Diapausing embryos encased within cladoceran ephippia result from sexual reproduction and increase genetic diversity. They are also important means by which species bypass harsh environmental conditions and disperse in space and time. Once released, ephippia usually sink to the benthos and remain there until hatching. Using the Sars’ method (incubating sediments to identify cladoceran hatchlings), ephippial egg bank biodiversity can be evaluated. Yet, even when samples are incubated under a variety of conditions, it is not possible to warrant that all have hatched. Few keys are available that facilitate the identification of cladocerans by using only ephippial morphology. Our goal was to analyze some cladoceran ephippia from Mexico, to develop a means to identify them using easily recognizable characteristics. Ephippia of 23 cladoceran species from waters in Aguascalientes (México) in 11 genera (Alona, Biapertura, Ceriodaphnia, Chydorus, Daphnia, Dunhevedia, Ilyocryptus, Macrothrix, Moina, Pleuroxus, and Simocephalus) were analyzed. In our analysis six morphological features were selected that permitted the identification of ephippia to species(-group) level. The results demonstrate that with a proper catalog of features, some ephippia can be identified.

Diapausing embryos, dormancy, ephippia, sediment, SEM, taxonomy, ultrastructure

Many aquatic micrometazoans produce diapausing embryos (DEs) that permit them to bypass adverse conditions in their habitat, including drought (Schröder et al. 2007), inadequate food conditions (Drillet et al. 2011), and the presence of predators (Hairston and Munns 1984; Slusarczyk 1995; Pijanowska and Stolpe 1996). Taxa capable of producing DEs or other dormant stages include protists (Perrigo et al. 2012), bryozoans (Figuerola et al. 2003), cladocerans (Kokkinn and Williams 1987), copepods (Uye 1985), gastrotrichs (Ricci and Balsamo 2000), nematodes (Ptatscheck and Traunspurger 2020), rotifers (Walsh et al. 2017), and tardigrades (Altiero et al. 2009). The resulting dormant stages in Cladocera usually sink to the bottom, although in some macrothricids, ephippia may be attached to macrophyte leaves (Fryer 1972) or algae (see appendix in Guerrero-Jiménez et al. 2020). Thus, DEs constitute a genetically diverse egg-bank for future generations and they may disperse passively by anemochory (Rivas Jr et al. 2018, 2019), anthropochory (Bailey et al. 2003; Panov et al. 2004; Perrigo et al. 2012), hydrochory (Ricci and Balsamo 2000), and zoochory (Moreno et al. 2019; Vanschoenwinkel et al. 2011). When conditions are favorable, these dormant stages may hatch and replenish the previous population or establish a new one elsewhere (Montero-Pau et al. 2017; Vargas et al. 2019; Odriozola et al. 2020; García-Zamorano and Jiménez-Contreras 2023).

Water fleas (Crustacea, Cladocera) are important components of freshwater ecosystems, passing energy and nutrients on to higher trophic levels (Gulati and DeMott 1998). Most cladocerans reproduce by cyclic parthenogenesis with gamogenetic reproduction happening occasionally. Gamogenetic females of the order Anomopoda Sars, 1865 produce diapausing embryos (resting eggs) covered by a protective, saddle-shaped structure called the ephippium (G., epi, on + hippos, horse), which usually contains one or two embryos (Fryer 1972); in some macrothricids more than two are observed (Scourfield 1902; Smyly 1956), as well as in Eurycercidae and some other lineages. Some taxa may reproduce dormant stages asexually (Decaestecker et al. 2009), such as an entire population of Daphnia pulex Leydig, 1860 from North America (Hebert and Crease 1983). Knowledge of the conditions that permit hatching of DEs from sediment samples allows researchers to incubate sediments and identify the hatchlings; from such studies one can develop species lists of the cladocerans in a habitat (Vandekerkhove et al. 2005b). This method is called “the Sars’ Method” (Van Damme and Dumont 2010). Nevertheless, one can never be certain that the DEs from all species present in a sediment sample have hatched, even when the samples have been incubated under a variety of settings that otherwise appear to mimic favorable natural conditions. In addition, hatching is not possible when sediments have been preserved in a fixative. Considering these obstacles, developing comprehensive species lists of a cladoceran community would be improved if researchers were able to identify unhatched ephippia based solely on their morphology. This method is already very widely used in paleolimnology, where cladoceran ephippial identification is one of the pillars of reconstructing past conditions of lakes; entire identification books exist on only remains of cladocerans and ephippia from Europe (Szeroczyfiska and Sarmaja-Korjonen 2007). One of the first advancements in the identification of unhatched ephippia came from drawings of different ornamentations in species of Bosmina longirostris complex O.F. Müller, 1785 by Scourfield (1901). Later, Scourfield (1902) expanded on this by illustrating the ephippia of what he called at the time the “Lynceid Entomostraca” (chydorids). Goulden (1966) further improved our knowledge of ephippial morphology by illustrating variation in four Moina species. Subsequently, Vanderkerkhove et al. (2004) recognized 29 ephippial morphotypes and Mergeay et al. (2005) observed clear differences in several Daphnia species from Kenya. Subfossil records are also a widely used tool for the identification of cladocerans in the field of paleolimnology. In particular, Szeroczyfiska and Sarmaja-Korjonen (2007) provide detailed descriptions of morphological traits of ephippia belonging to several species of Daphniidae and Chydoridae in particular in their ‘Atlas of Subfossil Cladocera from Central and Northern Europe’. More detailed studies were accomplished using SEM technology to illustrate ephippia ornamentations in fossil records, and in ephippia that were still attached to the female (Kotov 2013). Kotov et al. (2018) also demonstrated differences in six Ceriodaphnia species from central, northern European Russia, predominantly for their practical use in paleolimnological settings. More recently Guerrero-Jiménez et al. (2020) presented 11 morphotypes extracted from sediments from Spain and Mexico, of different species of water fleas.

Yet, despite the excellent progress that has been made in our ability to differentiate ephippia of different species, we still lack useful diagnostic features. Thus, the aim of our research was to revisit an old – yet important – and largely unstudied question posed more than 30 years ago by Kokkinn and Williams (1987): “is ephippial morphology a useful taxonomic descriptor in the Cladocera?” Studies like Kotov et al. (2018) have shown that it can indeed be useful even between close species, to some extent, yet sometimes large detail is needed. There are at least two major problems to the development of a key to cladoceran ephippia: (1) the information on their taxonomic characters is scattered in the literature and (2) we have insufficient information on their features as a standardized nomenclature of ephippial characters is largely lacking. This knowledge gap is acerbated by the fact that there are at least 620 described cladoceran species (Forró et al. 2007), and most ephippia are studied in Palearctic settings. Thus, we still lack sufficient information so that ephippial morphology can become a useful tool for researchers studying the contemporary ephippial egg bank.

Here we report our results in producing a first database that will hopefully improve our ability to identify cladoceran ephippia of some Mexican taxa based only on their morphology. This information could be an important tool to estimate cladoceran diversity in freshwaters in the region (Vandekerkhove et al. 2005a, b) and subsequently to appreciate their adaptation to the environment within an evolutionary context (Brendonck and De Meester 2003; Gueriau et al. 2016).

A total of 4017 ephippia belong to 23 cladoceran species in 11 genera: Alona Baird, 1843, Biapertura Smirnov, 1971, Ceriodaphnia Dana, 1853, Chydorus Leach, 1816, Daphnia O.F. Müller, 1785, Dunhevedia King, 1853, Ilyocryptus G.O. Sars, 1861, Macrothrix Baird, 1843, Moina Baird, 1850, Pleuroxus Baird, 1843, and Simocephalus Schoedler, 1858 were analyzed (see Figs 2–7, Suppl. materal 1: appendix B).

Figure 2. 

Micrographs of the ephippial ornamentation taken by A SEM B light microscope, and C the organisms hatched from the dormant embryos. 1. A. aguascalientensis Sinev & Silva-Briano, 2012 (immature individual in 1C), 2. Alona sp., 3. Biapertura ossiani Leydig, 1860, 4. Ceriodaphnia cornuta Sars, 1886. Arrows show the zoom of the specific ornamentation. Scale bars: 100 µm.

Figure 3. 

Micrographs of the ephippial ornamentation taken by SEM (A) light microscope (B) and organism hatched from the egg (C). 5. C. dubia Richard, 1894, 6. C. laticaudata P.E. Müller, 1867, 7. C. reticulata Jurine, 1820, 8. Chydorus sphaericus complex O.F. Müller, 1776. Arrows show the zoomed in image of the ornamentation. Scale bars: 100 µm.

Figure 4. 

Micrographs of the ephippial ornamentation of some Daphnia taxa, taken by SEM (A) and light microscope (B) and organism hatched from the dormant embryo (some immature) (C). 9. Daphnia (Ctenodaphnia) exilis Herrick, 1895, 10. D. leavis Birge, 1879, 11. D. parvula Fordyce, 1901, 12. D. pulex Leydig, 1860. Arrows show the zoomed in image of the ornamentation and some spinules or serrations. Scale bars: 100 µm.

Figure 5. 

Micrographs of the ephippial ornamentation taken by SEM (A) light microscope (B) and organism hatched from the egg (some immature) (C). 13. Dunhevedia crassa King, 1853, 14. Ilyocryptus agilis Kurz, 1878, 15. Macrothrix mexicanus Ciros-Pérez, Silva-Briano & Elías-Gutiérrez, 1996, 16. M. rosea Jurine, 1820. Arrows show the zoomed in image of the ornamentation. Scale bars: 100 µm.

Figure 6. 

Micrographs of the ephippial ornamentation taken by SEM (A) light microscope (B) and organism hatched from the egg (C). 17. Macrothrix smirnovi Ciros-Pérez & Elías-Gutiérrez, 1997, 18. M. macrocopa Straus, 1820, 19. M. micrura Kurz, 1875, 20. Pleuroxus denticulatus Birge, 1879. Arrows show the zoomed in image of the ornamentation. Scale bars: 100 µm.

Figure 7. 

Micrographs of the ephippial ornamentation of Simocephalus, taken by SEM (A) light microscope (B) and organism hatched from the egg (C). 21. Simocephalus mixtus Sars, 1903, 22. Simocephalus vetulus O.F. Müller, 1776, 23. Simocephalus sp. (embryo/adult absent). Scale bars: 100 µm. (*) Ephippium extracted from laboratory cultures without males being present; (**) no hatched organism.

1. Size. Within each genus, most species had ephippia of similar size; therefore, while size was useful in identification of the genus, it could not be used for species-level diagnosis (Table 1).
2. Shape. The shape of cladoceran ephippia varied among the genera we examined. Ephippial shape in some genera was consistent: all species of Simocephalus had a triangular shape (Fig. 7(21, 22, 23)), while a half oval shape was seen in Ceriodaphnia (Figs 2(4), 3(5, 6, 7)). On the other hand, ephippial shape varied in different Macrothrix species (Figs 5(15, 16), 6(17)). Another important feature was the margin of the ephippium. The margin of ephippia in Alona and Daphnia were similar in shape; but in some species the margins were rounded, while in others they were sharp. These differences are enough to distinguish between some species within a genus (Figs 2(1, 2, 3), 4(10, 11, 12)).
3. Color. The most common colors seen in the species studied were brown and dark (Gerrish and Cáceres 2003; Vandekerkhove et al. 2004); however, most ephippia were at least partially transparent. Thus, color was just used in cases that present patterns used to identify ephippia to the level of species: e.g., Simocephalus vetulus O.F. Müller, 1776 and Simocephalus sp. Within the ephippia, some embryos were orange in color (Fig. 7(22, 23)).
4. “Resting egg” (dormant embryo) number. The number of DEs within the ephippia of the species we studied is reported in Table 2.
5. Type of ornamentation. Five types of ornamentation were observed. (i) Depressions (“craters”) : These were commonly observed in Daphnia sp. and Moina macrocopa Straus, 1820. These structures were observed only under SEM (Fig. 6(18A, arrow)). (ii) Verrucae: Small spherical structures were observed only in Macrothrix mexicanus Ciros-Pérez et al. 1996 (Fig. 5(15A, arrow)). (iii) Striae: Pronounced marks in the surface of the ephippia were identified; these were linear in Biapertura ossiani Leydig, 1860 (Fig. 2(1A, arrow)), smooth and rounded in Ceriodaphnia dubia Richard, 1894 (Fig. 3(5A, arrow)), pronouncedly rounded in M. mexicanus (Fig. 5(15A), and small, interlocking linear in Simocephalus sp. (Fig. 7(23A)), no link or patterns between all these species were observed. (iv) Pores: Very small pores were detected on the surface of the ephippia of Dunhevedia crassa King, 1853 (Fig. 5(13A, arrow)). (v) Reticulation : Reticulations were seen in some genera. In Alona and Biapertura species, linear reticulations were evident (Fig. 2(1, 2, 3)); in Ceriodaphnia, the reticulation cells were oval (Figs 2(4), 3(5, 6, 7)); in Pleuroxus they were hexagonal in shape (Fig. 6(20A)). (vi) Scales: Different types of scales were observed. Small and irregular ones were present in Ceriodaphnia cornuta Sars, 1886 (Fig. 2(4A)); rounded scales were seen in the Macrothrix rosea Jurine, 1820 (see Dumont et al. 2002) (Fig. 5(16A)); oval scales were present in S. mixtus and S. vetulus (Fig. 7(21A, 22A)). Finally, in Moina micrura Kurz, 1875, two different types of scales were detected: rounded scales in the anterior portion of the ephippium and scales with irregular shape covering the section where the DE was located (Fig. 3(5A)).
6. Accessory structures. Only five species: Alona aguascalientensis Sinev and Silva-Briano, 2012, Daphnia (Ctenodaphnia) exilis Herrick, 1895, Daphnia leavis Birge, 1879, D. parvula Fordyce, 1901, and D. pulex Leydig, 1860 have accessory structures. Most of those were seen in Daphnia (Table 2, Fig. 4). (vii) Filamentous membranes: Filamentous membranes were observed only in A. aguascalientensis (Fig. 3(5A)). (viii) Ventral appendices: These were only observed in C. exilis (Fig. 3(5A)). (ix) Spinules: Spinules were specific to Daphnia, but they were also observed in C. exilis. In Daphnia, a long caudal spinule is present (Fig. 3(5A)), while in C. exilis, both caudal spinule and anterior projection were observed. In the posterior portion, a small spinule was attached to the ventral appendix while a longer spinule is observed in the anterior part of the ephippium (Fig. 3(5A)).

We found that ephippial morphology was relatively consistent and contained useful diagnostic features. Our results show useful differences in the morphology of ephippia in the taxa encountered, except in Ceriodaphnia; thus, morphological differences of ephippia in that genus remain a challenge. Because the six features we used in this study allowed us to achieve a useful identification of the ephippia we examined, we conclude that morphological characterization of ephippia is a sufficiently robust tool for the identification of ephippia, for certain taxa. Nevertheless, we recognize that a serious knowledge gap remains. Our analysis and the database we provide needs to be expanded to include many more species and additional stable characteristics. When this is achieved, ephippial morphology will be a convenient and practical means for cladoceran morphological identification.

We thank Biol. Roberto Viscalla who provided some samples used in the study and Ekaterina Retes Pruneda for identifying some species. We thank also the anonymous reviewers for their constructive remarks and comments on an earlier draft of the manuscript.