**2. Culture of glochidia in artificial media**

Glochidia were cultured in artificial media according to Kovitvadhi, (2000); Uthaiwan et al., (2002) and Kovitvadhi et al., (2006) until they transformed to 0-day-old juveniles. The details in each step of glochidia culture in artificial media were as follows:

<sup>\*</sup> Corresponding Author

#### **2.1 Composition of artificial media**

The composition of artificial media for culture of glochidia is shown in Table 1. Artificial media were based on those improved by Keller & Zam (1990), which consisted of a modification from the formulae of Isom & Hudson (1982). The main differences concerned the composition of the commercial media M199. While the protein source was exclusively horse serum in Keller & Zam (1990) and common carp, *Cyprinus carprio* fish plasma was used as an alternative support to the medium for cultured glochidia of *H.* (*L.*) *myersiana* (Uthaiwan et al., 2002; Kovitvadhi et al., 2006, 2007, 2008, 2009), *Hyriopsis* (*Hyriopsis*) *bialatus* (Areekijeree et al., 2006; Supannapong et al., 2008; Srakaew et al., 2010; Chumnanpuen et al., 2011; Kovitvadhi & Kovitvadhi., preparation), *Chamberlainia hainesiana* (Kovitvadhi et al., submitted) and *Anodonta cygnea* (Lima et al., 2006). Glochidia could transform into juvenile in the media containing common carp fish plasma as protein source. These glochidia were completely transformed within 8-11 days with a survival rate up to 93% except 34% of *A. cygnea*. All surviving larvae transformed into the juvenile stage except *A. cygnea* <34%. For these reasons, composition of artificial medium (Table 1) was according to Uthaiwan et al. (2002), based on Isom & Hudson (1982) and Keller & Zam (1990), was proposed for culture of glochidia.


Table 1. Composition of artificial medium for culture of glochidia

#### **2.2 Preparation of glochidia media**

#### **2.2.1 M199 preparation**

Dissolved one packet of M199 powder (Gibco, No. 6231100-035) in 1 liter volume of sterile distilled water and added 2 g of NaHCO3. Thereafter, M199 was filtered through 0.45 and 0.20 µm filter paper, respectively and kept at 4 °C.

#### **2.2.2 Fish plasma preparation**

Common carp was anesthetized with 50 mg/l of quinaldine. Fish blood was collected from the caudal vein, in the tail area, using a syringe needle no. 18 which was coated with sodium heparin at 1000 unit/ml. The blood sample was placed into sterile plastic test tube and centrifuged at 1000 and 3000 rpm for 10 min each. Plasma portion (clear yellow in colour) was separated and placed into the new test tube and centrifuged at 3000 rpm for 10 min. Then, plasma was separated and filtered through 0.45 and 0.20 µm filter paper, respectively and kept at -10 to -20 °C.

#### The composition of antibiotics and antimycotic chemicals (Isom & Hudson, 1982) is shown in Table 2. Compound Concentration (g / ml ) **Antibiotics**  Carbenicillin 100

100 100

**2.2.3 Antibiotics and antimycotic preparation** 

Amphotericin B 5

270 Aquaculture

The composition of artificial media for culture of glochidia is shown in Table 1. Artificial media were based on those improved by Keller & Zam (1990), which consisted of a modification from the formulae of Isom & Hudson (1982). The main differences concerned the composition of the commercial media M199. While the protein source was exclusively horse serum in Keller & Zam (1990) and common carp, *Cyprinus carprio* fish plasma was used as an alternative support to the medium for cultured glochidia of *H.* (*L.*) *myersiana* (Uthaiwan et al., 2002; Kovitvadhi et al., 2006, 2007, 2008, 2009), *Hyriopsis* (*Hyriopsis*) *bialatus* (Areekijeree et al., 2006; Supannapong et al., 2008; Srakaew et al., 2010; Chumnanpuen et al., 2011; Kovitvadhi & Kovitvadhi., preparation), *Chamberlainia hainesiana* (Kovitvadhi et al., submitted) and *Anodonta cygnea* (Lima et al., 2006). Glochidia could transform into juvenile in the media containing common carp fish plasma as protein source. These glochidia were completely transformed within 8-11 days with a survival rate up to 93% except 34% of *A. cygnea*. All surviving larvae transformed into the juvenile stage except *A. cygnea* <34%. For these reasons, composition of artificial medium (Table 1) was according to Uthaiwan et al. (2002), based on Isom & Hudson (1982) and Keller &

Composition of media The ratio of artificial medium

Dissolved one packet of M199 powder (Gibco, No. 6231100-035) in 1 liter volume of sterile distilled water and added 2 g of NaHCO3. Thereafter, M199 was filtered through 0.45 and

Common carp was anesthetized with 50 mg/l of quinaldine. Fish blood was collected from the caudal vein, in the tail area, using a syringe needle no. 18 which was coated with sodium heparin at 1000 unit/ml. The blood sample was placed into sterile plastic test tube and centrifuged at 1000 and 3000 rpm for 10 min each. Plasma portion (clear yellow in colour) was separated and placed into the new test tube and centrifuged at 3000 rpm for 10 min. Then, plasma was separated and filtered through 0.45 and 0.20 µm filter paper, respectively

**2.1 Composition of artificial media** 

Zam (1990), was proposed for culture of glochidia.

M199 2

Common carp fish plasma 1

Antibiotics and antimycotic 0.5

0.20 µm filter paper, respectively and kept at 4 °C.

**2.2 Preparation of glochidia media** 

**2.2.2 Fish plasma preparation** 

**2.2.1 M199 preparation** 

and kept at -10 to -20 °C.

Table 1. Composition of artificial medium for culture of glochidia

Table 2. Combination of antibiotics and antimycotic for culture of glochidia.

#### **2.3 Glochidia media preparation**

Gentamicin sulfate

Rifampin

**Antimycotic** 

Medium199 (see Section 2.2.1), fish plasma (see Section 2.2.2) and antibiotics/antimycotic (see Section 2.2.3) were mixed in the ratio 2:1:0.5, respectively (Table 1). The artificial media were divided into sterile plastic test tubes and kept at -10 to -20 °C for stocking culture media.

#### **2.4 Glochidia preparation**

Adult freshwater mussel were collected from the natural habitat. They were sexually identified by microscopic observation of sperms and eggs in fluid sucked from the gonads by use of a sterile syringe. Fifteen female and fifteen male adult mussels were cultured together in a cylinder net cage (diameter 50 cm × height 50 cm) in an earthen pond for the production of mature glochidia. They were allowed to feed freely on natural food. After 1-2 weeks, all females were observed marsupial colour by tongs to open the shell slightly, which marsupial colour indicates the development of larval stage. In the immature stage, glochidia was yellow in colour, while partially brown colour was at the beginning of maturity. Only completely brown marsupia of gravid mussel was selected in order to examine the strong and suitable glochidia for culturing in artificial media. Thereafter, the outer shell of gravid mussel with completely brown marsupia was washed with tap water and then sterile tap water. The glochidia were sucked by using a sterilized 1 ml syringe and discharged into depression or well slide with sterilized distilled water. Then, the glochidia were observed under a light microscope (×400). If their shells periodically closed, they were sucked according to abovementioned. Later, they were cleaned to eradicate tissue residues, mucus and glochidia shell fragments by spraying sterilized distilled water onto them. Complete cleaning and stronger glochidia were used for culture. Glochidia from gravid mussels should be cultured in artificial media within 5 h after harvesting.

#### **2.5 Glochidia culture**

Approximately 5000–6000 glochidia were transferred into a culture dish (90×15 mm) containing 10 ml of artificial medium (Table 1). The culture dishes were placed in a lowtemperature incubator at 25 C with 5% CO2. The culture medium was removed and replaced with fresh medium in the middle of cultured period. Finally, 4 ml of sterilized distilled water was added to the culture dish to stimulate the transformation when the mantle (Fig. 1A) was observed before 1 day of transformation from glochidia into juveniles. Juvenile transformation was observed under a light microscope (×400) for the movement of juvenile foot (Fig. 1B) and also juvenile movement as an indicator of the glochidia transformation success into juvenile stage.

Fig. 1. Light microscope of glochidial development to early juvenile. A; Glochidial development with the mantle edge bordering shell outside (arrow). B; Early juvenile with a foot (arrow). Bar = 100 m (Uthaiwan et.al., 2001).

#### **3. Selection of phytoplankton food species for juveniles**

Phytoplankton has proven to be a vital source of nutrient for several species of freshwater mussel juveniles (Hudson & Isom, 1984; Gatenby et al., 1996; Gatenby et al., 1997; ÓBeirn et al., 1998; Uthaiwan et al., 2001). Moreover, Kovitvadhi et al. (2000) reported that phytoplankton contributed to 99% of the gastrointestinal tract content of the adult freshwater pearl mussel, *H. (L.) myersiana* in natural habitat. Consequently, phytoplankton from the gastrointestinal tract of adult were cultured and selected for juvenile feeding. Collecting mussels of different sizes from natural habitat, which phytoplankton existed in the gastrointestinal tract then it was cutting and sucking to culture in sterilized water with f/2 media (Guillard & Ryther, 1962). Then, they were cultured under light not less than 10,000 Lux for 12 h as well as in mixed about 3% carbon dioxide. From there, phytoplankton to be cultured were in the process of sub-culture and purified every 2-10 days by streak plating technique (Hoshaw & Rosowski, 1973). Streak plates were placed under light until the single colonies of phytoplankton appeared which might last for 10-30 days. Thereafter, those phytoplankton were kept in slant tube. Whenever phytoplankton were required for feeding juveniles, those phytoplankton were multiplied in 1 liter bottle by using the same separated formulae. Kovitvadhi et al. (2006) as abovementioned found ten species of phytoplankton; *Ankistrodesmus gracilis*, *Chlamydomonas* sp., *Chlorella* sp.1, *Chlorella* sp.2, *Kirchneriella incurvata*, *Monoraphidium* sp., *Navicula* sp., *Scenedesmus* sp., *Stichococcus* sp. and *Coccomyxa* sp. Thereafter, all phytoplankton were selected for suitable cultured juvenile which should be considered based on 6 criteria according to Areekijseree et al. (2006); Kovitvadhi et al. (2006); Supannapong et al., (2008): (1) size (2) capability of filter into gastrointestinal tract by observing under microscope within 30 min and 1.30 h after giving phytoplankton (3) the movement of cilia at gill, mantle and foot by observing under microscope (4) color changes in gastrointestinal tract which should be from green to yellow or brown and shape from normal to debris (5) carbohydrate and protein contents of

distilled water was added to the culture dish to stimulate the transformation when the mantle (Fig. 1A) was observed before 1 day of transformation from glochidia into juveniles. Juvenile transformation was observed under a light microscope (×400) for the movement of juvenile foot (Fig. 1B) and also juvenile movement as an indicator of the glochidia

 Fig. 1. Light microscope of glochidial development to early juvenile. A; Glochidial

development with the mantle edge bordering shell outside (arrow). B; Early juvenile with a

Phytoplankton has proven to be a vital source of nutrient for several species of freshwater mussel juveniles (Hudson & Isom, 1984; Gatenby et al., 1996; Gatenby et al., 1997; ÓBeirn et al., 1998; Uthaiwan et al., 2001). Moreover, Kovitvadhi et al. (2000) reported that phytoplankton contributed to 99% of the gastrointestinal tract content of the adult freshwater pearl mussel, *H. (L.) myersiana* in natural habitat. Consequently, phytoplankton from the gastrointestinal tract of adult were cultured and selected for juvenile feeding. Collecting mussels of different sizes from natural habitat, which phytoplankton existed in the gastrointestinal tract then it was cutting and sucking to culture in sterilized water with f/2 media (Guillard & Ryther, 1962). Then, they were cultured under light not less than 10,000 Lux for 12 h as well as in mixed about 3% carbon dioxide. From there, phytoplankton to be cultured were in the process of sub-culture and purified every 2-10 days by streak plating technique (Hoshaw & Rosowski, 1973). Streak plates were placed under light until the single colonies of phytoplankton appeared which might last for 10-30 days. Thereafter, those phytoplankton were kept in slant tube. Whenever phytoplankton were required for feeding juveniles, those phytoplankton were multiplied in 1 liter bottle by using the same separated formulae. Kovitvadhi et al. (2006) as abovementioned found ten species of phytoplankton; *Ankistrodesmus gracilis*, *Chlamydomonas* sp., *Chlorella* sp.1, *Chlorella* sp.2, *Kirchneriella incurvata*, *Monoraphidium* sp., *Navicula* sp., *Scenedesmus* sp., *Stichococcus* sp. and *Coccomyxa* sp. Thereafter, all phytoplankton were selected for suitable cultured juvenile which should be considered based on 6 criteria according to Areekijseree et al. (2006); Kovitvadhi et al. (2006); Supannapong et al., (2008): (1) size (2) capability of filter into gastrointestinal tract by observing under microscope within 30 min and 1.30 h after giving phytoplankton (3) the movement of cilia at gill, mantle and foot by observing under microscope (4) color changes in gastrointestinal tract which should be from green to yellow or brown and shape from normal to debris (5) carbohydrate and protein contents of

transformation success into juvenile stage.

foot (arrow). Bar = 100 m (Uthaiwan et.al., 2001).

**3. Selection of phytoplankton food species for juveniles** 

phytoplankton (6) efficiency of digesting carbohydrate and protein of phytoplankton by using crude enzyme extracts from juveniles (*in vitro* digestibility). The major phytoplankton were suitable for culturing juvenile of freshwater mussel, namely *Chlorella* sp. 2 and *K. incurvata* (Table 3).


Table 3. Characteristic of phytoplankton isolation from gastrointestinal tract of adult freshwater mussel in natural habitat and *in vitro* digestibility for carbohydrate (µg maltose mg plankton-1) and protein (µg DL-alanine equivalent mg plankton-1) of ten algal species at seven days old, using crude enzyme extracts from 30-day-old juveniles. 1 Kovitvadhi et al. (2006), 2 Supannapong et al. (2008).

#### **4. Juvenile culture**

After glochidia transformed to juvenile in artificial medium, water of recirculating aquaculture system was added in the ratio of medium to water equaled 3:1, 1:1, 1:3 for 30 min, added *K. incurvata and Chlorella* sp. in ratio 1:1 at density of 1×105 cells per ml in ratio medium to water equaled 1:3. From there, cultured juvenile was transferred to culture units. Culture of juvenile stage could be divided into 3 stages, namely first stage beginning from 0 days old juvenile until two shell mussel completely closed. Kovitvadhi et al. (2008) cultured juvenile 0-120 days old of freshwater mussel, *H.* (*L.*) *myersiana* by recirculating aquaculture system (Fig. 2). This system comprised of three filter cabinets made of 6 mm thick acrylic (particulate filter cabinet, macrophytes filter cabinet and biological filter cabinet), one water resting cabinet (Fig. 2E) and nine plastic culture units (Fig. 2F). The size of particulate filter cabinet (Length×Width×Height×Water level = 46×35×51×42 cm) was divided into two equal parts, of which the first part was filled with a 30 cm thick nylon filter (Fig. 2B). Water flowed through this filter and via the second to the macrophytes filter cabinet. The size of macrophytes filter cabinet (Length×Width×Height×Water level = 80×40×51×42 cm) was divided into four equal units. Each unit contained 57 ambulia plants, *Limnophila heterophylla* (Raxb.) Bentham; these, 228 plants in total, were introduced when they were 6 cm in height and had an average weight of 2.69±0.13 g (Fig. 2C). The plants were removed and replaced when their tips reached the water surface. The upper parts of the cabinets were equipped with three fluorescent lamps (each 20 W) 25 cm above the water surface (light intensity at water surface equal to 5320 lux; 24 h) (Fig. 2G). The water then flowed into the biological filter cabinet (60×34×51×42 cm) filled with bioball to full capacity (Fig. 2D) and then to the resting cabinet (46×41×51×42 cm). In the resting cabinet there were two water pumps: The first returned water to the particulate filter cabinet at the rate of 1 l per minute continuously and the second pumped water at 20 ml per minute to nine plastic culture units (each 84×14×15×7 cm). This pump was stopped for 1 h after phytoplankton was introduced into the culture unit. The bottom of the culture unit was filled with sand (<120 µm) at 0.27 g/cm2. The inside of the culture unit was divided into two section, as described previously, but of different sized (section 1-66.1×14×15×7 cm; section 2-17.9×14×15×7 cm). The first section also consisted of five acrylic sheets jutting from the walls on alternate sides. Juveniles were fed *Chlorella* sp. and *K. incurvata*. Each species of algae was collected from the 100 l by being pumped through 0.3 µm ceramic filters and then separated from the water by centrifuging at 8000 ×g. The sediments of the two algal species were mixed at a ratio of 1:1 wet weight and kept in a freezer. When required, the mixture was brought to room temperature then sucked by Pasteur pipette into the all plastic culture unit to an algae density of 1×105 cells per ml. Algae were supplied twice a day (06.00 h and 18.00 h), and the frozen stock was usually used within 7 days of collection.

Second stage, thirty-five juveniles (120 days old) were transferred to culture units (20×12×72 cm). The culture units had all four vertical sides lined with nylon net (0.42 mm mesh size) and each had a plastic lid with holes to cover the top. The lower part of the culture unit consisted of a section 2 cm in height, which fitted snugly into the culture unit, from which it could be removed (Fig. 3). This lower part contained 400 g of sand (<425 µm in size). The juveniles were placed directly on the sand. The culture unit was then hung in the earthen pond; the base of the culture unit was adjusted to a position approximately 50 cm below the water surface. The juveniles fed by filtering phytoplankton from the water in the earthen pond. All mussels from the culture unit were rinsed every 10 days.

After glochidia transformed to juvenile in artificial medium, water of recirculating aquaculture system was added in the ratio of medium to water equaled 3:1, 1:1, 1:3 for 30 min, added *K. incurvata and Chlorella* sp. in ratio 1:1 at density of 1×105 cells per ml in ratio medium to water equaled 1:3. From there, cultured juvenile was transferred to culture units. Culture of juvenile stage could be divided into 3 stages, namely first stage beginning from 0 days old juvenile until two shell mussel completely closed. Kovitvadhi et al. (2008) cultured juvenile 0-120 days old of freshwater mussel, *H.* (*L.*) *myersiana* by recirculating aquaculture system (Fig. 2). This system comprised of three filter cabinets made of 6 mm thick acrylic (particulate filter cabinet, macrophytes filter cabinet and biological filter cabinet), one water resting cabinet (Fig. 2E) and nine plastic culture units (Fig. 2F). The size of particulate filter cabinet (Length×Width×Height×Water level = 46×35×51×42 cm) was divided into two equal parts, of which the first part was filled with a 30 cm thick nylon filter (Fig. 2B). Water flowed through this filter and via the second to the macrophytes filter cabinet. The size of macrophytes filter cabinet (Length×Width×Height×Water level = 80×40×51×42 cm) was divided into four equal units. Each unit contained 57 ambulia plants, *Limnophila heterophylla* (Raxb.) Bentham; these, 228 plants in total, were introduced when they were 6 cm in height and had an average weight of 2.69±0.13 g (Fig. 2C). The plants were removed and replaced when their tips reached the water surface. The upper parts of the cabinets were equipped with three fluorescent lamps (each 20 W) 25 cm above the water surface (light intensity at water surface equal to 5320 lux; 24 h) (Fig. 2G). The water then flowed into the biological filter cabinet (60×34×51×42 cm) filled with bioball to full capacity (Fig. 2D) and then to the resting cabinet (46×41×51×42 cm). In the resting cabinet there were two water pumps: The first returned water to the particulate filter cabinet at the rate of 1 l per minute continuously and the second pumped water at 20 ml per minute to nine plastic culture units (each 84×14×15×7 cm). This pump was stopped for 1 h after phytoplankton was introduced into the culture unit. The bottom of the culture unit was filled with sand (<120 µm) at 0.27 g/cm2. The inside of the culture unit was divided into two section, as described previously, but of different sized (section 1-66.1×14×15×7 cm; section 2-17.9×14×15×7 cm). The first section also consisted of five acrylic sheets jutting from the walls on alternate sides. Juveniles were fed *Chlorella* sp. and *K. incurvata*. Each species of algae was collected from the 100 l by being pumped through 0.3 µm ceramic filters and then separated from the water by centrifuging at 8000 ×g. The sediments of the two algal species were mixed at a ratio of 1:1 wet weight and kept in a freezer. When required, the mixture was brought to room temperature then sucked by Pasteur pipette into the all plastic culture unit to an algae density of 1×105 cells per ml. Algae were supplied twice a day (06.00 h and 18.00 h), and the

frozen stock was usually used within 7 days of collection.

pond. All mussels from the culture unit were rinsed every 10 days.

Second stage, thirty-five juveniles (120 days old) were transferred to culture units (20×12×72 cm). The culture units had all four vertical sides lined with nylon net (0.42 mm mesh size) and each had a plastic lid with holes to cover the top. The lower part of the culture unit consisted of a section 2 cm in height, which fitted snugly into the culture unit, from which it could be removed (Fig. 3). This lower part contained 400 g of sand (<425 µm in size). The juveniles were placed directly on the sand. The culture unit was then hung in the earthen pond; the base of the culture unit was adjusted to a position approximately 50 cm below the water surface. The juveniles fed by filtering phytoplankton from the water in the earthen

**4. Juvenile culture** 

Fig. 2. Photographs of the recirculating used to rear freshwater pearl mussel juveniles. A; Recirculating aquaculture system, B; Particulate filter cabinet, C; Macroplants filter cabinet, D; Biological filter cabinet, E; Water resting cabinet, F; Plastic culture unit, G; Fluorescent box.

Fig. 3. Rearing container of juveniles (Kovitvadhi et al., 2006).

Third stage, juveniles (180 days old) were transferred to culture into natural habitat or the earthen pond by a cylinder net cage until adult.

#### **5. Adult culture**

At present, adult of some species freshwater pearl mussels had been successfully cultured in an earthen pond and natural habitat. They had high survival and could produce glochidia stage such as *H.* (*L.*) *myersiana* (Uthaiwan et al., 2002; Kovitvadhi et al., 2006, 2007, 2008, 2009), *H.* (*H.*) *bialatus* (Areekijeree et al., 2006; Supannapong et al., 2008; Srakaew et al., 2010; Chumnanpuen et al., 2011; Kovitvadhi & Kovitvadhi., preparation), *C. hainesiana* (Kovitvadhi et al., submitted). Fifteen female and fifteen male adult mussels, were cultured together in a cylinder net cage (diameter 50 cm × height 50 cm). Then, it was hung under the raft (Fig. 4) at 1.5-2 m deep from water surface which phytoplankton were plenty at this level. The cage was shaken every week for protecting biofouling attachment which could mass mortality.

Fig. 4. Raft for adult freshwater pearl mussel culture. (Kovitvadhi, 2008).

#### **6. Water analysis**

Prior to culturing mussel, water quality in habitat, was studied in water improvement suitable to growth and survival which conformed to Kovitvadhi et al. (2006, 2008) that juvenile cultured in the laboratory and the earthen pond nearby natural habitat (Kovitvadhi et al., 1998) (Table 4). Water quality parameters should be analyzed for freshwater pearl mussel culture: water temperature, pH, tubidity, conductivity, dissolved oxygen, total alkalinity, free carbon dioxide, total hardness, total ammonia nitrogen, nitrite, nitrate, phosphorus, silica and calcium. In this connection, juvenile stage had been more sensitive to environmental changes than another stages, particularly water qualities suitable and rather stable; water temperature, pH, free carbon dioxide, dissolved oxygen, nitrate and phosphorus and decreasing values; total alkalinity, total hardness, total ammonia nitrogen, silica, and calcium except nitrite that had increasing value (Fig. 5). When averaged water quality value was calculated to relationship with averaged survival value and shell length with equation: Y=b0+b1X+b2X2+b3X3 where Y is the survival or shell length, X is age (days), and b0, b1, b2 and b3 are parameters. It was found that survival of 0-120-day-old juveniles would have direct relationship with pH, total alkalinity, total hardness, silica and calcium with highly significant difference (P<0.01) and with reverse relationship to free carbon dioxide and nitrite (Table 5) (Kovitvadhi et al., 2008).


Third stage, juveniles (180 days old) were transferred to culture into natural habitat or the

At present, adult of some species freshwater pearl mussels had been successfully cultured in an earthen pond and natural habitat. They had high survival and could produce glochidia stage such as *H.* (*L.*) *myersiana* (Uthaiwan et al., 2002; Kovitvadhi et al., 2006, 2007, 2008, 2009), *H.* (*H.*) *bialatus* (Areekijeree et al., 2006; Supannapong et al., 2008; Srakaew et al., 2010; Chumnanpuen et al., 2011; Kovitvadhi & Kovitvadhi., preparation), *C. hainesiana* (Kovitvadhi et al., submitted). Fifteen female and fifteen male adult mussels, were cultured together in a cylinder net cage (diameter 50 cm × height 50 cm). Then, it was hung under the raft (Fig. 4) at 1.5-2 m deep from water surface which phytoplankton were plenty at this level. The cage was

shaken every week for protecting biofouling attachment which could mass mortality.

Fig. 4. Raft for adult freshwater pearl mussel culture. (Kovitvadhi, 2008).

dioxide and nitrite (Table 5) (Kovitvadhi et al., 2008).

Prior to culturing mussel, water quality in habitat, was studied in water improvement suitable to growth and survival which conformed to Kovitvadhi et al. (2006, 2008) that juvenile cultured in the laboratory and the earthen pond nearby natural habitat (Kovitvadhi et al., 1998) (Table 4). Water quality parameters should be analyzed for freshwater pearl mussel culture: water temperature, pH, tubidity, conductivity, dissolved oxygen, total alkalinity, free carbon dioxide, total hardness, total ammonia nitrogen, nitrite, nitrate, phosphorus, silica and calcium. In this connection, juvenile stage had been more sensitive to environmental changes than another stages, particularly water qualities suitable and rather stable; water temperature, pH, free carbon dioxide, dissolved oxygen, nitrate and phosphorus and decreasing values; total alkalinity, total hardness, total ammonia nitrogen, silica, and calcium except nitrite that had increasing value (Fig. 5). When averaged water quality value was calculated to relationship with averaged survival value and shell length with equation: Y=b0+b1X+b2X2+b3X3 where Y is the survival or shell length, X is age (days), and b0, b1, b2 and b3 are parameters. It was found that survival of 0-120-day-old juveniles would have direct relationship with pH, total alkalinity, total hardness, silica and calcium with highly significant difference (P<0.01) and with reverse relationship to free carbon

earthen pond by a cylinder net cage until adult.

**5. Adult culture** 

**6. Water analysis** 


1Kovitvadhi et al. (2006), 2Kovitvadhi et al. (1998).

Table 4. Water quality during culturing of 0-360 day-old juveniles and the adult mussel habitat of *H.* (*L.*) *myersiana* in the Mae Klong River, Kanchanaburi Province.


Table 5. Coefficient of correlation between average survival rate and water quality; average growth rate and water quality of juvenile *H.* (*L.*) *myersiana* cultured in recirculating aquaculture system every 10 days. (Kovitvadhi et al., 2008) (\* = *P*<0.05, \*\* = *P*<0.01, *ns* = not significant difference, *P*>0.05).

Fig. 5. Water quality during culture for 0-120 days of *Hyriopsis* (*Limnoscapha*) *myersiana* juveniles in recirculating aquaculture system (Kovitvadhi et al., 2008).
