly-Dillon and Sharma [40] reported that female sticklebacks experience changes in red sensitivity during the developmental transition. They also mentioned that the same changes are observed in fish which life cycle involves migrating between sea and freshwater environments. As such mi- grations occurred during the evolution of sticklebacks, this observation suggests developmental transition might be, at least partly, an exaptation, i.e., red sensitivity could have originally evolved as a by-product to varying salinity levels [206,207].

Metamorphosis in vertebrates are classical example of developmental transitions. They involve changes in morphology and functions. Interestingly, the way these changes occur (or not) has deep consequences on evolution of vertebrates. In his review on this topic, Laudet [208] showed that variations in metamorphosis, or more broadly developmental transitions, are very important to explain the origine of species diversity. This means they are not only the reflect of evolution, but also one of its mechanisms.

As discussed previously, NO and MAPK pathways are involved in developmental transition in a variety of organisms but are in no way specific to this type of process. Rather, they are ubiquitous and are found in many living beings [209–212], underlying a vast array of functions. This makes exaptation processes possible explanations for the molecular mechanisms of developmental transitions. NO and MAPK could have been repurposed during evolution, causing their implications within distinct functions. Similar reasoning could be performed for other molecular pathways, for instance, steroid hormones [213]. While much work remains to lead this type of topic beyond hypotheses, setting developmental transitions in an evolutionary frame will help formalize future research.

Thus, the material covered in this review points to the universal presence and similar processes of developmental transition among living beings, but it is not plausible that this characteristic was inherited from a common ancestor. Rather developmental transitions have appeared several times in evolution and would be a convergent trait. Such convergence points to similar pressures, likely experienced by all living beings. As a result, developmental transitions would be a motor in evolution by promoting diversification and ultimately apparition of new species.

### CRediT authorship contribution statement

**Evan Force:** Writing – review & editing, Writing – original draft, Validation, Supervision, Conceptualization. **Dominique Lamy:** Writing – review & editing, Writing – original draft, Validation, Conceptualization. **Stéphane Debernard:** Writing – review & editing, Writing – original draft, Validation, Conceptualization. **Arnould Savouré:** Writing – review & editing, Writing – original draft, Validation, Conceptualization. **Matthieu Dacher:** Writing – review & editing, Writing – original draft, Validation, Supervision, Conceptualization.
**Data availability statement**

No data was used for the research described in the article.

**Declaration of competing interest**

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

**Acknowledgments**

Thanks to Thomas Duchateau (École Normale Supérieure de Lyon), Sébastien Duperron and Hervé Tostivint (Muséum National d'Histoire Naturelle), and Jean-François Mauffrey (Aix-Marseille Université) for thoughtful feedback on earlier drafts.

**References**

[1] F. Cohn, Untersuchungen über bacterien, IV. Beiträge zur biologie der Bacillen, Beiträge zur biologie der Pflanzen 7 (1877) (1913) 249-276.

[2] M. Chailakhyan, New facts in support of the hormonal theory of plant development, Comptes Rendus Acad. Sci. URSS 12 (1936) 443-447.

[3] H. Lacaze-Duthiers, Recherches sur l'organisation et l'embryogénie des Ascides, Évolution de la *Molgula tubulosa*, Comptes rendus de l'Académie des Sciences Paris 70 (1870) 1154-1157.

[4] H. Lacaze-Duthiers, On the organization and embryogeny of the *Ascidia*, Development of *Molgula tubulosa*, Ann. Mag. Nat. Hist. 6 (1870) 109-111.

[5] I. Bäurle, C. Dean, The timing of developmental transitions in plants, Cell 125 (2006) 655-664.

[6] A.C. Oliveira, A.R. Rebelo, C.C.F. Homem, Integrating animal development: How hormones and metabolism regulate developmental transitions and brain formation, Dev. Biol. 475 (2021) 256-264.

[7] G. Carrera-Castaño, J. Calleja-Cabrera, M. Pernas, L. Gómez, L. Onate-Sánchez, An updated overview on the regulation of seed germination, Plants 9 (2020) 703.

[8] E. Huq, B. Al-Sady, M. Hudson, C. Kim, K. Apel, P.H. Quail, Phytochrome-interacting factor 1 is a critical bHLH regulator of chlorophyll biosynthesis, Science 305 (2004) 1937-1941.

[9] J.-H. Ha, S.-H. Han, H.-J. Lee, C.-M. Park, Environmental adaptation of the heterotrophic-to-autotrophic transition: the developmental plasticity of seedling establishment, Crit. Rev. Plant Sci. 36 (2017) 128-137.

[10] R.F.A. Moritz, S. Fuchs, Organization of honeybee colonies: characteristics and consequences of a superorganism concept, Apidologie 29 (1998) 7-21.

[11] J. Tautz, D.C. Sandeman, H.R. Heilmann. The Buzz About Bees: Biology of a Superorganism, Springer, Berlin, 2008, p. 284.

[12] B.R. Johnson, T.A. Linksvayer, Deconstructing the superorganism: social physiology, groundplans, and sociogenomics, Q. Rev. Biol. 85 (2010) 57-79.

[13] R.E. Page, J. Erber, Levels of behavioral organization and the evolution of division of labor, Naturwissenschaften 89 (2002) 91-106.

[14] G.V. Amdam, S.W. Omholt, The hive bee to forage transition in honeybee colonies: the double repressor hypothesis, J. Theor. Biol. 223 (2003) 451-464.

[15] D. Ongeng, A.H. Geeraerd, D. Springael, J. Ryckeboer, C. Muyanja, G. Mauriello, Fate of Escherichia coli O157:H7 and Salmonella enterica in the manure-amended soil-plant ecosystem of fresh vegetable crops: a review, Crit. Rev. Microbiol. 41 (2015) 273-294.

[16] H. Sun, M. Wang, Y. Liu, P. Wu, T. Yao, W. Yang, Q. Yang, J. Yan, B. Yang, Regulation of flagellar motility and biosynthesis in enterohemorrhagic Escherichia coli O157:H7, Gut Microbes 14 (2022) 2110822.

[17] J.H. Cummings, G. Macfarlane, The control and consequences of bacterial fermentation in the human colon, J. Appl. Bacteriol. 70 (1991) 443-459.

[18] B. Dalile, L. Van Oudenhove, B. Vervliet, K. Verbeke, The role of short-chain fatty acids in microbiota-gut-brain communication, Nat. Rev. Gastroenterol. Hepatol. 16 (2019) 461-478.

[19] T. Lackraj, J.I. Kim, S. Tran, D.E. Barnett Foster, Differential modulation of flagella expression in enterohaemorrhagic Escherichia coli O157: H7 by intestinal short-chain fatty acid mixes, Microbiology 162 (2016) 1761-1772.

[20] T. Tobe, N. Nakanishi, N. Sugimoto, Activation of motility by sensing short-chain fatty acids via two steps in a flagellar gene regulatory cascade in enterohemorrhagic Escherichia coli, Infect. Immun. 79 (2011) 1016-1024.

[21] K. Flärdh, D.M. Richards, A.M. Hempel, M. Howard, M.J. Buttner, Regulation of apical growth and hyphal branching in Streptomyces, Curr. Opin. Microbiol. 15 (2012) 737-743.

[22] M.J. Bush, N. Tschowri, S. Schlimpert, K. Flärdh, M.J. Buttner, c-di-GMP signalling and the regulation of developmental transitions in streptomycetes, Nat. Rev. Microbiol. 13 (2015) 749-760.

[23] J.R. McCormick, K. Flärdh, Signals and regulators that govern Streptomyces development, FEMS Microbiol. Rev. 36 (2012) 206-231.

[24] M.J. Bibb, A. Domonkos, G. Chandra, M.J. Buttner, Expression of the chaplin and rodlin hydrophobic sheath proteins in Streptomyces venezuelae is controlled by σ(BldN) and a cognate anti-sigma factor RsbN, Mol. Microbiol. 84 (2012) 1033-1049.

[25] K. Flärdh, M.J. Buttner, Streptomyces morphogenetics: dissecting differentiation in a filamentous bacterium, Nat. Rev. Microbiol. 7 (2009) 36-49.

[26] S. Kodani, M.E. Hudson, M.C. Durrant, M.J. Buttner, J.R. Nodwell, J.M. Willey, The SapB morphogen is a lantibiotic-like peptide derived from the product of the developmental gene ramS in Streptomyces coelicolor, Proc. Natl. Acad. Sci. U S A 101 (2004) 11448-11453.

[27] A. Dance, What is a cell type, really? The quest to categorize life's myriad forms, Nature 633 (2024) 754-756.

[28] S.N. Freytes, M. Canelo, P.D. Cerdán, Regulation of flowering time: when and where? Curr. Opin. Plant Biol. 63 (2021) 102049.

[29] Y. Hyun, R. Richter, C. Vincent, R. Martinez-Gallegos, A. Porri, G. Coupland, Multi-layered regulation of SPL15 and cooperation with SOC1 integrate endogenous flowering pathways at the Arabidopsis shoot meristem, Dev. Cell 37 (2016) 254-266.

[30] C. Liu, Z. Thong, H. Yu, Coming into bloom: the specification of floral meristems, Development 136 (2009) 3379-3391.

[31] E. Coen, Goethe and the ABC model of flower development This paper is based on chapter 4 from the Art of Genes (1999) by Enrico Coen, with permission of Oxford University Press, Comptes Rendus Académie Sci. - Ser. III - Sci. Vie 324 (2001) 523-530.

[32] F. Delsuc, H. Brinkmann, D. Chourrout, H. Philippe, Tunicates and not cephalochordates are the closest living relatives of vertebrates, Nature 439 (2006) 965-968.

[33] P. Lemaire, Evolutionary crossroads in developmental biology: the tunicates, Development 138 (2011) 2143-2152.

[34] R.A. Cloney, Ascidian metamorphosis: Review and analysis, in: F-S Chia, ME Rice, editors (Eds.), Adhesion and metamorphosis of marine invertebrate larvae, Elsevier, Amsterdam, 1978, pp. 255-282.

[35] A. Rosner, E. Moiseeva, Y. Rinkevich, Z. Lapidot, B. Rinkevich, Vasa and the germ line lineage in a colonial urochordate, Dev. Biol. 331 (2009) 113-128.

[36] M. Kowarsky, C. Anselmi, K. Hotta, P. Burighel, G. Zaniolo, F. Caicci, B. Rosental, N.F. Neff, K.J. Ishizuka, K.J. Palmeri, J. Okamoto, T. Gordon, I.L. Weissman, S.R. Quake, L. Manni, A. Voskoboynik, Sexual and asexual development: two distinct programs producing the same tunicate, Cell Rep. 34 (2021) 108681.

[37] A.R. Holyoak, Morphogenetic movements and assembly of strobilae into zooidal systems in early colony development of the compound Ascidian Polyclinum planum, Biol. Bull. 183 (1992) 432-439.

[38] N. Tinbergen. The Study of Instinct, Clarendon Press/Oxford University Press, 1951.

[39] A.H. Brush, H.M. Reisman, The carotenoid pigments in the three-spined stickleback, Gasterosteus aculeatus, Comp. Biochem. Physiol. 14 (1965) 121-125.
[40] J. Cronly-Dillon, S.C. Sharma, Effect of season and sex on the photopic spectral sensitivity of the three-spined stickleback, J. Exp. Biol. 49 (1968) 679-687.

[41] W.J. Rowland, C.L. Baube, T.T. Horan, Signalling of sexual receptivity by pigmentation pattern in female sticklebacks, Anim. Behav. 42 (1991) 243-249.

[42] U. Candolin, Changes in expression and honesty of sexual signalling over the reproductive lifetime of sticklebacks, Proc. Biol. Sci. 267 (2000) 2425-2430.

[43] R.A. Cloney, Ascidian larvae and the events of metamorphosis, Am. Zool. 22 (1982) 817-826.

[44] R.A. Cloney, Larval tunic and the function of the test cells in ascidians, Acta Zool. 71 (1990) 151-159.

[45] Y. Sasakura, A. Hozumi, Formation of adult organs through metamorphosis in ascidians, Dev. Biol. 7 (2018) e304.

[46] A. Nakayama, N. Satoh, Y. Sasakura, Tissue-specific profile of DNA replication in the swimming larvae of Ciona intestinalis, Zoolog. Sci. 22 (2005) 301-309.

[47] J.-P. Chambon, J. Soule, P. Pomies, P. Fort, A. Sahuquet, D. Alexandre, P.-H. Mangeat, S. Baghdiguian, Tail regression in Ciona intestinalis (Prochordate) involves a Caspase-dependent apoptosis event associated with ERK activation, Development 129 (2002) 3105-3114.

[48] A. Jacqmard, I. Gadisseur, G. Bernier, Cell division and morphological changes in the shoot apex of Arabidopsis thaliana during floral transition, Ann. Bot. 91 (2003) 571-576.

[49] D. Kwiatkowska, Flowering and apical meristem growth dynamics, J. Exp. Bot. 59 (2008) 187-201.

[50] F. Fornara, A. de Montaigu, G. Coupland, SnapShot: control of flowering in Arabidopsis, Cell 141 550 (550) (2010) e1, 2.

[51] G. Bernier, The control of floral evaporation and morphogenesis, Annu. Rev. Plant Physiol. Plant Mol. Biol. 39 (1988) 175-219.

[52] A.V. Klepikova, M.D. Logacheva, S.E. Dmitriev, A.A. Penin, RNA-seq analysis of an apical meristem time series reveals a critical point in Arabidopsis thaliana flower initiation, BMC Genom. 16 (2015) 466.

[53] K.F. Chater, S. Biró, K.J. Lee, T. Palmer, H. Schrempf, The complex extracellular biology of Streptomyces, FEMS Microbiol. Rev. 34 (2010) 171-198.

[54] S.G. Kang, K.J. Lee, Kinetic analysis of morphological differentiation and protease production in Streptomyces albidoflavus SMF301, Microbiol. Read. Engl. 143 (1997) 2709-2714.

[55] A. Manteca, D. Claessen, C. Lopez-Iglesias, J. Sanchez, Aerial hyphae in surface cultures of Streptomyces lividans and Streptomyces coelicolor originate from viable segments surviving an early programmed cell death event, FEMS Microbiol. Lett. 274 (2007) 118-125.

[56] E.M. Migüeléz, C. Hardisson, M.B. Manzanal, Hyphal death during colony development in Streptomyces antibioticus: morphological evidence for the existence of a process of cell deletion in a multicellular prokaryote, J. Cell Biol. 145 (1999) 515-525.

[57] H. Wildermuth, Development and organization of the aerial mycelium in Streptomyces coelicolor, J. Gen. Microbiol. 60 (1970) 43-50.

[58] J.W. Costerton, P.S. Stewart, E.P. Greenberg, Bacterial biofilms: a common cause of persistent infections, Science 284 (1999) 1318-1322.

[59] H.-C. Flemming, J. Wingender, U. Szewzyk, P. Steinberg, S.A. Rice, S. Kjelleberg, Biofilms: an emergent form of bacterial life, Nat. Rev. Microbiol. 14 (2016) 563-575.

[60] H.-C. Flemming, T.R. Neu, D.J. Wozniak, The EPS matrix: the "house of biofilm cells.", J. Bacteriol. 189 (2007) 7945-7947.

[61] D. López, H. Vlamakis, R. Kolter, Biofilms, Cold Spring Harbor Perspect. Biol. 2 (2010) a000398.

[62] G. O'Toole, H.B. Kaplan, R. Kolter, Biofilm formation as microbial development, Annu. Rev. Microbiol. 54 (2000) 49-79.

[63] D. López, H. Vlamakis, R. Kolter, Generation of multiple cell types in Bacillus subtilis, FEMS Microbiol. Rev. 33 (2009) 152-163.

[64] V.L. Marlow, F.R. Cianfanelli, M. Porter, L.S. Cairns, J.K. Dale, N.R. Stanley-Wall, The prevalence and origin of exoprotease-producing cells in the Bacillus subtilis biofilm, Microbiol. Read. Engl. 160 (2014) 56-66.

[65] J.-W. Veening, W.K. Smits, O.P. Kuipers, Bistability, epigenetics, and bet-hedging in bacteria, Annu. Rev. Microbiol. 62 (2008) 193-210.

[66] D.B. Kearns, R. Losick, Cell population heterogeneity during growth of Bacillus subtilis, Genes Dev. 19 (2005) 3083-3094.

[67] K.M. Blair, L. Turner, J.T. Winkelman, H.C. Berg, D.B. Kearns, A molecular clutch disables flagella in the Bacillus subtilis biofilm, Science 320 (2008) 1636-1638.

[68] S.S. Branda, J.E. González-Pastor, S. Ben-Yehuda, R. Losick, R. Kolter, Fruiting body formation by Bacillus subtilis, Proc. Natl. Acad. Sci. USA. 98 (2001) 11621-11626.

[69] M.M. Nakano, R. Magnuson, A. Myers, J. Curry, A.D. Grossman, P. Zuber, srfA is an operon required for surfactin production, competence development, and efficient sporulation in Bacillus subtilis, J. Bacteriol. 173 (1991) 1770-1778.

[70] M.M. Nakano, L.A. Xia, P. Zuber, Transcription initiation region of the srfA operon, which is controlled by the comP-comA signal transduction system in Bacillus subtilis, J. Bacteriol. 173 (1991) 5487-5493.

[71] D. López, M.A. Fischbach, F. Chu, R. Losick, R. Kolter, Structurally diverse natural products that cause potassium leakage trigger multicellularity in Bacillus subtilis, Proc. Natl. Acad. Sci. USA. 106 (2009) 280-285.

[72] D. López, H. Vlamakis, R. Losick, R. Kolter, Paracrine signaling in a bacterium, Genes Dev. 23 (2009) 1631-1638.

[73] J. Dworkin, R. Losick, Developmental commitment in a bacterium, Cell 121 (2005) 401-409.

[74] P. Eichenberger, M. Fujita, S.T. Jensen, E.M. Conlon, D.Z. Rudner, S.T. Wang, C. Ferguson, K. Haga, T. Sato, J.S. Liu, R. Losick, The program of gene transcription for a single differentiating cell type during sporulation in Bacillus subtilis, PLoS Biol. 2 (2004) e328.

[75] L. Kroos, The Bacillus and Myxococcus developmental networks and their transcriptional regulators, Annu. Rev. Genet. 41 (2007) 13-39.

[76] P.J. Piggot, D.W. Hilbert, Sporulation of Bacillus subtilis, Curr. Opin. Microbiol. 7 (2004) 579-586.

[77] R.D. Monds, G.A. O'Toole, The developmental model of microbial biofilms: ten years of a paradigm up for review, Trends Microbiol. 17 (2009) 73-87.

[78] P.S. Stewart, M.J. Franklin, Physiological heterogeneity in biofilms, Nat. Rev. Microbiol. 6 (2008) 199-210.

[79] H. Zheng, J. Kim, M. Liew, J.K. Yan, O. Herrera, J.W. Bok, N.L. Kelleher, N.P. Keller, Y. Wang, Redox metabolites signal polymicrobial biofilm development via the NapA oxidative stress cascade in Aspergillus, Curr. Biol. 25 (2015) 29-37.

[80] A.S. Blix, Adaptations to polar life in mammals and birds, J. Exp. Biol. 219 (2016) 1093-1105.

[81] M. Zimova, K. Hackländer, J.M. Good, J. Melo-Ferreira, P.C. Alves, L.S. Mills, Function and underlying mechanisms of seasonal colour moulting in mammals and birds: what keeps them changing in a warming world? Biol. Rev. 93 (2018) 1478-1498.

[82] G. Prota, The chemistry of melanins and melanogenesis, in: J.B. Barrera, H.J.M. Gijsen, A.G. González, Ae de Groot, G. Prota, J.B.P.A. Wijnberg, W. Herz, G. W. Kirby, R.E. Moore, W. Steglich, Ch Tamm (Eds.), Progress in the Chemistry of Organic Natural Products, Springer, Vienna, 1995, pp. 93-148.

[83] A. Slominski, D.J. Tobin, S. Shibahara, J. Wortsman, Melanin pigmentation in mammalian skin and its hormonal regulation, Physiol. Rev. 84 (2004) 1155-1228.

[84] C.-F. Chen, J. Foley, P.-C. Tang, A. Li, T.X. Jiang, P. Wu, R.B. Widelitz, C.M. Chuong, Development, regeneration, and evolution of feathers, Annu. Rev. Anim. Biosci. 3 (2015) 169-195.

[85] R.B. Payne, Chapter 3 - mechanisms and control of molt, in: D.S. Farner, J.R. King (Eds.), Avian Biology, Academic Press, Amsterdam, 1972, pp. 103-155.

[86] E.O. Höhn, C.E. Braun, Hormonal induction of feather pigmentation in parrmigan, The Auk 97 (1980) 601-607.

[87] T. Yoshimura, Thyroid hormone and seasonal regulation of reproduction, Front. Neuroendocrinol. 34 (2013) 157-166.

[88] C.-M. Chuong, V.A. Randall, R.B. Widelitz, P. Wu, T.-X. Jiang, Physiological regeneration of skin appendages and implications for regenerative medicine, Physiology 27 (2012) 61-72.

[89] I. Mayer, B. Borg, R. Schulz, Seasonal changes in and effect of castration/androgen replacement on the plasma levels of five androgens in the male three-spined stickleback, Gasterosteus aculeatus L., Gen. Comp. Endocrinol. 79 (1990) 23-30.

[90] Z. Li, Y. He, Roles of brassinosteroids in plant reproduction, Int. J. Mol. Sci. 21 (2020) 872.

[91] G. Vert, J.L. Nemhauser, N. Geldner, F. Hong, J. Chory, Molecular mechanisms of steroid hormone signaling in plants, Annu. Rev. Cell Dev. Biol. 21 (2005) 177-201.

[92] Z. Li, Y. Ou, Z. Zhang, J. Li, Y. He, Brassinosteroid signaling recruits histone 3 lysine-27 demethylation activity to FLOWERING LOCUS C chromatin to inhibit the floral transition in Arabidopsis, Mol. Plant 11 (2018) 1135-1146.

[93] Q. Ye, W. Zhu, L. Li, S. Zhang, Y. Yin, H. Ma, X. Wang, Brassinosteroids control male fertility by regulating the expression of key genes involved in Arabidopsis anther and pollen development, Proc. Natl. Acad. Sci. USA. 107 (2010) 6100-6105.

[94] L. Keller, M.G. Surette, Communication in bacteria: an ecological and evolutionary perspective, Nat. Rev. Microbiol. 4 (2006) 249-258.
[95] C.M. Waters, B.L. Bassler, Quorum sensing: cell-to-cell communication in bacteria, Annu. Rev. Cell Dev. Biol. 21 (2005) 319–346.

[96] D.G. Davies, M.R. Parsek, J.P. Pearson, B.H. Iglewski, J.W. Costerton, E.P. Greenberg, The involvement of cell-to-cell signals in the development of a bacterial biofilm, Science 280 (1998) 295–298.

[97] J.W. Costerton, Z. Lewandowski, D.E. Caldwell, D.R. Korber, H.M. Lappin-Scott, Microbial biofilms, Annu. Rev. Microbiol. 49 (1995) 711–745.

[98] T. Danhorn, C. Fuqua, Biofilm formation by plant-associated bacteria, Annu. Rev. Microbiol. 61 (2007) 401–422.

[99] H.-C. Flemming, J. Wingender, The biofilm matrix, Nat. Rev. Microbiol. 8 (2010) 623–633.

[100] T.F. Mah, G.A. O'Toole, Mechanisms of biofilm resistance to antimicrobial agents, Trends Microbiol. 9 (2001) 34–39.

[101] M.R. Parsek, E.P. Greenberg, Sociomicrobiology: the connections between quorum sensing and biofilms, Trends Microbiol. 13 (2005) 27–33.

[102] R. Lami, Quorum Sensing in Marine Biofilms and Environments, 2019, p. 55.

[103] P. Stoodley, K. Sauer, D.G. Davies, J.W. Costerton, Biofilms as complex differentiated communities, Annu. Rev. Microbiol. 56 (2002) 187–209.

[104] D.L. Milton, Quorum sensing in vibrios: complexity for diversification, Int. J. Med. Microbiol. 296 (2006) 61–71.

[105] M. Urvoy, R. Lami, C. Dreanno, D. Daudé, M.A.S. Rodrigues, M. Gourmelon, S. L'Helguen, C. Labry, Quorum sensing disruption regulates hydrolytic enzyme and biofilm production in estuarine bacteria, Environ. Microbiol. 23 (2021) 7183–7200.

[106] M. Colasanti, G. Venturini, Nitric oxide in invertebrates, Mol. Neurobiol. 17 (1998) 157–174.

[107] A. Palumbo, Nitric oxide in marine invertebrates: a comparative perspective, Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 142 (2005) 241–248. Papers From: Nitric Oxide: Comparative Aspects of Respiratory and Cardiovascular Homeostasis, an SEB symposium, organised by Prof. Bruno Tota (University of Calabria, Italy) and Prof. Tobias Wang (University of Aarhus, Denmark) - Capri, Italy, September 2004.

[108] I. Heckler, E.M. Boon, Insights into nitric oxide modulated quorum sensing pathways, Front. Microbiol. 10 (2019).

[109] S. Honma, S. Ito, S. Yajima, Y. Sasaki, Nitric oxide signaling for aerial mycelium formation in Streptomyces coelicolor A3(2) M145, Appl. Environ. Microbiol. 88 (2022) e01222, 22.

[110] K.A. Gallagher, M.A. Schumacher, M.J. Bush, M.J. Bibb, G. Chandra, N.A. Holmes, W. Zeng, M. Henderson, H. Zhang, K.C. Findlay, R.G. Brennan, M.J. Buttner, c-di-GMP arms an anti-σ to control progression of multicellular differentiation in streptomyces, Mol. Cell. 77 (2020) 586–599.

[111] N. Tschowri, M.A. Schumacher, S. Schlimpert, N. babu Chinnam, K.C. Findlay, R.G. Brennan, M.J. Buttner, Tetrameric c-di-GMP mediates effective transcription factor dimerization to control Streptomyces development, Cell 158 (2014) 1136–1147.

[112] S. Jeandroz, D. Wipf, D.J. Stuehl, R. Lamattina, M. Melkonian, Z. Tian, Y. Zhu, E.J. Carpenter, G.K.-S. Wong, D. Wendehenne, Occurrence, structure, and evolution of nitric oxide synthase-like proteins in the plant kingdom, Sci. Signal. 9 (2016) re2.

[113] E. Planchet, K. Jagadis Gupta, M. Sonoda, D. M.A.S. Rodrigues, Nitric oxide emission from tobacco leaves and cell suspensions: rate limiting factors and evidence for the involvement of mitochondrial electron transport, Plant J. Cell Mol. Biol. 41 (2005) 732–743.

[114] Y. He, R.-H. Tang, Y. Hao, R.D. Stevens, C.W. Cook, S.M. Ahn, L. Jing, Z. Yang, L. Chen, F. Guo, F. Fiorani, R.B. Jackson, N.M. Crawford, Z.-M. Pei, Nitric oxide represses the Arabidopsis floral transition, Science 305 (2004) 1968–1971.

[115] G.G. Simpson, NO flowering, Bioessays 27 (2005) 239–241.

[116] A. Heyland, L.L. Moroz, Signaling mechanisms underlying metamorphic transitions in animals, Integr. Comp. Biol. 46 (2006) 743–759.

[117] H.-T. Chung, H.-O. Pae, B.-M. Choi, T.R. Billiar, Y.-M. Kim, Nitric oxide as a bioregulator of apoptosis, Biochem. Biophys. Res. Commun. 282 (2001) 1075–1079.

[118] W.R. Jeffery, Programmed cell death in the ascidian embryo: modulation by FoxA5 and Manx and roles in the evolution of larval development, Mech. Dev. 118 (2002) 111–124.

[119] C.D. Bishop, W.R. Bates, B.P. Brandhorst, Regulation of metamorphosis in ascidians involves NO/cGMP signaling and HSP90, J. Exp. Zool. 289 (2001) 374–384.

[120] S. Comes, A. Locascio, F. Silvestre, M. d'Ischia, G.L. Russo, E. Tosti, M. Branno, A. Palumbo, Regulatory roles of nitric oxide during larval development and metamorphosis in Ciona intestinalis, Dev. Biol. 306 (2007) 772–784.

[121] N. Ueda, S.M. Degnan, Nitric oxide acts as a positive regulator to induce metamorphosis of the ascidian herdmania momus, PLoS One 8 (2013) e72797.

[122] M. Dacher, M. Gauthier, Involvement of NO-synthase and nicotinic receptors in learning in the honey bee, Physiol. Behav. 95 (2008) 200–207.

[123] M. Cikala, B. Wilm, E. Hobmayer, A. Böttger, C.N. David, Identification of caspases and apoptosis in the simple metazoan Hydra, Curr. Biol. 9 (1999) 959–962.

[124] N.A. Thornberry, Y. Lazebnik, Caspases: enemies within, Science 281 (1998) 1312–1316.

[125] J.-P. Chambon, A. Nakayama, K. Takamura, A. McDougall, N. Satoh, ERK- and JNK-signalling regulate gene networks that stimulate metamorphosis and apoptosis in tail tissues of ascidian tadpoles, Development 134 (2007) 1203–1219.

[126] Y.-F. Chen, S.-J. Shin, S.-R. Lin, Ets1 was significantly activated by ERK1/2 in mutant K-ras stably transfected human adrenocortical cells, DNA Cell Biol. 24 (2005) 126–132.

[127] Y. Deng, X. Ren, L. Yang, Y. Lin, X. Wu, A JNK-dependent pathway is required for TNFalpha-induced apoptosis, Cell 115 (2003) 61–70.

[128] A. Lin, B. Dibling, The true face of JNK activation in apoptosis, Aging Cell 1 (2002) 112–116.

[129] M. Gionti, F. Ristoratore, A. Di Gregorio, F. Aniello, M. Branno, R. Di Lauro, Cihox5, a new Ciona intestinalis/Hox-related gene, is involved in regionalization of the spinal cord, Dev. Gene. Evol. 207 (1998) 515–523.

[130] T. Ikuta, N. Satoh, H. Saiga, Limited functions of HOx genes in the larval development of the ascidian Ciona intestinalis, Development 137 (2010) 1505–1513.

[131] M.W. Jacobs, S.M. Degnan, R. Woods, E. Williams, K.E. Roper, K. Green, B.M. Degnan, The effect of larval age on morphology and gene expression during ascidian metamorphosis, Integr. Comp. Biol. 46 (2006) 760–776.

[132] T. Ikuta, N. Yoshida, N. Satoh, H. Saiga, E.H. Davidson, Ciona intestinalis Hox gene cluster: its dispersed structure and residual colinear expression in development, Proc. Natl. Acad. Sci. USA. 101 (2004) 15118–15123.

[133] M. Primon, K.D. Hunter, H.S. Pandha, R. Morgan, Kinase regulation of HOX transcription factors, Cancers 11 (2019) 508.

[134] A. Wilmerding, L. Bouteille, L. Rinaldi, N. Caruso, Y. Graba, M.-C. Delfini, HOXB8 counteracts MAPK/ERK oncogenic signaling in a chicken embryo model of neoplasia, Int. J. Mol. Sci. 22 (2021) 8911.

[135] G. Komis, O. Šamajová, M. Ovečka, J. Šamaj, Cell and developmental biology of plant mitogen-activated protein kinases, Annu. Rev. Plant Biol. 69 (2018) 237–265.

[136] K.L. Johnson, S. Ramm, C. Kappel, S. Ward, O. Leyser, T. Sakamoto, T. Kurata, M.W. Bevan, M. Lenhard, The tinkerbell (tink) mutation identifies the dual-specificity MAPK phosphatase INDOLE-3-BUTYRIC ACID-RESPONSE5 (IBR5) as a novel regulator of organ size in Arabidopsis, PLoS One 10 (2015) e0131103.

[137] E. Guttes, S. Guttes, H.P. Rusch, Morphological observations on growth and differentiation of Physarum polycephalum grown in pure culture, Dev. Biol. 3 (1961) 588–614.

[138] A. Chapman, J.G. Coote, Sporulation competence in Physarum polycephalum CL and the requirement for DNA replication and mitosis, J. Gen. Microbiol. 128 (1982) 1489–1501.

[139] R. Kroneder, A.R. Cashmore, W. Marwan, Phytochrome-induced expression of lig1, a homologue of the fission yeast cell-cycle checkpoint gene hus1, is associated with the developmental switch in Physarum polycephalum plasmodia, Curr. Genet. 36 (1999) 86–93.

[140] T. Nakagaki, S. Umemura, Y. Kakiuchi, T. Ueda, Action spectrum for sporulation and photoavoidance in the plasmodium of Physarum polycephalum, as modified differentially by temperature and starvation, Photochem. Photobiol. 64 (1996) 859–862.

[141] H.W. Sauer, K.L. Babcock, H.P. Rusch, Sporulation in Physarum polycephalum: a model system for studies on differentiation, Exp. Cell Res. 57 (1969) 319–327.

[142] C. Starostzik, W. Marwan, Kinetic analysis of a signal-transduction pathway by time-resolved somatic complementation of mutants, J. Exp. Biol. 201 (1998) 1991–1999.

[143] C. Starostzik, W. Marwan, Functional mapping of the branched signal transduction pathway that controls sporulation in Physarum polycephalum, Photochem. Photobiol. 62 (1995) 930–933.

[144] G. Golderer, E.R. Werner, S. Leitner, P. Gröbner, G. Werner-Felmayer, Nitric oxide synthase is induced in sporulation of Physarum polycephalum, Genes Dev. 15 (2001) 1299–1309.
[145] R. Martel, A. Tessier, D. Pallotta, G. Lemieux, Selective gene expression during sporulation of Physarum polycephalum, J. Bacteriol. 170 (1988) 4784-4790.

[146] J. Bailey, Building a plasmoidum: development in the acellular slime mould Physarum polycephalum, Bioessays 19 (1997) 985-992.

[147] I. Barrantes, G. Glockner, S. Meyer, W. Marwan, Transcriptomic changes arising during light-induced sporulation in Physarum polycephalum, BMC Genom. 11 (2010) 115.

[148] S. Jin, Z. Nasim, H. Susila, J.H. Ahn, Evolution and functional diversification of FLOWERING LOCUS T/TERMINAL FLOWER 1 family genes in plants. Semin. Cell Dev. Biol. 109 (2021) 20-30.

[149] L. Corbesier, C. Vincent, S. Jang, F. Fornara, Q. Fan, I. Searle, A. Giakountis, S. Farrona, L. Gissot, C. Turnbull, G. Coupland, FT protein movement contributes to long-distance signaling in floral induction of Arabidopsis, Science 316 (2007) 1030-1033.

[150] H. Takagi, A.K. Hempton, T. Imaizumi, Photoperiodic flowering in Arabidopsis: multilayered regulatory mechanisms of CONSTANS and the florigen FLOWERING LOCUS T, Plant Commun. 4 (2023) 100552.

[151] D.D. Brown, L. Cai, Amphibian metamorphosis, Dev. Biol. 306 (2007) 20-33.

[152] C.C. Helbing, K. Werry, D. Crump, D. Domanski, N. Veldhoen, C.M. Edwards, Expression profiles of novel thyroid hormone-responsive genes and proteins in the tail of Xenopus laevis tadpoles undergoing precocious metamorphosis, Mol. Endocrinol. Baltim. 17 (2003) 1395-1409.

[153] M.L. Wright, Melatonin, diel rhythms, and metamorphosis in anuran amphibians, Gen. Comp. Endocrinol. 126 (2002) 251-254.

[154] M.L. Edwards, E.B. Pivorun, The effects of photoperiod and different dosages of melatonin on metamorphic rate and weight gain in Xenopus laevis tadpoles, Gen. Comp. Endocrinol. 81 (1991) 28-38.

[155] V.B. Eichler, Jr.L.S. Gray, The influence of environmental lighting on the growth and prometamorphic development of larval Rana pipiens, Dev. Growth Differ. 18 (1976) 177-182.

[156] M.L. Wright, S.T. Jorey, Y.M. Myers, M.L. Fieldstad, C.M. Paquette, M.B. Clark, Influence of photoperiod, daylength, and feeding schedule on tadpole growth and development, Dev. Growth Differ. 30 (1988) 315-323.

[157] H.W. Korf, A. Oksche, 4 - The pineal organ, in: P.K.T. Pang, M.P. Schreibman (Eds.), Morphological Considerations, Academic Press, 1986, pp. 105-145.

[158] M.L. Wright, K.L. Cuthbert, M.J. Donohue, S.D. Solano, K.L. Proctor, Direct influence of melatonin on the thyroid and comparison with prolactin, J. Exp. Zool. 286 (2000) 625-631.

[159] M.L. Wright, A. Pikula, L.J. Cykowski, K. Kuliga, Effect of melatonin on the Anuran thyroid gland: follicle cell proliferation, morphometry, and subsequent thyroid hormone secretionin vitroafter melatonin treatmentin vivo, Gen. Comp. Endocrinol. 103 (1996) 182-191.

[160] B. Capel, Vertebrate sex determination: evolutionary plasticity of a fundamental switch, Nat. Rev. Genet. 18 (2017) 675-689.

[161] Y. Nagahama, T. Chakraborty, B. Paul-Prasanth, K. Ohta, M. Nakamura, Sex determination, gonadal sex differentiation, and plasticity in vertebrate species, Physiol. Rev. 101 (2021) 1237-1308.

[162] M. Stöck, L. Kratochvíl, H. Kuhl, M. Rovatsos, B.J. Evans, A. Suh, N. Valenzuela, F. Veyrunes, Q. Zhou, T. Gamble, B. Capel, M. Schartl, Y. Guiguen, A brief review of vertebrate sex evolution with a pledge for integrative research: towards ‘sexomics.’, Philos. Trans. R. Soc. B Biol. Sci. 376 (2021) 20200426.

[163] Y.S. De Mitcheson, M. Liu, Functional hermaphroditism in teleosts, Fish Fish. 9 (2008) 1-43.

[164] T. Kuwamura, T. Sunobe, Y. Sakai, T. Kadota, K. Sawada, Hermaphroditism in fishes: an annotated list of species, phylogeny, and mating system, Ichthyol. Res. 67 (2020) 341-360.

[165] S. Pla, F. Maynou, F. Piferrier, Hermaphroditism in fish: incidence, distribution and associations with abiotic environmental factors, Rev. Fish Biol. Fish. 31 (2021) 935-955.

[166] R.C. Francis, K. Soma, R.D. Fernald, Social regulation of the brain-pituitary-gonadal axis, Proc. Natl. Acad. Sci. 90 (1993) 7794-7798.

[167] C.M.St Mary, Sex allocation in a simultaneous hermaphrodite, the blue-banded goby (Lythrypnus dalli): the effects of body size and behavioral gender and the consequences for reproduction, Behav. Ecol. 5 (1994) 304-313.

[168] L. Fishelson, Protogynous sex reversal in the fish anthias squamipinnis (teleostei, anthiidae) regulated by the presence or absence of a male fish, Nature 227 (1970) 90-91.

[169] R.H. Reavis, M.S. Grober, An integrative approach to sex change: social, behavioural and neurochemical changes in Lythrypnus dalli (Pisces), Acta Ethol. 2 (1999) 51-60.

[170] D.R. Robertson, Social control of sex reversal in a coral-reef fish, Science 177 (1972) 1007-1009.

[171] D.Y. Shapiro, Intragroup behavioural changes and the initiation of sex reversal in a coral reef fish in the laboratory, Anim. Behav. 29 (1981) 1199-1212.

[172] D.Y. Shapiro, Size, maturation and the social control of sex reversal in the coral reef fish Anthias squamipinnis, J. Zool. 193 (1981) 105-128.

[173] T. Chakraborty, Y. Yamamoto, S. Hanai, M. Hirano, S. Mohapatra, A. Yamaguchi, T. Takeda, M. Matsuyama, K. Ohta, Divulging the social sex change mechanism in a unique model system for studying the sexual plasticity of protogynous hermaphrodite fish, three bamboo leaf wrasse (Pseudolabrus sieboldi), Front. Mar. Sci. 9 (2022).

[174] M. Liu, Y. Sadovy, Early gonadal development and primary males in the protogynous epinepheline, Cephalopholis boenak, J. Fish. Biol. 65 (2004) 987-1002.

[175] D. Policansky, Sex change in plants and animals, Annu. Rev. Ecol. Systemat. 13 (1982) 471-495.

[176] W.A. Müller, T. Leitz, Metamorphosis in the Cnidaria, Can. J. Zool. 80 (2002) 1755-1771.

[177] C.C.C.R. de Carvalho, Marine biofilms: a successful microbial strategy with economic implications, Front. Mar. Sci. 5 (2018).

[178] M.G. Hadfield, Biofilms and marine invertebrate larvae: what bacteria produce that larvae use to choose settlement sites, Ann. Rev. Mar. Sci 3 (2011) 453-470.

[179] J.D. Zardus, B.T. Nedved, Y. Huang, C. Tran, M.G. Hadfield, Microbial biofilms facilitate adhesion in biofouling invertebrates, Biol. Bull. 214 (2008) 91-98.

[180] A.M. Cantley, J. Clardy, Animals in a bacterial world: opportunities for chemical ecology, Nat. Prod. Rep. 32 (2015) 888-892.

[181] T. Wichard, C. Beemelmanns, Role of chemical mediators in aquatic interactions across the prokaryote-eukaryote boundary, J. Chem. Ecol. 44 (2018) 1008-1021.

[182] A. Woznica, N. King, Lessons from simple marine models on the bacterial regulation of eukaryotic development, Curr. Opin. Microbiol. 43 (2018) 108-116.

[183] J.M. Sneed, K.H. Sharp, K.B. Ritchie, V.J. Paul, The chemical cue tetrabromopyrrole from a biofilm bacterium induces settlement of multiple Caribbean corals, Proc. R. Soc. B Biol. Sci. 281 (2014) 20133086.

[184] J. Tebben, D.M. Tapiolas, C.A. Motti, D. Abrego, A.P. Negri, L.L. Blackall, P.D. Steinberg, T. Harder, Induction of larval metamorphosis of the coral Acropora millepora by tetrabromopyrrole isolated from a Pseudoalteromonas bacterium, PLoS One 6 (2011) e19082.

[185] C.F. Ericson, F. Eisenstein, J.M. Medeiros, K.E. Malter, G.S. Cavalcanti, R.W. Zeller, D.K. Newman, M. Pilhofer, N.J. Shikuma, A contractile injection system stimulates tubeworm metamorphosis by translocating a proteinaceous effector, Elife 8 (2019) e46845.

[186] N.J. Shikuma, I. Antoshechkin, J.M. Medeiros, M. Pilhofer, D.K. Newman, Stepwise metamorphosis of the tubeworm Hydroides elegans is mediated by a bacterial inducer and MAPK signaling, Proc. Natl. Acad. Sci. 113 (2016) 10097-10102.

[187] N.J. Shikuma, M. Pilhofer, G.L. Weiss, M.G. Hadfield, G.J. Jensen, D.K. Newman, Marine tubeworm metamorphosis induced by arrays of bacterial phage tail-like structures, Science 343 (2014) 529-533.

[188] S. Kelly, J.T. Sullivan, Y. Kawaharada, S. Radutoiu, C.W. Ronson, J. Stougaard, Regulation of Nod factor biosynthesis by alternative NodD proteins at distinct stages of symbiosis provides additional compatibility scrutiny, Environ. Microbiol. 20 (2018) 97-110.

[189] S. Bensmihen, F. de Billy, C. Gough, Contribution of NFP LysM domains to the recognition of Nod factors during the Medicago truncatula/Sinorhizobium meliloti symbiosis, PLoS One 6 (2011) e26114.

[190] D. Khokhani, C. Carrera Carriel, S. Vayla, T.B. Irving, C. Stonoha-Arther, N.P. Keller, J.-M. Ané, Deciphering the chitin code in plant symbiosis, defense, and microbial networks, Annu. Rev. Microbiol. 75 (2021) 583-607.

[191] G.E.D. Oldroyd, J.D. Murray, P.S. Poole, J.A. Downe, The rules of engagement in the legume-rhizobial symbiosis, Annu. Rev. Genet. 45 (2011) 119-144.

[192] F. Xie, J.D. Murray, J. Kim, A.B. Heckmann, A. Edwards, G.E.D. Oldroyd, J.A. Downie, Legume pectate lyase required for root infection by rhizobia, Proc. Natl. Acad. Sci. USA. 109 (2012) 633-638.

[193] K.M. Jones, H. Kobayashi, B.W. Davies, M.E. Taga, G.C. Walker, How rhizobial symbionts invade plants: the Sinorhizobium-Medicago model, Nat. Rev. Microbiol. 5 (2007) 619-633.
[194] A. Kereszt, P. Mergaert, J. Montiel, G. Endre, É. Kondorosi, Impact of plant peptides on symbiotic nodule development and functioning, Front. Plant Sci. 9 (2018) 1026.

[195] C.R. Schopfer, M.E. Nasrallah, J.B. Nasrallah, The male determinant of self-incompatibility in Brassica, Science 286 (1999) 1697-1700.

[196] P. Mergaert, T. Uchiumi, B. Alunni, G. Evanno, A. Cheron, O. Catrice, A.-E. Mausset, F. Barloy-Hubler, F. Galibert, A. Kondorosi, E. Kondorosi, Eukaryotic control on bacterial cell cycle and differentiation in the Rhizobium-legume symbiosis, Proc. Natl. Acad. Sci. USA. 103 (2006) 5230-5235.

[197] A.F. Haag, M. Baloban, M. Sani, B. Kerscher, O. Pierre, A. Farkas, R. Longhi, E. Boncompagni, D. Hérouart, S. Dall'Angelo, E. Kondorosi, M. Zanda, P. Mergaert, G.P. Ferguson, Protection of sinorhizobium against host cysteine-rich antimicrobial peptides is critical for symbiosis, PLoS Biol. 9 (2011) e1001169.

[198] A.C. Love, Evolution and Development: Conceptual Issues, Cambridge University Press, 2024.

[199] G.B. Müller, Evo-devo: extending the evolutionary synthesis, Nat. Rev. Genet. 8 (2007) 943-949.

[200] W.A. Cresko, K.L. McGuigan, P.C. Phillips, J.H. Postlethwait, Studies of threespine stickleback developmental evolution: progress and promise, Genetica 129 (2007) 105-126.

[201] C.L. Peichel, D.A. Marques, The genetic and molecular architecture of phenotypic diversity in sticklebacks, Phil Trans R Soc B 372 (2017) 20150486.

[202] D.A. McLennan, Phylogenetic relationships in the gasterosteidae: an updated tree based on behavioral characters with a discussion of homoplasy, Copeia 1993 (1993) 318-326.

[203] D.A. McLennan, Integrating phylogenetic and experimental analyses: the evolution of male and female nuptial coloration in the stickleback fishes (gasterosteidae), Syst. Biol. 45 (1996) 261-277.

[204] J.W. Boughman, Divergent sexual selection enhances reproductive isolation in sticklebacks, Nature 411 (2001) 944-948.

[205] R.J. Scott, Assortative mating between adjacent populations of threespine stickleback (Gasterosteus aculeatus), Ecol. Freshw. Fish 13 (2004) 1-7.

[206] A. Casinos, From Cuénot's préadaptation to Gould and Vrba's exaptation: a review, Biol. J. Linn. Soc. 121 (2017) 239-247.

[207] S.J. Gould, E.S. Vrba, Exaptation-A missing term in the science of form, Paleobiology 8 (1982) 4-15.

[208] V. Laudet, The origins and evolution of vertebrate metamorphosis, Curr. Biol. 2 (2011) R726-R737.

[209] K. Kuida, D.M. Boucher, Functions of MAP kinases: insights from gene-targeting studies, J. Biochem. 135 (2004) 653-656.

[210] S. Torii, K. Nakayama, T. Yamamoto, E. Nishida, Regulatory mechanisms and function of ERK MAP kinases, J. Biochem. 136 (2004) 557-561.

[211] I. Sharina, E. Martin, Cellular factors that shape the activity or function of nitric oxide-stimulated soluble guanylyl cyclase, Cells 12 (2023) 471.

[212] M. Shepherd, D. Giordano, C. Verde, R.K. Poole, The evolution of nitric oxide function: from reactivity in the prebiotic earth to examples of biological roles and therapeutic applications, Antioxidants 11 (2022) 1222.

[213] G.F. Pauli, J.B. Friesen, T. Gödecke, N.R. Farnsworth, B. Glodny, Occurrence of progesterone and related animal steroids in two higher plants, J. Nat. Prod. 73 (2010) 338-345.