The cultivation of the freshwater pearl mussel, Margaritifera margaritifera. A. McIvor and D. Aldridge. CCW Science Report No. 849

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1 The cultivation of the freshwater pearl mussel, Margaritifera margaritifera. A. McIvor and D. Aldridge CCW Science Report No. 849 CCGC/CCW 2008 You may reproduce this document free of charge for non-commercial and internal business purposes in any format or medium, provided that you do so accurately, acknowledging both the source and Countryside Council for Wales/Environment Agency copyright, and do not use it in a misleading context. This is a report of research commissioned by the Countryside Council for Wales and the Environment Agency. However, the views and recommendations presented in this report are not necessarily those of the Council or Agency and should not, therefore, be attributed to either party.

2 Report series: Report number: 849 CCW Science Report Publication date: October 2008 Contract number: FC Contractor: Contract Manager: Title: Author(s): Restrictions: St Catharine s College, University of Cambridge A.P. Fowles The cultivation of the freshwater pearl mussel, Margaritifera margaritifera A. McIvor & D. Aldridge None Distribution list (core): CCW HQ Library, Bangor CCW North Region Library, Mold CCW North Region Library, Bangor CCW S&E Region Library, Cardiff CCW S&E Region Library, Llandrindod CCW West Region Library, Aberystwyth Distribution list (others): CCW, Bangor Environment Agency National Library of Wales British Library Welsh Assembly Government Library Joint Nature Conservation Committee Library Scottish Natural Heritage Library Natural England Library A.P. Fowles, Senior Invertebrate Ecologist Dr J. Taylor (ten copies) Recommended citation for this volume: McIvor, A. & Aldridge, D The cultivation of the freshwater pearl mussel, Margaritifera margaritifera. CCW Contract Science Report No: 849, Countryside Council for Wales/Environment Agency, Bangor.

3 CONTENTS CONTENTS...iii LIST OF FIGURES...v LIST OF TABLES...vi CRYNODEB GWEITHREDOL...vii EXECUTIVE SUMMARY...ix 1 Introduction The parent mussels Collecting adult mussels Transporting adults Holding the adults and maintaining them in good health Holding tanks Holding mussels in a river or stream Holding densities Water quality, filtration and what to do about silt Flow and water depth Substrate Feeding mussels Affect of temperature on mussels Affect of light and photoperiod on mussels Handling mussels What to do if adults are in poor condition Reproduction and glochidial release Timing of spawning and glochidial release Ensuring that fertilization occurs Monitoring glochidial development Predicting glochidial release Glochidial release Artificially triggering glochidial release Possible reasons for mussels not releasing glochidia Fish infection and juvenile collection Method 1 Mussels and fish held together Method 2 - Mussel outflow enters fish tanks Method 3 - Glochidia placed in non-flowing fish tank Method 4 Laboratory scale infection of fish Summary of fish infection methods Estimating excystment dates of juveniles from fish Altering encystment duration Proportion of encysted juveniles surviving to excystment Collecting the juveniles after they have excysted from fish...19 iii

4 5 Rearing juveniles Releasing infected fish into rivers and streams Natural rearing in Lower Saxony, Germany with river restoration Buddensiek s cages Rearing mussels in an artificial stream or mill race Hruška s method Feeding juveniles Avoiding domestication of juveniles Growth and survival of juveniles in hatcheries Juvenile biology, ecology and behaviour Juvenile habitat Food When do pearl mussels reach reproductive age? Relationship of juveniles with adult mussels Relationships with other species Juvenile behaviour Feeding behaviour Substrate and use of byssal threads Pearl mussel rearing in Wales Mawddach Hatchery Adult survival Glochidial production Fish infection Juvenile collection Rearing juveniles Cynrig Hatchery Maerdy Hatchery Glasbury Future directions Rearing juveniles beyond 1 year Monitoring juvenile survival and growth Testing different conditions to optimise rearing conditions Improving propagation and rearing conditions up to 1 year More intensive rearing methods such as those trialled at Glasbury The wider perspective rearing mussels for return to rivers Other areas for future research Acknowledgements References...50 iv

5 LIST OF FIGURES Figure 1. Locations and tanks where adult mussels are held. (a) Mawddach hatchery; (b) Ballinderry hatchery; (c) Windermere research station; (d) the depot just in front of the trees in the River Lutter, Lower Saxony, Germany...4 Figure 2. Adult holding tank at Ballinderry showing direction of flow...9 Figure 3. Method used to infect fish by Roger Sweeting and Louise Miles of the FBA at Lake Windermere. Figure shows a tray containing gravel and mussels, and fish swimming in, over and around it in a flowing circular fish tank Figure 4. Adult mussels tank (round black tank at far left), first fish infection tanks (two of them) and in the distance the second and third infection tanks...15 Figure 5. Second and third fish infection tanks. The second tank is the round tank on the left and the third tank is the square tank on the right Figure 6. Recirculating fish tank used at Heinerscheid pearl mussel rearing station, Luxembourg, to keep fish warm to speed up encystment duration. The top container is the fish tank, and the bottom container contains a pump, which pumps water through a water filter and heater. The white tray is used to catch juveniles from the fish tank outflow...19 Figure 7. The River Lutter in Lower Saxony, Germany: two different types of silt trap used in river restoration, including a box silt trap set into the substrate (top left) and a very large artificial settling channel through which water flows before entering the main river (bottom left); a site where juvenile pearl mussels are doing well (right) Figure 8. The cages used by Buddensiek (1995) to hold juvenile mussels in rivers. Each cage consists of two covering plates (A) and a thicker 9mm central plate (C) into which 92 holes, each 6mm in diameter, have been drilled. The covering plates hold 2 pieces of mesh gauze in place (B), with mesh size 200µm. The juveniles are placed in the individual cells (D). Diagram from Buddensiek (1995) Figure 9. (a) The artificial stream where juveniles have been successfully reared, showing the wooden baffles that force the water to flow in a zigzag through the system; (b) the pool near the inflow which allows some sediment to drop out, also showing the large rocks which are used to create habitat diversity; (c) an island at the channel s edge and the netting which surrounds the whole channel; (d) the second stream, used by Preston, Keys and Roberts (2007) and recently altered to be more similar to the other raceway Figure 10. Diagrammatic view of the baffles showing how the water is forced to run through gaps that create stronger turbulent flows...28 Figure 11. Juvenile mussels reared by Alan Keys et al. in the artificial stream at Ballinderry Fish Hatchery. Mussels range in age from 5 to 10 years old, reaching 50mm length at 10 years...28 Figure 12. The circular tank in which the 9-10 year old juveniles are being held...29 Figure 13. The other tank holding 9-10 year old juveniles (left) and juvenile mussels visible in the sediment of this tank, showing how some of them are clustering around the larger rocks, which would presumably provide stability in times of high flow if they were in a river channel Figure 14. The siphons of mussels visible in the silt in this tank (15 in this picture)...30 Figure 15. (a) The mill race; (b) the fish frightening device, which is continually turning powered by the water flowing past; (c) one of the trays containing sediment and young mussels which is being held in the mill race...31 v

6 Figure 16. The mesh sieves and petri dishes (top right and left), plastic container used to hold juveniles (mid-left), cage (bottom left) and artificial ditch (bottom right, not yet operational in this picture) used to rear juvenile mussels according to Hruška s method, as seen at the Luxembourg rearing station Figure 17. Growth of juveniles reported from different hatcheries. Juveniles are assumed to be 400µm in length immediately after excysting from fish when not otherwise stated in source paper. Mean values are plotted where possible...35 Figure 18. Small juvenile mussel raceway, similar to the artificial stream used by Alan Keys et al. at Ballinderry Fish Hatchery in Northern Ireland Figure 19. Method for preparing sediment using sieves, to ensure that juveniles are a different size from all other sediment particles and can easily be refound by wet-sieving them out...46 LIST OF TABLES Table 1. Methods of fish infection listing advantages and disadvantages of each method...16 Table 2. Current and previous work culturing pearl mussels (excluding Welsh work)...22 Table 3. The advantages and disadvantages of the various culture methods available Table 4. Possible conditions for rearing 1+ juveniles in a raceway...43 vi

7 CRYNODEB GWEITHREDOL Mae r adroddiad hwn yn dwyn ynghyd y wybodaeth bresennol am fagu misglod perlog. Mae n cynnwys adrannau ar sut i hel a chadw misglod perlog aeddfed, sut i sicrhau cyflenwad da o glochidia (larfae misglod), sut i heintio pysgod a hel misglod ifanc, a sut i fagu misglod ifanc. Mae hefyd yn trafod bioleg, ecoleg ac ymddygiad misglod ifanc. Mae dwy adran ar y diwedd yn gwerthuso r gwaith sy n cael ei wneud ar hyn o bryd yng Nghymru ac yn awgrymu cyfeiriad i r gwaith hwn at y dyfodol. I ddechrau magu, argymhellir hel rhwng ugain a hanner cant o fisglod aeddfed, i sicrhau bod misglod gwrywaidd a benywaidd yn bresennol, i gynyddu amrywiaeth genetig y misglod ifanc ac fel nad oes raid i rai misglod atgenhedlu pob blwyddyn. Dylid cadw r misglod mewn tanciau llifo gyda dŵr o ansawdd da, crynodiad uchel o ocsigen a minimwm o laid yn y dŵr, mewn digon o ddwysedd i ganiatáu i r wyau ffrwythloni, a chyda system o dymheredd naturiol a ffotogyfnod. Dylid cynnwys sediment fel y medrant gladdu. O dan yr amodau hyn, gall misglod sydd mewn cyflwr gwael gael adferiad iechyd. Mae gollwng sbermau (gan y rhai gwryw i ffrwythloni r rhai benyw) fel arfer yn digwydd tua mis cyn i r glochidia gael eu rhyddhau rhwng canol a diwedd yr haf. I achub y blaen ar amseriad rhyddhau r glochidia, gall diwrnodau gradd roi syniad i rywun yn fras, neu gellir monitro r glochidia n ffurfio o fewn y misglod. Gellir hefyd defnyddio dyddiadau rhyddhau o ddeorfeydd cyfagos i ragfynegi r amseriad pan fo r gwahaniaeth amser rhwng gwahanol ddeorfeydd yn wybyddus. Mae glochidia n debygol o gael eu rhyddhau ar yr un pryd mewn gwahanol fisglod yn yr un tanc, fel y gellir trosglwyddo glochidia o nifer o fisglod i danciau pysgod i w heintio. Gellir cyflawni cyfraddau heintio uchel drwy ostwng lefel y dŵr, atal y llif, ac awyru r tanciau pysgod ar ôl ychwanegu r glochidia. Gellir amcanu parhad y systio gyda physgod drwy ddefnyddio diwrnodau gradd, ac mae r misglod ifanc yn debygol o ddeori o bysgod dros gyfnod o ddwy neu dair wythnos. Gellir eu hel drwy ddefnyddio sgrîn rwydog. Mae r dulliau a ddefnyddiwyd yn Neorfa Mawddach yng Nghymru i fagu misglod ifanc yn y flwyddyn gyntaf wedi bod yn hynod lwyddiannus, gyda mwy ohonynt yn byw yno na mewn gorsafoedd magu eraill. Mae tri dull o fagu misglod ifanc ar ôl y cyfnod hwn ar gael: eu rhoi mewn caetsys bychain yn ôl yn yr afonydd, fel a wneir yng Ngweriniaeth Tsiec a r Almaen; eu rhoi n syth mewn i swbstrad naturiol mewn nant artiffisial, fel yn Neorfa Bysgod Ballinderry yng Ngogledd Iwerddon; neu adfer yr afonydd fel bod misglod ifanc yn gallu byw ynddynt, fel sy n digwydd yn Afon Lutter, Sacsoni Isaf yn yr Almaen. Mae cryn risg o ddefnyddio caetsys, gyda nifer uchel o gaetsys a misglod ifanc yn cael eu colli, ond twf da. Mae rhoi r misglod ifanc mewn nant artiffisial yn gwneud monitro n anodd iawn hyd nes eu bod yn ddigon mawr i w gweld yn bedair neu n bump oed. Fodd bynnag, mae r dull hwn wedi bod yn llwyddiant mawr yng Ngogledd Iwerddon, gyda rhai misglod ifanc yn cyrraedd deg oed, a dyma sy n cael ei argymell ar gyfer deorfeydd yng Nghymru. Mae adfer afonydd hefyd yn angenrheidiol dros y tymor hir, a chyn gynted ag yr adferir afonydd Cymreig yn foddhaol i sicrhau y gall y misglod ifanc fyw, gellir gollwng pysgod wedi eu heintio i w canol i hadu r swbstrad â misglod. Wrth greu nant artiffisial i fagu misglod ifanc, dylid ystyried llif a hidlad y dŵr, y math o sediment a i ddyfnder, trefniant y bafflau i sicrhau amrywiaeth llif, a r system golau a thymheredd. Dylai r pethau hyn ddynwared yr amodau sydd i w cael mewn natur cymaint â phosibl. Gall defnyddio hidlydd dŵr leihau faint o ronynnau mân sy n mynd mewn i r system, a dylai r bafflau sicrhau bod peth o r sediment yn aros yn ddi-laid. Bydd llif uchel a gronynnau mawr o sediment (h.y. gro) yn sicrhau digon o ocsigen yn y sediment lle mae r misglod ifanc yn byw. vii

8 Mae angen i waith ar gyfer y dyfodol ganolbwyntio ar optimeiddio r amodau magu ar gyfer y misglod ifanc, ac ar ddulliau o adfer afonydd. Mae angen i fagu ac adfer ddigwydd ochr yn ochr, er mwyn gweld a ydyw r gwaith adfer yn creu amodau addas fel bo r misglod ifanc yn gallu byw. Mae cwestiynau pwysig yn cynnwys: pa mor hen y mae angen i r misglod ifanc fod i oroesi mewn afonydd sydd wedi eu hadfer i wahanol lefelau? Pa ffactorau y mae r misglod ifanc yn fwyaf sensitif iddynt, ac sy n eu hatal rhag byw mewn afonydd heb eu hadfer ar hyn o bryd? Gall gwaith i r dyfodol adeiladu ar gryfderau presennol y dull Cymreig; mae r rhain yn cynnwys amrywiaeth o ddulliau a ddefnyddir mewn gwahanol ddeorfeydd, cyfathrebu da rhwng y deorfeydd, dull pragmataidd, trosglwyddo arbenigedd mewn ffermio pysgod i ofalu am fisglod, monitro pysgod a misglod ifanc yn rheolaidd, ynghyd â diddordeb brwd ac angerdd tuag at y gwaith. Os bydd amser a gweithlu digonol yn caniatáu, mae r system yn Glasbury yn ddelfrydol ar gyfer arbrofi o r fath. Mae cadw cofnodion trylwyr hefyd yn hanfodol i ail-greu amodau yn y blynyddoedd i ddod; mae hefyd yn golygu y gellir cadw cofnod ysgrifenedig o ddulliau llwyddiannus fel y gall eraill eu dynwared. viii

9 EXECUTIVE SUMMARY This report brings together current knowledge about the rearing of freshwater pearl mussels. It includes sections on how to collect and hold adult pearl mussels, how to ensure a good supply of glochidia, how to infect fish and collect the juveniles, and how to rear juveniles on. It also covers juvenile biology, ecology and behaviour. Two final sections evaluate the current work in Wales and suggest future directions for this work. To begin rearing, it is recommended to collect between twenty and fifty adult mussels in order to ensure that both male and female mussels are present, to increase the genetic diversity of offspring and to allow for some mussels to not reproduce each year. Mussels should be held in flowing tanks with good water quality, high oxygen concentrations and a minimum of silt in the water, at densities that permit fertilization of eggs, and with a natural temperature regime and photoperiod. They should be provided with sediment in which they can bury. Mussels in poor condition can recover when held in this way. Spawning (release of sperm by males to fertilize females) usually occurs approximately a month before glochidial release in mid to late summer. To anticipate the timing of glochidial release, degree days can provide an approximate guide, or alternatively glochidial formation within mussels can be monitored. Release dates from nearby hatcheries can also be used as predictors when the lag between different hatcheries is known. Glochidial release is likely to occur simultaneously in different mussels in the same tank, allowing glochidia from several mussels to be transferred to fish tanks for infection. High infection rates can be achieved by lowering water levels, stopping the flow, and aerating the fish tanks after the glochidia have been added. Encystment duration on fish can be estimated using degree days, and juveniles are likely to excyst from fish over a period of two to three weeks. They can be collected using a mesh screen. The methods used at Mawddach Hatchery in Wales to rear juveniles for the first year have been highly successful, with better survival rates than those reported from other rearing stations. Three methods of rearing juveniles beyond this stage are available: placing them in tiny cages back in rivers, as used in the Czech Republic and Germany; placing them directly into natural substrate in an artificial stream, as in Ballinderry Fish Hatchery, Northern Ireland; or restoring rivers such that juveniles can survive, as in the River Lutter, Lower Saxony, Germany. The use of cages is risky, with high loss rates both of cages and juveniles, but good growth. Placing juveniles in an artificial stream makes monitoring very difficult until juveniles are large enough to see at four to five years old. However this method has been highly successfully in Northern Ireland, with some juveniles reaching ten years of age, and is recommended for the Welsh hatcheries. Restoring rivers is also necessary over the longer term, and as soon as Welsh rivers have been restored sufficiently for juvenile survival, infected fish can be released into them to seed the substrate with mussels. When constructing an artificial stream for juvenile rearing, consideration should be given to flow rates, filtration of water, sediment type and depth, arrangement of baffles to ensure a variety of flows, and the light and temperature regime. As much as possible, these should mimic conditions found in nature. Use of a water filter can reduce the amount of fine particles entering the system, and the baffles should ensure that some areas of sediment remain silt-free. High flows and large sediment particles (i.e. gravel) will ensure good oxygenation of the sediment in which the juveniles live. Future work needs to focus both on optimising rearing conditions for juveniles and on river restoration methods. Rearing and restoration need to take place side by side, in order to test ix

10 whether the restoration work is creating suitable conditions for juvenile survival. Important questions include: how old do juveniles need to be in order to survive in rivers with different levels of restoration? What factors are juveniles most sensitive to, which are currently preventing juveniles surviving in unrestored rivers? Future work can build on the current strengths of the Welsh approach; these include a diversity of methods used in the different hatcheries, good communication between the hatcheries, a pragmatic approach, expertise in fish husbandry transferred across to caring for mussels, regular monitoring of fish and juveniles, and the keen interest and passion of those involved in the work. If time and man-power allow, a more experimental approach can contribute to optimising rearing conditions, and the set-up at Glasbury is ideal for such experimentation. Good record keeping is essential in order to recreate conditions in future years; it also allows successful methods to be written up for others to replicate. x

11 1 INTRODUCTION The freshwater pearl mussel, Margaritifera margaritifera, is listed as Endangered on the IUCN Red List (IUCN, 2008) and is included in Annex II and V of the EC Habitats & Species Directive. It has a holarctic range, with populations in Europe, Russia, and North America. However it is declining rapidly and is threatened with extinction across much of its range (Young et al., 2001). Major threats include sedimentation, pollution, pearl fishing, loss of the host fish species, eutrophication, habitat destruction, river bed compaction and surrounding land use changes (Bauer et al., 1980; Bauer, 1988; Beasley and Roberts, 1996; Cosgrove et al., 2000; Young et al., 2001; Roberts, 2005; Hartmut and Gerstmann, 2007; Jensen, 2007). Early juvenile life is likely to be the part of the life cycle that is most vulnerable to threats, particularly sedimentation and pollution, because juveniles live interstitially within the sediment and therefore have different and more specific habitat requirements than adults. It is estimated that large populations with active recruitment are present in less than 50 rivers in Canada, northwest Russia, northeast Scandinavia, Scotland and a small number of sites in Austria, Bavaria, the Czech Republic and Eire (Young et al., 2001; Skinner et al., 2003). Many populations exist only as relic populations, consisting of older adults but with no evidence of recruitment of young mussels to the population for some years (Bauer, 1980; Bauer et al, 1980; Beasley et al., 1998; Rudzite, 2004; Cosgrove et al., 2000). For these populations, the culture of juveniles is a priority in order to ensure that young individuals survive until habitat restoration can allow reproduction in situ. Conservation programmes are now underway in several European countries to build up captive-bred stocks of juveniles for subsequent release. Ideally the juveniles will be maintained in captivity until they are old enough to filter-feed and are no longer dependent upon interstitial water quality. In the UK, pearl-fishing was one of the major threats to pearl mussel populations but this has substantially reduced since the pearl mussel was given full protection on Schedule 5 of the Wildlife & Countryside Act in The main problem now is the lack of recruitment in the majority of British rivers, resulting from the inability of juvenile mussels to survive their first three to five years buried within the interstitial gravels of the riverbed. Siltation, leading to anaerobic conditions, is believed to be the primary cause, and only in some Scottish rivers are pearl mussel populations still recruiting. In response to this, the Countryside Council for Wales and Environment Agency Wales have had a conservation strategy in place since November 2004, and mussels from seven Welsh rivers are currently being held in EAW fish hatcheries. Breeding has taken place and many thousands of juvenile mussels are being reared from these populations, with a small number reaching more than two years of age. However, little experimental research has been undertaken to determine optimal conditions for rearing juveniles in Wales. Mortality rates are high and, given the plight of the freshwater pearl mussel, the objective of captive breeding should be to rear as many juveniles as possible to an age where they are able to survive in their native streams and rivers or to reproduce in captivity until rivers have been restored. This report aims to bring together recent work on freshwater pearl mussel rearing in order to suggest ways to maintain high survival and growth of juveniles in the Welsh hatcheries. Sections 2 to 5 focus on the different phases of the propagation and rearing process, and sections 6, 7 and 8 cover juvenile biology, the rearing work in Wales, and recommended future directions. 1

12 2 THE PARENT MUSSELS The first stage in any rearing attempt is to locate the adult mussels that will be the brood stock and will provide the glochidia to infect the fish. Generally people have brought these adults into a holding facility where they are provided with good quality water and from where the glochidia can be used to infect the fish. 2.1 Collecting adult mussels Often adult mussels have been collected from a waterway because their decline was so fast that taking them into captivity provided the only hope of keeping them alive. In this case all mussels are often collected. Where there is not an imminent threat or where the mussel populations are declining only slowly, it may be sensible to leave some mussels in situ, and take only enough for the propagation work. It is recommended to collect between 20 and 50 mussels to act as parents, in order to ensure that both male and female mussels are collected (the sex ratio is usually 1:1; Bauer, 1987a), to allow for some mortality after collection (there is often a low level of mortality following transit), to ensure a large enough gene pool for future generations reared from the mussels, and to allow for the fact that not all mussels will reproduce in all years. Both Bauer (1987a) and Dick Neves (pers. comm.) have found that each year some mussels do not reproduce and this is probably normal even for mussels in their native rivers and steams. To increase the genetic diversity of mussels collected, it is sensible to collect a few mussels from several different areas in a single stream. However if there are obvious differences in the ecology or morphology of upstream and downstream populations, it is wise to breed these separately as they could be quite different genetically, and interbreeding them could cause outbreeding depression (Jones et al., 2006). To increase the likelihood of the remaining mussels in the river breeding, it may be sensible to collect them up and place them together in one location in the river, preferably in or near a place where suitable host fish are known to congregate and where there is good habitat for juveniles. The distances between the mussels should be short to increase the chance that sperm will reach females, and if males are placed slightly upstream of females, this should increase fertilization of eggs. Males and females can be identified using tongs to gently prise open the shell; females usually have thicker gills than males, and this is most obvious during the gravid period when females are brooding glochidia. Österling et al. (2008) found that in Swedish streams, the glochidial load on fish was related to adult mussel density; this implies that creating areas with high densities of mussels will increase fish infection rates and the number of juveniles excysting into nearby habitat. In order to increase genetic diversity in juveniles and to maintain good health in adults, the parent mussels can be cycled between the captive population and the river population (Jones et al., 2006; Dick Neves, pers. comm.). This is only sensible if the river populations are doing well, and the main reason for the decline in populations is that juvenile habitat is no longer available. As the conditions required to keep adult mussels in good health are poorly known (in particular what to feed them), this may help to maintain healthy parent mussels for glochidial production. 2

13 2.2 Transporting adults Many species of freshwater mussels are considered to be tolerant to handling and disturbance during transportation (Cope and Waller, 1995). However inevitably it will result in a certain level of stress for the animals, and if they are already in poor condition, may cause some mortality. It is recommended that mussels should be moved quickly and carefully, keeping them moist during transit, and holding them out of water for the shortest time possible (Waller et al., 1995). Good planning will ensure that mussels can be moved directly from the river to their new home. Holding them in buckets of water or wrapped in wet tissue paper or towels is easiest. Extremes of temperature should be avoided (Dunn and Sietman, 1997), although some people recommend cooling them during transport (Bishop et al., 2007). The ideal temperature is probably the water temperature of the river that they have come from. It is advisable not to move mussels during their gravid period as they are likely to eject eggs or glochidia from their gills due to the reduced oxygen concentrations (Aldridge and McIvor, 2003). 2.3 Holding the adults and maintaining them in good health In order to maintain adults in good health, they need flowing water, with good water quality, high oxygen concentrations, an adequate supply of organic particulate matter for food, and low silt levels. They need a substrate in which to bury themselves, and suitable annual temperature regimes for them to breed. A natural photoperiod may also be necessary for the breeding cycle. These requirements are discussed in more detail below. 2.4 Holding tanks Holding tanks vary between hatcheries, with equally good results. Rectangular tanks with linear flow work well in Mawddach Hatchery in Wales (Figure 1a), while very shallow circular tanks with inwardly spiralling flow provide a good supply of glochidia in Ballinderry Fish Hatchery in Ireland (Alan Keys, pers. comm., Figure 1b). Trays of gravel placed at the bottom of flowing circular fish tanks are in use at Windermere research station (Louise Miles and Roger Sweeting, pers. comm., Figure 1c). In all these places, adult survival is high with good glochidial production. An important issue is ensuring a good water supply even when things go wrong, such as a pipe getting blocked or a pollution event upstream. Alarm systems can be used to ensure this, either measuring flow or dissolved oxygen. Using more than one water supply to the adult tanks can also provide some insurance (Alan Keys has 5 pipes leading into the adult tank at Ballinderry Fish Hatchery). Having the outflow a little higher than the mussels also ensures that even if inflow stops, the mussels will not be out of the water. Mussels are able to close up for periods, so can survive short intervals with poor water quality or no water supply. However the time they are able to survive will depend on the temperature and time of year; during the heat of summer, it will be much reduced. 2.5 Holding mussels in a river or stream Another possibility is to hold the parent mussels in a river or stream where they are known to do well and are very unlikely to get washed away or be affected by pollution, and then to bring them in only when they are expected to release glochidia, as Altmüller and Dettmer do in the River Lutter, Lower Saxony, Germany (Rainer Dettmer, pers. comm., Figure 1d). This has the advantage of there being no need for holding tanks for mussels for most of the year. They judge when glochidia are about to be released by monitoring nearby populations that are 3

14 known to produce glochidia a week or two before the ones in the depot stream. The mussels are taken to a tank to release their glochidia, which are then collected and used to infect wild fish caught by electrofishing and immediately released back into the stream. This approach is very time and cost effective, requiring no permanent tanks or husbandry of the mussels and fish. However it is only feasible in rivers where the habitat has been restored sufficiently for fish and juvenile mussel survival, and where there are few risks to the adult mussels. Figure 1. Locations and tanks where adult mussels are held. (a) Mawddach hatchery; (b) Ballinderry hatchery; (c) Windermere research station; (d) the depot just in front of the trees in the River Lutter, Lower Saxony, Germany. 2.6 Holding densities Holding densities can be quite high. Dick Neves recommends a density of at least 12 mussels per square metre to ensure fertilization of eggs by sperm. At most hatcheries, mussels are held at higher densities than this with good survival. It is likely that high flows are needed to hold mussels at high densities in order to ensure a good supply of oxygen and food to all mussels. However if only a few mussels are available, mussels should still be able to reproduce. Bauer (1987a) found that fecundity (i.e. number of eggs produced) was independent of density of mussels in Bavarian streams, and that pearl mussels became hermaphrodites when present at low densities. 4

15 2.7 Water quality, filtration and what to do about silt Good water quality is considered essential for the long-term holding of mussels. In particular, high oxygen concentrations, low silt concentrations, low nutrient levels and very low levels of pollutants are required. Acidity and calcium carbonate concentrations should be kept as close as possible to the levels in the streams where the adults were taken from. Oligotrophic waters, with a ph less than 7 to 7.5 and low overall conductivity are generally considered to be most suitable. Skinner et al. (2003) review this topic in detail. When high levels of silt are entering the system, the use of a high capacity filter is recommended. High silt levels are likely to cause mussels to close for extensive periods and to reduce their filtering efficiency. There are some concerns about whether such a filter will remove the mussels food. To avoid this problem, the filter could be used only during and after spates when the silt load is highest, and at other times mussels could receive unfiltered water (this is only feasible if the silt load is generally very low except during spates). Some groups have also opted for using a recirculating system with a suitable biological filter to maintain high water quality, which they only connect up during periods of high silt loads. If silt begins to accumulate on the surface of mussels, it is a good idea to wash it off the mussels using a hose or brush it off gently so it gets carried away in the flow. 2.8 Flow and water depth High flows are recommended as these are closest to what mussels would experience in nature. They will also ensure an ample supply of oxygen and food and should reduce the build up of silt on the substrate. Hastie et al. (2000) estimate that optimum water depths are m and optimum current velocities are m/s at intermediate water levels, based on computed habitat suitability curves. However, much higher flows during spates help to keep the substrate clear of silt in rivers and streams, and artificial increases in flow from time to time might serve the same purpose for mussels held in hatcheries. 2.9 Substrate Mussels prefer to bury, and sufficient substrate should be provided for them to do this. Approximately 20cm depth of mixed gravel should be adequate, and it is sensible to use the same or similar substrate to that found in the river where the mussels were taken from, possibly providing a mix of substrates so that mussels can choose their preferred substrate type Feeding mussels Dick Neves (pers. comm.) and others working on North American species and other European species consider it necessary to feed most species of mussels; possibly pearl mussels will receive enough food if the water comes from a river and contains some organic particulate matter of small particle size, but research is needed to confirm this. The size of particles ingested by pearl mussels are not known, but are likely to be larger than 0.8µm as this is what other mussel species are known to filter out and ingest (Jørgensen et al., 1984, Vanderploeg et al., 1995). With the current trend to install filter systems that remove particles larger than 30µm, it is important to check that food particles are still getting through. The diet of adult freshwater mussels is poorly known and it has generally been assumed that they feed primarily on phytoplankton, in the same way as marine bivalves (Ziuganov et al., 5

16 1994; Gosling, 2003). A variety of particles have been found in their guts, such as phytoplankton, detrital particles and small animals such as rotifers and crustaceans. However Nichols and Garling (2000) showed that freshwater mussels gained most of their carbon from bacterial sources. It is possible that mussels gain their nutrition from the bacterial films that cover other particles such as detritus. This is most likely to be the case for pearl mussels as they live in the upper parts of catchments where phytoplankton and zooplankton will only be present at low densities Affect of temperature on mussels Ensuring a natural temperature regime is important, as all mussel processes are temperature regulated, and the timing of spawning and duration of glochidial development are probably controlled by temperature. Therefore constant temperature rooms and places which are unusually hot or cold are likely to have negative effects on these processes, potentially inhibiting reproduction Affect of light and photoperiod on mussels It is unknown how important light and photoperiod are for mussels. However it seems likely that these form an important cue for seasonal cycles such as breeding, alongside water temperature. There is no doubt that mussels are sensitive to light, as they respond to passing shadows by temporarily closing their valves (pers. obs.). Gethin Thomas (pers. comm.) has also shown that mussels spend more time open and presumably filtering at night. It is recommended to use a natural photoperiod, either by having mussels in a naturally lit space or by putting lights on timers and changing the settings with the seasons. Natural light is preferable in order to keep conditions as close as possible to river conditions. The amount of shading is also important, as it often noted that pearl mussels are found only in shaded areas of rivers (e.g. Gittings et al., 1998; Álvarez-Claudio et al., 2000; Skinner et al., 2003; Outeiro et al., 2008). Direct sunlight with low to medium flows is likely to result in heating of water and mussels, which will affect the timing of spawning and glochidial production. Sunlight also encourages filamentous algal growth, which can form algal mats that will smother mussels (Skinner et al., 2003) Handling mussels While most people avoid handling their pearl mussels as much as possible, they may be quite resilient to some handling, as other mussel species are. The use of tongs to look inside mussels to check their sex and to monitor glochidial development is probably not too invasive providing it is done gently and mussels are not opened too wide. However repeated handling or opening of mussels with tongs should be avoided What to do if adults are in poor condition Adult mussels brought in from the river may be in poor condition, and may continue to gape open even when touched (normally healthy mussels immediately close on contact). Mussels brought into hatcheries in poor condition have been known to recover: Roger Sweeting and Louise Miles (pers. comm.) received pearl mussels from several English rivers, and many of these were showing signs of poor health. The mussels were placed in large flowing circular fish tanks at the FBA research station by Lake Windermere, with well-oxygenated water from the lake itself. The mussels now look healthy and are producing glochidia, which are infecting 6

17 fish. However Keith Scriven (pers. comm.) reports that if mussels have been held in hatcheries for extended periods, and then show signs of ill-health, they rarely recover; how to help these mussels recover or to avoid them losing condition is an important area for future research. Possible reasons for mussels being in poor condition include starvation, pollution, sickness and old age. Starvation is a likely cause, for example if there have been high silt loads or poor water quality causing mussels to keep their valves shut for extended periods. High silt loads in the water may also interfere with the mussels filtering apparatus, clogging up the gills, preventing the collection of food and possibly reducing oxygen transport. Alternatively land use changes in streams and rivers may have reduced the amount of suitable organic particulate matter entering a stream or river (e.g. if different species of trees have been planted or forests have been removed). If mussels have been exposed to high levels of pollutants such as DDT, cadmium and other heavy metals, these can accumulate in their tissues possibly affecting calcium metabolism, causing brittle shells and other mal-effects (Hartmut and Gerstmann, 2007). Pollutants such as nitrates and phosphates are also known to affect mussel survival (Bauer, 1983, 1988); Bauer (1988) found a significant correlation between mussel mortality and nitrate concentration, with 20-80% mortality at NO 3 -N concentrations of 1-3 parts per million. It is unknown if mussels can recover from long-term exposure to pollutants, and this requires further study. If many mussels die after being taken into a hatchery, this could be caused by disease. In such cases, it is wise to remove mussels from tanks as soon as they show signs of ill-health, to reduce the chance of other mussels becoming infected. They can be placed in a quarantine tank, and potentially given extra food or other measures taken to increase their health. As few such mussels recover, it is a good time to experiment with anything that might help them to do so, as doing nothing will probably result in their loss. As mussels can live for more than 100 years, it is unlikely that old age is a cause of poor health in whole populations as most populations should contain individuals of different ages. For other mussel species, Dick Neves recommends feeding them to help them recover (pers. comm.). Algae are usually the preferred food, although other foods have been used, such as blended lettuce leaves (Kelly Poole, pers. comm.). However most people believe that this is not necessary for pearl mussels providing there is some natural food in the water. Some mortality usually occurs after mussels have been moved, but mortality should then settle down to a very low level. If it remains higher than one individual per several months, it is worth thinking about changing the holding conditions or moving the mussels to another site. 7

18 3 REPRODUCTION AND GLOCHIDIAL RELEASE 3.1 Timing of spawning and glochidial release Spawning, which is the release of sperm by males to fertilize the eggs held in the females gills, usually occurs in early to mid-summer, followed by glochidial release (sometimes called spatting) in mid to late summer (Hastie and Young, 2003). The timing of both is related to temperature. It is not possible to observe spawning in rivers. As a proxy, people often use the first date that glochidia are seen forming in the gills of females (Hastie and Young, 2003). The glochidia are visible as a cream coloured mass within the translucent brown gill (Young and Williams, 1983). Spawning presumably occurs over several weeks, as the number of mussels brooding glochidia increases gradually until glochidial release (Hastie and Young, 2003). In mussel tanks, spawning can be observed when mussels are seen to release white clouds of sperm into the water column. Spawning may be associated with behaviours such as mussels moving closer together or burying themselves less deeply (as seen in other species; Amyot and Downing, 1998; Schwalb and Pusch, 2007). Glochidial release can be detected in rivers by monitoring plankton nets that trap the glochidia (Hastie and Young, 2003) or by monitoring glochidia on fish e.g. in hatcheries receiving the water. In mussel tanks glochidia are often visible when mussels release pink or white clumps of glochidia (conglutinates), which can be seen resting on the substrate next to the mussels. Glochidial release is often highly synchronised in rivers, with the majority of glochidia being released over one or two days (Hastie and Young, 2003) although longer release periods have been observed in hatcheries. Young and Williams (1984a) found that glochidial release occurred more during the day, with twice as many glochidia being released as overnight. This is presumably triggered either by daylight or small changes in water temperature as the sun heats the water. The timings of spawning and glochidial release vary in different rivers, in different countries and in different years. In Scottish rivers spawning occurred in June and July in 1997, and west coast populations were approximately one month earlier than east coast populations (Hastie and Young, 2003). In Austria, spawning occurs in early August in most years, but in 2007 it also occurred in early July (there were two periods of glochidial production that year, but this appears to be unusual) (Scheder and Gumpinger, 2008). Glochidial release varies in a similar way and usually follows about a month after spawning (Hastie and Young, 2003; Scheder and Gumpinger, 2008). Hastie and Young (2003) found that there were about 3000 to 3600 degree days between successive glochidial release dates, with earlier release in warmer years. They suggest that glochidial release may be controlled both by the summed water temperature (expressed in degree days) and a critical water temperature that must be reached before it can occur. They also note that it often coincides with a sudden change in water temperature (more than two degrees Celsius) and / or river level (more than 0.1m), which might result in impaired respiration by mussels, causing them to eject their glochidia. 8

19 The best method to estimate when spawning and glochidial release will occur is to keep records of previous release dates and daily temperatures, so that the number of degree days can be calculated and used to estimate future release dates. Even with this method there is likely to be some uncertainty, so that monitoring of mussels is recommended from about a month before the predicted glochidial release date. 3.2 Ensuring that fertilization occurs There are several requirements to ensure a healthy supply of glochidia, and the main one is that fertilization occurs. The mussels need to be of both sexes, or to be hermaphrodites (these sometimes occur in low density populations; Bauer, 1987a). As most populations are dioecious with separate males and females, collecting at least 20 mussels from a site should ensure that both are present. The tanks need to be set up in such a way that the sperm from the males reach the females. To achieve this, the distance between males and females needs to be short, and the circulation of the water must allow the sperm to travel to the females, so the water must be moving and there should be good mixing of water before it leaves the tanks. Without flow the sperm will not be carried away from the males, and with very high flows they may be washed out of the tank without reaching the females. Ideally males should be placed upstream of females if the sex of mussels is known (it can be checked by partially opening mussels using tongs and then examining the gills; females have thicker gills, and this is easiest to see when mussels are gravid). To achieve high fertilization, Alan Keys (pers. comm.) has installed very shallow circular tanks with medium flows, with the water circling in from the outside of the tank to a drain hole in the middle (Figure 2). This ensures that the water circles round several times before it leaves the tank, increasing the chance that sperm from a male will reach a female. Alan also moves the females towards the centre of the tank, both to increase the chance that sperm reach them, and to shorten the distance that glochidia must travel to get washed out of the drain in the middle into the fish tanks. The rectangular tanks used at Mawddach also provide a good supply of glochidia so fertilization must be occurring well in these tanks. Figure 2. Adult holding tank at Ballinderry showing direction of flow. 9

20 3.3 Monitoring glochidial development Glochidial development can be monitored in mussels by using tongs to gently prize open the valves and then using a syringe to take a small sample from inside the swollen demibranchs (gills) (see pictures in Scheder and Gumpinger, 2008 for how to do this). The bore of the syringe needs to be larger than the expected size of an egg or glochidium, and a needle with bore of approximately 100µm should be adequate, as glochidia are usually 50-80µm in length (Ziuganov et al., 1994; Pekkarinen and Valovirta, 1996); only a small sample is needed. This sample can be examined under a dissecting microscope to check glochidial development. This should not harm the mussels, but it is worth doing it to one or two mussels and then monitoring them to make sure they are OK before doing it to all mussels. Christian Scheder et al. (of Consultants in Aquatic Ecology and Engineering, Wels, Austria; Scheder and Gumpinger, 2008) use this technique to monitor glochidial development, and have identified five stages of development as follows: 1. The egg appears as an amorphous mass of cells within a membrane 2. First constrictions visible, i.e. you can make out the line where the two glochidial valves are going to split 3. Glochidium appears fully formed within membrane 4. Glochidial snapping visible within membrane 5. Glochidium is free from membrane and moves around snapping freely. Pictures of glochidia at these different stages are available in their presentation from the 2008 Luxembourg conference ( In Austria this development takes about a month, and usually starts at the beginning of August and finishes towards the end of August (e.g. in 2005 and 2006). However the dates vary between years as discussed in section 3.1. Hastie and Young (2003) report that in Scottish rivers, gravid mussels were usually present for approximately two months during the period June to September. Mature glochidia (i.e. glochidia in stage 5) can be identified by their snapping movements, which should increase in the presence of salt, fish mucus, blood, gill tissue or fin tissue (Young and Williams, 1984b). The glochidial valves should close following a direct touch or when in contact with fish tissue. The addition of salt to a subsample of the glochidia is a standard test to see if the glochidia are healthy and suitable for use with fish; healthy glochidia snap vigorously. Christian Scheder has found that in some years, the glochidia never reach the 5 th stage and are not released. He correlates this with abnormal temperatures: in the hot summer of 2007 there were two periods of glochidial development, both only reaching the fourth stage, and in the cold summer of 2006 glochidia only reached the fourth stage. This suggests that a change in the temperature regime can prevent glochidia reaching maturity, resulting in no glochidial release. 10

21 3.4 Predicting glochidial release The timing of glochidial release is difficult to predict. The best estimates are likely to be gained from degree days, in combination with other factors (see discussion in Section 3.1). Another way of predicting release is to monitor one population that is known to release slightly earlier than other nearby populations, and use it to predict release dates for the other populations. For example, Rainer Dettmer (pers. comm.) knows when the mussels in his stream depot (section 2.5) are likely to release as it is always approximately 5 days after another nearby population. If the different groups working on mussels tell each other when release is starting, it may be possible to work out which population releases first and how many days later the other populations are likely to follow. Glochidial development can also be monitored (as described in section 3.3) and when mussels reach stage 4 or 5, then release is imminent. If there have been problems with release before, it is possible to force mussels to expel stage 4 glochidia (section 3.6), which should still be able to infect fish. While they may not be as healthy at this stage as if they had reached stage 5, it is better to have a low level of infection than no infection. 3.5 Glochidial release Glochidial release occurs over several days (Young and Williams, 1984a). This release is almost certainly triggered by a change in water temperature or flow level and is highly synchronised even between sites one kilometre apart (Hastie and Young, 2003; section 3.1). Approximately 2-10 million glochidia are released by each mussel (Bauer, 1987a). In hatchery tanks, glochidial release is usually seen when mussels release conglutinates onto the surface of the sediment. With high enough flows these conglutinates can be washed straight out of the mussel tank into the fish infection tank. However most people find that flows are not high enough to achieve this and either give the glochidia a gentle push towards the outflow of the tank (Alan Keys, pers. comm.), or pick them up with a syringe and place them directly into the tank with fish (Keith Scriven, pers. comm.). The advantage of picking them up is that all glochidia can be transferred at the same time into a tank with lowered water levels and no flow in order to increase the infection rates on fish. 3.6 Artificially triggering glochidial release The easiest way to trigger glochidial release is to place mussels in a bucket of warm water without oxygenation (Scheder and Gumpinger, 2008). The low oxygen levels stress the mussels, causing them to eject glochidia from their gills (also seen in other mussel species, Aldridge and McIvor, 2003). The glochidia will be released in clumps or conglutinates, which break up easily when picked up in a syringe, and can then be placed in tanks with fish for infection. The danger of doing this is that the glochidia may be expelled when they are not mature, so it is advisable to check for glochidial maturity using Christian Scheder s method above (section 3.3). The other alternative is to get one mussel to release and do a test infection on one fish to make sure they can attach before causing all mussels to expel their glochidia. 11

22 3.7 Possible reasons for mussels not releasing glochidia If no mussels in a hatchery produce glochidia, possible reasons include: that they are all of one sex; that the positioning of mussels and mixing in the tank is insufficient to transport sperm to females; that mussels are in too poor body condition to divert energy to reproduction; that the temperature or light regime have not triggered reproduction; that the mussels are diseased (some freshwater mussel diseases castrate males; Jokela et al., 1993); or that the glochidial release has gone undetected. To encourage reproduction, the following is recommended: to ensure an adequate food supply; to use tanks that encourage mixing of water; to provide a natural temperature regime and photoperiod; to increase the number of mussels being held; to check their sex; and to monitor glochidial development to check if eggs are being produced and fertilized and if they are developing normally and becoming mature glochidia. It is likely that only a proportion of mussels release glochidia each year, with some taking a rest. Bauer (1987a) found that in Bavarian populations, between 10 and 70% of mussels were gravid each year. This might be explained by female mussels needing to reach a threshold soft body mass before they can reproduce (Bauer, 1998). If insufficient food is present, females may delay reproduction until another year. This would be an effective strategy for a mussel species with such a long life span (more than 80 years). Therefore feeding mussels may encourage reproduction. 12

23 4 FISH INFECTION AND JUVENILE COLLECTION Freshwater pearl mussels are known to use salmon (Salmo salar) and brown trout (S. trutta) as hosts for their parasitic glochidia (Young and Williams, 1984b; Bauer, 1987b). Which of these species is a better host is likely to vary with the mussel populations. Rainbow trout (S. gairdneri) and minnows (Phoxinus phoxinus) are unsuitable as hosts, with the glochidia being lost from fish within 48 hours of infection (Young and Williams, 1984b; Bauer, 1987b). Brown trout are better hosts for use in hatcheries to avoid problems with salmon smolting before juveniles have excysted from the fish (Roger Sweeting and Louise Miles, pers. comm.). It is generally recommended to use fish taken from the same river as the mussels where possible, although good infection rates are achieved with hatchery-reared fish sourced elsewhere. It is important to use young fish that have not previously been exposed to mussel glochidia, as fish can acquire immunity to glochidia (observed in other fish and mussel species; Dodd et al., 2006). It is also important to use glochidia from several mussels to infect the fish in order to ensure high genetic diversity in juveniles. The glochidia attach to the gills of fish, and most glochidia attach to the middle and ventral gill sections (Young and Williams, 1984b). The glochidia grow during their time on fish and are usually 200 to 400µm in length when they excyst from the fish (Ziuganov et al., 1994). A variety of methods have been used to infect fish. These methods are described briefly here; the methods used in Mawddach hatchery are working well. 4.1 Method 1 Mussels and fish held together Possibly the easiest way of infecting fish is to hold the fish in a tank with the mussels and allow natural infection to occur. This method is being used by Roger Sweeting and Louise Miles at the Freshwater Biology Association research station by Lake Windermere. Mussels are held in gravel in wire baskets, and the baskets are placed in circular fish tanks with water flowing in near the edge and out from the middle (Figure 3). A hose is used to wash silt off the mussels regularly. While the flow pattern in tanks is altered by the mussel baskets, the fish survive well there and can be seen swimming directly over the mussels. In 2007 approximately 1000 salmon were successfully infected with up to 2000 glochidia per fish (Roger Sweeting and Louise Miles, pers. comm.). The advantages of this method include that it is not necessary to monitor when mussels release glochidia, and that possibly mussels are more likely to release glochidia if fish are present. The disadvantages are that it is not possible to control the infection rate on the fish. 4.2 Method 2 - Mussel outflow enters fish tanks Another simple method for infecting fish is to connect the outflow from the mussel holding tank into the fish tank so that glochidia are washed into the fish tank and infect the fish when they come into contact with them. As with the previous method, this does not require constant monitoring of mussels to check when they are releasing glochidia. This method was originally used in Mawddach Fish Hatchery. 13

24 Figure 3. Method used to infect fish by Roger Sweeting and Louise Miles of the FBA at Lake Windermere. Figure shows a tray containing gravel and mussels, and fish swimming in, over and around it in a flowing circular fish tank. One problem is that the mussels often release the glochidia in clumps (conglutinates) and there may not be enough flow to wash these out of the mussel tanks into the fish tanks. Another problem with this method is that infection rates on fish may not be very high as the residence time of the water and glochidia in the fish tanks is relatively low, so that many glochidia leave the tank without having come into contact with fish. In Ballinderry Fish Hatchery in Northern Ireland, Alan Keys et al. (pers. comm.) hold the parent mussels in very shallow circular tanks with a central outflow (Figure 2), so that flow over the mussels is fast enough to wash the conglutinates into the outflow where the turbulence breaks them up before they reach the fish tank. Any conglutinates not flowing out naturally can be manually swept into the outflow. When mussels are seen to be releasing glochidia, they are moved into the central part of the tank where flow is highest and conglutinates have the shortest distance to reach the outflow. Water from the mussels then flows sequentially into three different stages of fish infection, so that the same glochidia have three chances to infect fish. The first tanks are long thin tanks, the second tanks are circular and finally there is a third square tank. The water from the mussel tank is supplemented with other water as it enters the fish tanks, to ensure high enough flows and oxygen levels to keep fish healthy. Fish become infected in all tanks, and the infection rates are highest in the second circular tanks, perhaps because the circular flow ensures a long residence time of glochidia and several opportunities for contact with fish (Figures 4 and 5). There has been some discussion over which type of tank fish should be held in to achieve the highest infection levels. Possibly square tanks have more dead space where fish and glochidia can come into contact with each other. However high infection rates have been achieved in the circular tanks at Ballinderry Fish Hatchery. 4.3 Method 3 - Glochidia placed in non-flowing fish tank An effective way to ensure high infection rates is to stop the flow of water through the fish tank, reduce the water level, provide aeration for the fish, and transfer the glochidial conglutinates manually from the mussel tank into the fish tank. The conglutinates can be moved using a syringe to suck them up, when they immediately break up into a suspension of 14

25 glochidia. High levels of infection can be achieved in 20 to 30 minutes; it is not advisable to leave fish for longer than this as their health may deteriorate without a good flow of water. This method has been shown to produce high infection rates at Mawddach hatchery (Keith Scriven, pers. comm.), and is widely used, also with other mussel species. It is important when using this method to ensure that conglutinates from several parent mussels are used so as to obtain high genetic diversity in the juveniles produced. This is usually possible because mussels often release their glochidia simultaneously, so that conglutinates from several mussels can be transferred to fish tanks at the same time. Flow direction Second and third fish tanks First fish tanks Adult mussels Figure 4. Adult mussels tank (round black tank at far left), first fish infection tanks (two of them) and in the distance the second and third infection tanks Figure 5. Second and third fish infection tanks. The second tank is the round tank on the left and the third tank is the square tank on the right. 15

26 The only disadvantage of this method is that mussels have to be checked daily for release of conglutinates, as release only occurs over a relatively short period of a few days. However if it is not possible to monitor mussels this closely, they can be made to release glochidia by placing them in a bucket of warm deoxygenated water (see section 3.6). The danger is that the glochidia may not be fully mature or ready to attach to fish, and this might result in lower infection rates. It is also recommended that tanks are plumbed together as described in section 4.2, with the outflows from mussel tanks entering the fish tanks, so that if some mussels release glochidia unnoticed, these glochidia still have a chance to infect fish. This method of fish infection is also suitable for bank-side use; the right species of host fish are electro-fished out, placed in a bucket with a good supply of air or oxygen, and glochidia are added (Rainer Dettmer, pers. comm.) These glochidia can either be obtained directly from mussels placed in a bucket of warm water, or from conglutinates released naturally into a tank and transported to the fish. 4.4 Method 4 Laboratory scale infection of fish A final method of fish infection is to infect fish individually by pipetting glochidia onto fish while holding them in small containers. This has been done with other mussel and fish species (e.g. McIvor, 2004), and can achieve very high levels of infection, but may require a Home Office Licence and fish may need to be anaesthetized. There is some evidence that very high levels of infection can cause an immune response in fish resulting in all glochidia being sloughed off. Only small numbers of fish can be infected in this way. This method is not recommended except for the laboratory-scale infection of fish and production of small numbers of glochidia for research. 4.5 Summary of fish infection methods The four methods described above are summarised in Table 1 with a brief description of their advantages and disadvantages. Table 1. Methods of fish infection listing advantages and disadvantages of each method. Infection method Advantages Disadvantages Mussels and fish held in same tank Requires little monitoring or intervention Not possible to control infection rate on fish Source of information on this method Roger Sweeting and Louise Miles (FBA, Windermere) Mussel tank outflow enters fish tank Requires little monitoring or intervention May not achieve high levels of infection Keith Scriven (Mawddach Hatchery, Wales); Alan Keys (Ballinderry Fish Hatchery, Northern Ireland) Glochidia placed in non-flowing fish tank Pipetting glochidia onto fish held in small containers Very quick and achieves good levels of infection High levels of infection Mussels need to be monitored regularly for glochidial release Keith Scriven (Mawddach Hatchery, Wales); Rainer Dettmer (Lower Saxony, Germany) May need a Home Office McIvor 2004 and refs Licence. Fish may need to therein. be anaesthetized and only small numbers can be infected. 16

27 4.6 Estimating excystment dates of juveniles from fish Glochidia usually remain encysted on fish for several months, and encystment duration is related to temperature. The juveniles excyst from fish (i.e. they fall off the fish) over a period of five to ten days, with the majority of juveniles falling off during two to three days in the middle. Therefore it is essential that the mesh screens to collect the juveniles from the outflow water of fish tanks are in place in time to catch these juveniles, and a method of calculating how long juveniles will remain encysted is needed in order to estimate when the juveniles will start to drop off. Hruška (1992) proposes the use of degree days to estimate the duration of encystment on fish, with an adjustment relating to water temperature during the last 2 weeks of encystment. He calculates degree days as the sum of average daily water temperatures. Under artificial conditions with fish held in tanks at 15.5 to 17 C, juveniles remained on fish for 84 ± 4 days i.e to 1430 degree days. However under natural conditions in the River Blanice, south Bohemia, in the Czech Republic, encystment lasted 11 months corresponding to 1760 to 1860 degree days. This varied between years: degree days in 1987, and 1818 to 1860 degree days in A possible explanation for this variation is that during the final days, the average water temperature needs to be at or above 15 C (accounting for about 225 degree days of the time encysted), as observed in both 1987 and To test this requirement for a warm period just prior to excystment, Hruška moved infected fish to colder temperatures either one month or one week before juveniles were due to excyst. Fish moved one month before showed either delayed release of juveniles or no release at all at the lowest temperatures; this could imply that glochidial development was incomplete when the fish were moved, and could not reach completion at the lower temperatures. However fish moved a week before the due date all released their glochidia within 14 days, suggesting that these glochidia were fully developed and had begun the excystment process when moved, and that once started there was no going back. He suggests that the development of glochidia is continuous during the early to mid stage of encystment, but later on (after approximately 1100 degree days) the development slows down as if the glochidia were waiting for a longer period during which the average water temperature would remain at or above 15 C. If the temperature goes down before the glochidia have finished their metamorphosis, their development slows down or stops, but if it gets colder at or after the onset of glochidial release from the cysts, there is no retardation. Bauer (1979) found differences in encystment duration in five German populations. In two populations the glochidia overwintered on fish with an encystment duration of seven to nine months, while in other populations the glochidia completed development in days. In one population both strategies were seen. The only river where young mussels were found was the one where all juveniles overwintered on fish, and this river also supported the highest densities of mussels. This is possibly because juveniles excysting in late summer are obliged to spend the winter in the sediment, and given Buddensiek s (1995) observation that the smallest juveniles generally do not survive the winter, most of them are likely to die. Warmer summer water temperatures, resulting from e.g. a loss of forest cover from upstream sections, may be causing this premature release of juveniles before the winter. 17

28 Ziuganov et al. (1994) found significant variation in encystment duration within a single population in the rivers of north-west Russia. In one year glochidia metamorphosed in 18 days while in another year, encystment lasted 11 months. He believes that temperature is the main determinant of these different encystment times. Young and Williams (1984b) report that the encystment time in Scotland was approximately 290 days. In , juveniles excysted days post-encystment, and in after approximately 293 days. About 12% of the glochidia that originally encysted completed metamorphosis to be released as juveniles. To summarize, only Hruška (1992) has calculated degree days. Other authors note the variability in encystment duration, with some juveniles excysting after a very short time period (<2 months); however there have been no reports of this in British populations. It is likely that Hruška s degree day method will also be applicable to British populations, although the precise number of degree days may vary due to genetic differences between populations. The adjustment relating to the temperature during the final weeks of encystment may explain observed variability in the total degree days seen in individual populations. 4.7 Altering encystment duration It is possible to either speed up or slow down encystment by altering the temperature at which fish are kept. Hruška and various others use this to ensure a more constant supply of juveniles, which makes possible experimentation on factors affecting juveniles survival. Fish have to be kept in a recirculating tank with a good biological filter to ensure high fish survival and to allow heating up of the water (Figure 6). 4.8 Proportion of encysted juveniles surviving to excystment Throughout encystment there is a gradual loss of glochidia from fish, so that final numbers of juveniles are much less than the number of glochidia that originally attached to fish. Young and Williams (1984b) report that less than 12% of the glochidia that originally encysted on fish survived to be released as juveniles. They found that the losses from fish were different between brown trout and salmon. By day 40, most of the trout had lost their glochidia, with a few fish retaining large numbers. With the salmon, a significant number of glochidia were lost from all fish by day 40 but the losses were evenly spread among the fish. There was a second period of glochidial loss after 110 days that carried on until excystment. The unevenness in long-term fish infection rates seen in the brown trout makes it possible to select out and keep only those fish with high numbers of glochidia, while releasing fish with lower infection rates back into the river; this procedure is used at Ballinderry Fish Hatchery (see section 4.9 below; Alan Keys et al., pers. comm.). 18

29 Figure 6. Recirculating fish tank used at Heinerscheid pearl mussel rearing station, Luxembourg, to keep fish warm to speed up encystment duration. The top container is the fish tank, and the bottom container contains a pump, which pumps water through a water filter and heater. The white tray is used to catch juveniles from the fish tank outflow. 4.9 Collecting the juveniles after they have excysted from fish Juveniles may excyst from fish over several weeks, so mesh screens need to be in place in preparation for the start of excystment and then throughout that time. The water from the fish tank is passed through a filter with small enough mesh size to catch the juveniles, as used at Mawddach Hatchery. The mesh needs to be kept underwater so that the juveniles do not dry out, and needs to be emptied daily to collect the juveniles. Increasing the mesh area and not feeding the fish during excystment can reduce problems associated with the mesh becoming clogged and overflowing. However some debris from the fish tank will inevitably get mixed up with juveniles. The juveniles and debris can be sieved again and just the fraction containing the juveniles collected. However the matter remaining on the larger sieves should also be checked as many juveniles may get caught with the debris in these sieves (Keith Scriven and John Taylor, pers. comm.). If there are only small numbers of juveniles, it may be feasible to individually pick out the juveniles from the debris to transfer them to trays, and this is the best option when possible as it reduces the chances of transferring other small organisms, such as rotifers and planktonic crustaceans, that may prey on the juveniles. It also allows the number of juveniles to be counted, which is essential in order to estimate juvenile survival later on. 19

30 For larger numbers of juveniles, it is possible to put them in a bucket with plenty of water and swirl the water, so that the juveniles settle in the middle (Keith Scriven, pers. comm.). The juveniles can then be taken out with a syringe and their number estimated volumetrically. However many juveniles remain behind with the other debris; to avoid losing these juveniles, Keith has been transferring the juveniles and debris to the rearing trays. Although this debris may decrease juvenile survival, this may be offset by the larger numbers of juveniles placed in the trays. Further work is needed to decide the best course of action. An alternative and easier option is to place the fish in a flowing tank or artificial stream with suitable substrate for juvenile survival, and allow the juveniles to drop off straight into the substrate (Alan Keys, pers. comm.). This is the principle used in Ballinderry Fish Hatchery in Northern Ireland, where the fish with the most glochidia attached are selected by a quick visual examination of their gills; the glochidial cysts are clearly visible on the gills once the glochidia have encysted. These fish are then moved into an artificial stream with suitable substrate, so that juveniles can excyst straight into the sediment. The fish are moved just before the juveniles are due to excyst, and are not fed their normal food once in the stream to avoid increasing the nutrient load in the sediment. Aquamats that have been left in a river and have become infested with aquatic invertebrates are provided for the fish to feed on along with some maggots. The disadvantage of this method is that juveniles are very difficult to find until they are 4 to 5 years old as the substrate is mixed gravel. Also the raceway needs to be sufficiently long, with some slower flowing areas, to ensure that juveniles have a chance to settle onto the sediment before being washed out of the tank. 20

31 5 REARING JUVENILES Various methods are in use to rear juvenile pearl mussels across Europe. These include highly intensive methods with juveniles cleaned and checked regularly (e.g. Jaroslav Hruška in the Czech Republic) and very low intensity approaches where juveniles are allowed to grow unchecked in natural sediment until they are large enough to be located visually, four to five years later (e.g. Alan Keys in Ballinderry Fish Hatchery, Northern Ireland). The different methods and approaches are described here, with the lowest intensity approaches described first. These approaches are summarized in Table 2, and Table 3 lists the options for rearing juveniles with comments on their advantages and disadvantages. 5.1 Releasing infected fish into rivers and streams The lowest intensity approach to rearing juvenile freshwater pearl mussels involves infecting fish with glochidia and releasing them into streams so that the juveniles fall straight into the substrate (Hruška, 1999; Hastie and Young, 2003). This approach has been used numerous times but with questionable and often unmeasurable success, because of the difficulty of monitoring juvenile survival and growth; juvenile mussels are unlikely to be found until they are 4 to 6 years old because of their small size. It is only likely to work where the limiting factor for pearl mussel reproduction is something other than juvenile survival, such as loss of host fish or low densities of adult mussels. In such cases it may be a highly effective method of repopulating streams with pearl mussels; Österling et al. (2008) found that in streams with on-going recruitment (i.e. with adequate conditions for juveniles), juveniles mussel density was positively related to the number of glochidial infections per unit of stream area, suggesting that increasing the infection rate and number of fish will result in greater recruitment of juveniles. However mostly juvenile survival is believed to be the limiting factor, as non-recruiting adult populations exist in rivers where suitable host fish are still present and the adults are still producing glochidia (Österling et al., 2008). Even if large numbers of juveniles excyst from fish in such a place, they will probably all die if the substrate is unsuitable for them to survive in, e.g. because of higher silt loads or organic content with resultant low oxygen concentrations in the substrate. Therefore in order for this approach to work, it is necessary to understand what factors are limiting juvenile survival and to restore rivers such that these factors are corrected. It is also necessary to find ways of monitoring juvenile survival in these waterways to ensure that the restoration has been adequate for their survival. The most successful work on this has been carried out on the River Lutter in Lower Saxony, Germany, described in the next section. 21

32 Who Where Fish Methods Volker Buddensiek Jaroslav Hruška North Germany River Blanice, Czech Republic Brown trout Brown trout Juveniles placed in cages consisting of a thick perspex plate with holes drilled through, and a mesh covering on either side. 5 juveniles were placed in the holes, and the mesh allowed flow of water past mussels. The plates were placed in 5 rivers fastened above the bottom so that they were facing the current. Juveniles initially held in non-flowing tanks, then placed in special cages in rivers or in waters connected to rivers. Young juveniles are fed an organic detrital suspension produced in the riparian zones immediately adjacent to the rivers. Reference Measures of success Juveniles have been kept alive for up to 52 months (as of 1995). Buddensiek, Survival declined, reaching 5 to 20% after 1 to 2 years, and <5% after months (calculated as proportion of excysted juveniles alive). The maximum shell lengths varied from 2.1mm to 6.4mm after 36 to 52 months. 30,000 juveniles reared so far, with maximum size of 5-year old juveniles reaching 24mm. Hruška, 1999 Lee Hastie and Mark Young River Salmon Moidart and River Dee, Scotland Juvenile mussels were held in 200μm mesh baskets with sediment held in gravity-fed flowing raceways using unfiltered river water. Baskets and raceways were cleaned every 2 to 4 weeks removing silt. 2 separate hatcheries on different rivers. Juvenile mussels placed in cages following Buddensiek (1995). Some cages were placed in River Moidart attached to pegs, and cleaned by gentle brushing every 2 months. Other cages were kept in a large tank at Dinnet Hatchery on the River Dee, with a flow rate of 10 litres/minute, with cages being cleaned regularly. No juveniles found in Kinlochmoidart hatchery. 2 live juveniles found in Dinnet hatchery, suggesting a maximum 40% survival rate after 10 months. Mussels were 1.13 and 1.25mm in length. Eight of the ten cages in River Moidart were lost in a flood. In remaining two cages, there was 11% survival after 7 months (27 juveniles), 3% survival after 12 months (7 juveniles) and 1% survival after 16 months (3 juveniles, lengths: 2.11mm, 2.25mm. 1.75mm). In the hatchery, no juveniles survived past 11 months. Hastie and Young, 2003 Schmidt and Wenz Germany Brown trout Juvenile trout infected with glochidia and released into rivers Survival of farm-reared fish is low, and no evidence of establishment of juvenile mussels. Schmidt and Wenz, in Hastie and Young, 2003 Michael Lange Vogtland, Saxony, Germany Followed the methods of Hruška and Buddensiek. Fed detritus from wet meadows with ground animal protein. After 3 months, reached 1.2mm; reached 6mm in 2 to 3 growth seasons (= years) Lange, 2005 Alan Keys, Ballinderry Jane Preston, River, Dai Roberts Northern et al. Ireland Brown trout Semi-natural system. Juvenile mussels excyst from fish directly into gravel sediment in an artificial stream. Juveniles aged 5-10 years old now present, with younger ones probably growing in the substrate but not yet visible. Oldest juveniles have reached 50mm in length. Preston, Keys and Roberts, 2007 Table 2. Current and previous work culturing pearl mussels (excluding Welsh work). 22

33 Least energy intensive Most energy intensive Juvenile culture Advantages Disadvantages Use with M. margaritifera Collect juveniles and place in nonflowing tanks with food. Collect juveniles and place in recirculating tanks with sediment and feed regularly. Collect juveniles and place in recirculating tanks with sediment and a water filter, and feed continuously. Collect juveniles and place in trays in flowing tanks fed with river water. Collect juveniles and place in trays in flowing tanks fed with river water, with options to filter the water first and to add food to the water or sediment. Collect juveniles and place in cages or other closed containers which allow flow, then put cages in rivers Allow infected fish to release juveniles into large raceways fed with river water and with a natural substrate. Release infected fish into river for juveniles to excyst naturally onto river substrate. This is often sufficient for rearing juveniles for the first few days or weeks post-excystment and requires little equipment or space. Good success has been achieved with such tanks in North America for a wide range of species. Usually quite economical of space depending on size of tanks. This allows for better control of algal concentrations, reducing the risk of overdosing with algae and causing eutrophication. Depending on water source, this can ensure very high water quality with a natural temperature regime. This is as close as it is possible to get to putting juveniles directly into rivers, while still allowing them to be monitored over time. This is like creating an artificial river with natural substrate. After building the raceway and infecting fish, little further work is required until you need to find the mussels again. Unsuitable for rearing most species of juveniles long term. Very energy intensive as juveniles need checking and cleaning regularly. Species requiring very high water quality, such as the pearl mussel, may not do well in these tanks. This may require very large amounts of algal food as the algae are continually being lost in the water filter. Algae are often expensive to buy in these quantities, and are time-consuming and space-intensive to culture in large quantities. Raceways will need to be plumbed into rivers (usually associated with fish hatcheries, otherwise very expensive to build). Suitable large water filters and copious quantities of algae are expensive. The mesh may prevent food particles entering the cages, or sediment from leaving the cages. The cages can get washed away during floods. Such large raceways are very expensive to build and take a lot of space. Must be located by a suitable water source. Very difficult to find mussels once released (similar to releasing directly into river) during first years. The success or failure will not be known for some years, Low intensity approach in relation to mussel infection until juveniles have reached a size where they can be (but high intensity in terms of river restoration that may found. If there has been a lack of recruitment caused by be required for juveniles to survive). Work period is pollution, siltation, low food availability or changes in during glochidial release period of mussels, when fish hydrology or temperature, the juveniles may not be able must be infected. to survive unless efforts are simultaneously made to restore habitats. Used by Hruška (1999) and others for rearing pearl mussels for the first three months. Kennedy (2006) achieved good growth with recirculating tanks over the first month postexcystment. Scriven has achieved long-term survival and growth (2 years+) using a raceway fed with river water with juveniles on mesh trays (pers. comm.). Buddensiek (1995), Lange (2005) and Kennedy (2006) have used this technique successfully. Hastie and Young (2003) lost many of their cages. Preston, Keys and Roberts (2007) used this approach, achieving good growth, but they had difficulty refinding the juveniles. They now have a large cohort of 9-10 year old juveniles. Long-term project in the River Lutter in Germany has done this successfully, alongside a large river restoration project. Others (e.g. Hastie and Young, 2003) have also tried it with unknown success. Table 3. The advantages and disadvantages of the various culture methods available. 23

34 5.2 Natural rearing in Lower Saxony, Germany with river restoration Work on restoring pearl mussel populations in the River Lutter in Lower Saxony began several decades ago. Young mussels have been reared in situ for some years now, and this has been achieved by releasing infected trout into restored habitats (Altmüller and Dettmer, 2001; Rainer Dettmer, pers. comm.). There has been extensive river restoration work throughout this part of the catchment, with a number of silt traps installed and several channel restoration projects (Figure 7). In conjunction with this, fish are caught by electrofishing and artificially infected with glochidia and then immediately returned to the river (see section 4.3). The success of this work is demonstrated by the fact that the young mussels have reached reproductive age and are infecting fish themselves, thereby completing the cycle (Rainer Dettmer, pers. comm.). While this work is not intensive from the mussel rearing side, overall it is very intensive and costly because of the enormous scale of restoring a whole river catchment. Figure 7. The River Lutter in Lower Saxony, Germany: two different types of silt trap used in river restoration, including a box silt trap set into the substrate (top left) and a very large artificial settling channel through which water flows before entering the main river (bottom left); a site where juvenile pearl mussels are doing well (right). 5.3 Buddensiek s cages Working in the same catchment, the River Lutter in Lower Saxony, Buddensiek (1995) devised a method of holding juveniles in small cages so that their survival could be monitored in the rivers and streams. This provided an opportunity to study some of the factors affecting juvenile survival. Buddensiek made these cages from perspex plates with holes drilled through them to make the individual cells in which juveniles were placed, with mesh on either side to hold the juveniles in the cells (Figure 8). 24

35 Figure 8. The cages used by Buddensiek (1995) to hold juvenile mussels in rivers. Each cage consists of two covering plates (A) and a thicker 9mm central plate (C) into which 92 holes, each 6mm in diameter, have been drilled. The covering plates hold 2 pieces of mesh gauze in place (B), with mesh size 200µm. The juveniles are placed in the individual cells (D). Diagram from Buddensiek (1995). Buddensiek s cages proved an efficient way of placing juveniles in rivers without losing them. Survival was generally low over the first winter post-excystment (<50%) and after one year between 5 and 20% of mussels had survived, reaching a mean length of approximately 900µm. Larger juveniles survived better: all individuals less than 700μm at 3 months postexcystment had died 4 months later, while 45% of juveniles larger than 900μm survived. This suggests that the early nutrition of juveniles is very important, to ensure good growth before the first winter. The most important factor affecting survival and growth of juveniles was temperature. Growth increased with temperature and only occurred during the warm period of each year, decreasing to almost zero from October to March. However high temperatures also correlated with high mortality in 3 of the 4 rivers where juveniles were placed, and the reasons for this are unknown. Growth in Buddensiek s cages was generally slower than in Scottish rivers (as estimated from growth rings on shells), except in the most eutrophic of his 4 rivers, where growth was similar to Scottish mussels. This is surprising as it is well-known that eutrophication usually results in juvenile death. Possibly holding mussels in cages in the flow reduced the build-up of organic matter in the interstitial spaces, preventing oxygen depletion that can otherwise kill juveniles. The increase in food availability due to the higher nutrient levels presumably facilitated higher growth. This emphasizes the impact that different source waters can have on juvenile survival and growth, whether in a river or hatchery. 25

36 Buddensiek also found that both organic and inorganic material got into the individual cages or perspex cells. Juveniles in cells that had become one to two thirds full of material had higher survival than those in cells with little or no extra material. There was also lower survival in cells that were more than two thirds full, but this was not statistically significant. When Hastie and Young (2003) used Buddensiek s cages in Scottish rivers, they also found live juveniles only in cells with sediment. This shows that providing juveniles with sediment (containing both organic and inorganic material) is likely to increase growth and survival. Surprisingly, Buddensiek found that cells that had been colonised by chironomid larvae supported higher juvenile survival than those without, perhaps because the larvae produced faecal pellets that may have provided food for the mussels, or because the larvae ate other organisms that would have preyed on the juveniles. Finally, he found that juveniles could be transferred between catchments with good survival (unlike adults, which usually have low survival (<50%) when transferred to new catchments, Valovirta, 1990), so the rearing of juveniles in water from rivers other than their parent river is possible. These cages make excellent opportunities to study juvenile survival in different rivers. They are used widely as they remain the best way of replacing juveniles in rivers while still being able to monitor their growth and survival. They are mostly used to place slightly older juveniles in waterways after intensive early rearing for the first 3-4 months in the laboratory (see method used by Hruška and others in section 5.5 below). However some groups have encountered problems with these cages. Hastie and Young (2003) tried a variety of rearing methods, including a semi-intensive culture system with juveniles in cages similar to those used by Buddensiek (1995). Losses were extremely high: out of ten cages placed in the River Moidart, eight were washed away during a 10-year-return flood. In the 2 remaining cages, there was low survival (11% after 7 months). Schmidt and Wenz (in Hastie and Young, 2003) also used these cages; they had inconsistent results with adjacent cages in one river having very different success rates, and with a survival of less than 1% after 2 years. 5.4 Rearing mussels in an artificial stream or mill race At Ballinderry Fish Hatchery in Northern Ireland, Alan Keys et al. are working both on river restoration and the hatchery rearing of juveniles. They have created an artificial stream using mixed sediment, plants and baffles to ensure a variety of substrate types, into which juveniles are allowed to excyst directly off fish (Figure 9). They are also experimenting with using a mill race in the same way, after adding gravel of suitable size to the areas that tend to remain free of silt. They are keen to get both mussels and fish reproducing in the rivers as soon as possible, and their restoration work also focuses on creating suitable fish habitats, as the decline in host fish populations may have contributed to the decline of the pearl mussels. Their approach is intermediate between placing infected fish back into rivers and streams as described above, and intensively rearing juveniles in flowing tanks or trays. The process starts just before the juveniles are due to drop off the fish, when they are moved to the artificial stream. Only fish with a large number of juveniles are moved; infection rates are estimated with a quick visual examination of fish gills. The fish are not fed their usual food in the stream, in order to avoid adding extra nutrients to the sediment. They are fed maggots, insects that fall in naturally, and aquatic invertebrates from the aquamats that have been colonised by stream invertebrates. In the future, Alan Keys (pers. comm.) is thinking 26

37 about trapping insects intentionally to act as food, by perhaps putting a light to attract insects at night that would then get caught within the netting which covers the stream. The netting is there to stop birds coming in to eat the fish and to stop the fish leaping out. It also provides shade to the stream, and will buffer temperature extremes. Figure 9. (a) The artificial stream where juveniles have been successfully reared, showing the wooden baffles that force the water to flow in a zigzag through the system; (b) the pool near the inflow which allows some sediment to drop out, also showing the large rocks which are used to create habitat diversity; (c) an island at the channel s edge and the netting which surrounds the whole channel; (d) the second stream, used by Preston, Keys and Roberts (2007) and recently altered to be more similar to the other raceway. 27

38 Figure 10. Diagrammatic view of the baffles showing how the water is forced to run through gaps that create stronger turbulent flows. 50mm Figure 11. Juvenile mussels reared by Alan Keys et al. in the artificial stream at Ballinderry Fish Hatchery. Mussels range in age from 5 to 10 years old, reaching 50mm length at 10 years. The stream contains river gravel i.e. natural sediment. There have been some problems with siltation, so they have installed a series of wooden baffles to increase the flow (Figures 9 and 10). The effect of the baffles is clearly visible as the water becomes turbulent and the sediment beneath is clear of silt. There are also larger rocks and small islands with plants, creating a variety of substrate types and flows. This diversity is likely to ensure that at least some of the sediment is suitable for young juveniles to survive in. Just downstream of the inflow, there is a pool where the water flows more slowly to act as a silt trap. Young juveniles live within the substrate. Juveniles can first be found when they are about 4 years old, and are visible either because their shells poke above the surface of the sediment or as two holes in the sediment surface, corresponding to the inhalant and exhalent siphons of the mussels (Figures 11 to 14). Up to 4 years, there is no way of knowing how many juveniles are surviving as they are very hard to find in amongst the sediment. Due to a silt problem a few years ago, the first juveniles reared in this system were taken out and transferred to other tanks and these juveniles are now 9-10 years old. Younger juveniles unexpectedly survived this silt episode, and some 5 to 8 year old juveniles have been found there recently (Figure 11). Younger juveniles are hopefully living in the sediment too. 28

39 The 700 oldest juveniles, which are 9 to 10 years old, are now kept in two tanks, a circular one that is very similar to the tank that the adults are held in (Figure 12), and a long thin tank (Figure 13). The circular tank is approximately 1m across and is filled with river sediment. Water flows in at the edge and spirals into the middle. Some silt settles out, but because the flow is variable there are areas of sediment that are silt-free and many mussels are visible there. However some juveniles are also sitting in the middle of the silt and seem to be doing well. To get rid of the silt, it can be gently disturbed so that it gets carried away by the flow. Additionally there is a pipe leading into the bottom of the tank that ends in a coil of pipe with holes in it. This can be connected to a pressurised air or water supply in order to clean the gravel from underneath from time to time. The mussels are large enough not to get washed out. The water is about 5cm deep, and flows out through the centre into a fish tank, because it is likely that these mussels will soon be old enough to reproduce, and the fish will be monitored for glochidia to check this. The long thin tank (Figure 13) also contains gravel with some larger rocks, and the juveniles are clearly visible, both above the substrate and also buried in it. Figure 12. The circular tank in which the 9-10 year old juveniles are being held. 29

40 Figure 13. The other tank holding 9-10 year old juveniles (left) and juvenile mussels visible in the sediment of this tank, showing how some of them are clustering around the larger rocks, which would presumably provide stability in times of high flow if they were in a river channel. Figure 14. The siphons of mussels visible in the silt in this tank (15 in this picture). The latest project is to release infected fish into a mill race to grow up naturally in a place where they can still be monitored. If this works it will be easy to replicate in other sites as mill races are still common, while artificial streams are expensive to build. The Ballinderry mill race is a channel approximately 2-3m across and 0.3m deep, with natural gravel substrate augmented by adding gravel to the areas where the silt does not tend to settle (Figure 15a). Because the channel has gentle bends, some areas are slower flowing and the silt settles here; these areas have been left without gravel as it is assumed that juvenile mussels will not survive there, and the areas would soon get silted up again if new gravel were added. 30

41 The infected fish are confined within a section of the mill race by a fish-frightening device at the top end (consisting of a water wheel structure, Figure 15b) and a grating at the other end. Even if some fish escape, the juveniles will be released directly into the Ballinderry river, which still contains adult mussels, so the juveniles may be able to survive. Originally the fish were held in wire and mesh cages, to protect them from predators. However this year the cages have not been used as the fish are staying in the mill race and predation levels do not seem to be too high. Infected fish were first released into the mill race 2 years ago, so they will know whether it has been successful in 2 to 3 years time, when juveniles are large enough to find. Another experiment which is being tried in this mill race is the use of trays of gravel (Figure 15c), which were initially held in the artificial stream and juveniles allowed to excyst off fish into them, and were then moved into the mill race. This occurred 2-3 years ago, so the juveniles will not be big enough yet to judge whether this was successful, but if it works this would be another easy way of rearing. The trays are about 60cm long and 40cm wide, with mesh bottoms and river gravel and sand inside (approx 3cm depth). Figure 15. (a) The mill race; (b) the fish frightening device, which is continually turning powered by the water flowing past; (c) one of the trays containing sediment and young mussels which is being held in the mill race. 5.5 Hruška s method Hruška (1999) has had success rearing juveniles from the River Blanice in the Czech Republic. Juveniles have reached more than 5 years of age and 20mm in length. His method is now used widely by other groups in Europe, including Michael Lange and Heidi Zeltman in Germany (Lange, 2005; Zeltman, pers. comm.) and Frankie Thielen et al. in Luxembourg (Thielen, pers. comm.). The description below has been compiled from Hruška (1992, 1999), Lange (2005), a visit to the Luxembourg hatchery, Heidi Zeltman s talk at the Luxembourg 2008 conference and a short visit to her lab. 31

42 Juveniles are initially reared in small containers (plastic boxes) where they are fed with detritus and ground up animal protein (Figure 16). Approximately 100 to 200 juveniles are placed in each container with approximate 500ml of water. The detritus is collected from either the wet grassy meadows that border the river or from organic matter settling out in the river. It is sieved to be smaller than the juveniles (i.e. less than µm) and this also removes potential predators. The ground up animal protein is from chironomid larvae (available as fish food), which are blended in water and sieved as with the detritus. This mixture of detritus and animal protein is kept suspended in aerated water until ready for use. A small amount is added to the water in the plastic tubs so that there is a thin layer at the bottom of the tubs when it has settled out (up to 2mm thick). The water is usually spring water or bore water. The boxes are held in a constant temperature refrigerator at 16 C. Hruška (1999) reports that it is advantageous to include a small number of water fleas (he uses Pleuroxus truncata) in the tubs as they clean the juvenile shells, removing mucus and faeces. For the first two to three weeks, juveniles are checked daily; this involves pouring the water they are in through a mesh sieve (Figure 16) to catch them, placing them in a petri dish with clean water, and using a dissecting microscope to remove any debris and dead juveniles which are gaping open. They are then returned to the box with fresh water and food. After approximately 3 weeks, these checks take place every 2 to 3 days, and then after another 3-4 weeks, they are weekly. Survival is approximately 50% over these three months. Figure 16. The mesh sieves and petri dishes (top right and left), plastic container used to hold juveniles (midleft), cage (bottom left) and artificial ditch (bottom right, not yet operational in this picture) used to rear juvenile mussels according to Hruška s method, as seen at the Luxembourg rearing station. 32