After fertilization, we redefined 26 stages for embryogenesis of A. aspersa (Supplementary Figure S1; Table 1) based on the staging definition used for Ciona (Hotta et al., 2007). The developmental stages describe the process from fertilization to hatching in cases without chorion (Figure 1). Table 1 shows the time required to reach each developmental stage at 20°C as well as tail and trunk length ratios (T/H ratio) and time ratio to hatch (% hatch). The hatching time (avg. 16 h and 6 min post fertilization) was examined by time-lapse imaging of the embryos with the chorion. Embryogenesis was separated into six periods (Table 1): periods of zygote (Figure 2), cleavage (Figures 2, 3), gastrula (Figure 4), neurula (Figure 5), tailbud (Figures 6, 7), and larva (Figure 8).

The criteria for staging up to Stage 10 depends on the embryo shape and the number of cells. The later criteria are based primarily on the shape of the embryo and the length of the tail and trunk (after Stage 15) as well as angular tail bend according to Hotta’s criteria (Hotta et al., 2007).

One of staging criteria was the “standard development time” (in hours)—the normalized time for development when incubated at 20°C. Previously reported optimal incubation temperatures range from 16°C to 20°C for Ciona staging (Hotta et al., 2007). The hatching time after fertilization is slightly later than Ciona (16.1 hpf and 15.6 hpf at 20°C in A. aspersa and Ciona, respectively; Supplementary Figure S2).

The zygote period (0–1.2 h post-fertilization at 20°C) was composed of one stage: Stage 1 extends from fertilization to the end of the first mitotic cell division (Figure 2). Figures 2B,B′ (optical section) show the first cell division in progress. The boundary between the two daughter cells is constricted but not yet fully separated. The egg diameter is relatively larger (175.4 ± 3.3 μm, N = 10) than that of Ciona (145.1 ± 1.8 μm, N = 10).

The cleavage period (1.2–4.4 h post-fertilization at 20°C) was composed of eight stages: Stages 2–9 (Figures 2C–3G″). The cleavage period is characterized by the formation of blastomeres through sequential mitosis. At these stages, the embryo is bilaterally symmetrical. In ascidians, unequal cell division (UCD) forms two small cells at the 16-cell stage followed by two successive UCDs. From the 16-cell stage, the mitotic cycle becomes asynchronous with the vegetative half dividing before the animal half producing the 24-, 32-, 44-, and 64-cell stages (McDougall et al., 2019). The criteria for staging during the cleavage period are not focused only on the number of cells but also on the embryo shape. During the early cleavage period (e.g., 16- and 32-cell stages) embryos change shape dynamically and quickly due to compaction. Thus, the 16-cell stage and 32-cell stage were divided into two sub-stages (16-cell stage: Stage 5a and Stage 5b; 32-cell stage: Stage 6a and Stage 6b, respectively).

Gastrulation is a nearly universally conserved developmental process in animal embryogenesis during which dramatic morphological changes occur. During the seventh cell division, gastrulation begins when the large endoderm cells on the vegetal side of the embryo invaginate (Swalla, 1993). Ascidian gastrulation is initiated by the invagination of 10 endodermal precursor cells between the 64-cell stage and the late 112-cell stage. This process is driven in two steps by a myosin-dependent contraction of the actomyosin network in the absence of endodermal cell division (Sherrard et al., 2010; Winkley et al., 2019; Fiuza et al., 2020).

Neurulation occurs during the neurula period. This is one of the defining events of chordate morphogenesis in which the neural tube forms and separates from a surface epidermis to form the rudiment of the future nervous system. The neurula period (Figures 5A–C′′) includes Stages 14–16.

Stages 17–25 is defined as the tailbud period (Figures 6, 7; Table 1; Supplementary Figures S2) and is subdivided into four phases: initial tailbud, early tailbud, mid tailbud, and late tailbud stages. All tailbud stages, besides late tailbud, include two sub-stages representing the start and end of the stage. In every case, there is a clear visual marker at the start of the stage.

Images of A. aspersa embryos were exported as a series of image files so that both cross-sectional and 3D images could be easily viewed. These were visible via a web-browser. Images were linked to information on developmental stage, developmental nomenclature, hour post-fertilization (hpf), % hatch, cell lineage, and time-lapse movies. We integrated these data into a web-based database, “RAMNe.” The user can observe the 3D images interactively and for each z-section image online (Figure 9; https://www.bpni.bio.keio.ac.jp/RAMNe/latest/index.html). When selecting any developmental stage from the developmental table (Figure 9A), users can easily view 3D- (Figure 9C) and Z-section (Figure 9B) images of the entire embryo at that stage. In the “z-section module,” users can display all the section images between the selected focus ranges using a slider (Figure 9B). The “rotation module” allows one to view the 3D image of the embryo rotated along the Y-axis in 1° angle steps. One of the unique features of the RAMNe is that corresponding images from Ciona can also be displayed (Hotta et al., 2007, Hotta et al., 2020). This can then be interactively compared with A. aspersa (Figure 9D). In the download section, users can freely download resource files under the terms of the Creative Commons Attribution License (CC BY). All pages are adaptive for viewing on various devices and are free to access anytime; anyone can freely access the ascidian morphology resource.

Although Ciona and A. aspersa shared stereotyped development with conserved cleavage patterns and developmental timing, our detailed observation shows few differences between them. A. aspersa has a relatively shorter tail and larger trunk with Ciona. This indicates different body shapes in some stages. For example, the embryo was ovular in A. aspersa at Stage 14 (Figure 5A; Supplementary Figure S1) but diamond-shaped in Ciona. The initial tailbud embryos from Stage 17—Ciona have an hourglass-like epithelial bending between trunk and tail regions. This is similar to “KUBIRE” in Japanese (Nakamoto and Kumano, 2020) whereas no KUBIRE is seen in A. aspersa (7.9 h, Figures 6A,A′, 10). This KUBIRE shape was made by different orientation of cell division in the transition between trunk and tail. To understand how such different body shapes are created, it will be interesting to investigate the orientation of cell division responsible for making KUBIRE in A. aspersa. However, the ratio of the trunk and the tail length in the tailbud period was smaller than that of Ciona indicating a relatively short tail (Figure 11B). At Stage 22 and Stage 26, the tail length is 1.9 and 4.2 times of the trunk, respectively (Hotta et al., 2007). These values are 1.8 and 3.0 times, respectively, in A. aspersa.

The two species in the tailbud stages show tail curving in the tailbud embryo, but the timing of the relaxation of the tail bending is relatively earlier than that of Ciona (Figure 11A), and the maximum curvature of A. aspersa in tailbud stages is weaker than that of Ciona (Figure 6; see RAMNe using dual view). Tailbud embryo tail bending is controlled by asymmetric actomyosin localization in notochord at early tailbud stages (Lu et al., 2020); Admp regulates the tail bending (Kogure et al., 2021, submitted). The differences in the tail bending curvature among the two species might be regulated by the different spatio-temporal expression of these molecules.

Some morphological differences are also observed between the two species in the larva period. The ventral papilla in A. aspersa is located more ventral than Ciona (Figures 10O,O′). Three papillae in Ciona form an equilateral triangle from the frontal view. On the other hand, A. aspersa forms an isosceles triangle (Figures 10P,P′). A. aspersa larva have around 30 lateral epidermal cells [fewer than Ciona (Figures 10O,O′)]. This lower number of tail epidermis cells might reduce the surface area of the tail and lead to a shorter tail in A. aspersa. In A.aspersa, two atrial siphon primordia are located in the dorsal trunk, but they are located in the lateral trunk in Ciona (Figures 10Q,Q′).

The short tail in A.aspersa compared to Ciona may reflect divergence of ascidian larva morphology and ecology. Short tails related to the reduction of the surface area of the tail can lead to a decline in the tadpole swimming (Van Buskirk and Mccollum, 2000). Upward and downward swimming with optical and gravity sensors can offer important insight into habitat and selection of substratum (Kajiwara and Yoshida, 1985; Bostwick et al., 2020): Weak swimming predicts poor dispersion despite the considerable mobility and invasiveness of this species (Lynch et al., 2016). However, short tail ascidian may have advantages in terms of metamorphosis speed as in no-tail ascidian, Molgula (Fodor et al., 2021). Investigations of the divergence in larval morphology can encourage a better understand of ecology-evolution-development studies in ascidians.

The egg diameter of A. aspersa is relatively larger (175.4 ± 3.3 μm, N = 10) than that of Ciona (145.1 ± 1.8 μm, N = 10). The timing of cleavage (∼Stage 12) and subsequent morphogenesis, gastrulation, neurulation, tail elongation, and the hatching time of A. aspersa after fertilization were nearly the same as Ciona (Supplementary Figure S2) if incubated at the same temperature. The growth curves of the two species are also very similar at the same temperature (20°C; Supplementary Figure S2) confirming that the developmental stages of both species correspond to each other. The ratio of each period was also conserved among both species. A previous dwarf embryo experiment was produced by artificial egg size reduction (Matsumura et al., 2020) and indicated that ascidian developmental speed was robust regardless of embryonic size. At least two Ascidian species have an evolutionarily robust mechanism for development speed regardless of the embryo size.

A. aspersa has a big advantage in bioimaging research because of its extremely transparent embryo (Shito et al., 2020). This feature is unique to Ascidians of the family Ascidiidae such as Phallsuia and A. aspersa embryos, which are not found in Ciona and can help visualize embryos in three dimensions (Supplementary Movie S2). Moreover, remarkable feature of A. aspersa is the ability to do quick translation of external mRNA in the unfertilized egg (Yasuo and McDougall, 2018). To demonstrate this, we show the Ca²⁺ elevation at the fertilization by using genetic Ca²⁺ sensor, GCaMP6s (Supplementary Movie S3). As previously reported in Ciona savignyi and A. aspersa by using chemical Ca²⁺ sensor (Yoshida et al., 1998; Levasseur and McDougall, 2000), two series of Ca²⁺ oscillations were also observed in A. aspersa (Supplementary Figure S3). This demonstrates usefulness as a model animal for bioimaging and the potential benefits of using a variety of genetic sensors to visualize biological events early in their development.

However, there are still several points to note. Unlike Ciona, eggs are considerably more sensitive when dechorionated; fertilization is more difficult, and they are considerably more seasonal. The reason for the sensitivity in fertilization may be contamination of highly acidic body fluids. Isolation of gametes should involve gonoduct flushing with ASW to remove acidic fluids as shown by Bolton and Havenhand (Bolton and Havenhand, 1996). A. aspersa is continuous breeder year-round, but individuals with mature gonads are most abundant in the winter and spring and lowest in the summer (Lynch et al., 2016). Improved methods of handling and egg availability will promote bioimaging research with A. aspersa.

Our precise description of anatomy and developmental staging for A. aspersa in this study, along with the ontology for anatomy and development, leads to a standard developmental table of A. aspersa for the scientific community including evo-devo, developmental biology, ecology, and cell biology. The embryos imaged in this study did not have chorion for the clear observation but the dechorionation was known to cause a mild effect on the morphogenesis (Oonuma et al., 2016; Kourakis et al., 2021). Thus, we expect chorionated specimens of A. aspersa development in subsequent versions of our RAMNe database.

Our embryo-imaging resource has value in creating a 3D standard model based on the real stack images for quantitative approach (Robin et al., 2011a; Nakamura et al., 2012; Veeman and Reeves, 2015; Matsumura et al., 2020). Ciona and Phallusia have good developmental 3D image resources that are very useful to understand morphometrical information of each embryo/blastomere (e.g., FABA (https://www.bpni.bio.keio.ac.jp/chordate/faba/1.4/top.html), TunicAnatO (https://www.bpni.bio.keio.ac.jp/tunicanato/3.0/index.html), and ANISEED database (https://www.aniseed.cnrs.fr/) (Hotta et al., 2007; Brozovic et al., 2016; Brozovic et al., 2018; Hotta et al., 2020).

The morphology in the stages before tailbud of A. aspersa is similar to that one of Ciona. Guignard suggested considerable embryonic reproducibility with early embryonic cell lineages conserved between distantly related ascidian species may play a role in contact area-dependent cell inductions during animal embryogenesis instead of morphogen gradients. These may relax constraints on genome evolution in ascidians (Guignard et al., 2020). Resource data of multiple ascidian embryo morphology can verify these hypotheses and provide valuable insight into the mechanism of deep conservation of morphology as well as the evolution of system-level morphogenesis in embryonic development.

We hope that the developmental staging and anatomy of A. aspersa via RAMNe will facilitate the use of this fascinating animal as a model animal for SLUMEICA.

A. aspersa adults were obtained from Onagawa Field Center or Hakodate Fisheries Research Institute. Eggs and sperm were isolated from the gonoducts. Chorion surrounding eggs were removed enzymatically according to a previous protocol (Hotta et al., 1999). Dechorionation of eggs was performed with the solution 0.05% actinase-E and 1% mercaptoacetic acid sodium salt in ASW as in Ciona (Hotta et al., 1999). The time of the dechorionation was relatively longer and took 10–20 min. Eggs and embryos were incubated with Millipore-filtered seawater (MFSW) on the 0.1% gelatin-coated dishes. Embryos were incubated at 20°C after fertilization.

A Peltier-based thermo-stage (TOKAI-Hit) was placed on the microscope stage (OLYMPUS SZX16); the temperature was stabilized and maintained. The embryo was placed on a thermal plate and maintained during observation to stabilize the temperature at 20°C to acquire images. Images were acquired every one or 3 min.

We fixed embryos every 10–30 min from fertilized eggs to hatching larva stage. The method of the fixation was described previously (Hotta et al., 2007). Distinct representative embryos in each stage were chosen based on the Ciona staging criteria (Hotta et al., 2007) from CLSM data (Figure 1). We used an 40× oil-immersion objective lens (NA: 0.75) and set the confocal aperture to 140 μm. The size of the images was set to 1,024 px × 1,024 px. The size of the pixels was 0.129–0.31 μm/pixel in X and Y directions. The scanning interval in the z-axis direction was set to 1 μm. Cortical actin filaments was stained by using Alexa 546 phalloidin. All 3D images were reconstructed from around 100 sectioned images.

The same method of visualization of Ca²⁺ for Ciona embryo was used (Akahoshi et al., 2017). In brief, the pSPE3- GCaMP6s plasmid was linearized using SfiIor DraIII, and GCaMP6s mRNA was produced and precipitated using the mMESSAGE mMACHINE T3 kit (Life Technologies, Carlsbad, CA, United States) following the manufacturer’s protocol. GCaMP6s mRNA was injected into dechorionated A. aspersa eggs at 0.5 μg/μl. The mRNA-injected egg was incubated in a 3-cm glass base dish at 20°C for more than 3 hours. Spontaneous fertilization was observed under fluorescence microscopy after adding one drop of sperm.

FM4-64 (Invitrogen) was used to visualize plasma membranes. Embryos were incubated in seawater with the final concentration of FM4-64 10 μM. Under CLSM, Laser excitation at 559 nm was used to visualize the signals in FM4-64. The size of the images was set to 512 px × 512 px. The size of the pixels was 0.564 μm/pixel in X and Y directions. The scanning interval in the z-axis direction was set to 4 μm. The z-stack consists of 36 images was taken every 2 min.