Introduction
Sea urchins have long served as important model organisms for studies of fertilization and early development, providing fundamental insights into sperm–egg interactions and early embryogenesis (e.g., Wolpert and Gustafson, Reference Wolpert and Gustafson1961; McClay, Reference McClay2011). Sea urchins are also important model organisms for fields that require reproductive experiments using gametes or embryos, such as biomedical studies (e.g., Bernardo and Carlo, Reference Bernardo, Carlo and Agnello2017) and ecotoxic studies (e.g., Dorey et al., Reference Dorey, Martin, Oberhänsli, Teyssié, Jeffree and Lacoue-Labarthe2018; Kiyomoto, Reference Kiyomoto2019). Therefore, the development of reliable and practical methods for sperm preservation is essential, as it allows efficient and sustainable use of these valuable resources, particularly for experiments involving genetically modified or otherwise rare individuals.
There is a significant market for sea urchins in fisheries, and demand has been increasing for years; however, global catches are declining due to overfishing, making the protection of natural populations a critical issue (e.g., Andrew et al., Reference Andrew, Agatsuma, Ballesteros, Bazhin, Creaser, Barnes, Botsford, Bradbury, Campbell, Dixon, Einarsson, Gerring, Hebert, Hunter, Hur, Johnson, Juinio-Meñez, Kalvass, Miller, Moreno, Palleiro, Rivas, Robinson, Schroeter, Steneck, Vadas, Woodby, Xiaoqi, Gibson, Barnes and Atkinson2002; Baião et al., Reference Baião, Moura, Rocha, Valente and Cunha2021). Sea urchin aquaculture and seed production are conducted in various countries (e.g., Eddy et al., Reference Eddy, Brown, Harris, Brown and Eddy2015; Unuma et al., Reference Unuma, Sakai, Agatsuma, Kayaba, Brown and Eddy2015), and effective and easy gamete preservation methods support basic research in aquaculture and increase the chances of successful seed production. Sea urchins also occupy important ecological niches and contribute to maintaining marine ecosystems (e.g., Steneck, Reference Steneck and Lawrence2013). Thus, the development of standardized methods for gamete preservation in sea urchins is necessary from both ecological and economic perspectives (Fabbrocini et al., Reference Fabbrocini, Silvestri and D’Adamo2021).
There are three main methods for preservation of the sperm of sea urchins. The first method is cryopreservation. Detailed protocols for cryopreservation of sperm and embryos have been established in several sea urchin species (reviewed in Paredes et al., Reference Paredes, Adams, Vignier, Foltz and Hamdoun2019). Cryopreservation is an effective method since it allows the stable long-term storage of sperm for years, but in many cases, it needs expensive and large-scale equipment such as a programmed freezer, and there is a limitation for shipping since the cryopreservation utilizes liquid nitrogen. It is also known that adjustment of the multiple parameters for the programmed freezing is needed due to differences among genera or species (Vacquier and Hamdoun, Reference Vacquier and Hamdoun2024). Thus, cryopreservation is relatively suitable for experimental usage of specific species. The second method is the chilled preservation (usually at 4°C) of dry sperm. It is traditionally known that the sea urchin sperm collected without the contamination of seawater (called ‘dry sperm’) can be preserved under chilled conditions and are activated when diluted in seawater (e.g., Rahman et al., Reference Rahman, Tsuchiya and Uehara2009). This method is quite easy and effective for the short term, but in many cases, the fertilization capability is lost in a few days (Fernández et al., Reference Fernández, Di Marco, Rubilar, Cledón and Pisani2024). The third method is chilled preservation in the preservation medium, and it has intermediate features of these two methods, i.e., it needs only a refrigerator or ice and preservation media containing simple chemicals and enables preservation for several weeks. It is known that sea urchin eggs can be stored in seawater containing antibiotics (Epel et al., Reference Epel, Vacquier, Peeler, Miller and Patton2004), and we previously utilized this method for the sperm of sea urchin Hemicentrotus pulcherrimus and succeeded in the storage of competent sperm at 4°C for 60 days (Kiyomoto et al., Reference Kiyomoto, Hamanaka, Hirose and Yamaguchi2014). Recently, it was confirmed that storage at 0°C using seawater containing HEPES and additional potassium chloride as medium enabled the preservation of competent sperm of Strongylocentrotus purpuratus for over 50 days and Lytechinus pictus for over 30 days (Vacquier and Hamdoun, Reference Vacquier and Hamdoun2024). Thus, these methods are thought to be potentially applicable to other echinoid species, but whether these methods are applicable to various species, and the effects of preservation on sperm features, are still unclear.
In this study, we employed the method used by Kiyomoto et al. (Reference Kiyomoto, Hamanaka, Hirose and Yamaguchi2014) for various sea urchins in Japan to examine whether the protocol is universally effective for echinoid species other than H. pulcherrimus. In addition, to evaluate in detail the effects of preservation on sperm competency, features such as fertilization rate, development of fertilized eggs, sperm motility, and the shape of flagella beat were investigated in detail. We investigated five species, including four regular echinoid species that have different breeding seasons and one irregular echinoid that is phylogenetically distant from regular echinoids (Figure 1). We reinvestigated H. pulcherrimus, which was used by Kiyomoto et al. (Reference Kiyomoto, Hamanaka, Hirose and Yamaguchi2014) and is the major echinoid used for developmental studies in Japan (Kinjo et al., Reference Kinjo, Kiyomoto, Yamamoto, Ikeo and Yaguchi2018). The other three regular echinoids we studied, Mesocentrotus nudus, Pseudocentrotus depressus, and Heliocidaris crassispina, are edible and commercially important species in Japan (reviewed in Agatsuma, Reference Agatsuma and Lawrence2020a, Reference Agatsuma2020b). We also studied Clypeaster japonicus, a common irregular echinoid in Japan that is also used in developmental studies (Hamaguchi and Hamaguchi, Reference Hamaguchi and Hamaguchi2001; Hibino et al., Reference Hibino, Minokawa, Yamazaki, Foltz and Hamdoun2019).
Echinoid species used in this study and their breeding season. Echinoids used in this study were collected from various sites and kept in outdoor flow-through tanks at Tateyama Marine Laboratory (Chiba, Japan) until sexual maturation (see also materials and methods). The columns represent months, and grey bars indicate the breeding season of each species at the laboratory.
Figure 1 Long description
The image contains one diagram and five photos of echinoids. The diagram displays a timeline with months labeled from 4 to 12 and 1 to 3, representing the breeding seasons of different echinoid species. Grey bars indicate the breeding season for each species. The five photos, arranged vertically, show different species of echinoids: Hemicentrotus pulcherrimus, Mesocentrotus nudus, Pseudocentrotus depressus, Heliocidaris crassispina, and Clypeaster japonicus. Each species has arrows pointing to specific months, indicating their respective breeding seasons.
Materials and methods
Gamete collection
Sea urchins used in this study were collected from ocean depths of 0 to 20 m during the breeding season of each species (Figure 1). The habitats and the months of sperm collection for each species were as follows:
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• H. pulcherrimus: Choshi and Tateyama in Chiba Prefecture and Asamushi in Aomori Pref. (January)
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• M. nudus: Otsuchi in Iwate Pref. (October)
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• P. depressus: Tateyama in Chiba Pref. (November)
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• H. crassispina: Tateyama in Chiba Pref. and Usa in Kochi Pref. (June)
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• C. japonicus: Tateyama in Chiba Pref. and Usa in Kochi Pref. (June)
The detailed procedure for gamete collection is described in Kiyomoto et al. (Reference Kiyomoto, Hamanaka, Hirose and Yamaguchi2014). In brief, mature sperm and eggs were obtained by injecting 0.1–1 mL of 0.5 M potassium chloride or 2 mM acetylcholine into the body cavity of urchins. Sperm was collected as dry sperm. Before sperm collection, a male individual was wrapped in paper towels and gently shaken to remove the seawater on the body surface to prevent the contamination with seawater. Injected males were placed directly on a glass dish without seawater, with the gonopore facing downward. Semen containing little seawater (dry sperm) was then collected from the dish using a glass pipette and used for preservation. Eggs were collected and washed several times with Millipore-filtered natural seawater (MFSW) before the experiments.
Preparation for sperm preservation
Collected dry sperm was diluted with 100 times the volume of MFSW containing 200 μg/mL sulfamethoxazole and 10 μg/mL trimethoprim (ST-SW) according to Epel et al. (Reference Epel, Vacquier, Peeler, Miller and Patton2004). Diluted sperm was dispensed in 1.5 mL Eppendorf tubes and stored at 4°C during preservation period.
Measurement of fertilization rate and observation of development
To quantify the fertilization capability of preserved sperm, the fertilization rate was measured using a sperm dilution series. Sperm stopped swimming under the storage condition and was settled on the bottom of tubes, so the tube was gently inverted to mix sperm just before the experiment. For the measurements, serial 10-fold dilutions of the preserved sperm were made using MFSW. Sperm was preserved at 10−2 dilution, and serial dilutions of the preserved sperm were made from 10−2 to 10−7, so the final concentration was diluted 10−4 to 10−9 times compared to dry sperm. Dilution series were made in 24-well plates with ultra-low attachment surface (Corning 3473), and around 100 unfertilized eggs were dropped quickly into each well after the dilution. The fertilization rate was measured by counting the eggs with fertilization envelope under a stereo microscope (Olympus SZX7) 5 min after the egg addition. Each measurement included 3 biological replicates. Measurement was started from the day of the sperm collection (0 weeks) and conducted every 2 weeks, up to 4–8 weeks after the collection. Eggs used in the measurement were freshly collected on every measurement day. The sperm dilution that achieved 50% fertilization was calculated using a curve fitting function for the logistic curve in ImageJ software (https://imagej.nih.gov/ij/). Regression curves for Figure 2F were calculated using Microsoft Excel. Fertilized eggs obtained after the measurement were washed with MFSW and kept in 10 mL dishes until they became pluteus larvae. The embryos in each developmental stage were placed on a hole-slide glass and observed under a microscope (Olympus BX53) equipped with a camera (Olympus DP74). Incubation temperatures for fertilization and embryos were approximately 18°C for H. pulcherrimus, 20°C for M. nudus and P. depressus, 24°C for H. crassispina and C. japonicus.
Fertilization assays using dilution series of preserved sperm. (A–E) Fertilization rate by preserved sperm for each species. The horizontal axis of each panel represents the final concentration calculated from dry sperm. Sperm was preserved at 10−2 dilution in ST-SW, and dilution series were prepared by 10−2–10−7 dilution of preserved sperm. The vertical axis of each panel represents the fertilization rate of fresh eggs by preserved sperm. The fertilization rates were obtained by counting the eggs with fertilization envelopes. Each value is the mean of 3 biological replicates. The fertilization rates were measured on the day of the sperm sampling (0 w) and every 2 weeks during the preservation period. Individual data of each week is presented in supplementary figure 1. The standard deviations for each time point are provided in the Supplementary Figure 1. (F) Changes in fertilization competency of sperm during preservation. Dilution for 50 % fertilization was plotted for each biological triplicate for each species and preservation period. The regression curve for each species is presented as a line. Regression curves are y = 7E-08e0.5893x (R 2 = 0.2979) for H. pulcherrimus, y = 1E-07e0.3277x (R 2 = 0.3884) for M. nudus, y = 5E-07e0.538x (R 2 = 0.5228) for P. depressus, y = 6E-08e1.1325x (R 2 = 0.6322) for H. crassispina and y = 3E-07e0.602x (R 2 = 0.9451) for C. japonicus.
Figure 2 Long description
The image contains five line graphs labeled A to E and one scatter plot labeled F. Each line graph shows the fertilization rate of fresh eggs by preserved sperm for different sea urchin species over various preservation periods. The horizontal axis of each panel represents the final concentration calculated from dry sperm, while the vertical axis represents the fertilization rate. The species and preservation periods are color-coded: H. pulcherrimus (A), M. nudus (B), P. depressus (C), H. crassispina (D), and C. japonicus (E). Graph F shows the changes in fertilization competency of sperm during preservation, with dilution for 50 percent fertilization plotted for each biological triplicate for each species and preservation period. The regression curves for each species are presented as lines. The data indicates how fertilization rates decrease over time and with increasing sperm dilution for each species.
Measurement of flagellar beat frequency
Sperm flagellar beat frequency was measured by the previously described method (Wessel et al., Reference Wessel, Wada, Yajima and Kiyomoto2021). The preserved sperm was transferred using a toothpick to prevent co-transfer of surrounding liquid onto MFSW containing 0.01% (w/v) Bovine Serum Albumin (Wako, Japan) and placed between a glass slide and a coverslip with two strips of plastic tape (Scotch, 3 M) as spacers. The frequency of flagellar beating was measured by synchronizing the frequency of custom-made stroboscopic LED illumination equipped on a microscope (Olympus CHBS) to sperm flagellar beating. Around 30 sperm from more than three microscopic fields were measured for each preserved batch.
Analysis of flagellar beat form
After measurement of beat frequency, the observation chamber was removed from the darkfield microscope and placed upside down onto an inverted phase contrast microscope for flagellar beat form analysis. Images of motile spermatozoa were observed under a phase contrast microscope (UPlan Fl objective lens, IX2-LWUCD condenser on Olympus IX-51 inverted microscope) and recorded at 90 fps with a camera (UI-3130CP-M-GL, iDS, Germany) driven by control software (Boh Navi, Bohboh software, Japan) on a LAVIE Direct N15 (NEC, 64 bit, Windows 11). The flagellar beat waveforms were analyzed using motion analysis software (Bohboh ver. 4.94; BohbohSoft, Tokyo, Japan; Ohmuro et al., Reference Ohmuro, Mogami and Baba2004). First, using an automatic tracking function of the motion analyzer, the XY-coordinates along the centre line of the flagella were obtained for each spermatozoa image (total 100 frames for 1.1 s). Then, using that coordinate data, we obtained the curvature of each 1 μm flagellar segment (for details, see Wada et al. (Reference Wada, Baba and Kamimura2015)). We used these curvature values to evaluate how long the original waveform was maintained in each preserved suspension.
Results
Fertilization capability of preserved sperm
In every species, the preserved sperm were capable of fertilization for at least 4 weeks after the gamete collection (Figure 2). Overall, there was a tendency for the fertilization rate at a given sperm concentration to decrease as the preservation period increased. In other words, a higher concentration of sperm was necessary to obtain the same fertilization rate after preservation.
For H. pulcherrimus, the fertilization rate was almost 100% at 10−6 or lower dilution on the day of sperm collection (0 weeks, Figure 2A). At 10−6 dilution, the fertilization rate was maintained at around 100% until 2 weeks after the sperm collection (2 weeks), but the fertilization capability was gradually reduced, and the fertilization rate was decreased to approx. 50% at 4–6 weeks. Finally, the fertilization capability was lost at this concentration, and the fertilization rate was approx. 0% at 8 w preservation. At 10−4 dilution, the fertilization rate was almost 100% even at 6 weeks and around 70% at 8 weeks. For M. nudus, the fertilization rate was approx. 90% at 10−6 dilution at 0 w and decreased to approx. 80% at 2 weeks, 70% at 4 weeks, 50% at 6 weeks and 0% at 8 weeks (Figure 2B). At 10−4 dilution, the fertilization rate was almost 100% until 8 weeks. For P. depressus, the fertilization rate was almost 100% at 10−5 dilution or higher concentration at 0 weeks (Figure 2C). At 10−5 dilution, the fertilization rate was approx. 100% at 2 weeks, 80% at 4 weeks and 30% at 6 weeks. At 10−4 dilution, the fertilization rate was almost 100% until 6 weeks. For H. crassispina, the fertilization rate was almost 100% at 10−6 dilution at 0 weeks and decreased to approx. 80% at 2 weeks and 10% at 4 weeks (Figure 2D). At 10−4 dilution, the fertilization rate was almost 100% at 2 weeks and around 60% at 4 weeks. For C. japonicus, the fertilization rate was approx. 80% at 10−6 dilution at 0 weeks and decreased to approx. 70% at 2 weeks and 20% at 4 weeks (Figure 2E). At 10−4 dilution, the fertilization rate was almost 100% until 4 weeks.
Focusing on the concentration of sperm which leads to a substantial fertilization rate (50%), the required sperm concentration increased as the preservation period was extended (Figure 2F). The rate of decrease in fertilization competency was steepest for H. crassispina. The competency was reduced to 1/10th in 2 weeks. For the other species, although the sperm dilution that produced 50% fertilization varied among species, the rate of decrease was approximately the same and less steep than that for H. crassispina. The competency was reduced to 1/10th in 4 weeks in these species.
Morphology of embryos fertilized with preserved sperm
As shown in Figure 2, the fertilization capability was gradually reduced as preservation periods increased, but fertilized eggs were obtained even after long preservation, so we observed the development of those embryos. The sperm preservation period for the observation was 8 weeks for H. pulcherrimus and M. nudus, 6 weeks for P. depressus, and 4 weeks for H. crassispina and C. japonicus. The morphology of the embryos fertilized by the preserved sperm was normal, and the developmental process proceeded without any defects (Figure 3). In each species, the fertilization envelope was raised after fertilization (Figure 3A–E). After the fertilization, cleavage proceeded (Figure 3F–J), the embryo became a blastula (Figure 3K–O), and then became a normal pluteus larva (Figure 3P–T). There was no recognizable difference in morphology and progression of development between embryos fertilized with preserved sperm and embryos fertilized with normal, fresh sperm.
Development of embryos obtained by fertilization with preserved sperm. The morphology of embryos obtained by fertilization with preserved sperm was normal at each presented stage for the 5 investigated species. The columns correspond to the species, and the rows correspond to the developmental stages. Embryos of H. pulcherrimus (A, F, K, P) and M. nudus (B, G, L, Q) were obtained by fertilization with sperm preserved for 8 weeks. Embryos of P. depressus (C, H, M, R) were obtained by fertilization with sperm preserved for 6 weeks. Embryos of H. crassispina (D, I, N, S) and C. japonicus (E, J, O, T) were obtained by fertilization with sperm preserved for 4 weeks. Scale bars: 100 µm. The time point for each developmental stage is shown in the lower left corner of each panel. hpf, hours post fertilization; dpf, days post fertilization.
Figure 3 Long description
The image displays the development of embryos from five different species of sea urchins at various stages. The species are H. pulcherrimus, M. nudus, P. depressus, H. crassispina, and C. japonicus. Each row represents a different developmental stage: fertilized egg, cleavage, blastula, and pluteus. The embryos of H. pulcherrimus and M. nudus were fertilized with sperm preserved for eight weeks, while P. depressus embryos were fertilized with sperm preserved for six weeks. The embryos of H. crassispina and C. japonicus were fertilized with sperm preserved for four weeks. The time points for each developmental stage are indicated in the lower left corner of each panel, with hpf standing for hours post fertilization and dpf for days post fertilization. Scale bars are set at one hundred micrometers.
Flagellar beat frequency
To evaluate the effect of preservation on sperm motility, we next investigated the frequency of sperm flagellar beat. Figure 4 shows the average sperm flagellar beat frequency of each sea urchin species measured every 2 weeks. In general, the beat frequency was 30–40 Hz at the date of sperm collection and then decreased as the preservation period increased. For H. pulcherrimus, the beat frequency was around 35 Hz and decreased by approx. 5 Hz every 2 weeks (Figure 4A). After 10 weeks of preservation, the frequency dropped to 10–20 Hz, but the sperm maintained competency for flagellar beating. In M. nudus, the frequency was around 35 Hz at 0 weeks, decreased by approx. 5 Hz every 2 weeks, and finally dropped to 10–20 Hz at 8 weeks (Figure 4B). For P. depressus, the frequency was 35–40 Hz at 0 weeks, decreased by approx. 10 Hz every 2 weeks, and was 10–25 Hz at 6 weeks, though the difference among samples was relatively large (Figure 4C). For H. crassispina, the frequency was around 40 Hz at 0 weeks, decreased by approx. 5 Hz every 2 weeks, and was around 30 Hz at 4 weeks (Figure 4D). For C. japonicus, the frequency was around 35–40 Hz at 0 weeks, decreased by approx. 10 Hz every 2 weeks, and was around 25 Hz at 4 weeks (Figure 4E). The rate of decrease of flagellar beat during the preservation period varied among species, but the frequency at the end of the period was 10–20 Hz in H. pulcherrimus, M. nudus, and P. depressus (species that mature in autumn to winter) and was higher, around 20–35 Hz, in H. crassispina and C. japonicus, species that mature in summer (Figures 1 and 4F). The average frequency at 0 w was also higher in species that mature in summer (Figure 4F).
Changes in sperm flagellar beat frequency during preservation. (A–E) Flagellar beat frequency of preserved sea urchin sperm. Vertical axes indicate the beat frequency measured, and horizontal axes indicate the preservation periods. Each dot represents the mean value of approx. 30 sperm with circular swimming path in stable wave form, and standard deviations for each dot are shown. Sea urchin species are indicated for each graph. Three biological replicates were measured at each time point, and data from the same storage tube were connected by lines. (F) Superimposed plots of 5 species. The average frequency of three replicates is presented.
Figure 4 Long description
The image contains six line graphs showing changes in sperm flagellar beat frequency during preservation for five sea urchin species: H. pulcherrimus, M. nudus, P. depressus, H. crassispina, and C. japonicus. Each graph plots the beat frequency on the vertical axis and the preservation period in weeks on the horizontal axis. Graph A shows data for H. pulcherrimus over ten weeks, Graph B for M. nudus over eight weeks, Graph C for P. depressus over six weeks, Graph D for H. crassispina over four weeks, and Graph E for C. japonicus over four weeks. Graph F superimposes the data from the five species, presenting the average frequency of three replicates. Each dot represents the mean value of approximately thirty sperm with circular swimming paths in stable wave form, with standard deviations shown for each dot. The data points from the same storage tube are connected by lines. The graphs illustrate a general decline in beat frequency over time for all species, with varying rates of decline. All values are approximated.
The beat frequency of preserved sperm could be measured until 10 weeks after sperm collection for H. pulcherrimus, 4 weeks for H. crassispina and C. japonicus, 6 weeks for P. depressus and 8 weeks for M. nudus (Figure 4A–E). After these periods, the beat frequency could not be measured due to the unstable swimming of preserved sperm (see Discussion for details). In other words, stable swimming of sperm could not be observed beyond these periods, and the periods vary among species. At the date of sperm collection, almost all sperm swam in a circle with stable beat, but the number of sperm that swam in a circle gradually decreased, and finally, the flagellar beat frequency could not be measured at the end of the preservation period since almost all sperm swam in an unstable manner. Some examples of the types of unstable swimming were three-dimensional swimming, swimming with flagellar beat vertical to the focal plane, intermittent swimming in a typical ‘quiescent’ manner (Gibbons, Reference Gibbons1980), and so on.
There was also a tendency for the duration of stable sperm swimming to decrease as the preservation period increased. Sperm in ST-SW were thought not to be moving and started swimming when diluted with MFSW. In every species, at 0 weeks almost all sperm swam in circles for more than 10 min, but the duration of stable swimming became shorter in later periods, and at the end of the preservation period, flagellar beat ceased soon after the dilution. For example, in P. depressus after 6 weeks storage, about half of sperm swam in circles just after dilution, though most sperm stopped in 1–2 min (Movie S2).
Flagellar beat form
In addition to the beat frequency, we also observed the sperm flagellar beat form and calculated curvature of flagella along the whole length during the preservation. Figure 5 shows representative flagellar curvature and trace of a sperm in each species at the corresponding preservation period. Overall, flagellar beat form was well maintained during all preservation periods, and both the trace of flagella and the profile of curvature showed little change during storage.
Changes in sperm flagella wave form during preservation. Superimposed traces of flagella (upper panel) and curvature profile(;lower panel);taken from 100 video frames are presented. For the flagellar trace, the shape of flagella in each video frame was superimposed, fixing the position of the sperm head. Sperm head position is indicated by an arrowhead. For the curvature profile, the bending curvatures obtained from the XY-coordinates of sperm tails were plotted. Vertical axes indicate the curvatures, and horizontal axes correspond to the distance from the proximal end of the flagellum. For H. pulcherrimus, data from every 2nd week are presented (A), and for other species, data from the beginning and latest period are shown (B–E).
Figure 5 Long description
The image contains multiple graphs showing changes in sperm flagella wave form during preservation for different sea urchin species over time. Each species is represented by a set of flagellar traces and curvature profiles. The upper panels display superimposed traces of flagella from 100 video frames, with the sperm head position indicated by an arrowhead. The lower panels show curvature profiles, where bending curvatures obtained from the XY-coordinates of sperm tails are plotted. Vertical axes indicate the curvatures, and horizontal axes correspond to the distance from the proximal end of the flagellum. For H. pulcherrimus, data from every 2nd week are presented, while for other species, data from the beginning and latest period are shown. The graphs illustrate how the flagellar wave form changes over the preservation period for each species.
For H. pulcherrimus, the trace of flagella and the curvature profile were almost unchanged from 0 w to 8 w. The trace of flagella indicated that flagella of preserved sperm bent uniformly from the proximal part to the distal tip at every preservation period (Figure 5A, upper row). Curvature plots showed the same tendency (Figure 5A, lower row). The width of the curvature plot was the same from the proximal part to the distal tip, indicating that the flagella curvature changed symmetrically during the measurement, and the curvature was constant regardless of the distance from the base of the flagella. This means that the bending of flagella propagated properly from the sperm head to the end of the flagella, even after the long preservation. Movie S1 shows side-by-side movies of H. pulcherrimus sperm swimming at 0 weeks and 8 weeks of ST-SW storage. Sperm swimming became somewhat slower due to the lower beating frequency, while the basic swimming pattern did not change much. In M. nudus, the overall shape of the flagella trajectory and the curvature plot did not change between 0 weeks and 8 weeks (Figure 5B). Similarly, no definitive change was observed between 0 weeks and 4 weeks in P. depressus (Figure 5C), 0 weeks and 2 weeks in H. crassispina, or 0 weeks and 4 weeks in C. japonicus (Figure 5E). On the other hand, although the overall shape of the curvature plot was almost the same throughout the preservation, as described above, at the end of the preservation period, the width of plot for the distal portion was slightly narrowed in some species. This tendency was observed most obviously in H. crassispina at 2 weeks (Figure 5D), and was also observed in other species such as H. pulcherrimus at 8 weeks (Figure 5A), and P. depressus at 4 weeks (Figure 5C). This change in shape indicates that flagella bending became attenuated at the distal part.
Discussion
Our results showed that our sperm preservation method at 4°C using ST-SW was applicable to each of the five investigated species (Figure 2A–E). In general, the fertilization rate decreased as the preservation period increased, but over 50 % fertilization rate was maintained at 10−4 dilution throughout all preservation periods. In addition, preservation did not affect the larval morphology (Figure 3). These results indicate that our method is widely applicable to various echinoid species, including irregular echinoids, and competent sperm is potentially preservable for 4–8 weeks.
As for the fertilization rate, decreased fertilization rate with increasing preservation period was obviously observed at 10−6 dilution (Figure 2A–E). The reduction in fertilization competency occurred the fastest in H. crassispina, which matures in summer, and occurred at almost the same rate in the other four species (Figure 2F). For these four species, fertilization competency was reduced to approximately 1/10th of the initial value after 4 weeks of preservation. This indicates that the number of active and competent sperm in preserved populations was decreased to about 10% from the beginning of preservation. On the other hand, at 10−4 dilution, the fertilization rate using preserved sperm was almost 100 % throughout all preservation periods. So, in general, a 10−4 dilution of sperm (i.e., 10−2 dilution of the preserved sperm) resulted in enough sperm to obtain a substantial fertilization rate even after 4–8 weeks of preservation. At this concentration, the number of competent sperm was sufficient to obtain high fertilization rates even after long preservation, since the whole population was large. In addition, these findings suggest that the addition of preserved sperm at higher concentrations to eggs would potentially be successful for sperm with longer preservation periods. Incidentally, in our experiment, the effective concentration of sperm varied even at 0 w (Figure 2F). This phenomenon is thought to be due to the difference in the number of sperm in dry sperm preparations and the inevitable contamination of seawater during the sampling procedure. In future studies, normalization by counting of the actual number of sperm will be necessary to obtain a more accurate value of the concentration of effective preserved sperm.
Observation of the embryos fertilized by preserved sperm revealed that sperm preservation did not affect the larval morphology. This result is consistent with previous research (Kiyomoto et al., Reference Kiyomoto, Hamanaka, Hirose and Yamaguchi2014; Vacquier and Hamdoun, Reference Vacquier and Hamdoun2024). The results showed that our preservation methods were also effective for obtaining normal embryos, not only for the fertilization assay. In H. pulcherrimus, the method using ST-SW is also applicable for unfertilized egg preservation (Vacquier and Hamdoun, Reference Vacquier and Hamdoun2024). Testing whether this method is applicable for eggs of other species in detail will be important for effective gamete utilization.
The flagellar beat frequency was gradually reduced during preservation (Figure 4). In addition, the ratio of sperm with a stable manner of swimming decreased, and the duration of flagellar beating became shorter, as the preservation period increased. These observations show that sperm gradually lost their activity with increasing preservation period, and the reduction of fertilization rate with the increasing preservation period (Figure 2) might reflect the gradual loss of sperm activity.
On the other hand, we observed a substantial average beat frequency of sperm flagella of all species investigated in this study throughout the preservation period. This means that stably swimming sperm were present in the preserved populations since the measurement of flagellar beat frequency requires manual synchronization of strobe frequency to the stable flagellar beat. These stably swimming sperm are thought to contribute to the substantial fertilization rate after long preservation (Figure 2). There was almost no change in sperm flagella beating form during preservation (Figure 5), so it can be said that the propagation of flagella beating was not damaged during preservation. The slight constriction of the curvature plot in the distal portion observed in H. crassispina (2 weeks) indicates attenuation of the curvature of flagella bending at the distal portion, and this might be due to partial damage of the flagella during preservation. It is known that flagellar beat frequency is determined by the outer arm dynein in flagella, and beat form (beat amplitude) is determined by the inner arm dynein (Brokaw and Kamiya, Reference Brokaw and Kamiya1987), so one possibility is that the activity of outer arm dynein in flagella is lost faster than that of inner arm dynein under the preservation conditions of this study. Detailed motility assays focusing on sperm with unstable beat to extract changes in beat form during preservation may offer new insights into the mechanisms of sperm deterioration during preservation.
In further studies, we will also need to accurately quantify the number of active sperm in the whole population. Our observations mainly focused on the surviving, active, and normal sperm in the preservation population, and showed that competent sperm are included in the preserved sperm population, so practically it can be said that the sperm preservation method using ST-SW is very effective. On the other hand, to know the number of inactive or damaged sperm during the preservation and to understand the mechanism of deterioration should also be important for improvement of the preservation method. Interestingly, it is known that the preservation of sperm motility is also decreased with time in cryopreserved sperm (Fabbrocini et al., Reference Fabbrocini, Silvestri and D’Adamo2023). Comparison of damage of the sperm by different preservation strategies will also help us to understand the relation between sperm activity and preservation condition.
In this study, we show that sperm obtained from various sea urchin species can be stored for up to 8 weeks at 4°C using ST-SW, and that our simple method is practically applicable across multiple species. This method allows a longer preservation period than chilled storage of ‘dry sperm’, thereby reducing the frequency of sperm collection from parental individuals and conserving biological resources in research, such as seed production, larval embryology, and ecotoxicological studies. It is particularly advantageous for rare species, specific strains or genetically manipulated individuals with limited opportunities for sperm collection. Although storage for several months or years cannot be achieved, as is possible with cryopreservation, our method requires only simple equipment and materials and does not require species-specific condition optimization. Therefore, it is especially suitable for repeated experiments over several weeks using multiple species and is useful for researchers as well as for applications in aquaculture and education.
Supplementary material
The supplementary material for this article can be found at https://doi.org/10.1017/S0967199426100458.
Acknowledgements
This work was partly performed as a project of the Joint Usage/Education Center by the Ministry of Education, Culture, Sports, Science & Technology in Japan and partly supported by the Nippon Foundation in Japan and by the Cooperative Program (No. 106, 2023) of the Atmosphere and Ocean Research Institute, The University of Tokyo, in Japan.
We thank the staff of Usa Marine Biological Institute, Kochi University, International Coastal Research Center, Atmosphere and Ocean Research Institute, The University of Tokyo, Research Center for Marine Biology, Tohoku University and Tateyama Marine Laboratory, Institute for Marine and Coastal Research, Ochanomizu University for the animal collections.
We also thank Dr. Shoji A. Baba for providing flagellar analyzing tool Bohboh and Dr. Yoshihiro Mogami for arranging the flagellar recording system.
Competing interests
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.
