New Technology to Support Culturing of Freshwater Mussels for Research and Restoration
Freshwater mussels are one of the most critically imperiled groups of organisms worldwide. Encouraging mussel reproduction in a controlled setting has become an area of interest due to their widespread population declines and regional extinctions (Williams et al., 1993; Lydeard et al., 2004; Strayer et al., 2004; Haag 2012; Lopes-Lima et al., 2017). With constantly evolving techniques, rearing healthy populations of mussels in a laboratory is a promising option. These laboratory-raised mussels are used for studies to better understand population declines and in reintroduction efforts (Patterson et al., 2018).
The Issue:
Young freshwater mussels are so small and delicate that captive culture presents challenges for handling, feeding, avoiding predation, and maintaining a suitable habitat. Despite advances in captive propagation, the rearing of mussels between newly metamorphosed juveniles (approximately the size of tip of pencil) and those that are 2- to 4-months-old (approximately the size of pencil eraser) is often hindered by low survival or growth (Patterson et al., 2018). Low growth rates increase the time and resources required to obtain mussels of a target size for restoration.
Addressing the Issue:
The objective of the study was to develop a system for large-scale laboratory culture. This included starting with newly transformed mussels of different species, two of which are commonly cultured (L. siliquoidea and Villosa iris) and two that have been historically challenging to culture (Anodonta californiensis and Margaritifera falcata). Newly transformed mussels are the earliest life stage of the mussel and start to resemble the shape of the adult. These small young mussels have historically been more challenging to culture (O’Brien et al., 2013; Wang et al., 2017). To optimize survival and good growth rates we developed an auto feeding system.
- The auto-feeding system is used to deliver 1 L of fresh culture water (A) into each mixing cell (B) every hour. Just before the mixing cells fill with water, a pump (C) is triggered to deliver 16 ml of food (algal mixture) into each mixing cell.
- Food (algal mixture) is maintained at <12°C in an aerated container in a cooler with ice packs (D).
- Water and food flow from the mixing cells (B) into the water delivery tubes (E) that evenly divide water into each culture beaker. Boxes of beakers are maintained in a temperature-controlled water bath (F).
- Juvenile mussels are held in 300-ml beakers with a thin layer of sand substrate (~150-µm particles). The water in the beakers is static except for pulses of food and water delivered every hour. Excess water flows out through the screen-covered hole and is not reused.
- Beakers and sand are replaced weekly or biweekly. Mussels are separated from the sand by sieving through a 250-µm mesh screen.
Results/Future Results:
During the development of the pulsed flow-through auto-feeding system, five elements emerged as critical to the success of the system.
One of our keys to success was found in the presence of a sand substrate in culture chambers. Sand allows for a more uniform distribution of food on the bottom of test chambers and limits the accumulation of debris, fungi, and algae on mussel shells. The advantages of using silica sand, rather than fine sediment, in the present system are that (1) silica sand is commercially available, (2) small juveniles can be easily isolated from fine sand particles, and (3) sand has less potential impact on water quality.
Isolating of smaller batches of mussels in individual beakers also proved to be crucial to our progress. This isolation increases the likelihood of detecting problems early (e.g., signs of disease or blooms of competitors or predators) and helps prevent problems from affecting the entire batch of mussels.
We found that it was important to frequently replace culture chambers and sand as well. The weekly replacement removes excess fecal matter and unconsumed food that might otherwise contribute to poor water quality, for example elevated concentrations of ammonia.
Frequent water renewal was another important piece of our feeding system’s efficiency. As with the chamber replacement, periodically renewal of the culture water helps maintain optimal water quality.
The final element for success is ensuring that feeding is regulated and occurs often. New food is provided alongside water into the chamber every 1 or 2 hours. Considering these factors, the pulsed flow auto-feeding system can be modified and used for large- or small-scale mussel cultures.
The system was an effective means of generating large batches of robust juvenile mussels for reintroduction trials and restoration. The system is best suited for culture of juveniles during the first few weeks or months, which is a critical period. The system’s automation increases precision of feeding and water delivery and decreases the potential for human errors. Use of this system has also decreased the batch-to-batch variability in survival that was common in previous systems.
We are hopeful that the advancements we have made in mussel culturing can aid in future mussel research and restoration. At the USGS this technology has made it possible to investigate the effects of contaminants, sediments, and nutrients on the viability of mussels.
Cited References
Williams, J.D., Warren Jr., M.L., Cummings, K.S., Harris, J.L., and Neves, R.J., 1993, Conservation Status of Freshwater Mussels of the United States and Canada: Fisheries, v. 18, no. 9, p. 6-22.
Lydeard, C., Cowie, R.H., Ponder, W.F., Bogan, A.E., Bouchet, P., Clark, S.A., Cummings, K.S., Frest, T.J., Gargominy, O., Herbert, D.G. and Hershler, R., 2004, The global decline of nonmarine mollusks: BioScience, v. 54, no. 4, p. 321–330.
Strayer, D.L., Downing, J.A., Haag, W.R., King, T.L., Layzer, J.B., Newton, T.J. and Nichols, J.S., 2004, Changing perspectives on pearly mussels, North America's most imperiled animals: BioScience, v. 54, no. 5, p. 429–439.
Haag, W.R., 2012, North American freshwater mussels: natural history, ecology, and conservation. Cambridge University Press.
Lopes-Lima M, Sousa R, Geist J, Aldridge DC, Araujo R, Bergengren J, Bespalaya Y, Bodis E, Burlakova L, Van Damme D, Douda K, Froufe E, Georgiev D, Gumpinger C, Karatayev A, Kebapci U, Killeen I, Lajtner J, Larsen BM, Lauceri R, Legakis A, Lois S, Lundberg S, Moorkens E, Motte G, Nagel KO, Ondina P, Outeiro A, Paunovic M, Prie V, von Proschwitz T, Riccardi N, Rudzite M, Rudzitis M, Scheder C, Seddon M, Sereflisan H, Simic V, Sokolov, S, Stoeckl K, Taskinen J, Teixeira A, Thielen F, Trichkova T, Varandas S, Vicentini H, Zajac K, Zajac T, Zogaris S, 2017, Conservation status of freshwater mussels in Europe: state of the art and future challenges: Biological Reviews: v. 92, no. 1, p. 572–607.
Patterson, M.A., Mair, R.A., Eckert, N.L., Gatenby, C.M., Brady, T., Jones, J.W., Simmons, B.R. and Devers, J.L., 2018, Freshwater mussel propagation for restoration. Cambridge University Press.
O'Brien, C., Nez, D., Wolf, D. and Box, J.B., 2013, Reproductive Biology of Anodonta californiensis, Gonidea angulata, and Margaritifera falcata (Bivalvia: Unionoida) in the Middle Fork John Day River, Oregon: Northwest Science, v. 87, no. 1, p. 59–73.
Wang, N., Kunz, J.L., Ivey, C.D., Ingersoll, C.G., Barnhart, M.C. and Glidewell, E.A., 2017. Toxicity of chromium (VI) to two mussels and an amphipod in water-only exposures with or without a co-stressor of elevated temperature, zinc, or nitrate. Archives of Environmental Contamination and Toxicology, v. 72, no. 3, p. 449–460.
Return to Fish and Invertebrate Toxicology
Freshwater mussels are one of the most critically imperiled groups of organisms worldwide. Encouraging mussel reproduction in a controlled setting has become an area of interest due to their widespread population declines and regional extinctions (Williams et al., 1993; Lydeard et al., 2004; Strayer et al., 2004; Haag 2012; Lopes-Lima et al., 2017). With constantly evolving techniques, rearing healthy populations of mussels in a laboratory is a promising option. These laboratory-raised mussels are used for studies to better understand population declines and in reintroduction efforts (Patterson et al., 2018).
The Issue:
Young freshwater mussels are so small and delicate that captive culture presents challenges for handling, feeding, avoiding predation, and maintaining a suitable habitat. Despite advances in captive propagation, the rearing of mussels between newly metamorphosed juveniles (approximately the size of tip of pencil) and those that are 2- to 4-months-old (approximately the size of pencil eraser) is often hindered by low survival or growth (Patterson et al., 2018). Low growth rates increase the time and resources required to obtain mussels of a target size for restoration.
Addressing the Issue:
The objective of the study was to develop a system for large-scale laboratory culture. This included starting with newly transformed mussels of different species, two of which are commonly cultured (L. siliquoidea and Villosa iris) and two that have been historically challenging to culture (Anodonta californiensis and Margaritifera falcata). Newly transformed mussels are the earliest life stage of the mussel and start to resemble the shape of the adult. These small young mussels have historically been more challenging to culture (O’Brien et al., 2013; Wang et al., 2017). To optimize survival and good growth rates we developed an auto feeding system.
- The auto-feeding system is used to deliver 1 L of fresh culture water (A) into each mixing cell (B) every hour. Just before the mixing cells fill with water, a pump (C) is triggered to deliver 16 ml of food (algal mixture) into each mixing cell.
- Food (algal mixture) is maintained at <12°C in an aerated container in a cooler with ice packs (D).
- Water and food flow from the mixing cells (B) into the water delivery tubes (E) that evenly divide water into each culture beaker. Boxes of beakers are maintained in a temperature-controlled water bath (F).
- Juvenile mussels are held in 300-ml beakers with a thin layer of sand substrate (~150-µm particles). The water in the beakers is static except for pulses of food and water delivered every hour. Excess water flows out through the screen-covered hole and is not reused.
- Beakers and sand are replaced weekly or biweekly. Mussels are separated from the sand by sieving through a 250-µm mesh screen.
Results/Future Results:
During the development of the pulsed flow-through auto-feeding system, five elements emerged as critical to the success of the system.
One of our keys to success was found in the presence of a sand substrate in culture chambers. Sand allows for a more uniform distribution of food on the bottom of test chambers and limits the accumulation of debris, fungi, and algae on mussel shells. The advantages of using silica sand, rather than fine sediment, in the present system are that (1) silica sand is commercially available, (2) small juveniles can be easily isolated from fine sand particles, and (3) sand has less potential impact on water quality.
Isolating of smaller batches of mussels in individual beakers also proved to be crucial to our progress. This isolation increases the likelihood of detecting problems early (e.g., signs of disease or blooms of competitors or predators) and helps prevent problems from affecting the entire batch of mussels.
We found that it was important to frequently replace culture chambers and sand as well. The weekly replacement removes excess fecal matter and unconsumed food that might otherwise contribute to poor water quality, for example elevated concentrations of ammonia.
Frequent water renewal was another important piece of our feeding system’s efficiency. As with the chamber replacement, periodically renewal of the culture water helps maintain optimal water quality.
The final element for success is ensuring that feeding is regulated and occurs often. New food is provided alongside water into the chamber every 1 or 2 hours. Considering these factors, the pulsed flow auto-feeding system can be modified and used for large- or small-scale mussel cultures.
The system was an effective means of generating large batches of robust juvenile mussels for reintroduction trials and restoration. The system is best suited for culture of juveniles during the first few weeks or months, which is a critical period. The system’s automation increases precision of feeding and water delivery and decreases the potential for human errors. Use of this system has also decreased the batch-to-batch variability in survival that was common in previous systems.
We are hopeful that the advancements we have made in mussel culturing can aid in future mussel research and restoration. At the USGS this technology has made it possible to investigate the effects of contaminants, sediments, and nutrients on the viability of mussels.
Cited References
Williams, J.D., Warren Jr., M.L., Cummings, K.S., Harris, J.L., and Neves, R.J., 1993, Conservation Status of Freshwater Mussels of the United States and Canada: Fisheries, v. 18, no. 9, p. 6-22.
Lydeard, C., Cowie, R.H., Ponder, W.F., Bogan, A.E., Bouchet, P., Clark, S.A., Cummings, K.S., Frest, T.J., Gargominy, O., Herbert, D.G. and Hershler, R., 2004, The global decline of nonmarine mollusks: BioScience, v. 54, no. 4, p. 321–330.
Strayer, D.L., Downing, J.A., Haag, W.R., King, T.L., Layzer, J.B., Newton, T.J. and Nichols, J.S., 2004, Changing perspectives on pearly mussels, North America's most imperiled animals: BioScience, v. 54, no. 5, p. 429–439.
Haag, W.R., 2012, North American freshwater mussels: natural history, ecology, and conservation. Cambridge University Press.
Lopes-Lima M, Sousa R, Geist J, Aldridge DC, Araujo R, Bergengren J, Bespalaya Y, Bodis E, Burlakova L, Van Damme D, Douda K, Froufe E, Georgiev D, Gumpinger C, Karatayev A, Kebapci U, Killeen I, Lajtner J, Larsen BM, Lauceri R, Legakis A, Lois S, Lundberg S, Moorkens E, Motte G, Nagel KO, Ondina P, Outeiro A, Paunovic M, Prie V, von Proschwitz T, Riccardi N, Rudzite M, Rudzitis M, Scheder C, Seddon M, Sereflisan H, Simic V, Sokolov, S, Stoeckl K, Taskinen J, Teixeira A, Thielen F, Trichkova T, Varandas S, Vicentini H, Zajac K, Zajac T, Zogaris S, 2017, Conservation status of freshwater mussels in Europe: state of the art and future challenges: Biological Reviews: v. 92, no. 1, p. 572–607.
Patterson, M.A., Mair, R.A., Eckert, N.L., Gatenby, C.M., Brady, T., Jones, J.W., Simmons, B.R. and Devers, J.L., 2018, Freshwater mussel propagation for restoration. Cambridge University Press.
O'Brien, C., Nez, D., Wolf, D. and Box, J.B., 2013, Reproductive Biology of Anodonta californiensis, Gonidea angulata, and Margaritifera falcata (Bivalvia: Unionoida) in the Middle Fork John Day River, Oregon: Northwest Science, v. 87, no. 1, p. 59–73.
Wang, N., Kunz, J.L., Ivey, C.D., Ingersoll, C.G., Barnhart, M.C. and Glidewell, E.A., 2017. Toxicity of chromium (VI) to two mussels and an amphipod in water-only exposures with or without a co-stressor of elevated temperature, zinc, or nitrate. Archives of Environmental Contamination and Toxicology, v. 72, no. 3, p. 449–460.
Return to Fish and Invertebrate Toxicology