Abstract
Unionoids are in global decline, which may be associated with their complex life cycle. Their juveniles are unique because while hidden (burrowed deeply in bottom sediments) they undergo critical anatomical changes (also developing a characteristic juvenile shell sculpture). Currently, the juveniles’ period of life is believed to be both the least known and one of the most vulnerable—thus the possibility of obtaining any biological knowledge is essential for establishing conservation strategies and addressing functional or evolutionary questions. I propose two new methods for visualization of the burrowing behavior of unionoid juveniles within deposits that are cheap and easy: (1) laminated deposits of quartz–aragonite sand for time-stepped X-ray images of bivalve traces, and (2) silica gel serving as 'invisible sand' for direct observations and video recording of behavior within sediments. Both deposits in a pilot study were accepted by the juvenile unionoids—they were stable enough and penetrable, with no observable signs of harmful effects on animals’ behavior during trials. In both, juveniles were clearly visible, settled within the top 1 cm layer of deposits. Both methods are promising tools for future in situ within the deposits research on the biology of this much unexplored and vulnerable unionoids' life stage.
Similar content being viewed by others
Introduction
Unionoid populations today suffer from many, commonly anthropogenic factors and are in serious decline worldwide (e.g., Ferreira-Rodríguez et al., 2019), thus the possibility of obtaining any biological knowledge of their rarely observed juvenile life stage that is also believed to suffer high mortality is essential for both establishing and optimizing conservation strategies as well as addressing functional or evolutionary questions. Common freshwater mussels classified in the order Unionida (unionoids, naiads) form a monophyletic and biologically coherent taxonomic unit. They live today in lakes and rivers worldwide (Bogan & Roe, 2008; Lopes-Lima et al., 2018; Graf & Cummings, 2021). They are also currently regarded as one of the most threatened freshwater taxa in the world (e.g., Haag & Williams, 2014), with estimates of up to 985 living species worldwide (Bogan & Roe, 2008; Graf, 2013; Graf & Cummings, 2007, 2021, 2023), with their diversity decreasing rapidly on a global scale (Seddon et al., 2011; Haag & Williams, 2014; Pereira et al., 2014; Lopes-Lima et al., 2018; Van Tu et al., 2018; Ferreira-Rodríguez et al., 2019; Haag, 2019; Böhm et al., 2020). These bivalves are characterized by a unique life cycle: their larvae must pass through the parasitic stage on a fish host, to complete the metamorphosis into a juvenile mussel, what is a believed adaptation for dispersal (Kat, 1984; Graf & Cummings, 2006; Ferreira-Rodríguez et al., 2019) which probably appeared already before the Middle Jurassic when the modern lineages of Unionida already existed (Watters, 2001; Skawina, 2021). Adults are large (up to a dozen of cm in length) and live shallowly submerged in the deposits where they filter feed—in the 'siphoning position' of Archambault et al. (2014), Amyot & Downing (1991), Schwalb & Push (2007), Watters et al. (2001). On the other hand, juveniles, which commonly develop characteristic and species-specific shell sculpture (so-called umbonal sculpture, or umbonal rugae; Haas, 1969; Aldridge, 1999; Zieritz et al., 2015) live hidden within the bottom deposits.
Biology of early juveniles—what we know according to the scattered data
Larvae of unionoids are small [length from 60 to 360 µm, depending on the species; e.g., Wächtler et al. (2001)] and after absorbing nutrients from the body of their host they metamorphose and post-larval minute mussels, juveniles, escape from the host and sink to the bottom. If they find a suitable substratum, they burrow there and they undergo drastic morphological changes including the development of ciliated gills, which allows the transition from the juvenile's deposit feeding via the foot—to suspension feeding via gills. After reaching a size of about 3 cm—they emerge at the surface and begin adult life (Yeager et al., 1994; Wächtler et al., 2001; Schwalb & Ackerman, 2011; Schartum et al., 2017; Araujo et al., 2018; Irmscher & Vaughn, 2018). The post-larval or ‘early juvenile’ stage of life within sediments, as was introduced by Isely (1911) for mussels up to 15 mm in length (today the definition is understood as up to 3 cm in length) is currently believed to be the least known and one of the most vulnerable (Geist, 2010; Schartum et al., 2017; Patterson, 2018; Bílý et al., 2021). One of the reasons for the lack of basic biological knowledge is that minute juvenile unionoids are difficult to observe in the field e.g., (Lefevre & Curtis, 1910; Piechocki, 1969; Neves & Widlak, 1987; Amyot & Downing, 1991; Hastie & Cosgrove, 2002; Irmscher & Vaughn, 2018) and long were difficult to be grown in cultures to subadult age (Lefevre & Curtis, 1908; Dimock & Wright, 1993; Lima et al., 2012; Mair, 2018).
What we do know is that after their detachment from the fish host they are small (about 0.2–0.5 mm; Wächtler et al., 2001; Irmscher & Vaughn, 2018), may drift with the current (in streams) until they sink to the bottom of the stream/lake (where they can possibly exert some control over habitat selection; Schwalb & Ackerman, 2011; French & Ackerman, 2014; Irmscher & Vaughn, 2018) and immediately burrow themselves in well-aerated deposits (they are more mobile than older unionoids; Yeager et al., 1994; Wächtler et al., 2001; Kemble et al., 2020; Hyvärinen et al., 2021). Authors expect that general conditions are similar to those of the adults, but differ in microhabitats occupied (Neves & Widlak, 1987; Buddensiek et al., 1993; Strayer, 2008). They can live clustered (Neves & Widlak, 1987; Hastie & Cosgrove, 2002) or dispersed (Piechocki, 1969; Hastie & Cosgrove, 2002). More often they were found within fine deposits of small pools close to rivers' banks (Piechocki, 1969), in 'riffles, runs, and behind boulders' (but are rarer in pools) of the stream in pebbles, gravel, and sand (Neves & Widlak, 1987) or in ‘clean sand of lakes’ (James, 1985). Early juveniles of at least some species were observed to live byssally attached to submerged objects (Isely, 1911; Lavictoire et al., 2018). The depth to which early juveniles may burrow in nature is unknown, however, they were reported from a maximum depth up to 20 cm (Schwalb & Push, 2007), while in experiments the post–metamorphic early juveniles were recovered within the top 1 cm of sediment (Yeager et al., 1994), and more recently around 2–3 cm (Archambault et al., 2014; Kemble et al., 2020; Bílý et al., 2021). They are more commonly noticed when some tools [dredges, bottom scratch samplers, Bernatowicz’s grabs, or Günther’s sampler; see in Lewandowski & Kołodziejczyk (2014)] and intensive search are applied (Strayer, 1981; Amyot & Downing, 1991; Hastie & Cosgrove, 2002). Although field information is quite scarce, it is believed that they spend 2–3, up to 6 or even 20 years (Hochwald & Bauer, 1990; Amyot & Downing, 1991; Balfour & Smock, 1995; Strayer et al., 2004; Schwalb & Push, 2007; Simon et al., 2015) within the sediment in interstitial waters (hyporheal environment). There they pedal-feed [sensu Reid et al. (1992)] on unknown particles [possibly bacteria, detritus, different algae and diatom species (Gatenby et al., 1996; Jones et al., 2005; Barnhart, 2006)]. During this period, they undergo critical anatomical changes to establish the ability of suspension feeding.
The majority of the knowledge about these internal changes comes from laboratory cultures, designed for the conservation of a few dozen species at a maximum (e.g., Buddensiek, 1995; Gatenby et al., 1997; O'Beirn et al., 1998; Jones et al., 2005; Barnhart, 2006; Kovitvadhi et al., 2007; Schmidt & Vandré, 2010; Denic, 2018; with a detailed description of successful systems in Mair (2018)). In particular, Mair (2018) recently summarized that although mussel culturing has progressed and developed during the last two decades, there is no universal method for raising juvenile unionoids in the laboratory for conservation purposes. At the same time, quite common problems in culturing juvenile unionoids refer to their high mortality during the first months of life—in some cultures up to 100% (Lefevre & Curtis, 1908; Kovitvadhi et al., 2001; Jones et al., 2005; Eybe et al., 2013), but in more successful cultures the mortality dropped to 10–80% (Gatenby et al., 1996; Jones et al., 2005; Schmidt & Vandré, 2010; Eybe et al., 2013). This high mortality is interpreted as a result of diet or predation on juveniles (Kovitvadhi et al., 2001; Jones et al., 2005), or more recently as a result of possible food shortage—low filtering efficiency that accompanies organogenesis and the transition to filter–feeding [(Schartum et al., 2017; Araujo et al., 2018) but not confirmed by Lavictoire et al. (2018)]. The other described difficulties affecting survival of juveniles refer to their possible high sensitivity to environmental parameters in sediments [e.g., low oxygen concentration or high nitrate and ammonium levels; (Dimock & Wright, 1993; Mummert et al., 2003; Newton & Bartsch, 2007; Hyvärinen et al., 2022)]. This contributed to the idea to use early juveniles and their survival rates as a potentially useful biological indicator providing a measure of the quality of the streambed (Geist & Auerswald, 2007; Archambault et al., 2017; Kemble et al., 2020).
Juvenile shell ornamentation—biological role and evolution
In addition to the little-known biology, young unionoids (up to about 1 cm in length) typically bear pronounced surface protrusions [widely discussed by Zieritz et al. (2015)]. The sculpture’s functional role, while poorly understood, is often considered to be an adaptation to life within the sediment. It is believed to ease burying in the substrate and support anchoring in it (Stone et al., 1982; Watters, 1994; Hornbach et al., 2010), which represents one of the shell sculpture specializations typical for burrowing bivalves (Stanley, 1969; Seilacher, 1984; Savazzi & Yao, 1992). Nevertheless, the oldest Late Triassic unionoids did not have any prominent sculpture on the shell (Good, 1998; Skawina & Dzik, 2011) but it is known only since the Early or Middle Jurassic (Cai, 1986; Skawina & Dzik, 2011). The late emergence of prominent juvenile sculpture in the evolution of the unionoids may suggest that the discrepancy between the early juvenile and later stages of life characteristic of many extant unionoids emerged gradually during the Early Jurassic period. This interpretation of the unionoid phylogeny remains speculative until the actual functional meaning of the prominent juvenile (umbonal) sculpture is identified.
Aim of the experiment
The aim of this pilot experiment which was conducted as an observational study on a small number of animals was therefore to test two distinct methods of visualization of behavior within the deposits of unionoid juveniles of different, common in Poland (and in Europe) species. These species differ from the others by having prominent [Unio tumidus Philipson, 1788, U. pictorum (Linnaeus, 1758)] or delicate [Anodonta anatina (Linnaeus, 1758)] umbonal sculpture. My study also highlights the difficulties and feasibility of obtaining juvenile unionoids from the field in numbers that are sufficient for behavioral experiments. Limited access to the captive-breeding facilities at the time of this study on the one hand, together with the aim of tracing juveniles in the field in Central Europe decades after the last published attempt (Piechocki, 1969) on the other hand supported this endeavor. The scarce data from the literature cited above suggest that such minute individuals (preferred up to 10 mm, according to the common range of sculptured juvenile shell) have to be searched for with tools within the top 10 up to 20 cm layer of the bottom sediment. The limit of their penetration into sediments is presumably imposed by oxygen availability and contamination with toxic metabolites. They are also likely unable to penetrate the partly lithified clay that frequently underlies the sandy bottom of Polish rivers.
Methods
Fieldwork
The following techniques for collecting samples were applied, depending on the water depth and thickness of the oxygenated sediment layer:
-
1.
Bottom scratch sampler—to a water depth of about 1 m, if the thickness of suitable sediment was small; side length 20 cm; (Lewandowski & Kołodziejczyk, 2014).
-
2.
Spade—to a water depth of about 1 m, if the suitable sediment was more than 5 cm thick.
-
3.
Günther sampler from the boat at water depths from about 1 m to about 5 m; sample catchment surface 272 cm2; (Lewandowski & Kołodziejczyk, 2014; Jurkiewicz-Karnkowska et al., 2017).
More than 30 localities were available to collect juvenile unionids, identified from literature data (e.g., Lewandowski, 1996 and personal information (advice) from K. Lewandowski and A. Piechocki, as well as my own assessment. They belonged to the public river–lake system of Krutynia (the Masurian Lakes, NE Poland; (Jakubik & Lewandowski, 2011). Sampling was performed in June 2009 and its aim was to search the promising deposits until success. Each sample was sieved through a 1 mm mesh geological sieve; animals, if found, were identified and placed in a container for transit. The length of each animal was measured after transfer to the laboratory. Fifteen live juvenile individuals less than 10 mm long (range 6–10 mm) were collected (two A. anatina (Linnaeus, 1758), one Unio pictorum (Linnaeus, 1758) and 12 U. tumidus Philipsson, 1788). All small mussels came from sandy deposits both in lakes and in rivers (nevertheless, the majority of the collected specimens came from rivers, commonly from shallow run water and close to river banks; they appeared dispersed—if any, not more than one individual per site was found); they were collected with a bottom scratch sampler (commonly in rivers) or spade (usually in lakes), and no juvenile bivalve was collected with the Günther sampler. Additionally, more than 20 juveniles measuring 11–30 mm (three A. anatina and more than 20 U. tumidus) were collected. All gathered individuals were kept alive in the water tank, however, only the 11 smallest individuals of all species (together with one 21 mm long U. tumidus) were selected for this observational study (thus requiring a small number of animals)—a pilot experiment (Table 1).
All bivalves were sustained in a 60 l freshwater water tank with aerated water filtered with a carbon filter and were fed Scenedesmus sp. algae suspension (each 48 h) for 2 weeks before experiments. Each tank contained quartz sand deposits of about 1–2 cm in depth (sieved with a geological sieve, 0.25–1 mm fraction, following the field observation of common strata). The quartz sand was washed and sterilized after purchasing from a sand mine in the Vistula River (in Warsaw), in which adult unionoids are observed abundantly but where juveniles haven’t been gathered (pers. obs.).
X-ray method
To trace the dynamics of vertical behavior of juvenile unionoids within sediments, I applied an X-ray method that requires laminated deposits of different density contrast (following idea of Gingras et al., 2008). After the bivalve burrows deeply, it should leave a pathway of bioturbated, mixed sands of two kinds, visible on X-ray images. First, in an attempt to identify the best contrast of deposit lamination in X-ray pictures, several successions of laminations were prepared containing layers of quartz sand, sand with quartz grains stained with iron minerals, and aragonite sand (commercially prepared for marine coral–reef aquaria and purchased from the zoological shop; washed and sterilized prior to use). Glass test tubes (19.0 cm high and 1.5 cm in diameter) were filled with interbedded laminae of fine–grained sands (Fig. 1). The laminations were obtained by filling the tubes with water and sprinkling sand into them by hand. The thickness of a single layer was about 5 mm. To examine the suitability of the method for X-ray imaging empty juvenile shells of U. tumidus or A. anatina of similar size (about 8 mm) were placed in the middle of each test tube. The tubes were X-ray imaged at the Philips Duo Diagnost (SN: 7001487) diagnostic device in the Department of Radiology, Military Institute of Medicine, Warsaw. Files were produced as DICOM files. All shells were at least noticeable as blackish areas within lighter sands (Fig. 1c), and the best contrast between layers was achieved by using laminas of quartz and aragonite sands, thus this lamination was used to fill the glass aquariums.
Two flat, glass aquariums (dimensions 25.0 × 25.0 × 1.5 cm) were filled with 5 mm thick layers of fine–grained sand (alternately quartz layer and aragonite layer), following the procedure first described in Gingras et al. (2008), see above. Both aquariums were aerated, kept in a room temperature, and in a natural light-darkness regime of a long day (18 h light: 8 h darkness). Juvenile naiads were then placed in the aquariums on the surface of the top quartz sand layer. The first aquarium accommodated one 8 mm long individual of U. tumidus while the second one contained two 9 mm and 14 mm long individuals of A. anatina. Bivalves were fed with Scenedesmus sp. suspension every 48 h (around 25 ml of thoroughly mixed algae suspension each time, added directly to the aquarium water). At variable time steps—1, 25, 72 h and after a week, X-ray images were produced (Fig. 2). The sequences of X-ray images were then used to determine the amount of sediment disruption and the track of the animal.
Transparent sand
For direct observations and video–recording of the mussels’ behavior within the sediments, ‘transparent sand’ consisting of silica gel was tested. While the material is sold as a desiccant comprised of solid white beads 1–3 mm in diameter (product no. 908250460, POCH S.A.), it becomes immediately quite transparent after soaking in water. No harmful effects were observed on the animals’ behavior during the experiment. Because the grains in the packages were often crushed during transit, the dry silica gel was first sieved to exclude fractions smaller than 0.25 mm.
Two other flat glass aquariums of the same dimensions (25.0 × 25.0 × 1.5 cm) were filled with pre-washed silica gel (approximately 15 cm deep layer) of and used for observations and video recording of juvenile behavior. In the first aquarium two juvenile mussel individuals were introduced—U. tumidus and U. pictorum, both 8 mm long; in the second one—three individuals of A. anatina measuring 6 mm, 9 mm [the individual from the X-ray experiment was re-used here], and 14 mm in length. The two flat water tanks were continuously recorded on video for 4 and 5 days, respectively, with the Sony DC-RSR 37E camera. Bivalves were fed with Scenedesmus sp. suspension (around 25 ml, feeding procedure as described above) every 48 h, and aquariums were aerated, kept in a room temperature, and in constant light (due to the limitations of the recording possibilities of the camera). In addition, video–recording was performed for another two individuals of U. tumidus (8 and 10 mm long), after finishing the first experiment (in the same aquarium and after removing previous Unio individuals, with the same observing regime).
Shallow aquarium
To trace the horizontal movements and accompanying behavior of the bivalves light photographs of the surface of deposits with U. tumidus measuring 21 mm, 10 mm, and 8 mm in length were taken in a shallow aquarium (11 × 11 cm and 15 cm high) with standardized as above quartz sand (1.5 to 2.0 cm deep to not exceed the depth of the sand already verified as ensuring good quality of the planned X-ray images). Bivalves were fed with Scenedesmus algae suspension every 48 h, aquarium was aerated, kept in a room temperature and a natural light-darkness regime of long day (18 h light: 8 h darkness). The camera used was Olympus E-510. The aquarium was light photographed from the top two times—the next day after setting mussels and a week later. Then after a day, X-ray pictures were taken from the top to show the number and position of bivalves in the sediment.
Results
X-ray method
Both kinds of sediment (quartz and aragonite) for this method did not vary in grain size in nature—at least quartz sand, as a top layer, was supportive and penetrable for juveniles (they were able to settle within the top layer). Nevertheless, no—or only minimal sediment disruption was recorded and no animal’s trace was recorded. All animals rested in the top 0.5 cm layer of quartz sand after obtaining the ‘adult’ siphoning position, without an attempt to burrow into deeper layers during the experiment.
The activity of U. tumidus within the sediment demonstrated by X-ray photography was very low (Fig. 2, Online Resource 1). The naiad was passively lying on the sediment surface for the first hour. During the next few hours, it set itself in the siphoning position typical for adult unionoids: the anterior part of the body was buried in the sediment, the posterior exposed above its surface (Fig. 2e). This position was recorded after 25 h, and only after 72 h the images showed a small horizontal shift of the mussel (2 cm) that was now almost completely covered with sand. All movements were horizontal and near the surface of the deposit, within the first lamina of quartz sand that was occasionally mixed with the aragonite grains underlying it (left area of Fig. 2f, compared to Fig. 2e). Both specimens of A. anatina attained the ‘adult’ siphoning orientation during the first 25 h of the experiment and there were no changes in their position until the end (Fig. 2g, h).
Transparent sand
The silica-gel sediment was sufficiently stable, supportive, and penetrable for the bivalves, and was quite transparent (larger crystals ensured more transparency than smaller ones), allowing direct observations of mussel behavior within it. The observed behavior was similar between prominently sculptured juvenile Unio species and delicately sculptured A. anatina. Individuals from all three species behaved in a similar way (Fig. 3), initiating locomotory movements in minutes after their introduction to the aquarium, and finishing movements when settled in an adult position. Only the smallest A. anatina differed from them in both timespans to begin locomotory movements (longer) and the way of performing these movements.
The 8 mm long individual of U. tumidus needed 17 min to start locomotion after the initial placement in the aquarium; it then settled in the ‘adult’ position after about 15 min of movement. U. pictorum (8 mm long) started to move after 3 min and found the appropriate place to settle after about 2 h of mobility. The largest individual of A. anatina (14 mm long) required almost 2 min to start moving and settled after 21 min of moving while the 9 mm individual, began locomotory movements after 4 min and settled in an ‘adult’ position after 1 min. The smallest, 6 mm long individual stayed motionless for 9.5 h, then moved to settle down for about 1 h, in an unusual way (Fig. 4).
An 8 mm long individual of U. pictorum was observed to settle under the sediment surface (Fig. 3d) and a similar behavior was observed in all 3 individuals of U. tumidus (Fig. 5; Table 1). In each case, the sediment covering the animals was a few millimeters deep. No deeper burrows were observed, but the animals treated this position as suitable for settlement.
Shallow aquarium
The behavior of the three individuals of U. tumidus kept in the shallow aquarium differed by size (corresponding to age), but the sample size was too small to draw general conclusions. The larger bivalve was visible from above, while the smaller ones were completely covered with sand (Fig. 6; Table 1). Although the depth of the sand did not exceed the length of the largest bivalve, observations suggested that it would be possible for it to burrow itself completely inside the deposits because the position of the bivalve within the deposits is not vertical. X-ray photography (Fig. 6d) shows the position of all three animals.
Discussion
Despite more than a century of research, the hidden hyporheal period of the lives of juvenile mussels remains underexplored because early juveniles are difficult to trace in nature, as well as maintain and grow in artificial cultures (Lefevre & Curtis, 1908, 1910; Isely, 1911, Piechocki, 1969; Hastie & Cosgrove, 2002; Mair, 2018). Researchers most commonly refer to clear aerated sand and gravel in both the rivers and lakes as a habitat of juveniles (Piechocki, 1969; James, 1985; Neves & Widlak, 1987; Hastie & Cosgrove, 2002). In this study, the attempt to gather minute unionoids resulted in 15 individuals (< 10 mm in length) and all the collected juveniles of three species came from clear, aerated, sandy habitats in shallow water, both in rivers (majority of mussels) and lakes (several individuals). The better success in the catchment of riverine juveniles was possibly a result of a shallow layer of aerated sand (about 5–10 cm) with a partially lithified clay layer underneath, which was likely impenetrable to mussels. Therefore, it was easier to search the entire available mussels’ habitat. The most useful tool was a bottom scratch sampler, supported by a spade, while the Günther sampler appeared less useful likely because this tool sampled only a few top centimeters of the deposits with quite small square dimensions, and a thorough search was unlikely (insufficient and inefficient sampling). The importance of sampling efficiency and sufficiency is underlined by Neves & Widlak (1987) and Hastie et al. (2010). Additionally, although Neves & Widlak (1987) and Newton et al. (2008) described juveniles as clumped in distribution, my study does not support this observation in line with previous studies from mussel populations in Scotland (Hastie & Cosgrove, 2002) and Poland (Piechocki, 1969).
Gingras et al. (2008) proposed time–stepped X-ray images of laminated deposits in which they determined the amount of sediment disruption made by marine invertebrates. In their method, the interbedded quartz with heavy mineral sand gave a sufficient result. I supplemented this method with aragonite sand, which is easy and cheap to provide, and gave a clear visual contrast when interbedded with quartz grains. The mobility of marine invertebrates reported by Gingras et al. (2008) was related to the mode of food gathering—suspension feeders disrupted the sediment by burrowing 10 to 100 times less than deposit–feeding taxa. Scientific interest in the diet and the way the food is gathered by juvenile unionoids has grown since an influential study by Yeager et al. (1994), which described the feeding and burrowing behavior of minute Villosa iris (Lea, 1829) individuals and documented their pedal–sweep, pedal–locomotory and interstitial suspension feeding behaviors. Araujo et al. (2018) specified that juveniles first feed with their ciliated foot and then filter suspension by developing gills, but it is not yet determined what type of food is ingested during the pedal– and filter–feeding periods of juvenile life.
Although juvenile unionoids are regarded as both deposit and suspension feeders (Yeager et al., 1994), my pilot study shows that they were not burrowing deeply and that there was no deep reworking of any of the deposits. All bivalves rested in an adult-like siphoning position after a short period of mobility and were covered with quartz sand within the top layer during the X-ray experiment (however, a slightly reworked area on the previous border of the first quartz and aragonite layers is visible after 72 h and a week of trial, in the left area of the aquarium of U. tumidus, Fig. 2d, f) and were placed a little deeper, but still within the top 1 cm,—in silica gel (Figs. 2, 3 and 5). Although a number of studies indicate that juveniles are supposed to burrow at least to a depth of several cm (e.g., Neves & Widlak, 1987; Wächtler et al., 2001; Schwalb & Push, 2007), similar results were previously obtained in laboratory experiments conducted by Yeager et al. (1994) (with early V. iris juveniles that were recovered from the top 1 cm of sediment), Kemble et al. (2020) (commonly up to 3.4 mm only, and not deeper than 5.1 mm), or Archambault et al. (2014) (not deeper than 2.5 cm). However, as expected, small (8–10 mm long) U. tumidus were observed to burrow deeper in the sediment than larger individuals (Figs. 3, 5 and 6; Table 1). Juveniles of relatively advanced age may maintain the typical adult siphoning orientation (Figs. 3, 6; Table 1). The shallow burrowing depth may be the result of unnatural experimental conditions (e.g., the sterilized quartz sand did not contain detritus and both aragonite sand and beads of silica gel were not natural to them), or of biological factors as for example experimental animals might have been collected for experiments after reaching the age at which they actually begin the post-juvenile life mode in close proximity to epibenthic environments. Hyvärinen et al. (2021) observed that the size of the grains within the deposit is important for the early juveniles of Margaritifera margaritifera—they burrowed well when grains were at least 0.25 mm (but much worse, when smaller), and the best, when grain size was above 0.5 mm in diameter, which is similar to the grain size used in my experiment. Nevertheless, Kemble et al. (2020) did not consider grain size an important determinant of the depth of burrowing of juveniles. Schwalb & Push (2007) suggest that burrowing behavior may depend on a variety of environmental factors. As laboratory conditions by definition exclude many of them, the juvenile mussels might lack the right environmental trigger for deep burrowing. Similarly, the supply of washed and sterilized deposits might have prevented bivalves from their presumed search for food in the interstitial water (Gatenby et al., 1996; Archambault et al., 2014). Feeding on the sediment may provide a significant portion of the total energy for locomotion (Vaughn & Hakenkamp, 2001).
Sparks & Strayer (1998) described a ‘stress surface behavior’ when a bivalve does not burrow itself in the deposits as a result of a stress reaction to unfavorable conditions. My results, however, do not support this explanation as the experimental bivalves did burrow quickly, albeit not deeply. Finally, Archambault et al. (2014) considered that the burrowing depth might be linked to the season because unionoids tend to burrow deeper in winter than in summer (Amyot & Downing, 1997; Perles et al., 2003; Saarinen & Taskinen, 2003; Schwalb & Push, 2007), which cannot be excluded in the above trial as the experiment was conducted during the summer months.
If the anatomical development of gills and the subsequent transition from pedal feeding to filter feeding are sufficient to trigger a shift to an adult mode of life outside sediments, the emergence of juveniles to the surface of the deposits could occur earlier. This idea may be weakened by the recent observations of Schartum et al. (2017) who showed that the critical size for a juvenile M. margaritifera to start switching from juvenile pedal feeding to adult filter feeding is about 2.2 mm in length and is related to size only, not age. Additionally, M. margaritifera that exceeds 4.5 mm in size must filter feed because pedal feeding is not sufficient. At the same time, a 1.5 cm long juvenile examined by the authors had underdeveloped gills. Mair (2018) noticed that, in general, mortality in cultures usually decreases when the juvenile mussels exceed 1 mm in length. This is consistent with the timing of the transition from pedal feeding to filter feeding recognized in juveniles of M. margaritifera and U. mancus Lamarck, 1819 (Araujo et al., 2018). A proper transition from pedal feeding to gill feeding during juvenile ontogenesis has recently been identified as essential for survival (Schartum et al., 2017; Araujo et al., 2018) but not by Lavictoire et al. (2018). Unfortunately, the timing of this organogenesis has been studied in a small number of species, and it varies between species, e.g., in Hyriopsis (Limnoscapha) myersiana (Lea, 1856) the development of gills is quicker than in Anodonta (Kovitvadhi et al., 2007). In this context, a single observation of an unusual behavior in a 6 mm long A. anatina (Fig. 4; Online Resource 2) could be interpreted as related to food intake behavior [e.g., pedal–locomotory feeding; (Yeager et al., 1994)] even though the bivalve was not inside the sediment, especially when Kaestner (1967) describes the development of the outer demibranchs of Anodonta which is not complete until its shell size reaches 3 to 5.7 mm. Normally, any locomotory movement of an adult naiad requires (1) a slight opening of the shell valves with protrusion of the foot into substrate until fully (or almost fully) extended, (2) anchoring the tip of the foot by dilation of its distal end, and (3) contraction of the pedal retractor that results in pulling the shell towards the anchored foot. After such movement, the shell is usually slightly tilted, its anterior end being oriented downward, and the ventral margin is horizontal (see, e.g., Figs. 2, 3a—the larger individual). The behavior of the smallest A. anatina (6 mm) was unusual in this respect. Although it was not buried in the sediment (as would be predicted for a juvenile), it moved in a manner different from that of the larger juvenile bivalves. Pulling off its foot rotated the shell to an upright position, with the anterior end towards the bottom, but the ventral part of the shell was oriented in the direction of movement (Fig. 4; Online Resource 2). The sculpture on the juvenile shell of unionoids is a feature that—with the hyporheal period of early juvenile life—may represent an adaptation to life inside the sediment (Seilacher, 1984; Savazzi, 1991). There are many possible reasons for evolving and supporting the burrowing behavior of juveniles including protection from larger predators and possible parasites or competitors [summarized by (Strayer, 2008), but also exampled by (Nichols & Wilcox, 1997; Taskinen & Saarinen, 2006; Schwalb & Push, 2007)]. However, the design of the current experiment does not permit further discussion of the functional aspects of juvenile sculpture based on empirical evidence. Due to the difficulties in obtaining minute juvenile unionoids from the field (resulting from their biology) in sufficient numbers to achieve statistical significance, the study shows potential for continuation in the future, possibly supplemented by incorporating juveniles from the captive-breeding facilities (like in Černá et al., 2017).
Conclusions
The rapid and global decline of unionoid freshwater mussels, as well as many outstanding questions about their biology during one of the most vulnerable and little-known stages of their life, led me to design two new experimental methods for visualizing the behavior of juvenile unionoids within sediments: (1) time-stepped X-ray images of bivalve traces within laminated deposits of quartz–aragonite sand and (2) silica gel serving as ‘transparent sand’ for direct video recording of the behavior within sediments. The methods were cheap and easy to set up. The deposits were stable, supportive, and penetrable (no signs of harmful effects were observed during the experiments) and were readily used for burrowing by juvenile unionoids. Although the experiment was conducted as an observational study on a small number of animals to test the methods, the findings may be informative for more substantial research in the future.
All tested in this study juvenile mussels settled in the top 1 cm layer of deposits with no attempts to burrow deeper in both methods. More data are needed, possibly with the use of younger stages of mussels (e.g., from captive-breeding facilities), to give a more conclusive answer if such behavior was the result of biological factors (e.g., they were collected for the experiment after they reached the post-juvenile stage) or the unnatural experimental conditions. However, consistent with my predictions, smaller mussels were observed to burrow deeper and were more thoroughly covered by the deposits than larger individuals.
Data availability
Electronical supplementary materials include a PowerPoint slideshow containing all X-ray pictures of experimental aquariums (Online Resource 1) and a short video recording mp4 file of A. anatina unusual behavior (Online Resource 2). All data generated or analysed during this study are included in this published article (and its supplementary information files); the video-recorded raw data are available from the corresponding author on reasonable request.
References
Aldridge, D. C., 1999. The morphology, growth and reproduction of Unionidae (Bivalvia) in a fenland waterway. Journal of Molluscan Studies 65: 47–60. https://doi.org/10.1093/mollus/65.1.47.
Amyot, J.-P. & J. A. Downing, 1991. Endo- and epibenthic distribution of the unionid mollusc Elliptio complanata. Journal of the North American Benthological Society 10: 280–285. https://doi.org/10.2307/1467601.
Amyot, J.-P. & J. A. Downing, 1997. Seasonal variation in vertical and horizontal movement of the freshwater bivalve Elliptio complanata (Mollusca: Unionidae). Freshwater Biology 37: 345–354. https://doi.org/10.1046/j.1365-2427.1997.00158.x.
Araujo, R., M. Campos, C. Feo, C. Varela, J. Soler & P. Ondina, 2018. Who wins in the weaning process? Juvenile feeding morphology of two freshwater mussel species. Journal of Morphology 279(1): 4–16. https://doi.org/10.1002/jmor.20748.
Archambault, J. M., W. G. Cope & T. J. Kwak, 2014. Survival and behaviour of juvenile unionid mussels exposed to thermal stress and dewatering in the presence of a sediment temperature gradient. Freshwater Biology 59(3): 601–613. https://doi.org/10.1111/fwb.12290.
Archambault, J. M., C. M. Bergeron, W. G. Cope, P. R. Lazaro, J. A. Leonard & D. Shea, 2017. Assessing toxicity of contaminants in riverine suspended sediments to freshwater mussels. Environmental Toxicology and Chemistry 36(2): 395–407. https://doi.org/10.1002/etc.3540.
Balfour, D. L. & L. A. Smock, 1995. Distribution, age structure, and movements of the freshwater mussel Elliptio complanata (Mollusca: Unionidae) in a headwater stream. Journal of Freshwater Ecology 10(3): 255–268. https://doi.org/10.1080/02705060.1995.9663445.
Barnhart, M. C., 2006. Buckets of muckets: a compact system for rearing juvenile freshwater mussels. Aquaculture 254(1–4): 227–233. https://doi.org/10.1016/j.aquaculture.2005.08.028.
Bílý, M., O. Simon, V. Barák & V. Jahelková, 2021. Occurrence depth of juvenile freshwater pearl mussels (Margaritifera margaritifera) in a river bed tested by experimental mesh tubes. Hydrobiologia 848(12–13): 3127–3139. https://doi.org/10.1007/s10750-020-04298-8.
Bogan, A. E. & K. J. Roe, 2008. Freshwater bivalve (Unioniformes) diversity, systematics, and evolution: status and future directions. Journal of the North American Benthological Society 27(2): 349–369. https://doi.org/10.1899/07-069.1.
Böhm, M., N. I. Dewhurst-Richman, M. Seddon, S. E. H. Ledger, C. Albrecht, D. Allen, A. E. Bogan, J. Cordeiro, K. S. Cummings, A. Cuttelod, G. Darrigran, W. Darwall, Z. Fehér, C. Gibson, D. L. Graf, F. Köhler, M. Lopes-Lima, G. Pastorino, K. E. Perez, K. Smith, D. van Damme, M. V. Vinarski, T. von Proschwitz, T. von Rintelen, D. C. Aldridge, N. A. Aravind, P. B. Budha, C. Clavijo, D. Van Tu, O. Gargominy, M. Ghamizi, M. Haase, C. Hilton-Taylor, P. D. Johnson, Ü. Kebapçı, J. Lajtner, C. N. Lange, D. A. W. Lepitzki, A. Martínez-Ortí, E. A. Moorkens, E. Neubert, C. M. Pollock, V. Prié, C. Radea, R. Ramirez, M. A. Ramos, S. B. Santos, R. Slapnik, M. O. Son, A.-S. Stensgaard & B. Collen, 2020. The conservation status of the world’s freshwater molluscs. Hydrobiologia. https://doi.org/10.1007/s10750-020-04385-w.
Buddensiek, V., 1995. The culture of juvenile freshwater pearl mussels Margaritifera margaritifera L. in cages: a contribution to conservation programmes and the knowledge of habitat requirements. Biological Conservation 74(1): 33–40. https://doi.org/10.1016/0006-3207(95)00012-S.
Buddensiek, V., H. Engel, S. Fleischauer-Rössing & K. Wächtler, 1993. Studies on the chemistry of interstitial water taken from defined horizons in the fine sediments of bivalve habitats in several northern German lowland waters II: Microhabitats of Margaritifera margaritifera L., Unio crassus (Philipsson) and Unio tumidus Philipsson. Archiv Fur Hydrobiologie 127: 151–166. https://doi.org/10.1127/archiv-hydrobiol/127/1993/151.
Cai, S.-Y., 1986. Freshwater fossil bivalves from Maanshan Member of Ziliujing Formation in Weiyuan, Sichuan. Acta Palaeontologica Sinica 25: 560–567.
Černá, M., O. P. Simon, M. Bílý, K. Douda, B. Dort, M. Galová & M. Volfová, 2017. Within-river variation in growth and survival of juvenile freshwater pearl mussels assessed by in situ exposure methods. Hydrobiologia 810(1): 393–414. https://doi.org/10.1007/s10750-017-3236-x.
Denic, M., 2018. Comparison of two different field cages for semi-natural rearing of juvenile freshwater pearl mussels, Margaritifera margaritifera (Linnaeus, 1758) (Bivalvia: Unionoidea: Margaritiferidae). Folia Malacologica 26(4): 189–195. https://doi.org/10.12657/folmal.026.018.
Dimock, R. V. & A. H. Wright, 1993. Sensitivity of juvenile freshwater mussels to hypoxic, thermal and acid stress. Journal of the Elisha Mitchell Scientific Society 109(4): 183–192.
Eybe, T., F. Thielen, T. Bohn & B. Sures, 2013. The first millimetre—rearing juvenile freshwater pearl mussels (Margaritifera margaritifera L.) in plastic boxes. Aquatic Conservation: Marine and Freshwater Ecosystems. https://doi.org/10.1002/aqc.2384.
Ferreira-Rodríguez, N., Y. B. Akiyama, O. V. Aksenova, R. Araujo, M. Christopher Barnhart, Y. V. Bespalaya, A. E. Bogan, I. N. Bolotov, P. B. Budha, C. Clavijo, S. J. Clearwater, G. Darrigran, V. T. Do, K. Douda, E. Froufe, C. Gumpinger, L. Henrikson, C. L. Humphrey, N. A. Johnson, O. Klishko, M. W. Klunzinger, S. Kovitvadhi, U. Kovitvadhi, J. Lajtner, M. Lopes-Lima, E. A. Moorkens, S. Nagayama, K.-O. Nagel, M. Nakano, J. N. Negishi, P. Ondina, P. Oulasvirta, V. Prié, N. Riccardi, M. Rudzīte, F. Sheldon, R. Sousa, D. L. Strayer, M. Takeuchi, J. Taskinen, A. Teixeira, J. S. Tiemann, M. Urbańska, S. Varandas, M. V. Vinarski, B. J. Wicklow, T. Zając & C. C. Vaughn, 2019. Research priorities for freshwater mussel conservation assessment. Biological Conservation 231: 77–87. https://doi.org/10.1016/j.biocon.2019.01.002.
French, S. K. & J. D. Ackerman, 2014. Responses of newly settled juvenile mussels to bed shear stress: implications for dispersal. Freshwater Science 33(1): 46–55. https://doi.org/10.1086/674983.
Gatenby, C. M., R. J. Neves & B. C. Parker, 1996. Influence of sediment and algal food on cultured juvenile freshwater mussels. Journal of the North American Benthological Society 15: 597–609. https://doi.org/10.2307/1467810.
Gatenby, C. M., B. C. Parker & R. J. Neves, 1997. Growth and survival of juvenile rainbow mussels, Villosa iris (Lea, 1829) (Bivalvia: Unionidae), reared on algal diets and sediment. American Malacological Bulletin 14: 57–66.
Geist, J., 2010. Strategies for the conservation of endangered freshwater pearl mussels (Margaritifera margaritifera L.): a synthesis of conservation genetics and ecology. Hydrobiologia 644(1): 69–88. https://doi.org/10.1007/s10750-010-0190-2.
Geist, J. & K. Auerswald, 2007. Physicochemical stream bed characteristics and recruitment of the freshwater pearl mussel (Margaritifera margaritifera). Freshwater Biology 52(12): 2299–2316. https://doi.org/10.1111/j.1365-2427.2007.01812.x.
Gingras, M. K., S. G. Pemberton, S. Dashtgard & L. Dafoe, 2008. How fast do marine invertebrates burrow? Palaeogeography, Palaeoclimatology, Palaeoecology 270(3): 280–286. https://doi.org/10.1016/j.palaeo.2008.07.015.
Good, S. C., 1998. Freshwater bivalve fauna of the Late Triassic (Carnian-Norian) Chinle, Dockum, and Dolores Formations of the Southwest United States. In Newell, N. D., P. A. Johnston & J. W. Haggart (eds), Bivalves: An Eon of Evolution—Paleobiological Studies Honoring Norman D Newell University of Calgary Press, Calgary: 223–249.
Graf, D. L., 2013. Patterns of freshwater bivalve global diversity and the state of phylogenetic studies on the Unionoida, Sphaeriidae, and Cyrenidae. American Malacological Bulletin 31: 135–153. https://doi.org/10.4003/006.031.0106.
Graf, D. L. & K. S. Cummings, 2006. Palaeoheterodont diversity (Mollusca: Trigonioida + Unionoida): what we know and what we wish we knew about freshwater mussel evolution. Zoological Journal of the Linnean Society 148(3): 343–394. https://doi.org/10.1111/j.1096-3642.2006.00259.x.
Graf, D. L. & K. S. Cummings, 2007. Review of the systematics and global diversity of freshwater mussel species (Bivalvia: Unionoida). Journal of Molluscan Studies 73(4): 291–314. https://doi.org/10.1093/mollus/eym029.
Graf, D. L. & K. S. Cummings, 2021. A ‘big data’ approach to global freshwater mussel diversity (Bivalvia: Unionoida), with an updated checklist of genera and species. Journal of Molluscan Studies 87(1): eyaa034. https://doi.org/10.1093/mollus/eyaa034.
Graf, D. L. & K. S. Cummings, 2023. The Freshwater Mussels (Unionoida) of the World (and other less consequential bivalves). http://www.mussel-project.net/. Accessed 18 Apr 2023.
Haag, W. R., 2019. Reassessing enigmatic mussel declines in the United States. Freshwater Mollusk Biology and Conservation 22(2): 43. https://doi.org/10.31931/fmbc.v22i2.2019.43-60.
Haag, W. R. & J. D. Williams, 2014. Biodiversity on the brink: an assessment of conservation strategies for North American freshwater mussels. Hydrobiologia 735(1): 45–60. https://doi.org/10.1007/s10750-013-1524-7.
Haas, F., 1969. Superfamilia Unionacea. In Martens, R. & W. Hennig (eds), Das Tiererich Lieferung 88 Walter de Gruyter and Co., Berlin: 663.
Hastie, L. C. & P. J. Cosgrove, 2002. Intensive searching for mussels in a fast-flowing river: an estimation of sampling bias. Journal of Conchology 37: 309–316.
Hastie, L. C., E. C. Tarr, B. al-Mousawi & M. R. Young, 2010. Medium-term recruitment patterns in Scottish freshwater pearl mussel Margaritifera margaritifera populations. Endangered Species Research 11: 21–33. https://doi.org/10.3354/esr00262.
Hochwald, S. & G. Bauer, 1990. Untersuchungen zur Populationsökologie und Fortpflanzungsbiologie der Bachmuschel Unio crassus (Phil.) 1788. Schriftenreihe Bayerisches Landesamt Für Umweltschutz 97: 31–49.
Hornbach, D. J., V. J. Kurth & M. C. Hove, 2010. Variation in freshwater mussel shell cculpture and shape along a river gradient. The American Midland Naturalist 164(1): 22–36. https://doi.org/10.1674/0003-0031-164.1.22.
Hyvärinen, H., M. Saarinen-Valta, E. Mäenpää & J. Taskinen, 2021. Effect of substrate particle size on burrowing of the juvenile freshwater pearl mussel Margaritifera margaritifera. Hydrobiologia 848(5): 1137–1146. https://doi.org/10.1007/s10750-021-04522-z.
Hyvärinen, H. S. H., T. Sjönberg, T. J. Marjomäki & J. Taskinen, 2022. Effect of low dissolved oxygen on the viability of juvenile Margaritifera margaritifera: hypoxia tolerance ex situ. Aquatic Conservation: Marine and Freshwater Ecosystems 32(8): 1393–1400. https://doi.org/10.1002/aqc.3859.
Irmscher, P. & C. C. Vaughn, 2018. Effects of juvenile settling and drift rates on freshwater mussel dispersal. The American Midland Naturalist 180(2): 258–272. https://doi.org/10.1674/0003-0031-180.2.258.
Isely, F. B., 1911. Preliminary note on the ecology of the early juvenile life of the Unionidae. Biological Bulletin 20(2): 77–80. https://doi.org/10.2307/1536037.
Jakubik, B. & K. Lewandowski, 2011. Molluscs of the Krutynia River (Masurian Lakeland). Folia Malacologica 19(1): 19–29. https://doi.org/10.2478/v10125-011-0003-x.
James, M. R., 1985. Distribution, biomass and production of the freshwater mussel, Hyridella menziesi (Gray), in Lake Taupo, New Zealand. Freshwater Biology 15(3): 307–314. https://doi.org/10.1111/j.1365-2427.1985.tb00203.x.
Jones, J. W., R. A. Mair & R. J. Neves, 2005. Factors affecting survival and growth of juvenile freshwater mussels cultured in recirculating aquaculture systems. North American Journal of Aquaculture 67(3): 210–220. https://doi.org/10.1577/A04-055.1.
Jurkiewicz-Karnkowska, E., B. Jakubik & K. Lewandowski, 2017. Long-term changes in the malacofauna of the pond-type experimental lake Warniak (Mazurian Lakeland, North-Eastern Poland). Folia Malacologica 25(1): 27–36. https://doi.org/10.12657/folmal.025.005.
Kaestner, A., 1967. Invertebrate Zoology, Wiley, New York:
Kat, P. W., 1984. Parasitism and the Unionacea (Bivalvia). Biological Reviews 59(2): 189–207. https://doi.org/10.1111/j.1469-185X.1984.tb00407.x.
Kemble, N. E., J. M. Besser, J. Steevens & J. P. Hughes, 2020. Assessment of burrowing behavior of freshwater juvenile mussels in sediment. Freshwater Mollusk Biology and Conservation 23(2): 69–81. https://doi.org/10.31931/fmbc.v23i2.2020.69-81.
Kovitvadhi, U., K. Chatchavalvanich, N. V. Noparatnaraporn & J. Machado, 2001. Scanning electron microscopy of glochidia and juveniles of the freshwater mussel. Hyriopsis Myersiana. Invertebrate Reproduction & Development 40(2–3): 143–151. https://doi.org/10.1080/07924259.2001.9652714.
Kovitvadhi, S., U. Kovitvadhi, P. Sawangwong & J. Machado, 2007. Morphological development of the juvenile through to the adult in the freshwater pearl mussel, Hyriopsis (Limnoscapha) myersiana, under artificial culture. Invertebrate Reproduction & Development 50(4): 207–218. https://doi.org/10.1080/07924259.2007.9652248.
Lavictoire, L., A. D. Ramsey, E. A. Moorkens, G. Souch & M. C. Barnhart, 2018. Ontogeny of juvenile freshwater pearl mussels, Margaritifera margaritifera (Bivalvia: Margaritiferidae). PLoS ONE 13(3): e0193637. https://doi.org/10.1371/journal.pone.0193637.
Lefevre, G. & W. C. Curtis, 1908. Experiments in the artificial propagation of freshwater mussels. Bulletin of the Bureau of Fisheries 28: 616–626.
Lefevre, G. & W. C. Curtis, 1910. Reproduction and parasitism in the Unionidae. The Journal of Experimental Zoology 9: 79–115.
Lewandowski, K. B., 1996. Występowanie Dreissena polymorpha (Pall.) oraz małży z rodziny Unionidae w systemie rzeczno-jeziornym Krutyni (Pojezierze Mazurskie). In Hilbricht-Ilkowska, A. & R. J. Wiśniewski (eds), Funkcjonowanie systemów rzeczno-jeziornych w krajobrazie pojeziernym: rzeka Krutynia (Pojezierze Mazurskie) Zeszyty Naukowe Komitetu “Człowiek i środowisko” PAN, Warszawa: 173–185.
Lewandowski, K. & A. Kołodziejczyk, 2014. Long-term changes in the occurrence of unionid bivalves in a eutrophic lake. Folia Malacologica 22(4): 301–309. https://doi.org/10.12657/folmal.022.028.
Lima, P., M. L. Lima, U. Kovitvadhi, S. Kovitvadhi, C. Owen & J. Machado, 2012. A review on the “in vitro” culture of freshwater mussels (Unionoida). Hydrobiologia 691(1): 21–33. https://doi.org/10.1007/s10750-012-1078-0.
Lopes-Lima, M., L. E. Burlakova, A. Y. Karatayev, K. Mehler, M. Seddon & R. Sousa, 2018. Conservation of freshwater bivalves at the global scale: diversity, threats and research needs. Hydrobiologia 810(1): 1–14. https://doi.org/10.1007/s10750-017-3486-7.
Mair, R., 2018. Juvenile mussel culture. In Patterson, M., et al., (ed), Freshwater Mussel Propagation for Restoration Cambridge University Press, Cambridge: 159–222.
Mummert, A. K., R. J. Neves, T. J. Newcomb & D. S. Cherry, 2003. Sensitivity of juvenile freshwater mussels (Lampsilis fasciola, Villosa iris) to total and un-ionized ammonia. Environmental Toxicology and Chemistry 22(11): 2545–2553. https://doi.org/10.1897/02-341.
Neves, J. R. & J. C. Widlak, 1987. Habitat ecology of juvenile freshwater mussels (Bivalvia: Unionidae) in a headwater stream in Virginia. American Malacological Bulletin 5: 1–7.
Newton, T. J. & M. R. Bartsch, 2007. Lethal and sublethal effects of ammonia to juvenile Lampsilis mussels (Unionidae) in sediment and water-only exposures. Environmental Toxicology and Chemistry 26(10): 2057–2065. https://doi.org/10.1897/06-245r.1.
Newton, T. J., D. A. Woolnough & D. L. Strayer, 2008. Using landscape ecology to understand and manage freshwater mussel populations. Journal of the North American Benthological Society 27(2): 424–439. https://doi.org/10.1899/07-076.1
Nichols, S. J. & D. A. Wilcox, 1997. Burrowing saves Lake Erie clams. Nature 389: 921. https://doi.org/10.1038/40039.
O’Beirn, F. X., J. R. Neves & M. B. Steg, 1998. Survival and growth of juvenile freshwater mussels (Unionidae) in a recirculating aquaculture system. American Malacological Bulletin 14: 165–171.
Patterson, M. A., 2018. Biology of freshwater mussels. In Patterson, M., et al., (ed), Freshwater Mussel Propagation for Restoration Cambridge University Press, Cambridge: 25–57.
Pereira, D., M. C. D. Mansur, L. D. S. Duarte, A. S. de Oliveira, D. M. Pimpão, C. T. Callil, C. Ituarte, E. Parada, S. Peredo, G. Darrigran, F. Scarabino, C. Clavijo, G. Lara, I. C. Miyahira, M. T. R. Rodriguez & C. Lasso, 2014. Bivalve distribution in hydrographic regions in South America: historical overview and conservation. Hydrobiologia 735: 15–44. https://doi.org/10.1007/s10750-013-1639-x.
Perles, S. J., A. D. Christian & D. J. Berg, 2003. Vertical migration, orientation, aggregation, and fecundity of the freshwater mussel Lampsilis siliquoidea. Ohio Journal of Science 103: 73–78.
Piechocki, A., 1969. Obserwacje biologiczne nad małżami z rodziny Unionidae w rzece Grabi. Acta Hydrobiologia 11: 57–67.
Reid, R. G. B., R. F. McMahon, D. Ó. Foighil & R. Finnigan, 1992. Anterior inhalant currents and pedal feeding in bivalves. Veliger 35: 93–104.
Saarinen, M. & J. Taskinen, 2003. Burrowing and crawling behaviour of three species of Unionidae in Finland. Journal of Molluscan Studies 69(1): 81–86. https://doi.org/10.1093/mollus/69.1.81.
Savazzi, E., 1991. Burrowing sculptures as an example in functional morphology. Terra Nova 3(3): 242–250. https://doi.org/10.1111/j.1365-3121.1991.tb00141.x.
Savazzi, E. & P. Yao, 1992. Some morphological adaptations in freshwater bivalves. Lethaia 25: 195–209.
Schartum, E., S. Mortensen, K. Pittman & P. J. Jakobsen, 2017. From pedal to filter feeding: ctenidial organogenesis and implications for feeding in the postlarval freshwater pearl mussel Margaritifera margaritifera (Linnaeus, 1758). Journal of Molluscan Studies 83(1): 36–42. https://doi.org/10.1093/mollus/eyw037.
Schmidt, C. & R. Vandré, 2010. Ten years of experience in the rearing of young freshwater pearl mussels (Margaritifera margaritifera). Aquatic Conservation: Marine and Freshwater Ecosystems 20(7): 735–747. https://doi.org/10.1002/aqc.1150.
Schwalb, A. N. & J. D. Ackerman, 2011. Settling velocities of juvenile Lampsilini mussels (Mollusca:Unionidae): the influence of behavior. Journal of the North American Benthological Society 30(3): 702–709. https://doi.org/10.1899/11-023.1.
Schwalb, A. & M. T. Push, 2007. Horizontal and vertical movements of unionid mussels in a lowland river. Journal of the North American Benthological Society 26: 261–272. https://doi.org/10.1899/0887-3593(2007)26[261:HAVMOU]2.0.CO;2.
Seddon, M., C. Appleton, D. Van Damme & D. Graf, 2011. Freshwater molluscs of Africa: diversity, distribution, and conservation. In Darwall, W. R. T., K. G. Smith, D. J. Allen, R. A. Holland, I. J. Harrison & E. G. E. Brooks (eds) The diversity of life in African freshwaters: under water, under threat. An analysis of the status and distribution of freshwater species throughout mainland Africa. IUCN, Cambridge, UK and Gland, Switzerland, 92–119.
Seilacher, A., 1984. Constructional morphology of bivalves: evolutionary pathways in primary versus secondary soft-bottom dwellers. Palaeontology 27: 207–237.
Simon, O. P., I. Vaníčková, M. Bílý, K. Douda, H. Patzenhauerová, J. Hruška & A. Peltánová, 2015. The status of freshwater pearl mussel in the Czech Republic: several successfully rejuvenated populations but the absence of natural reproduction. Limnologica 50: 11–20. https://doi.org/10.1016/j.limno.2014.11.004.
Skawina, A., 2021. Evolutionary history of bivalves as parasites. In De Baets, K. & J. W. Huntley (eds), The Evolution and Fossil Record of Parasitism: Identification and Macroevolution of Parasites Springer, Cham: 153–207.
Skawina, A. & J. Dzik, 2011. Umbonal musculature and relationships of the Late Triassic filibranch unionoid bivalves. Zoological Journal of the Linnean Society 163: 863–883. https://doi.org/10.1111/j.1096-3642.2011.00728.x.
Sparks, B. L. & D. L. Strayer, 1998. Effects of low dissolved oxygen on Juvenile Elliptio complanata (Bivalvia:Unionidae). Journal of the North American Benthological Society 17: 129–134. https://doi.org/10.2307/1468057.
Stanley, S. M., 1969. Bivalve mollusk burrowing aided by discordant shell ornamentation. Science 166: 634–635.
Stone, N. M., R. Earll, A. Hodgson, J. G. Mather, J. Parker & F. R. Woodward, 1982. The distributions of three sympatric mussel species (Bivalvia: Unionidae) in Budworth Mere, Cheshire. Journal of Molluscan Studies 48: 266–274.
Strayer, D. L., 1981. Notes on the microhabitats of unionid mussels in some Michigan streams. The American Midland Naturalist 106: 411–415.
Strayer, D. L., 2008. Freshwater Mussel Ecology. A Multifactor Approach to Distribution and Abundance, University of California Press, Berkeley:
Strayer, D. L., J. A. Downing, W. R. Haag, T. L. King, J. B. Layzer, T. J. Newton & S. J. Nichols, 2004. Changing perspectives on pearly mussels, North America’s most imperiled animals. BioScience 54(5): 429–439. https://doi.org/10.1641/0006-3568(2004)054[0429:CPOPMN]2.0.CO;2.
Taskinen, J. & M. Saarinen, 2006. Burrowing behaviour affects Paraergasilus rylovi abundance in Anodonta piscinalis. Parasitology 133(5): 623–629. https://doi.org/10.1017/S0031182006001077.
Van Tu, D., T. Le Quang & E. B. Arthur, 2018. Freshwater mussels (Bivalvia: Unionida) of Vietnam: diversity, distribution, and conservation status. Freshwater Mollusk Biology and Conservation 21(1): 1–18. https://doi.org/10.31931/fmbc.v21i1.2018.1-18.
Vaughn, C. C. & C. C. Hakenkamp, 2001. The functional role of burrowing bivalves in freshwater ecosystems. Freshwater Biology 46: 1431–1446. https://doi.org/10.1046/J.1365-2427.2001.00771.X.
Wächtler, K., M. C. Dreher-Mansur & T. Richter, 2001. Larval types and early postlarval biology in naiads (Unionoida). In Bauer, G. & K. Wächtler (eds), Ecology and Evolution of the Freshwater Mussels Unionoida. Ecological Studies, Vol. 145. Springer, Berlin: 93–125.
Watters, G. T., 1994. Form and function of unionoidean shell sculpture and shape (Bivalvia). American Malacological Bulletin 11: 1–20.
Watters, G. T., 2001. The evolution of the Unionacea in North America, and its implications for the worldwide fauna. In Bauer, G. & K. Wächtler (eds), Ecology and Evolution of the Freshwater Mussels Unionoida Springer, Berlin: 281–307.
Watters, G. T., S. H. O’Dee & S. Chordas, 2001. Patterns of vertical migration in freshwater mussels (Bivalvia: Unionoida). Journal of Freshwater Ecology 16(4): 541–549. https://doi.org/10.1080/02705060.2001.9663845.
Yeager, M. M., D. S. Cherry & R. J. Neves, 1994. Feeding and burrowing behaviors of juvenile rainbow mussels, Villosa iris (Bivalvia: Unionidae). Journal of the North American Benthological Society 13(2): 217–222.
Zieritz, A., A. F. Sartori, A. E. Bogan & D. C. Aldridge, 2015. Reconstructing the evolution of umbonal sculptures in the Unionida. Journal of Zoological Systematics and Evolutionary Research 53(1): 76–86. https://doi.org/10.1111/jzs.12077.
Acknowledgements
The study was supported by the Polish Ministry of Science and Higher Education, through the Faculty of Biology, University of Warsaw Intramural Grant BW 14 0000/501/68–183125. I thank my supervisor Jerzy Dzik for his advice during the preparation of the first version of this manuscript. I am grateful to Krzysztof Lewandowski and Andrzej Piechocki for their advice in selecting habitats for the search for juvenile unionoids, and Reinhard Altmüller for providing small M. margaritifera shells. I thank to Arthur Bogan, Nile E. Kemble and Simon Schneider for supplying additional literature and discussions. Finally, I am very grateful to Karina Vanadzina for her advice and help in English editing and to Kenneth De Baets for his support and advice during the preparation of the final version of this work. The submitted version of this manuscript was prepared thanks to Project PARADIVE, I.3.4 Action of the Excellence Initiative—Research University Programme at the University of Warsaw. Funding body: Ministry of Education and Science, Poland.
Author information
Authors and Affiliations
Corresponding author
Ethics declarations
Conflict of interest
The author declares no conflict of interest.
Additional information
Handling editor: Manuel Lopes-Lima
Publisher's Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Supplementary Information
Below is the link to the electronic supplementary material.
10750_2023_5362_MOESM1_ESM.pptx
Online Resource 1. X-ray documentation: PowerPoint slideshow with original X-ray pictures of experimental aquariums (corresponding to Figure 2). (PPTX 5390 kb)
Online Resource 2. A. anatina 6 mm long - 16 minutes of crawling in an unusual manner (corresponding to Figure 4). (MP4 79713 kb)
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Skawina, A. X-rays and invisible sand: two new methods for designing burrowing behavioral experiments with juvenile unionoids. Hydrobiologia 851, 649–665 (2024). https://doi.org/10.1007/s10750-023-05362-9
Received:
Revised:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s10750-023-05362-9