Embryonic Development in the Peppermint Shrimp, Lysmata boggessi (Caridea: Lysmatidae).
Among marine invertebrates, decapod crustaceans (Arthropoda: Crustacea: Multicrustacea: Malacostraca: Eumalacostraca: Eucarida: Decapoda) are recognized for their astonishing anatomical, ecological, physiological, and behavioral diversity (Martin and Davis, 2001; Ng et al., 2008; Schwentner et al., 2017). This impressive extant diversity likely implies the existence of equally diverse modes of development, including early embryogenesis (Martin and Davis, 2001 ; Scholtz and Wolff, 2013). Although developmental modes and their diversity in some decapods are relatively well known, the cellular and molecular mechanisms of early embryogenesis in most decapods remain poorly explored. Widely available tools for transgenesis and genome editing are available only for other non-decapod crustaceans such as the branchiopod Daphnia magna and the amphipod Parhyale hawaiensis, which also have sequenced genomes (Browne et al., 2005; Mittmann et al., 2014; Kao et al., 2016; Martin et al., 2016; Stamataki and Pavlopoulos, 2016; Ye et al., 2017). Within Decapoda, only a few species have a sequenced genome (e.g., the astacidean Procambarus virginalis, Gutekunst et al., 2018).
In this study, we are interested in the embryonic development of shrimps belonging to the infraorder Caridea, a species-rich clade of decapods (De Grave et al., 2015). For this purpose, we used the peppermint shrimp, Lysmata boggessi Rhyne & Lin, 2006, as a model species. Modes of development in the Decapoda vary grossly, from indirect to direct. At one extreme of this continuum, penaeid shrimps (Decapoda: Dendrobranchiata) do not provide any form of parental care, spawning small fertilized eggs into the pelagic environment that develop into a small lecithotrophic nauplius larva (Martin et al., 2014; Hertzler, 2015). This nauplius larva can use the first and second pair of antennae and mandibles for swimming. The nauplius also bears a simple medial and frontal non-ommatidial eye (Gurney, 1942). At the other extreme, some species with direct development produce large, yolky eggs that are brooded and hatch as juveniles (e.g., in some freshwater crabs [Brachyura] and crayfish [Astacidea], Thiel, 2003; Martin et al., 2014). In between these extremes, many species exhibit indirect development, and parental care is restricted to the protection of embryos incubated in bodily chambers of varying complexity, as in various clades within the decapod suborder Pleocyemata, including stenopodidean and caridean shrimps (Bauer and Newman, 2004). In the decapod suborder Caridea, the naupliar stage is "embryonized" and gives rise to a zoea larva that swims efficiently after hatching by using well-developed thoracic appendages (i.e., the maxillipeds) (Gurney, 1942; Martin et al., 2014).
Similarly, early embryogenesis in the Decapoda is also variable. The patterns of early cleavage can be holoblastic or incomplete (Weygoldt, 1961: Anderson, 1973; Scholtz and Wolff, 2013; Wolff and Gerberding, 2015). In some species (e.g., in the Caridea), blastomeres progressively become superficial and surround an acellular yolk, forming a superficial blastoderm from which a germ disk develops (Anderson, 1973: Dohle et al., 2004; Browne et al., 2005; Klann and Scholtz, 2014; Hertzler, 2015). In Dendrobranchiata, cleavage remains holoblastic, and no blastoderm forms. Whether the embryo forms a blastoderm or not, gastrulation mechanisms will invariantly be the result of various events of cell involution, delamination, and cell invasion that result in the correct positioning of the endoderm, mesoderm, and germ cells (Anderson, 1973; Price et al., 2010; Klann and Scholtz, 2014; Haug and Haug, 2015).
Caridean shrimps belonging to the genus Lysmata Risso, 1816 exhibit a wide diversity of lifestyles, mating systems, symbiotic partnerships, and colorations (Baeza et al., 2009). All species studied so far are protandric simultaneous hermaphrodites (Baeza et al., 2013). In these species, juveniles develop into males (or "male-phase" shrimps) with typical caridean external male morphology (Bauer, 2000). These male shrimps reproduce as males; however, the gonads are ovotestes with undeveloped ovaries. When shrimps reach larger body sizes, the ovarian portion of the ovotestes matures, and shrimps turn into functional simultaneous hermaphrodites capable of reproducing as both male and female (Bauer and Holt, 1998; Bauer and Newman, 2004; Baeza and Anker, 2008).
Shrimps belonging to the genus Lysmata offer opportunities for experimentation and genetic manipulation, considering that they are widely traded species in the ornamental aquarium industry, and various rearing protocols have been developed in recent years (Wabnitz et al., 2003; Calado et al., 2005; Figueiredo and Narciso, 2006). While a recent study in Lysmata amboinensis described generalities of embryo external morphology (Zhang et al., 2013), next to nothing is known about cellular behavior and morphogenesis during development in Lysmata. In addition, there is no consensus on a staging method within the Caridea (and other decapods), and the pattern of post-naupliar embryonic development is poorly known (Nazari et al., 2000; Muller et al., 2003, 2004; Garcia-Guerrero and Hendrickx, 2009; Habashy et al., 2012; Romney and Reiber, 2013; Zhang et al., 2013). From studies in Caridina multidentata, Crangon crangon, Atyephyra compresa, and various species from the genus Alpheus, early cleavage is known to vary from holoblastic to incomplete (Brooks and Herrick, 1891; Weygoldt, 1961; Anderson, 1973; Klann and Scholtz, 2014). The blastomeres ultimately form a blastoderm and a germ disk, from which the embryonic nauplius stage develops (Ishikawa, 1885; Weldon, 1887; Brooks and Herrick, 1891; Nair, 1949; Klann and Scholtz, 2014). Additional studies are needed to determine whether a general pattern of development occurs in the Caridea. The present study provides a baseline methodology for future comparative studies focusing on the developmental biology of caridean shrimps and other decapod crustaceans.
Materials and Methods
Maintenance and reproduction in Lysmata boggessi
Individuals of Lysmata boggessi Rhyne & Lin, 2006 were collected between June and July 2016 from Florida Bay, Florida, by K&P Aquatics (Tavernier, FL). Collected specimens were placed in large plastic bags containing aerated seawater and were shipped overnight to the laboratory at Clemson University, Clemson, South Carolina. Individuals were maintained in 40-1 glass aquaria containing artificial seawater (Instant Ocean. Blacksburg, VA), with a temperature of 24 [degrees]C, ~35 ppt salinity, and a light:dark cycle of 14 h:10 h. Shrimps were fed daily with shrimp pellets (Wardley, Secaucus, NJ).
From the available pool of shrimps, we selected "parturial" hermaphrodites: prespawning hermaphrodites with a gonad full of green mature oocytes that were likely to molt in the next (1-3) days. These parturial hermaphrodites were then paired with either a male or a second non-parturial (male-role) hermaphrodite and transferred to 40-1 glass aquaria (one pair per aquarium). In shrimps belonging to the genus Lysmata, including L. boggessi, copulation between sexual partners is not reciprocal. A newly molted prespawning hermaphrodite (parturial hermaphrodite) copulates as a female with a male or with another hermaphrodite acting exclusively as a male at that time. During copulation, which lasts a few seconds, a sperm mass is attached by the mating partner to the underside of the pre-spawning hermaphrodite. There is no long-term sperm storage in the genus Lysmata, and sperm from a mating is used to fertilize only the eggs released during the spawning, which occurs two or three hours later (Bauer and Holt, 1998).
Embryos were periodically removed from a total of 18 brooding hermaphrodites (see Results), using fine forceps, and kept in plastic petri dishes with seawater for direct observation under the stereoscope (Leica MZFLIII with a Leica MC170 HD camera. Wetzlar, Germany) while alive; and later they underwent fixation.
Fixation and nuclear staining of embryos in Lysmata boggessi
After extraction, embryos were initially fixed in seawater + 3.7% formaldehyde for 2 min before removing their chorions with the aid of fine forceps. Embryos with or without the inner membrane were then fixed in 3.7% formaldehyde in 1X phosphate-buffered saline (PBS) and stored at 4 [degrees]C until further use. For nuclear staining, fixed embryos were washed twice in 1X PBS and five times in PBS + 0.1% Triton-X (PBST). Next, embryos were incubated for 3 h in 4',6-diamidino-2-phenylindole (DAPI, 3.5 [micro]g [ml.sup.-1] in PBST). Whole embryos were mounted on glass slides for observation under a fluorescence microscope (Nikon TE2000. with a Nikon DS1 monochrome camera, Minato. Tokyo, Japan). For clearing and deep imaging, embryos were washed in 1X PBS and distilled water, then progressively dehydrated in ethanol. and finally transferred to methyl salicylate. Maximum projection and batch image processing were done using Fiji software (Schindelin et al., 2012). Image processing and figure assembly were done using Adobe Photoshop (Adobe Systems, Mountain View, CA).
Shrimp breeding and fertilization
From an initial pool of 40 shrimps, we examined embryos in a total of 18 successful fertilization events in the course of one month. In our laboratory, copulation and fertilization occurred only during the dark cycle. The time of fertilization, the time of egg harvesting, the time of fixation, and the time of hatching of the remaining eggs into the first larval stage (St.) were used to calculate developmental times, shown in Table 1. At 24 [degrees]C, Lysmata boggessi embryos completed their development in ~12 d, when first zoea larvae hatch. Herein, we propose a total of 17 stages during embryogenesis that depict major developmental landmarks, from egg fertilization to hatching of the first zoea larva (Table 1).
Early cleavage and blastoderm formation
Fertilized eggs are light green to the naked eye. Eggs are ellipsoid in shape, with a major axis average length of 0.58 mm [+ or -] 0.01 mm (SD) and a minor axis average width of 0.39 mm [+ or -] 0.01 mm (SD) (St. 1; Fig. 1A, D, G). The ends of the long axis will henceforth be referred to as embryo poles. After fertilization and zygote formation, mitosis occurs in the center of the yolk (Fig. 1C). After chromosome segregation, the plane of the first cleavage consistently occurs parallel to the minor axis and results in two equally sized blastomeres (St. 2; Fig. 1D-F). DAPI-positive superficial bodies are often spotted in variable positions along the surface of the fertilized egg, without any clear correlation between the latter and the first cleavage furrow. These elements could be polar bodies (Fig. 1B, C). During the first cell division, chromosomal segregation occurs centrally in the yolk (Fig. 1C, F). The second cell division forms a furrow perpendicular to the first plane and occurs slightly delayed on each of the two blastomeres. Before the third cell division, a partial rearrangement of the blastomeres is observed, resulting in two of the blastomeres relocating into a medial position, while the other two blastomeres relocate toward the poles (black arrows, St. 3; Fig. 1G, H). From the four-cell stage until the late blastoderm stage, cell divisions become superficial, and nuclei can be seen as white yolk-less spots under transmitted light (Figs. 1 H, 2A-H). At the eight-cell stage, it is difficult to distinguish stereotypical blastomere positions (Figs. 2A, A1). Indeed, an analysis of division planes in the eight-cell embryo showed a lack of parallel mitotic spindles between neighboring blastomeres (Figs. 2B; A1). We were also unable to detect stable cross-furrows (not shown). These results could suggest a lack of a stereotypical division pattern in this species.
The cellular blastoderm forms in the following cell divisions. Early blastoderm cell divisions occur from 10 to 24 h after fertilization, when the embryo goes from 16 cells to ~100 cells. By analyzing cell number, we estimate that there is a round of cell divisions every ~3.2 h (St. 5; Fig. 2C-F). By the end of this stage, mitoses become asynchronous, with visible differences in the progress of chromosome segregation (Fig. 2F). The late blastoderm stage is characterized by a reduction in the rate of cell proliferation, estimated from the total cell number (St. 6; Fig. 2G, H). During this stage, the embryo will add only about 200 cells to the blastoderm within the next 13 h, from 24 to 37 h postfertilization.
Gastrulation and germ disk formation
Gastrulation starts approximately 37 h after fertilization, when the embryo reaches ~300 cells (St. 7; Fig. 3A). At this point, cell membranes become less obvious (Fig. 3). The main cellular behavior observed during early gastrulation is the invasion of the internal yolk mass by an undetermined number of cells (Fig. 3A'). In most embryos, the site of delamination and yolk invasion occurred at one of the embryo poles and coincided with the place of future cellular aggregation and germ disk condensation (Fig. 3A', C). Nonetheless, during early gastrulation, we observed some embryos with cellular aggregates in variable positions across the blastoderm surface that did not necessarily match one of the poles. These sites of cell aggregation also showed underlying yolk invasion (Fig. 3B, B'). It is unknown whether these sites of cell ingression were blastopore-forming sites. During the late gastrula stage, however, most embryos consistently accumulated cells in one of the poles, where a small blastopore will form (St. 8; Fig. 3C). Invading cells might or might not be visible during this stage. Three days after fertilization, a small germ disk starts to form consistently at one of the embryo poles at the site of the blastopore (St. 9; Fig. 3D).
"Embryonized" nauplius stage
The egg-naupliar stage of L. boggessi is observed ~5 days after fertilization and is characterized by the presence of obvious head lobes, antennula (first antenna), antenna (second antenna), mandible primordia, and a caudal papilla (St. 10; Fig. 4A). In some embryos, we observed transient biramous antennules (Fig. 4A). Later, embryos will show biramous antennae, while antennules become uniramous (St. 10+; Fig. 4B). Other visible structures are the stomodeum and the labrum, located in the central portion of the naupliar embryo. The caudal papilla shows teloblastic growth as ectoteloblasts form conspicuous concentric rows (Fig. 4).
We have divided post-naupliar development into five stages, according to the presence and/or absence of appendages and the level of appendage, pleon, and eye differentiation. In other caridean shrimps such as Macrobrachium olfersii, Macrobrachium americanum, and Palaemonetes pugio, the first post-naupliar stage features head lobes, mandibles, maxillules, and maxillar primordia, but no maxilliped primordia (Table 1). We did not observe this first post-naupliar stage (St. 11) in any of the 18 egg clutches analyzed, possibly due to a short duration or a failure in our sampling method. Alternatively, the first post-naupliar stage may not be present in L. boggessi. In any case, we have kept this stage in Table 1 to ensure its applicability in other species.
The second post-naupliar stage embryo is comma shaped (curved), as a result of the outgrowth and ventral folding of the thoracopleonal process, derived from the caudal papilla (St. 12; Fig. 5A-A"). These embryos have large head lobes, with important retinal differentiation (Fig. 5A'). A distinctive feature of this stage is the 90[degrees] angle between the lateral protocerebrum and the eye growth zone, from which the optic stalk primordium will form in later stages. The stomodeum is also clearly visible, and there is an initial differentiation of the mandibules, maxillules, maxillae, and maxilliped primordia. The telson reaches the level of the maxillae (Fig. 5A', A").
The third post-naupliar stage bears a clearly visible labrum and a bipartite telson; both structures are located close to each other due to the sustained growth of the pleon. The optic stalk, attached to the lateral protocerebrum, can now be distinguished from the retinal growth zone (St. 13; Fig. 5B). At this point during development, the thoracic maxillipeds are obvious and continue growing ventrally (Fig. 5B'). Ommatidial differentiation is also detected at this stage by the presence of protoomatidial cell arrangements in the retina (Fig. 5B"). The fourth post-naupliar stage is characterized by additional development and growth of the maxillipeds that now cover the pleon and are segmented (St. 14; Fig. 5C, C). During this stage, further differentiation of the maxillipeds occurs, each now featuring an exopod and an endopod. The edge of the carapace is also discernible near the base of the maxillipeds.
In the fifth post-naupliar stage, yolk is distributed through about 80%-90% of the cephalothorax, and the pleon is 0.3-0.5 times the length of the cephalothorax (St. 15; Fig. 6A). Red chromatophores and initial eye pigmentation are also clearly observed externally. At this stage, the eyes are the most differentiated structures, showing a growth zone distinct from the optic stalk and ample ommatidial differentiation in the retina (Fig. 6A'). The limb outgrowth results in the folding of the maxillipeds that now cover the attached pleon (Fig. 6A").
The pre-zoeal stage is characterized by further reduction in yolk volume, now located dorsally and distributed through about 70%-80% of the cephalothorax. The pleon is as long as the cephalothorax (1.0-1.1 times) and shows sporadic movements. Dorsally, the optic stalk protrudes between the compound eyes and the carapace, to which the eyes are no longer attached (Fig. 6B'). At this stage, the maxillipeds are pigmented (St. 16; Fig. 6B, B"), and the telson reaches the space between the eyes (Fig. 6B").
Immediately after hatching, the first free-living zoea larval stage is characterized by three pairs of well-developed thoracic biramous maxillipeds located immediately posterior to the cephalic maxillary appendages (St. 17; Fig. 7A). Yolk is distributed through about 10%-20% of the cephalothorax. We observed neither first pereopod primordia nor a rostrum in the zoea larva after hatching. The pleon, 1.3-1.4 times longer than the cephalothorax, consists of six somites and lacks appendages (Fig. 7B). The fifth pleonal segment is particularly elongated and bears a telson with uropods. During this zoea stage, the differentiated eyes are not pedunculated but project away from the cephalothorax and carapace (Fig. 7C). Cephalic appendages show advanced differentiation. Antennulae and antennae are now central to the eyes, and the mandibulae show incisive processes, with further accessory teeth differentiation (Fig. 7D).
Comparative analyses of ontogenetic characters bear deep implications for the phylogenetic analysis and hypothesis of evolutionary change (Scholtz et al., 2009). In caridean shrimps, studies describing embryo development are limited and focus on only a few families: Alpheidae, Palaemonidae, and Atyidae (Ishikawa, 1885; Weldon, 1887; Brooks and Herrick, 1891; Nair, 1949; Weygoldt, 1961; Jalihal et al., 1993; Nazari et al., 2000; Habashy et al., 2012; Romney and Reiber, 2013: Klann and Scholtz, 2014). Only a single previous study has examined embryonic development in the family Lysmatidae (in Lysmata amboinesis, Zhang et al., 2013). In the present study, we focus on morphological aspects of embryonic development in Lysmata boggessi and propose a staging system to be used in the Caridea and other decapods. This is a tool for comparative analyses in the Caridea and Decapoda, in general, as we discuss below (Table 1).
Our results show that egg size, appearance, yolk distribution, and developmental rates in L. boggessi are very similar to those described for other caridean species (Fig. 1A-D; Table 1). The pattern of early cleavage shown for L. boggessi has been thoroughly described and reviewed for other caridean species (Ishikawa, 1885; Weldon, 1887; Brooks and Herrick, 1891; Anderson, 1973). In most caridean shrimps, including Crangon crangon, Caridina laevis, Paratya compressa, and Synalpheus minus, radial cleavage is maintained only until the third cell division, when blastomeres rearrange, as shown in Figure 2 (Ishikawa, 1885; Weldon, 1887; Brooks and Herrick, 1891; Nair, 1949). In other carideans, a high degree of plasticity in the direction of the cleavage planes has been described. For example, the first cleavage furrow in Hippolyte zostericola can form across the long egg axis or the minor egg axis or in a 45[degrees] angle (Gorham, 1895). To compare, in Caridina multidentata, plasticity was reported only on the second plane of cleavage, as a result of the rotation of the mitotic spindle (Nair, 1949; Klann and Scholtz, 2014). The mechanisms that cause non-teratogenic plasticity in the direction of cleavage are far from being understood and might be influenced by phylogenetic history, mechanical constraints imposed by the distribution of yolk (centrolecithal or isolecithal), and/or environmental conditions (Rappaport, 1986; Black and Vincent, 1988; Schulze and Schierenberg, 2011).
The process of formation of an acellular yolk mass is also poorly known and is apparently variable in caridean shrimps. In C. multidentata, holoblastic cell divisions continue well past the seventh cleavage, as shown by the formation of round blastomeres with less cellular contact with the yolk. In this species, acellular yolk is progressively deposited in the center of the embryo until the blastoderm is formed (Klann and Scholtz, 2014). To compare, in Palaemon varions, early cleavage forms incomplete furrows, and cellularization occurs later during blastomere formation (Weygoldt, 1961). This latter pattern, with high variability in the deepness of the incomplete furrows, has also been described in various alpheid carideans and in the anomuran Emerita talpoida (Brooks and Herrick, 1891). In L. boggessi, similar to other alpheids, blastomeres remain tightly attached to each other from cleavage to blastoderm formation, which could imply that L. boggessi also forms incomplete furrows after the third cell division; however, further analyses are still required to confirm that notion.
Cleavage in the Caridea invariably results in the formation of a blastoderm, a common feature in decapods with a centralized yolk distribution (centrolecithic) (Anderson, 1973; Arendt and Nubler-Jung, 1999; Scholtz and Wolff, 2013). In L. boggessi, we have divided the process of blastoderm formation into two stages that are distinguishable by the number of cells. During the early stage, the number of blastomeres increases to about 100 cells at 24 hours after fertilization. During the late stage, from 24 to 37 hours, cell divisions become asynchronous, as demonstrated by the limited change in cell number during the second day of development and the differences in cell cycle progression (Fig. 2F). Accordingly, the embryo adds only 200 cells during these 13 hours. These changes in proliferation and localized cell cycle arrest have also been described for Caridina multidentata and Crangon crangon (Weldon, 1887; Klann and Scholtz, 2014). Such changes in proliferation rate and synchrony could be associated with the maternal-zygotic transition, during which the embryo genome starts to be transcribed (Langley et al., 2014). These changes occur at the 32-cell stage in the amphipod Parhyale hawaiensis (Nestorov et al., 2013). Zygotic genome activation remains to be explored in L. boggessi.
Apart from changes in cell division rates, cell location and cleavage stereotypy during later cleavage stages might have significant consequences for cell fate determination in carideans. In C. multidentata, dividing blastomeres form two interlocking bands of clonally related cells that have the same mitotic spindle orientation (Klann and Scholtz, 2014). This particular arrangement of blastomeres is known as a stereotypical pattern of cell division. In several species of dendrobranchiate shrimps and the euphausiacean Meganyctiphanes norvegica, early cleavage stereotypy is associated with early cell fate determination, because each interlocking band limits movement and establishes, early in development, the future position and fate of the blastomeres (Hertzler and Clark, 1992; Alwes and Scholtz, 2004; Hertzler, 2005; Biffis et al., 2009: Pawlak et al., 2010; Scholtz and Wolff, 2013; Klann and Scholtz, 2014). A prehminary analysis of cleavage planes and the location of blastomeres at the eight-cell stage of L. boggessi showed that blastomere distribution varies across the long axis, and neighboring blastomeres do not necessarily have parallel mitotic spindles (Fig. A1). This lack of parallelism might also be associated with plasticity in the position of blastomeres and further cell rearrangement (Fig. A1). Compared to the pattern observed in C. multidentata, our results imply that there is diversity in the pattern of early cleavage in the Caridea; however, cell fate determination has not been demonstrated for either C. multidentata or L. boggessi. Lineage tracing using clones with similar mitotic spindle orientation is much more complex in C. multidentata and L. boggessi. given that gastrulation only starts after a large number of cells have been added to the blastoderm (Klann and Scholtz, 2014). In addition, the changes in proliferation rate, observed in both species, might mask the identification of clonal cells and interlocking bands when the blastoderm reaches more than 300 cells (Fig. 2F). Therefore, in vivo lineage tracing and functional analysis are needed to determine the mechanism of cell fate determination in the Caridea.
As for blastoderm formation, gastrulation in L. boggessi also follows a pattern that appears to be conserved in the Caridea. Internal yolk invasion by "wandering cells" has been reported before in every species of caridean shrimp in which gastrulation has been studied (Brooks and Herrick, 1891; Nair, 1949; Anderson, 1973; Klann and Scholtz. 2014). These invasive cells have been described as vitellophages, with some function in the formation of the midgut (Brooks and Herrick. 1891; Anderson, 1973). In Synalpheus minus, vitellophages are formed concomitantly with the formation of the germ disk (Brooks and Herrick, 1891). However, in other carideans, such as C. crangon and L. boggessi, yolk cell invasion occurs earlier (Weldon, 1887). Because of the limited availability of in vivo tracking systems, the ultimate role of this process in caridean shrimps is a matter of discussion (Anderson, 1973; Wolff and Gerberding, 2015). In carideans that form a blastoderm, the mesoderm and endoderm rise by subsequent cell ingression and delamination from a small blastopore located at one of the ends of the longest embryo axis. The germ disk also forms in this site (Nair, 1949; Oishi, 1959; Weygoldt, 1961; Anderson, 1973: Dohle et al., 2004; Klann and Scholtz, 2014). In contrast, in the Dendrobranchiata, gastrulation occurs concomitantly with early cleavage. Because of the invariant cell fate in the Dendrobranchiata, a pair of future mesendodermal cells undergoes arrest, ingresses to a yolk-free blastocoel, and organizes following gastrulation movements during later stages. These movements result in the internalization of the naupliar mesoderm and endoderm and in the localization of teloblasts in the most posterior part of the nauplius larva (Hertzler and Clark, 1992; Hertzler, 2002, 2005; Alwes and Scholtz, 2004; Biffis et al., 2009).
Distinctive eye and limb development patterns were documented during the naupliar and post-naupliar development of L. boggessi. After the egg-nauplius stage, L. boggessi embryos have large head lobes and show ommatidial differentiation during the third post-naupliar stage (Fig. 5B). In other Pleocyemata, such as Cherax destructor, retinal differentiation is visible only after the differentiation of the maxillipeds and pereopods (Sandeman and Sandeman, 1991). Similarly, the first zoea larvae of L. boggessi lack pereopods (Table 1; Fig. 7). The congeneric Lysmata amboinensis also lacks pereopods at hatching (Zhang et al., 2013). The genus Lysmata is fully marine. To compare, carideans adapted to freshwater habitats, such as C. multidentata, Neocaridina davidi, and many species of Macrobrachium, hatch with some pereopods or as fully formed decapodid larvae, with well-developed thoracic appendages (Magalhaes and Medeiros, 1998; Garcia-Guerrero and Hendrickx, 2009; Tracey et al., 2013; Pantaleao et al., 2017). Disparity in thoracic and pleonal appendage growth and differentiation is considerable in the Caridea, and it appears to be driven, at least partially, by habitat (marine vs. freshwater) (Scholtz, 2000; Martin et al., 2014). The genetic mechanisms that regulate such important differences in marine and freshwater carideans seem not to have been examined yet.
Outlook: a staging method for caridean embryos and future challenges
Embryonic development in decapods, including carideans, generally follows a conserved naupliar and post-naupliar pattern that could be synthesized in a single staging system. Yet, to date, very different staging systems have been proposed in the Caridea, and major discrepancies among these systems exist, even among members of the same family and genera (see Table 1; Nazari et al., 2000; Muller et al., 2004; Garcia-Guerrero and Hendrickx, 2009; Habashy et al., 2012; Romney and Reiber, 2013; Tracey et al., 2013; Zhang et al., 2013). In carideans, the use of percentages as stage identifiers increases the inaccuracy of stage identification and makes comparisons more difficult among both closely and distantly related species. This is particularly true, given that crustaceans differ greatly in post-naupliar embryonic development, which ranges from direct to indirect (Table 1). In an attempt to facilitate comparative analyses, here we have used distinctive developmental morphological landmarks to normalize the staging of embryo development in the Caridea. This staging system is similar to the much more detailed staging system previously proposed for the amphipod Parhyale hawaiensis (Browne et al., 2005). The adoption of a concise and accurate staging system will allow comparative studies not only in embryo morphology but also in evolutionary developmental biology across many groups of decapods and crustaceans (sensu lato).
Here we have demonstrated the relative amenability of L. boggessi to developmental biology studies. However, long-term in vitro embryo culture in caridean shrimps remains a challenge. Indeed, we failed to rear embryos for live tracing; and, after five cell divisions, in the case of early cleavage, development would stall and the embryos would die. This is likely an undesired consequence of the tight relationship between the parental female and her embryos in L. boggessi and other Pleocyemata, a clade defined by the incubation of embryos in abdominal "chambers" (Bauer and Newman, 2004). Therefore, further experimentation is required to standardize in vitro culture in Lysmata and other decapods (Ituarte et al., 2014). Here we ultimately argue in favor of further research focusing on lineage tracing and in vivo embryo culture methods to establish L. boggessi as a model species for experimental embryology. These future studies will improve our understanding of the evolution of developmental modes in the Decapoda and beyond.
We thank Rhonda Reigers Powell and the Clemson Light Imaging Facility for advice and support during the project. AR-C would like to thank to Mercedes Rodriguez. Hugo Navarrete, and the School of Biological Sciences of the Pontificia Universidad Catolica del Ecuador (PUCE) for administrative support. Image processing and manuscript writing were also financially supported by PUCE.
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ANDRES ROMERO-CARVAJAL (1), MATTHEW W. TURNBULL (2), AND J. ANTONIO BAEZA (2,3,*)
(1) Escuela de Ciencias Biologicas, Pontificia Universidad Catolica del Ecuador, Quito, Ecuador; (2) Department of Biological Sciences, 132 Long Hall, Clemson University, Clemson, South Carolina 29634; and (3) Smithsonian Marine Station at Fort Pierce, 701 Seaway Drive, Fort Pierce, Florida, and Departamento de Biologia Marina, Facultad de Ciencias del Mar, Universidad Catolica del Norte, Larrondo 1281, Coquimbo, Chile
Received 7 July 2017; Accepted 23 April 2018; Published online 23 May 2018.
(*) To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
Abbreviations: PBS, phosphate-buffered saline: PBST, PBS + 0.1% Triton-X.
Table 1 Lysmata boggessi stages of embryo development in comparison with other Caridea (Lysmata. Macrobrachium. Palaemonetes, and Alpheus), Pleocyemata (Cherax), and Amphipoda (Parhyale) staging systems Lysmata Macrobrachium Lysmata boggessi amboinensis (1) olfersi (2) Egg major axis ([micro]m) 580 600 600 Rearing temperature ([degrees]C) 24 26 25 Hatching time (days) 12 13 14 Hours Days L. boggessi stage 00:00 1 One cell 1 1 0-4 03:33 1 Two cells 2 2 4-7 06:43 1 Four cells 3 3 4-7 10:29 1 Eight cells 4 4 4-7 16:19 1 Early blastoderm 5 5 4-7 23:39 1 Late blastoderm 6 6 4-7 37:20 2 Early gastrula 7 6 7-14 54:10 3 Late gastrula 8 7 7-14 68:10 3 Germ disk 9 7 14-21 condensation 96:00 4 Egg-naupliar 10 7 21-29 ... Post-naupliar 1 (a) 11 8 29-36 (PNA) 5 Post-naupliar 2 (b) 12 9 36-50 (PNA) 7 Post-naupliar 3 (b) 13 9 36-50 (PNA) 8 Post-naupliar 4 (b) 14 10 50-64 (PNA) 9 Post-naupliar 5 (b) 15 11 64-93 (PNA) 10 Pre-zoea (b) 16 12-14 64-93 (PNA) 12 Zoea 1 (b) 17 15 93-100 (PNA) Macrobrachium Palaemonetes Lysmata boggessi americanum (3) pugio (4) Egg major axis ([micro]m) -350 691 Rearing temperature ([degrees]C) 24 20 Hatching time (days) 18 13 Hours Days L. boggessi stage Equivalent stage 00:00 1 One cell 0-10 I 03:33 1 Two cells 0-10 II 06:43 1 Four cells 0-10 II 10:29 1 Eight cells 0-10 II 16:19 1 Early blastoderm 0-10 II 23:39 1 Late blastoderm 0-10 II 37:20 2 Early gastrula 10-20 IIIa 54:10 3 Late gastrula 10-20 IIIa 68:10 3 Germ disk 10-20 IIIb condensation 96:00 4 Egg-naupliar 20-30 (a) IV ... Post-naupliar 1 (a) 30-40 (b) V (PNA) 5 Post-naupliar 2 (b) 30-40 (b) V-VIa (PNA) 7 Post-naupliar 3 (b) 40-50 (b) VI (PNA) 8 Post-naupliar 4 (b) 50-60 (c) VI-VIIa (PNA) 9 Post-naupliar 5 (b) 60-70 (c) VIIb (PNA) 70-80 (c) 10 Pre-zoea (b) 80-90 (c) VIIb (PNA) 12 Zoea 1 (b) 90-100 (c) VIII (pna) Alpheus Cherax Lysmata boggessi angulosus (5) destructor (6) Egg major axis ([micro]m) 750 2000-3000 Rearing temperature ([degrees]C) 21-22 19 Hatching time (days) 20-21 40 Hours Days L. boggessi stage 00:00 1 One cell 0-10 0 03:33 1 Two cells 0-10 5 06:43 1 Four cells 0-10 10-15 10:29 1 Eight cells 0-10 10-15 16:19 1 Early blastoderm 0-10 10-15 23:39 1 Late blastoderm 0-10 10-15 37:20 2 Early gastrula 0-10 10-15 54:10 3 Late gastrula 0-10 10-15 68:10 3 Germ disk 0-10 15-20 condensation 96:00 4 Egg-naupliar 0-10 20-35 ... Post-naupliar 1 (a) 0-10 (PNA) 35-40 (a) 5 Post-naupliar 2 (b) 10-20 (PNA) 40-45 (b) 7 Post-naupliar 3 (b) 10-20 (pna) 45-50 (c) 8 Post-naupliar 4 (b) 20-30 (PNA) 50-55 (c) 9 Post-naupliar 5 (b) 30-50 (PNA) 55-60 (c) 10 Pre-zoea (b) 50-70 (c) 60-75 (c) 12 Zoea 1 (b) 70-90 (c) 75-95 (c) 90-100 (c) Parhyale Lysmata boggessi hawaiensis (7) Egg major axis ([micro]m) 300 Rearing temperature ([degrees]C) 26 Hatching time (days) 10 Hours Days L. boggessi stage 00:00 1 One cell 1 03:33 1 Two cells 2 06:43 1 Four cells 3 10:29 1 Eight cells 4 16:19 1 Early blastoderm 5 23:39 1 Late blastoderm 6 37:20 2 Early gastrula 7 54:10 3 Late gastrula 7 68:10 3 Germ disk 8 condensation 96:00 4 Egg-naupliar 9-15 ... Post-naupliar 1 (a) 16-17 (b) 5 Post-naupliar 2 (b) 18-19 (c) 7 Post-naupliar 3 (b) 19-25 (c) 8 Post-naupliar 4 (b) 19-25 (c) 9 Post-naupliar 5 (b) 19-25 (c) 10 Pre-zoea (b) 19-25 (c) 12 Zoea 1 (b) 19-25 (c) 95-100 (d) Values for equivalent stages are shown as percentages, where applicable. Superscript numbers attached to species names indicate corresponding sources: 1 = Zhang et al., 2013; 2 = Muller et al., 2004; 3 = Garcia-Guerrero and Hendrickx, 2009: 4 = Romney and Reiber, 2013: 5 = Tracey et al., 2013: 6 = Sandeman and Sandeman. 1991; 7 = Browne et al., 2005. PNA = only reported as post-naupliar appendages. Appendage differentiation is indicated by superscript letters. (a) Maxillule and maxilla. (b) Maxilliped. (c) Pereopods. (d) Pleopods.
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|Author:||Romero-Carvajal, Andres; Turnbull, Matthew W.; Baeza, J. Antonio|
|Publication:||The Biological Bulletin|
|Date:||Jun 1, 2018|
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