The thick-shelled river mussel (Unio crassus) - host fish suitability and rearing of juvenile mussels Technical report LIFE10 NAT/SE/000046 ( The thick-shelled river mussel brings back life to rivers )
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UnioCrassusforLIFE (LIFE10 NAT/SE/000046) The thick-shelled river mussel (Unio crassus) - host fish suitability and rearing of juvenile mussels Technical report Within the project The thick-shelled river mussel brings back life to rivers LIFE10 NAT/SE/000046 Authors: Martin Österling & Lea D. Schneider Karlstad University Department of Environmental and Life Sciences - Biology 651 88 Karlstad Sweden E-mail: martin.osterling@kau.se 3
Introduction Unionoid mussels are one of the most threatened groups of organisms world-wide (Lydeard et al 2004). In Europe, Unio crassus is one of the most threatened mussel species and has been classified to have an unfavorable conservation status in the EC Habitats Directive. The decline of U. crassus is mainly due to habitat degradation and destruction negatively affecting the free-living mussels, and can also result in declines of host fish species in mussel rivers. The suitability and status of host fish populations are beginning to be explored in southern Europe (Stoeckl et al 2014), whereas in Sweden host fish suitability and distribution is poorly known. To be able to manage and design restoration measures, it is thus essential to improve the knowledge about host fish suitability all over the Swedish distribution of U. crassus. The LIFE+ Nature Demonstration and Best practice project The thick shelled river mussel brings life back to rivers aims to strengthen the conservation status of this highly endangered mussel species and to improve its ecological status in seven project rivers within the Swedish distribution area. The project is focused on habitat restoration measures such as re-meandering of channelized river parts, creation of bypass- channels, and improvement of sediment quality, which will favor both U. crassus and its host fish. To strengthen the conservation status of U. crassus, the restoration measures were intended to match the habitat requirements and the ecology of U. crassus and its host fish. However, identification of host fish used by U. crassus in the project rivers is a first step towards reaching this goal (Taeubert et al 2012). Therefore, a host fish inventory was performed in the mussels rivers, i.e. investigating the natural encystment rates of mussel larvae on wild fish. In parallel, host suitability tests were conducted in the projects laboratory aquaria facility and at the aquaria lab facility at Lund University. Furthermore, re-introduction of juvenile mussels and release of wild fish infested with mussel larvae aimed to re-introduce U. crassus in two project rivers, where U. crassus used to occur in high densities historically, but became extinct due to river channelization. Prior to mussel re-introduction, the original form and function of the rivers, which was broad meandering through the landscape, was re-established in two large scale habitat restorations, both part of the UC4LIFE project. To be able to discriminate among functional host fish species and to re-introduce juvenile mussels and mussel infested fish there is a need to a) know the timing of the reproduction season of U. crassus and its host fish species b) use non-destructive methods to collect gravid mussels such that they can release fit larvae that infest their host fish and c) use non-destructive and effective methods to test host fish suitability and d) to produce juvenile mussels for reintroduction. This includes the period from fishing in the rivers until juvenile mussels have been released from the fish. Lastly, there is also a need to know if the suitability of fish to mussels differs among strains of mussels and host fish, when choosing which combinations of mussels and fish strains to use in a rearing program. 4
Objectives The overall aim of this report is to present aspects essential for successful rearing activities of U. crassus, as well as tools for the practical implementation. The report therefore describes and evaluates the following aspects: 1) The tolerance of different host fish species to fishing, artificial infestation and maintenance in the lab until juvenile mussel collection. 2) The reproduction period of U. crassus, i.e. when gravid female mussels and host fish infested with mussel larvae are found, and methods to collect gravid mussels in the field and maintain them in the aquaria lab. 3) The host fish suitability, i.e. encystment rates and juvenile production of fish species inhabiting U. crassus rivers. In order to test the host fish suitability we started with a pilot investigation measuring encysted mussel larvae on the host fish. In the next step we performed investigations where we tested the suitability by collecting hatched juvenile mussels, which were later re-introduced into the rivers. Lastly, since re-introduction is an important part of the project and since fish are infested with mussel larvae from nonhome rivers, we also tested if the juvenile mussel hatching success differed depending on if fish were infested with mussels from its home (sympatric) or non-home (allopatric) rivers. 4) The survival of juvenile mussels in the laboratory facility after hatching from the fish. Methods 1. Fish handling 1.1 Fishing methods A variety of fishing methods was used to maximize the fish catch intended to cover the entire range of fish species in the rivers, thus catching all potential host fish available for U. crassus. Fishing was conducted at the project sites in River Klingavälsån, River Fyleån, River Bråån, River Tommarpsån, River Bräkneån, River Emån, River Storån, River Kilaån, and River Svärtaån during 2012-2015. We used electrofishing, four types of fyke nets (Fig. 1), minnow traps, stream nets and fly-fishing. The fyke nets were placed out in the rivers in the late afternoon, and the catch was checked in the following morning. Figure 1. Four types of fyke nets were used to catch fish. 5
Electrofishing was conducted using a 600 V LUGAB (L-600) (Fig. 2), both before and after any mussels started their annual glochidia release to catch wild fish with and without encapsulated mussel larvae. Stream nets were placed out in the rivers overnight. While electrofishing was conducted in wade through river stretches, fyke- and stream nets were placed out both at shallow and deeper river sites. Fish transport to the aquaria lab facility occurred with constant oxygen supply, where fish were kept separated according to species and stream of origin. Figure 2. Electrofishing was conducted by 2-3 persons. 1.2. Fish storage in the field To be able to catch fish over two to three days in rivers located far away from the laboratory aquaria facility, fish were stored in cages put out in the rivers overnight. Furthermore, selfconstructed fish cages were used as a method to re-introduce juvenile mussels. Here, fish, infested with mussel larvae were placed in the cages and held until the juveniles fell off the fish (Fig 3). Fish survival was almost 100 % when fish were maintained in the cages for about four weeks. Figure 3. Fish cages used for fish maintenance in the rivers. 6
1.3 Glochidia encystment check To ensure that fish caught in the rivers were not naturally infested with mussel larvae prior to our host suitability tests, one fourth of the fish captured were investigated for mussel larvae. Here, fish were anesthetized (Benzocaine) and their gills examined for glochidia using a microscope binocular (Leica MZ6). No fish died when carrying out this procedure in the field and in the laboratory, respectively. Also, mortality could not be related to the gill examination method over the three following weeks of fish maintenance in the aquarium lab. 1.4 Fish maintenance Fish were placed in 160 L glass aquaria in the aquarium laboratory and the water, which was exchanged weekly, was filtered using EHEIM classic 600. The fish were acclimatized successively to tap water and maintained until the glochidia infestation events. Fish were fed 4% of their body weight with frozen chironomids, gammarus or fish every 3 rd day. A day-night light cycle was installed in the aquarium laboratory. Water temperature was measured using loggers (Onset, Hobo pendant temp logger UA-002-64) and was set to 15 C. After fish infestation, fish were placed back to glass aquaria (year 2012) or in mussel hatchery tanks (40 L and 80 Liters), constructed after the descriptions of Eybe and Thielen (2010) and modified to our needs (Fig. 4). Survival and performance of fish were compared between the glass and the hatchery rearing tanks for each fish species. All fish were inspected for diseases. Figure 4. Mussel hatchery tanks in the aquarium laboratory at the Hemmestorps Mölla. To avoid fish to jump out, the tanks were covered with nets. The water was constantly pumped between the fish tank and a storing tank placed at a lower level (the floor). When the juvenile mussels had excysted from the host fish, they were transported via the pipes and caught in fine mesh in the storing tank. The sieve was emptied of juvenile mussels every day, which were then transferred to Petri Dishes. 7
2. Gravid mussels 2.1 Reproduction examination and gravid mussel period The timing of the U. crassus reproduction period was investigated in Tommarpsån (WGS84 55 33'7.7"N 14 8'40.7"E) through mussel gravity checks between April and August 2013, where adult mussels were examined non-destructively for gravidity. Tommarpsån holds good U. crassus populations with mussel densities up to 14 individuals m -2. Mussels were slightly opened (< 0.5 cm) using special opening tongs, whereupon their gills were visually inspected for swollen marsupia filled with glochidia (Österling, 2015). 2.2 Collection of gravid mussels and glochidia Gravid females from Bråån (WGS84: 55 48'3.5"N 13 38'33.1"E) and Tommarpsån (WGS84 55 33'7.7"N 14 8'40.7"E) were collected as their larvae were used for host fish suitability tests. Gravid mussels were transported to the aquarium lab facility at the Aquatic Ecology Unit of Lund University (2012-2013) and to the aquarium lab facility at Hemmestorps Mölla (2014-15). Individual mussels were kept in aerated 2 L stream water tanks, preferentially at 15 C. Water exchange and collection of released mussel larvae, using 50 µm sieves, was carried out daily. High numbers of glochidia were collected from the mussels from both rivers. Larval vitality, i.e. valve closure, was tested by the addition of a weak saline solution to a few glochidia. The larvae were stored in 4 C stream water until they were used for fish infestation. Vital larvae were then acclimatized to tap water. Glochidia from subsamples of 500 µl from the homogenized glochidia stock solutions were counted (Fig. 5). After glochidia collection, adult mother mussels were placed back in their streams of origin. Figure 4. U. crassus glochidia collected for fish infestation. 3. Parasitic stage and juvenile hatching 3.1 Pilot test parasitic stage To test whether fish from Klingavälsån and Fyleån function as potential hosts for future reintroduced mussels in the rivers, mussels from the same discharge area, Bråån and Tommarpsån, respectively, were used for fish infestation. The first investigation of fish suitability was performed in 2012 at Lund University. Eleven fish species were caught by electrofishing in 8
Klingavälsån and Fyleån, and were transported to the aquarium lab at Lund University. Fish were infested in separate infestation baths according to fish species and origin. Here, a volume of glochidia stock solution containing 350 glochidia per fish individual was added to each infestation bath. To improve the infestation success, the infestation was performed by constant gentle agitation of the oxygenated baths for 30 minutes (Fig. 6). Afterwards, fish were transferred to plastic and glass aquaria respectively. After 3 and 20 days post infestation fish were sacrificed (Benzocaine) and the gills dissected. The fish gills were then examined for mussel larvae and each larva counted using microscope binoculars. Figure 5. Fish mixed with glochidia larvae in an aerated infestation bath. 3.2 Parasitic stage and juvenile release summary of laboratory investigations 2013-2015 In total, nine infestation investigations were performed between 2013 and 2015. After the fish infestation events, fish were transferred to mussel hatchery tanks to be able to collect juveniles falling off their hosts. However, a subsample of fish individuals was sacrificed (Benzocaine) three days after the infestation to be able to evaluate early infestation rates. Here, the fish length [+/- 1 mm] and weight [+/- 0.1 g] were measured, whereupon the fish gills were dissected and the gill arches examined for mussel larvae using microscope binoculars. Likewise, in some of these investigations, a second fish sampling was conducted at 20 dpi. Using mussel hatchery tanks, juvenile mussels that fell off their fish hosts after successful metamorphosis could be collected in 50 µm nets adjacent to the fish tanks. The nets were checked for juvenile mussel on a daily basis and juveniles were transferred to Petri Dishes where they were examined for survival and counted using microscope binoculars. Living juveniles were measured in length [+/- 25 µm] and width [+/- 25 µm] using an ocular micrometer attached to a microscope. 3.3 Infestation of fish in a common garden experiment and juvenile collection In 2013, a so called common garden experiment was performed. The common garden approach is a design which is used to explore local adaptation patterns. Here, it was used to investigate if mussels are locally adapted to fish populations in its home streams, so called sympatric fish populations. Locally adapted mussel populations are, theoretically hypothesized to release higher 9
numbers of juvenile mussels from its sympatric fish populations than from its allopatric fish populations. This information is important when managing mussel streams, such as when reintroducing mussels or fish, or when restoring streams. Here, we used mussels and fish from two rivers. In River Bråån the minnow (P. phoxinus) is present, while in Tommarpsån, the minnow and bullhead (C. gobio) occur. Gravid mussels and fish were collected in late April 2013. The fish were brought to the aquaria laboratory at Lund University and were cross infested with their sympatric and allopatric mussels. Fish infestation with mussel larvae was carried out with fish separated according to fish and mussel origin. Therefore, up to 12 infestation baths were prepared. Up to 24 fish individuals were placed in every 8 L infestation bath. The infestation was carried out as described in section 3.1. No fish died during the infestation. The infestation was terminated by transferring the fish to the prepared fish tanks. The fish from each bath were divided into two groups of 12 individuals, whereupon each group was transferred to a randomly selected mussel hatchery tank. The water temperature was held constant at 16 C. Juvenile mussels falling off their host fish were collected and examined as described in section 3.2. 4 Duration of the parasitic stage and juvenile mussel survival All living juveniles collected were transferred to plastic boxes filled with 400 ml stream water. Juveniles from different streams and host origin/fish species were kept separated. Feeding (Nanno 3500, Shellfish diet 1800 and filtered detritus) and water exchange were performed weekly. Juvenile survival and size was investigated every second week. Results 1. Fish treatment 1.1 Fishing methods Electrofishing was generally an efficient method in terms of the time it took to catch the wanted number of fish individuals for the investigations in the aquaria lab. Also, electrofishing was more efficient in small than in large streams/rivers. Furthermore, electrofishing was more efficient for relatively stationary species/life stages such as young-of-the-year S. trutta, C. gobio and P. phoxinus than for more ephemeral species such as A. alburnus. The fishing efficiency was relatively low for benthic fish species such as L. lota. Fyke nets were generally less efficient compared to electrofishing, even if the fish survival was high. The fishing efficiency was lowest for P. phoxinus, G. aculeatus, B. barbatula and E. lucius. The fishing efficiency was relatively high for A. alburnus, L. lota, R. rutilus, P. fluviatilis, A. bjoerkna, T. tinca and S. erythrophthalmus. Stream nets was an efficient method for A. alburnus, and partly also for G. cernua and B. bjoerkna. The survival was however low, and were thus suitable for investigations when no alive fish was needed. Stream net fishing was not used regularly. Minnow traps had a very low fishing efficiency. Fish survival in the cages that were placed in the streams overnight was almost 100%, and thus considered a good method. 10
1.2 Fish maintenance The fish were tolerant to the infestation procedure, and no fish died during this treatment. Almost every fish species could be held in the two kinds of hatchery rearing aquaria (40 L and 80 L) and in the glass aquaria. The small hatcheries were less suitable than the large hatcheries for A. alburnus, S. trutta, R. rutilus and B. bjoerkna, probably because these fish species have high swimming abilities and need more space to thrive. Generally, when the space was too small, some fish species became injured or had diseases. P. phoxinus could be cultured in relatively high densities, whereas nets covering the fish tanks were needed to hinder them from jumping out of the tanks. C. gobio had an aggregated distribution in the glass aquaria, as fish were sitting on top of each other, and thus sometimes infected with fungal infections, particularly when water flow was too low. In the hatchery tanks the water was flowing, and C. gobio was not aggregated making them less susceptible to fungal infections on the skin, and could thus be kept in relatively high densities. A high swimming activity was observed for A. alburnus in the glass aquaria and hatchery tanks. Fish caught by means of fyke nets sometimes had fungal infections, as their skin is sensitive and therefore easily damaged. Electrofishing seems to be the more favorable fishing method for A. alburnus supposed to be held in aquaria tanks. Generally, survival of this fish species increased in the laboratory facility when the aquaria tank water was exchanged more frequently (> 2 times each week) and when the fish were treated in weak salt baths. R. rutilus and B. bjoerkna, sometimes were infected with fungus and white spot disease on their skin. Frozen chironomids were a good food provision for almost all fish species, particularly liked by P. phoxinus and A. alburnus. However, C. gobio, P. fluviatilis and G. cernua preferred frozen Gammarus, and E. lucius only ate fish as prey (Table 1 a-b). 11
Table 1a-b. Fishing efficiency and tolerance of fish species to catch and handling in the field and in the aquarium laboratory. 1 5 represent a grading, where 1 is the lowest efficiency or tolerance, and 5 is the highest efficiency or tolerance. Table 1a. Method Fishing Field Transport Food and feeding efficiency survival survival P. phoxinus; Eur. minnow Minnow traps, cages, e-fishing 1, 1, 4 5 5 5. Swim around and eat fast. Chironomidae C. gobio; Bullhead cages, e-fishing 1, 3 5 5 3. Long time to find food. Gammarus A. alburnus; Bleak E- and F-fishing, 4, 4, 4, 5 3 3, 4 5. Chironomidae cages, stream nets G. aculeatus; Three-spined cages, e-fishing 1,4 4. Chironomidae stickleback L. lota; Burbot cages, e-fishing 3, 3 5 5 3. Chironomidae S. trutta; Brown trout cages, e-fishing 1,4 5 Chironomidae R. rutilus; Roach cages, e-fishing 3, 4 4 4 5. Chironomidae P. fluviatilis; Perch cages, e-fishing 3, 4 4 5 5. Gammarus G. cernua; Ruffe cages, stream nets, 2, 3, 4 5 5 5. Gammarus e-fishing B. bjoerkna; White bream cages, stream nets, 3, 3-4, 4 4 4 5. Chironomidae e-fishing E. lucius; Pike cages, e-fishing 1, 4 5 3. Fish T. tinca; Tench cages, e-fishing 3, 4 5 5 5. Chironomidae V. vimba; Vimba bream cages, e-fishing 2, L. planeri; Brook lamprey cages, e-fishing 1, 3 5 S. erythrophthalmus; Common cages, e-fishing 3, 4 4 rudd B. barbatula; Stone loach cages, e-fishing 1, 3 5 3. Chironomidae Table 1b. Species P. phoxinus; Eur. minnow Glass aquaria Small hatchery Large hatchery General suitability Maintenance Comments 5 5 5 5 5 Can be cultured in higher densities than Cottus=better host; Can jump out with lids C. gobio; Bullhead 3 5 5 4-5 5 Glass aq: Low flow - close together - fungal infections. Suitability 4 gpaimmuno; slimy, water exchange 1x/week A. alburnus, Bleak 4 3; need space to swim 4 2-3 3 salt treatment, skin easily damaged, low densities, water exchange 2-3x /week G. aculeatus; Threespined stickleback 5 5 5 4 5 Low densities and not easy to catch; can escape e-fishing, L. lota; Burbot 3 5 5 Escape easily from net cages. Probably important locally pga high densities S. trutta; Brown trout 3 3 4 2 4 R. rutilus; Roach 3 2 3 1 (3) 3 Fungus; white spot disease; P. fluviatilis; Perch 4 5 5 2 5 May be in high densities and G. cernua; Ruffe 5 5 5 2 5 B. bjoerkna; White bream E. lucius; Pike 5 1 5 T. tinca; Tench 5 5 1 5 L. planeri; Brook lamprey B. barbatula; Stone loach 3 2 3 1 3 Fungus; white spot disease 4 5 2 5 12
2. Gravid mussels/reproduction season 2.1 Reproduction period of mussels Mussels became gravid between mid-april and early May, based on our investigations during 2012-2015, however particularly during 2012. The timing of gravidity was temperature dependent, such that mussels started to reproduce earlier in years when water temperatures rose early and later when water temperatures rose later in spring. The mussels reproduction period continued during the summer until late June to early July. 2.2 Naturally infested fish with mussel larvae The fish were found to be naturally infested with mussel larvae a couple of weeks after the first mussels became gravid. This pattern was consistent until the end of the reproduction season. Fish were thus found to have encysted mussel larvae on their gills around two weeks after the last gravid mussel was found. Fish were thus found to have encysted mussel larvae on their gills from late April and mid-may, until mid-july at latest. 3. Parasitic stage and juvenile release 3.1 Parasitic stage - pilot test Eleven of the fish species tested in the experimental pilot study, except for B. barbatula and R. rutilus, were found to be infested with mussel larvae 20 days post infestation. B. bjoerkna and E. lucius were both infested with larvae after 3 but not after 20 days. No individuals of A. alburnus and P. pungitus survived the 20 days, but both species were infested with larvae after 3 days. Table 2. Number of glochidia larvae on fish from Fyleån and Klingavälsån infested 3 and 20 days after infestation. Fyleån Klingavälsån 3 DPI Glochidia larvae fish -1 3 DPI 20 DPI Glochidia larvae fish -1 20 DPI 3 DPI Glochidia larvae fish -1 3 DPI 20 DPI Glochidia larvae fish -1 20 DPI B. bjoerkna - - 1.1 0 A. alburnus - - 38.5 - B. barbatula - - 0 0 C. gobio 37.2 4.2 54. 7 5.9 E. lucius - - 1 0 G. aculeatus 6 4.1 2.3 2 P. fluviatilis - - 0.33 0 P. phoxinus 12.2 25.3 - - P. pungitius - - 2 - R. rutilus - - 0 0 S. trutta 53 0.01 - - 13
3.2 Parasitic stage and juvenile release summary of laboratory investigations 2013-2015 Fish survival was generally high during the experiments, and only a few fish individuals of the 1313 fish individuals that were infested, died during the first three days. S. trutta had the lowest survival after three days, and was followed by A. alburnus, R. rutilus, P. phoxinus and C. gobio of which only a few individuals died. All individuals of the other eight species survived the first three days. After twenty days the survival was lowest for S. trutta, followed by A. alburnus, R. rutilus, P. phoxinus, G. aculeatus and C. gobio (Table 3). Five fish species were found to release juvenile mussels according to our investigations in the laboratory. P. phoxinus, C. gobio and G. aculeatus were the species with the highest juvenile hatching rates, whereas A. alburnus and P. fluviatilis had low juvenile hatching rates. In addition, S. trutta and B. barbatula were both found to be infested with mussel larvae after 20 days, although no juvenile mussels were released from these fish species. Generally, there was a high variation in juvenile excystment rates among different trials/experiments, albeit fish infestations were standardized. For example, juvenile excystment from C. gobio was 2.2 juveniles per gram fish in one experiment and 55.6 juveniles per gram fish in another experiment (Table 3). C. gobio represents, together with P. phoxinus, the fish species with the highest juvenile excystment rates. Table 3. Summary of the investigations 2012-2015 on fish survival, encystment rates, and juvenile excystment rates. Fish species Fish individuals Fish survival 3 DPI Fish survival 20 DPI Encystment rate 3 DPI Encystment rate 20 DPI Juvenile excystment rate (Nr. gram fish -1 ) A. alburnus 110 96.7 59.9 - - 0.71 (1) B. bjoerkna 26 100 100 0 (1) - 0 (2) B. barbatula 5 100 100 0 (1) 0.8 (1) 0 (1) C. gobio 569 98.7 97.4 56.0 (1) 6.0 (1) 2.2 55.6 (2) G. aculeatus 51 100 96.7 63.1 (2) - 0.3 106.5 (2) G. cernua 3 100 100 - - - L. lota 22 100 100 - - - L. idus 1 100 100 - - 0 P. fluviatilis 35 100 100 0 (1) - 0 0.50 (2) P. phoxinus 344 95.2 85.4 18.1 (1) 15.3 (1) 30.9 (1) R. rutilus 107 94.9 77.5 0 (1) - 0 (2) S. trutta 38 65.8 36.3 162.0 (1) 1.8 (1) 0 (1) T. tinca 2 100 100 - - - 3.2 Juvenile release - common garden experiment Different numbers of juveniles were excysted from different combinations of mussel and fish origins. For the minnows, the number of juvenile mussels that were released from the fish was highest from the sympatric fish mussel combination in one but not in the other mussel-fish combinations. For the bullheads, the numbers of released juvenile mussels was highest from the allopatric mussel-fish combination. Here, the juvenile mussels also held the highest survival rates and showed the highest growth rate after falling off the host fish. 14
There were no difference in encapsulated larvae on a host fish between 3 days post infestation and the juveniles successfully metamorphosed, i.e. no mussels seemed to die during parasitism. Also, the number of encysted mussel larvae was higher on larger host fish. 3.3 Juveniles excystment timing and survival The first juvenile mussels in the common garden experiment started to release from the host fish 15 days after the fish had been infested. Most juveniles were excysted between 17-18 and 26-27 days after infestation, and the last juveniles were excysted at 30-31 days after infestation. In total, 2412 juvenile mussels hatched from C. gobio and P. phoxinus together, which was 12 % of the initial number of glochidia used for fish infestation. Discussion 1. Fish storing and transport In summary, host fish suitability of 16 fish species were investigated in the aquarium facility. Generally, the results showed that depending on fish species and the aims of the investigations, different fishing methods can be used to effectively catch different fish species. Electrofishing was the method that almost always gave good results. At some river sites where it was too deep for wading, additional methods were however needed. Anyhow, when the goal is to catch relatively stationary but free-swimming fish species, electrofishing should be the preferred method. Fyke nets and stream nets were exceptionally successful for A. alburnus, occuring in high density in spring. However, the survival in the stream nets was low overnight, so if alive fish is needed, the nets must be emptied on a regular basis. Careful handling of fish is required as skin damages occur easily on the sensitive skin of A. alburnus. The survival of fish and mussels during transportation from the field to the aquaria laboratory facility was high just by keeping the water aerated and at the same temperatures as present in the stream. 2. Reproduction period of U. crassus Knowledge about the reproduction season of U. crassus is essential to match the timing of mussel rearing measures with available mussel brood for fish infestation. In Tommarpsån, mussels started to be gravid successively in mid-april, and up to 50 % of the individuals in a population were gravid in June. Fish can then be caught in mid-april, before mussels release their larvae. In this way, host suitability can be tested before any potential immune reactions have been established in the fish. The knowledge about the reproduction season of freshwater mussels can also be used to catch fish with larval encystment. Here, the natural larval encystment can be investigated and compared with encystment patterns between different fish species, which is useful for evaluating host suitability. Furthermore, the knowledge about the natural encystment period can also be used when fish are supposed to be transferred to the laboratory for juvenile mussel collection and reintroduction. Then, the fish should be taken from rivers with healthy mussel populations, i.e. with high mussel population densities. Regarding particularly small mussel populations, information about the reproduction season of freshwater mussels is essential as it informs about when not to disturb mussels. Likewise, 15
disturbance of fish carrying mussel larvae can be avoided when knowing the timing of mussel encystment, which seems a rather simple conservation measure for unionoids. 3. Pilot test parasitic stage According to our host suitability tests in 2013-2015, where juvenile mussels were collected after 17 days post infestation only, it may be reasonable to say that fish, that still held mussel larvae on their gills after 20 days post infestation in the pilot test, and kept at similar temperatures, can be considered functional hosts for U. crassus. This is supported by our findings of juvenile hatching in the common garden experiment, where the number of the juvenile mussels metamorphosed was as similar to the mussel infestation rates on fish sampled at three days post infestation. We therefore conclude that C. gobio, G. aculeatus, P. phoxinus and S. trutta are functional hosts for U. crassus in Klingavälsån, and C. gobio and G. aculeatus in Fyleån. Other studies investigating the duration of the parasitic stage of U. crassus reported that juveniles mussels hatched only later than 20 days post infestation (DPI). As mentioned previously, we collected juveniles earlier than this time point in our host suitability tests in 2013-2015. Therefore, fish that were not infested with mussel larvae at 20 DPI in our pilot study may still function as hosts, as juvenile mussels may then have detached from fish already, but could not be collected as fish were kept in glass aquaria. Anyhow, we conclude that also B. bjoerkna, A. alburnus, E. lucius and P. pungitus from Klingavälsån may be functional hosts for U. crassus (Table 2). 4. Summary of host fish suitability Based on all experiments and tests on host fish suitability, P. phoxinus, C. gobio and G. aculeatus were generally the most suitable host fish species not only when regarding infestation rates of fish individuals, but also on fish weight related numbers of larvae on the gills. A. alburnus and P. fluviatilis were also found to be functional host species for U. crassus, whereas S. trutta, B. bjoerkna, E. Lucius, B. barbatula and P. pungitus may only be considered potential hosts, depending on stream of origin and fish size. For instance, S. trutta seems to function as hosts in Fyleån, but only when fish are small, and much less quantitative than P. phoxinus or C. gobio. The results show that among all identified host fish species, P. phoxinus, C. gobio and G. aculeatus are the species to use when considering fish survival. G. aculeatus was however not found in high numbers, and the production of juvenile mussels from this fish species was relatively low. Nevertheless, it shall be remembered that local mussels may be adapted to local fish species. Such adaptations may however be lost when artificial infestation is undertaken in mussel breeding programs. 5. Common garden experiment juvenile release The results showed that different number of juveniles were excysted from different mussel-fish combinations. There was no consistent pattern of local adaptation for minnows, since the number of juvenile mussels that were released from the fish was highest from the sympatric fish mussel combination in one of the two mussel-fish combinations. For the bullheads, the numbers of released juvenile mussels was actually highest from the allopatric mussel-fish combination. Here, the juvenile mussels also had the highest survival and growth rate after hatched from the fish. 16
Another interesting finding was that successfully encapsulated mussel larvae did not die off during metamorphosis on P. phoxinus and C. gobio. This means that encapsulation rates after three days infestation resembled juvenile metamorphosis success. Therefore, host fish suitability can be determined just a few days after infestation. Furthermore, the numbers of encysted mussel larvae were higher on larger minnows and bullheads, which is beneficial for juvenile production by choosing the sizes of fish. In summary, the results show that testing host fish compatibility is essential prior to mussel reintroductions. For example, if one wants to produce high numbers of juveniles, tests can be carried out to find the most compatible mussel / fish combination. It can also be worthwhile to measure growth of juvenile mussel, since juveniles with higher growth rates may survive better. 6. Juvenile hatching timing The results show that the parasitic stage of U. crassus takes about two to four weeks depending on the temperature. However, preparations for juvenile collection should not be carried out later than two weeks after infestation when fish are held at 16 C water temperature. As described in other studies, lower water temperatures slow down the metamorphosis process. It is however not recommended to increase the temperature to speed up this process, as 15-16 C seems to be a critical threshold for fish diseases. In summary, the knowledge of the reproduction season of U. crassus and of host fish suitability in individual river systems gives important information about when gravid mussels can be collected in the field, and which fish species that are suitable for artificial infestation. In this way, repeated infestations may be achievable during one mussel reproduction season and increases the mussel conservation success at this early mussel life stage. References Eybe. T. and Thielen, F. 2010. Restauration des populations de moules perlières en Ardennes. Technical Report: Action A1 /D1 /F3 Mussel Rearing Station. LIFE05 NAT/L/000116. Fondation Hëllef fir d'natur. Heinerscheid, Luxembourg. Lydeard, C., Cowie, RH., Ponder, WF., Bogan, AE., Bouchet, P., Clark, SA., Cummings, KS., Frest, TJ., Gargominy, O., Herbert, DG., et al. 2004. The global decline of nonmarine molluscs. Bioscience, 54: 321 329. Österling, M. 2015. Timing, growth and proportion of spawners of the threatened unionoid mussel Margaritifera margaritifera: influence of water temperature, turbidity and mussel density. Aquatic Sciences, 77: 1-8. Stoeckl,K., Taeubert, J-E., Geist, J. 2014. Fish species composition and host fish density in streams of the thick-shelled river mussel (Unio crassus)- implications for conservation, Aquatic Conservation: Marine and Freshwater Ecosystems, 25, 2, 276. Taeubert, J-E., Gum, B. Geist, J. 2012. Host-specificity of the endangered thick-shelled river mussel (Unio crassus, Philipsson 1788) and implications for conservation. Aquatic Conservation: Marine and Freshwater Ecosystems, 22: 36-46. 17
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