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Vienneau Aquaculture has been working for several years to develop a technique for raising oysters in the water column rather than near the water surface. Initial results, although inconclusive in terms of growth (Lanteigne, 2008), highlighted constraints that could cause infestations of marine biofouling on culture structures, especially mussels. It sought to correct this situation by raising its culture structures to kill mussels through desiccation. Although this approach yielded positive results, it was affected by temperature variations, the intensity of the spat collection and the number of hydrozoans on the culture structures. This led to a 2007 research project on the use of a hot water bath to control biofouling. Based on the 2007 results, the company launched the second phase of its project, which aimed to validate a process for treating culture structures in hot water, perfecting a scalding process that is multipurpose in the sense that the company can treat a variety of its culture structures. This report presents the results of various trials performed in 2008 to perfect a scalding process better suited to the realities of the New Brunswick shellfish industry.
Bivalve aquaculture has grown considerably in Atlantic Canada over the past two decades. During this period, annual production has risen from 1,463 T (1986) to 28,039 T (2004) (Statistical Services, Department of Fisheries and Oceans Canada, DFO). In New Brunswick, the culture of the American oyster (Crassostrea virginica)dominates the aquaculture industry, with an estimated annual production of approximately 600 T per year, corresponding to 12% of commercial oyster production in the Atlantic. Although this number is relatively low, it is estimated that the New Brunswick shellfish industry will enter a significant growth phase in the near future (GTA Consultants en pêches, 2003). In 2005, there was a total of 671 leases in New Brunswick, 660 for oyster farming, specifically (Comeau et al., 2006). The various culture techniques currently used can be grouped under two categories: off-bottom culture techniques (i.e. suspended) and bottom culture techniques. The first technique is used for most of the commercial oyster production in New Brunswick (~90%). In short, these techniques are designed to suspend oysters in the upper water column to promote their growth, control predators and allow for easy access to the oysters. These techniques generally use Vexar or Durethene bags commonly referred to simply as "bags." However, one new technique that northeastern New Brunswick farmers have recently begun using involves lines of stacked cages suspended in the water column. The near-universal use of floating bags and cages is subject to various production constraints that differ from those of off-bottom culture production. Of these constraints, Vienneau Aquaculture Inc. believes that marine biofouling will be a priority issue from this point forward. It is currently developing a cage culture technique (Lanteigne, 2005; 2006; 2007). The presence of undesirable organisms on cultured oysters and suspended culture structures can reduce oyster growth rate and quality, and cause significant damage to equipment (increased abrasion, fragility and load). As well, communities of organisms that develop on cages slow the water current through the mesh, changing the drag force on structures. Although many methods have been developed to control marine biofouling, it remains an issue on culture sites worldwide (Braithwaite and McEvoy, 2005). A recent study suggests that cleaning oysters generates costs equal to 20% of the commercial value of the final product, and that marine biofouling can reduce oyster growth rate by 40% (Willemsen, 2005). According to Tenore et al., 1973, the Blue Mussel produces more biodeposition than the Oyster. This phenomenon, which is associated with the slowing of water currents through culture structures according to infestation intensity, could foster the accumulation of silt on oysters.
Recent observations suggest that the main species that fall in the category of marine biofouling on oyster culture in New Brunswick are the Mussel (Mytilus edulis), the Oyster (C. virginica (over-collection)) and the Barnacle (unidentified species). The collection of mussels on cultivated oysters and suspension culture systems currently appears to be the greatest biofouling problem, particularly in production areas in northeastern New Brunswick (Lanteigne, 2008). A recent study supporting these observations reveals that communities that develop on cages of fish farmed in seawater in the Gulf of Maine, south of New Brunswick, are always dominated by blue mussels (Greene and Grizzle, 2007). The same conclusions are reported in the Gaspé Peninsula and in northern New Brunswick on suspended longline systems of Sea Scallop (Placopecten magellanicus) culture, where the Blue Mussel and Wrinkled Rock Borer (Hiatella arctica) dominate biofouling communities (Claereboudt et al., 1994). The Mussel is a filter feeding bivalve mollusc that can compete with the Oyster for food, reducing the growth rate or meat yield of cultured oysters. Sea scallops cultivated in nets that are cleaned regularly have 68% greater muscle mass and 5% greater shell length than scallops cultivated in nets colonized by biofoulers (Claereboudt et al., 1994). Mussel spat is generally collected near the end of June or early July in the Shippagan area, and slightly earlier in Neguac Bay. Mussel collection in the Richibouctou area appears to extend over the entire seasonal period, which poses an even greater problem in terms of the management of culture structures and cages used for spat production, specifically (Daigle M., personal communication). This was also the case at the beginning of August 2006 in the Saint Simon area, where a significant collection of mussel spat was observed on Vienneau Aquaculture and Ferme Marine Lanteigne culture structures (Lanteigne, 2007). Mussels are frequently collected by scallop collectors set out in Baie des Chaleurs in mid-September, as well.
Although much less significant in number than mussels, a collection of oysters was observed on cultivated oysters in Neguac Bay, Saint Simon Bay and Bouctouche Bay, as well as on cultured oysters and lines of Dark Sea trays in Lamèque (Lanteigne, 2002; 2007). In our opinion, the growth of the industry and the increase in production volume in various bays in eastern New Brunswick may lead to an increase in the undesirable collection of oysters on culture structures and cultured oysters. Cleaning oysters infested with barnacles can be an arduous task because they must be removed by hand. Over-collection is prevalent in the Bouctouche area, where it is not unusual to see over 20 small oysters on one oyster cultivated in a floating bag. This can lead to a significant increase in operating costs.
Lastly, field observation suggests that the barnacle collection period corresponds well to that of oysters in New Brunswick, such that they could be removed at the same time. It is important to note that there are several species of barnacles, among them the Semibalanus balanoides, the Balanus balanus, and the B. crenatus, which are very well represented in the Gulf of St. Lawrence (Bourget, 1997). The seasonal trends of each species on the culture structures and the cultured oysters is as yet unknown. Large collections of barnacles can lead to significant operating costs and reduce the aesthetic quality of oysters. These species can also be very difficult to remove, which can also incur substantial additional costs during processing (Arakawa, 1980).
Marine biofouling issues are not limited to aquaculture. Several studies were conducted to establish measures to control marine biofouling in power plant cooling water, and are a significant source of information (Graham et al., 1975; Nel et al., 1996; Rajagopal, 2005). The marine transport industry also faces these types of constraints. However, unlike industries in which marine biofouling is an issue, few studies have examined the impact and use of competitive solutions in the aquaculture industry (Willemsen, 2005; Arakawa, 1980). The most widely-recognized control measures are mechanical removal or anti-fouling paint. Mechanical removal involves brushing and scratching the shells, or using powerful jets of water. These approaches require much time and effort (Hodson et al., 1997). In addition to cleaning, shells can also be immersed in warm or hot water, or chlorine, salt or lemon solutions (Arakawa, 1980). For example, calcium hydroxide dips, brine dips, acetic acid dips and hypochlorite solution dips successfully controlled growth of the Ascidian (Ciona intestinalis) on the Blue Mussel (Carver et al., 2003). A recent study revealed the fragility of oyster spat compared to mussel seed (André Mallet; pers. comm.). Two hours is sufficient to irradiate oyster spawn outside water, whereas it takes over 24 hours to do so with mussel seed. Exposure to air (desiccation) and heat has been widely tested to control marine biofouling on oysters in New Brunswick and elsewhere in the world. However, this technique kills the organisms without removing them (Willemsen, 2005). Coating the surface of the culture structures with biocide is an even more widespread approach used in aquaculture. However, the effectiveness of these substances, which are generally copper-based, is limited to one production season. It is also costly to apply them, and they are extremely polluting (Willemsen, 2005). Lastly, biofouling control measures that use grazers or predators have also been successfully developed recently (Lodeiros and Garcia, 2004; Hidu et al., 1981). However, the presence of the Rock Crab (Cancer irroratus) in floating bags seems to hinder measures to control biofouler colonization. The Crab has difficulty surviving in floating bags, likely because of the incessant movement of the culture structures (Lanteigne, 2003).
Major studies are currently being done in Europe to find effective, economic measures to control marine biofouling that apply to the aquaculture industry (Willemsen, 2005, projet CRAB : a collective approach to reduce the impact of biofouling for the aquaculture industry). Among the methods chosen, desiccation and hot water baths seem to be two effective approaches. French oyster farmers use the desiccation technique extensively, which lends itself well to the oyster table net technique. New Brunswick oyster farmers who use floating bags must turn them regularly to minimize the incidence of marine biofouling. However, according to Mallet et al. (2009), it is not necessary to turn the bags every month to manage biofouling. Vienneau Aquaculture Inc. and Aquaculture Acadienne Ltée tested desiccation in 2005 and 2006, respectively, on Dark Sea trays, bags and submersed cages to control marine biofouling (Lanteigne, 2006; 2007; Daigle M., person communication). Based on the weather conditions (humidity, rain) and specific composition of the attached organisms, up to three days of air exposure was required to remove mussel spat. However, this treatment may not be effective if the arborescent biofouler colony biomass (hydrozoans and macrophytic algae) is high. Arborescent organisms protect mussels and other undesirable organisms from desiccation (Lanteigne; 2007).
The Centre for the Experimentation and Development of Marine Aquaculture (CREAA) has been in operation in France since 2000. Its initial mandate was to help oyster farmers to choose proper temperatures and durations of oyster treatment in hot water. Experiments performed by the CREAA aimed to determine Cupped Oyster
(Crassostrea gigas) sensitivity to high temperatures used to remove undesirable spat attached to cultured oysters. Results achieved to date have enabled researchers to make certain recommendations. For example, it appears that oysters aged 12 to 18 months must be treated for 2 s at 80°C and gradually cooled, while oysters aged 30 months (40-60 g) can be treated at 85°C for 3 s (http://perso.orange.fr/creaa/doc/05_fiche_echaudage_
huitres.pdf). Although procedures were developed, it was deemed necessary to test them on species of interest at the culture site, which may host attached communities of organisms that differ from those prevalent on sites previously tested. For example, in Asia, treatment at 60°C for 10-15 s removes biofoulers without affecting the survival of the Cupped Oyster (Park et al., 1998), while in France, the CREAA suggests a shorter exposure time––85°C. Preliminary tests performed in P.E.I. reveal that this technique can be adapted to ostreaculture in New Brunswick. Based on results, large oysters resist 60°C water baths for up to 20 s, while spat can only resist water at this temperature for 5 s. The mussel collection was then successfully eradicated after 15 s at 60°C. Given that blue mussels cultivated by suspended culture have thinner shells than those of cupped oysters (and oysters cultivated on the bottom), they may be more sensitive to high-temperature baths. The following questions inevitably arise:
How does hot water treatment affect the size of the C. virginica?
What is the best strategy for removing biofoulers while minimizing oyster mortality or stress?
How does biodeposition impact the effectiveness of hot water treatments and, if applicable, can washing before treatment increase this effectiveness?
Studies led by Vienneau Aquaculture Inc. in 2007 confirmed the validity of this type of approach in controlling marine biofouling and in providing certain treatment measures for various oyster sizes, depending on the characteristics of the biofoulers (mussels, specifically), due to high-intensity mussel colonization (Lanteigne, 2008). However, the approach was based on the use of various treatment temperatures for set times due to the limitations of the scalding system available at the time. Based on the results, it was determined that treatment at 60°C would provide the best control of marine biofouling; this did not appear to affect the largest oysters. It would be more difficult to implement a measure in which water temperatures are varied according to size group. It would be easier to vary the duration of the treatment and maintain a set temperature, which involves validating treatment at 60°C for the various treatment durations.
The system used in 2007 for the first scalding trials was not flexible in terms of treatment duration. The development of a new scalding system should enable researchers to correct this limitation, which is deemed essential.
The New Brunswick oyster industry uses a variety of culture techniques. It is not easy to devise a scalding process that can use this approach to controlling biofoulers on all culture structures. Although there are machines available on the market, they are not adapted to the specific characteristics of cultivation activities practiced in New Brunwick and in Europe, which causes problems in terms of after-sales support.
One of the objectives in developing a new scalding system was to treat lines of 10-11 cages with better control of exposure time, which meant that treatment intensity for top cages would not be the same as for those on the bottom. The resulting differences between the various temperatures tested in 2007 revealed a narrow margin between oyster survival and that of biofoulers, particularly for very small oysters, which is another element that requires further investigation.
Research took place on Vienneau Aquaculture Inc. farms in Saint Simon Bay (in floating bags) and in Saint Simon Inlet, at the mouth of the bay (in Dark Sea trays). Trials using Dark Sea trays were also conducted in Neguac due to a significant barnacle infestation on the culture structures. Additional trials were conducted at Lamèque at the end of autumn because there had been a significant number of mortalities in stocks of 65-66 mm farmed oysters in Dark Sea trays on the Vienneau Aquaculture Inc. site.
The factorial experiment performed sought to determine the effect of brief exposure of culture structures and cultured oysters to 60°C water on the survival of mussels, oysters (including those collected during the second round) and barnacles attached to cultured oysters and culture structures from a prevention standpoint. The experiment also sought to determine the resistance of various sizes of oysters to hot water bath treatment. Specifically, the factors studied were the application period (3), the duration of treatment application (5), oyster size (3), the type of culture structure (2) and pre-treatment handling. Oysters belonging to various size groups were exposed to 60°C water for 5, 10, 15 and 20 seconds, and then compared to control structures.
Taking the seasonal evolution of marine biofouling properties into account, treatments were tested three times over the course of the growing season. Culture structures and various lots of oysters were identified at the beginning of the study to ensure that the structures remained operational. The first treatment was administered at the end of June, right after the peak mussel collection period. A second treatment was administered at the beginning of August, after the peak oyster spat and barnacle collection period (oyster spat and very small barnacles, mid-sized mussel spat and shells from the rims of very narrow oysters). The final treatment was administered at the end of September/mid-October, once the biofoulers had finished settling and before the over-wintering period. Oyster shells are thicker during this period. The impact of treatment on oyster size was also examined. Three size groups (25-35, 40-50 and 55-65 mm) of floating bags and two size groups (40-50 and 55-65 mm) of Dark Sea trays were treated.
Additional treatments were administered in August and October to determine whether or not washing the culture structures would improve the effectiveness of the treatment. A second set of culture structures with the same parameters as those presented in the previous section were treated after having been washed under a stream of water. However, due to the high incidence of death in the 55-65 mm oyster group in the Dark Sea trays during the fall, treatment was repeated for this size group in Lamèque in mid-November.
Given the low number of barnacles observed on culture structures in June and August, as well as the barnacle infestation reported by SaLant Aquaculture Inc. on its Dark Sea trays, a series of tests were performed in Neguac Bay. The Dark Sea trays were treated using the process used in Saint Simon, that is, water at 60°C for periods of 5, 10, 15 and 20 seconds. Only one size group was used––oysters of 55+ mm.
The sampling approach had to be adapted based on the material sampled, the targeted species and the targeted objectives. Six objectives were set and formed the basis of the sampling approach:
The first step involved emptying the bags and cages of oysters. The oysters were washed over a biofouler recovery system to remove all of the mussels. Biofoulers were harvested in a 1 mm mesh strainer. Due to the volume of the material, samples were divided to try to reduce the number of specimens in the samples to approximately 100 using a Fulton Splitter Counter. The cultivation bags and cages were then washed over the system to remove the mussels. Because of the volume of the material, the sample was divided using the same procedure to try to reduce the number of specimens in the samples to approximately 100. The mussel samples were then marked and stored to be analyzed later. A sample of approximately one tenth of the oyster volume per culture structure (15 oysters from the 55-65 mm size group, 20 oysters from the 40-50 mm size group and 50 oysters from the 25-35 mm size group) was removed and all of the barnacles and oysters from the second collection round were counted. Four quadrants with a surface area of 56.2 cm2 (7.5 x 7.5 cm) were analyzed on the sides of the bags and cages to determine the number of barnacles and oysters from the second collection round. Both living and dead oysters in the culture structures were counted.
Multivariate analyses (MANOVA) were performed to determine the effect of the date and the duration of treatment application and oyster size on the survival, growth and characteristics of communities of organisms attached to oysters.
The size of the oysters from the various size groups was measured at the beginning of the experiment and was 66.94 ± 6.54, 50.46 ± 5.94 and 37.85 ± 3.65 mm for the 55-65, 40-50 and 25-35 mm size groups, respectively.
No mortalities were observed following 5- and 10-second treatments for the various size groups. It was only after the 15-second treatment period that we began seeing this. Given the absence of mortalities during the 5- and 10-second treatments, they did not factor into any oyster analyses.
The only significant mortality levels were observed with oysters from the 25-35 mm group treated for 20 seconds. Only 55-65 mm oysters cultivated in Dark Sea trays and sampled in September were the exception. Given the particular nature of this group's results, the data for this group was not retained for this analysis; it was processed separately.
Although the mortality levels for the 40-50 mm and 60-65 mm groups were not significant, the numbers reveal certain correlations between oyster size and treatment intensity. For example, the mortality rate was greater for 20-second treatments and smaller oysters.
Mortality rates of approximately 85-95, 94-98 and 96-99%, respectively, were observed for the 25-35 mm, 40-50 mm and 55-65 mm groups for 20-second treatments, while mortality rates of approximately 97-99, 97-100 and 100% were observed for 15-second treatments. The mortality rate was significant for 25-35 mm oysters undergoing 20-second treatments, while some trials involving 40-50 mm oysters undergoing 20-second treatments revealed greater differences, however marginal. Although the mortality rates for 15-second treatments were less precise, they supported the trends observed for 20-second treatments. A 20-second bath in water at 60°C would therefore be the maximum for oysters of 40 mm or more.
Mortality rates varied between 5 and 15% for 20-second treatments of oysters from the 25-35 mm size group. These rates are too high to warrant treatment of this intensity for this size group. Despite an average survival rate of nearly 100% for 15-second treatments, this seems to be the maximum treatment duration for 25-35 mm oysters, and would be recommended for this size group.
Analyses to evaluate the effects of a water bath at 60°C on oysters were performed using only the data related to 20-second treatments because the only significant mortality levels observed were for this treatment intensity. The results reveal no differences between the months, with the exception of the smallest size group (25-35 mm). The lowest survival rates for this size group were observed in June, but gradually increased throughout the season. They increased from 86.6 ± 4.0% in June to 86.9 ± 5.0% in August, and 91.6 ± 4.4% in October. These variations are not significant, but an increase in survival rates over the course of the season for this size group is nonetheless understandable. The increase in oyster size during the summer season caused the shells to thicken, allowing for an increase in the oyster tolerance threshold.
In September, a significant accumulation of mud was observed in Dark Sea trays growing 55-65 mm oysters . Mortalities were also clearly observed in oysters of this size group in control cages at the time of sampling.
Researchers were conscious of the situation after making these observations and took samples of oysters from various size groups in the various culture structures tested (Dark Sea trays and submersed bags). They then analyzed these samples to determine meat yields and establish whether or not there were physiological differences between the oysters. The results reveal significantly lower meat yields for 55-65 mm oysters in Dark Sea trays compared to other groups of oysters. Meat yields for this group are approximately 5%, with yields as low as 2%, while yields for other groups are an average of approximately 10%.
The characteristics of the mussel infestations varied according to culture structure (site) and time of year. Mussel size was greater in June than in August and September; however, mussel density was much lower. Mussel density in June was much higher in culture structures, varying from 3 to 111 mussels, while on the exterior of culture structures, it varied from 0 to 26 mussels. Mussel density on culture structures in August was as high as 4,480 mussels, compared to 1,856 inside these structures. Intermediate densities, varying from 600 to 2,000 mussels on the exterior of culture structures and from 120 to 800 inside these structres were observed in September.
The effectiveness of the treatment seemed to vary according to period and position (on the exterior or interior of culture structures). Ten-second treatments were sufficient to remove mussel spat in August, while 15-second baths were required to irradiate mussels in September. Relatively high mussel survival rates were observed in trials conducted in June. It appears much easier to remove mussel spat off the exterior of culture structures; the spat was much more resistant to short treatments (5-10 seconds) inside cultivations structures).
Trials conducted in August and October show significant reductions in mussel density on culture structures after only 5 seconds immersion in water at 60°C. Mussels were generally small, measuring less than 10 mm. Survivalrates in Jne were still approximately 10% for 15-second treatments. Mussels were an average of approximately 18-19 mm in length. Specimens harvested after being immersed in hot water were generally over 25 mm in size.
The mussel mortality rate was calculated by comparing mussel density after treatment to mussel density on control structures. The term "drop in density," rather than survival, describes the impact of treating mussels on the exterior of culture structures particularly well. A drop in density may result from the cumulative effect of dead mussels and the reduction in number of remaining mussels because of the way they attach themselves to their substrate using byssal threads. The denser the colonization and the more the mussels grow, the more they attach to one another, which tends to weaken their attachment to the collection substrate. The weight of the dead mussels and shells likely caused some live mussels to fall from the culture structures. Mussels’ attachment to their substrate is very weak in mid-summer because the water temperature is too high.
It appears to be very easy to irradiate mussels on the exterior of culture structures. However, biofouling control measures are not as effective on mussels inside culture structures. In June, 10-second treatments removed mussels from the exterior of culture structures, while 15-second treatments were required to remove mussels from within these structures. Higher survival rates were also observed in August and September for mussels inside culture structures. Although relatively low in terms of percentage, remaining densities were nevertheless considerable when the initial densities are taken into account. Densities of approximately 200 to 500 mussels were observed in Dark Sea trays after a 10-second treatment, which is considerable compared to the densities of oysters grown in culture structures (150-250).
Measures taken to control barnacles on culture structures as part of trials conducted in June and August in the Shippagan area yielded relatively inconclusive results due to the size of the barnacles on the oysters studied during the trials in June and the limited amounts collected in August. Survival rates as high as 75% were observed in June for 20-second treatments. These rates were still high in August: more than 30% for 10-second treatments of barnacles on the exterior of culture structures and as high as 67% for barnacles on oysters in bags. However, the barnacles that survived were relatively large and must have come from the previous year's collection or before. Nevertheless, these results show that it would be possible to irradiate, or at least significantly reduce, barnacle densities on oysters raised in suspension using 15-second treatments at 60°C. A 10-second treatment would significantly lower the barnacle population, provided the barnacles are small.
Survival rates of less than 20% were noted for barnacles on oysters and inside cages undergoing 10-second treatments compared to 4% for barnacles on the exterior of the cages. However, the presence of two size groups of barnacles in the cages must be noted.
Washing oysters in hot water prior to treatment did not seem to affect oyster survival. We observed no difference between the survival rates of the various size groups of oysters washed prior to treatment and those of the size groups of oysters not washed for the two periods tested, with the exception of the 55-65 mm group cultivated in Dark Sea trays in October. The same is true for the barnacles. However, higher survival rates were observed for barnacles washed prior to 15-second treatments. Nevertheless, these differences are not significant. Given the size of the barnacles observed in August during trials performed in the Neguac area, it would have been interesting to evaluate this approach with culture cages.
Infestations of mussels on culture structures that had already been washed were irradiated using a 5-second treatment. Ten seconds were required to irradiate mussels on unwashed structures. The same amount of time was required to irradiate mussels from within culture structures after they had been washed and prior to scalding, although survival rates were relatively low after they had been treated for 5 seconds.
Pre-treatment washing in a hot water bath may therefore significantly improve the effectiveness of mussel infestation control measures involving scalding. Mussel densities on culture structures that had been washed lowered from 39-68 to 0.6-1.1% in August and from 54-67 to 2.4-20.6% in October. These differences may relate to the condition of the mussels during these periods. Mussels are much more fragile in summer because water temperatures are higher. They generally start to become stronger at the beginning of September, when water temperatures begin to drop. They are extremely resistant and grip their collection substrate strongly in October, when water temperatures are generally below 15°C.
Several oyster culture techniques have been developed since the first floating bag trials at the end of the 90s. Although most farmers currently use floating bags, some prefer surface cages and others, such as Vienneau Aquaculture Inc., use submersed bags and lines of stacked cages. One farmer recently developed cages (condos) that enabled him to handle 6,000 oysters glued onto ropes. Scalding systems available on the market could not meet the specific requirements of cultivation activities, such as those performed by Vienneau Aquaculture Inc. and other New Brunswick farmers. Machines used to scald bags, such as those used in France, have certain operational limitations because of buoys positioned beside or on top of bags. The Aquaculture Acadienne Ltée video that shows the scalding process reveals the system's shortcomings with respect to submersed bags. The flotation system built into the bags required an additional technician to ensure the bags remained submersed. Moreover, placing the culture structures in the treatment ank did not optimize performance for lines of stacked cages because they must be disassembled to allow for the individual treatment of the cages.
Vienneau Aquaculture Inc. wanted to develop a machine versatile enough to treat the various culture structures used in New Brunswick. The system should enable farmers to treat surface bags, submersed bags, lines of stacked cages and surface cages. With a few modifications to the condo structure, it will also be possible to treat oysters glued onto ropes using the scalding process. Farmers would therefore simply remove the conveyor for cages and condos.
The machine is comprised of a tank equipped with a heat exchanger, a heating unit and a conveyor. The tank is 1.2 x 1.2 x 3.0 metres, and two heat exchangers are installed at the bottom and on either side of the tank to allow for the complete submersion of Dark Sea tray lines. It is large enough to treat side bags and to submerse surface cages and condos. The heating unit comprises a 140,000 btu furnace that runs on oil and is equipped with a thermostat to control the treatment tank's water temperature. The conveyor is divided into four components: a conveyor to move the structures into the tank, an infeed conveyor and a chute at the other end. A deflector is installed over the top of the treatment tank to ensure that the bags are completely submersed.
The temperature of this new unit rose much faster than Aquaculture Acadienne Ltée's scalding system. While the Aquaculture Acadienne Ltée system required two hours to become operational, the new Vienneau Aquaculture Inc. system took only a half hour for the temperature to rise. Installation of the conveyor system was also faster and less cumbersome.
DarkSea trays. Trays are treated directly on the longline systems. With the cage treatment approach, lines of stacked cages are placed on the work platform and disassembled. Cages are submersed in the hot water bath individually. The lines are then reassembled and put back in the water. A team of six employees was able to treat approximately 1,400 cages in a 10-hour work period using this approach.
The system developed by Vienneau Aquaculture Inc. treated lines of stacked cages without disassembling them, which, in theory, should increase treatment speed and reduce the number of employees required. A hydraulic crane lifts lines of stacked cages out of the water and plunges them directly into the hot water bath. They are returned to the water immediately after treatment. Three technicians were able to treat nearly 250 lines of stacked cages (2,500 cages) in an eight-hour period. With a bit of practice and some minor modifications to the scalding system and the longline, it would be possible for only two employees to treat 2,500 lines or more.
This new approach eventually allowed for a reduction in staff of approximately 80% for one treatment period. Daily performance rose from 233 to 1,250 cages per technician.
Surface/submersed bags. The procedure used to treat bags differs slightly from that used to treat lines of stacked cages. The machine is installed on a work platform and bags are brought to the platform for treatment. The platform is set up near the culture sites, which reduces the need for employees to travel back and forth. The scalding machine is installed on a slant on the platform, with bags fed into one end of the platform and exiting the other. One team is responsible for picking up the bags and feeding them into the machine, while another works on the other end with a second vessel and is responsible for retrieving the bags and attaching them to the longlines. The Aquaculture Acadiennes Ltée machine was designed for the bags to be transferred from the work platform to the conveyor. Bags had to be fed length-wise and tended to slide under those ahead of them. The task became more cumbersome towards the end of the day, when staff had handled over 1,000 bags weighing between 20 and 35-40 pounds at shoulder height. A technician was also required to monitor the bags in the tank and ensure that they remained submersed.
Bag treatment could not be tested due to delays in the delivery of the machine. However, it was possible to determine the functioning and capacity of the machine based on trials conducted during this experiment and previous studies performed by the company and by Aquaculture Acadienne Ltée. The new machine will operate on the same principle, that is, one team will feed the machine, and a second team will return bags to the lines. The machine will be installed on a work platform with the two ends facing either side of the platform. However, like the first machine, the new machine will enable employees to load bags directly from the first vessel, rather than transfer them to the work platform. Bags will be loaded from the side at waist height, rather than from the end at shoulder height, which will reduce the employees' workload. Loading from the side will also increase the treatment rate because the bags are only 45 cm wide, compared to 92.5 cm long. It is no longer necessary for one technician to load the machine and another to monitor the bags to ensure that they remain submersed and do not slide under other bags.
A team of six technicians was able to treat 1,000 bags per day: two technicians to feed the machine, one to load it, one to monitorthe ags and two others to return the bags to the longlines. Daily yields could increase by 50% (to 1,500 bags), while allowing for a reduction in employees. Staffing needs per treatment period could eventually decrease by 56%. The rate at which the temperature increases (30 minutes, compared to 2 hours) could increase yields, reduce the amount of time required to treat one longline of bags (45 metres wide, compared to 92.5 metres for bags loaded length-wise) and reduce daily cumulative fatigue. It takes 11 minutes to treat one longline of bags loaded width-wise, compared to 23 minutes to treat bags loaded length-wise.
Although the scalding technique (hot water bath) has been used on a commercial scale in France for several years, it has not yet been fully mastered. In the early 2000s, the CREAA (http://www.creaa.com) did a study to validate all elements of the scalding technique. The Collective Research on Aquaculture Biofouling (CRAB) research program (http://www.crabproject.com), which brings together at least 10 research groups from various European countries, was created at roughly the same time to study all facets of marine biofouling colonization on culture structures and find innovative solutions to these production constraints.
For Vienneau Aquaculture Inc., the first objective in controlling marine biofouling is to address mussel infestations because of the consequences they can have on oyster growth and survival. Barnacle colonization and over-collection of oyster spat were also observed during operations involving suspended oyster culture in the Saint Simon area and throughout New Brunswick. However, with respect to the company's activities, these problems are not as common as mussel infestations, particularly on culture structures in deep waters. These culture structures can also quickly become infested with hydrozoans. Colonization begins early and continues throughout the growing season. A large sea grape tunicates (Molgula manhattensis) colony was also observed in the fall of 2008. At first glance, these last two species do not seem to pose any problems with respect to oyster growth performance, and are nevertheless easily irradiated through administration of a hot water bath treatment for a very short time. However, their recurrence, intensity and impact on cultured oysters must be evaluated and a decision must eventually be made on the frequency of treatment to determine whether or not these species should be targeted. Barnacle infestation, over-collection and mussel infestation in particular are more problematic. According to Mallet (2008), marine biofouling is not a major constraint to culture activities that involve the use of floating bags. The intensity of the infestations reported appeared less significant than those generally observed on structures in use on Vienneau Aquaculture Inc. culture sites, both using Dark Sea trays in the water column and bags. However, it should be noted that Vienneau Aquaculture Inc. uses submersed bags (buoys on top), rather than surface bags (buoys on the side), which fosters biofouler inestations. Therefore, differences observed between the culture sites could result from differences in oceanographic conditions and culture structure characteristics. The biofouling control approach will therefore be subject to these two elements.
Efforts have long been made to address production constraints associated with controlling marine biofouling on suspended culture structures. Arakawa's report (1980) refers to biofouling control measures tested in Japan that date back to the 70s. These methods were divided into two categories: preventive methods to avoid the problem and corrective methods to resolve a problematic situation. The first approach, as defined by Arakawa, does not apply to Vienneau Aquaculture Inc. sites because they have a history of significant marine biofouling. However, it is best not to wait until a problematic situation arises to react, as the company's managers realized in 2007, when the company faced a significant number of oyster mortalities and an increase in operation costs caused by major infestations of mussels (Lanteigne, 2008). Drawing on the knowledge it acquired, Vienneau Aquaculture seeks to implement a more prevention-oriented operation i.e. the company would like to develop a strategy to enable it to resolve the biofouling issue in a proactive manner. Assuming that these biofouling problems and mussel infestations, specifically, are recurring, it would like to treat its culture structures at strategic times to resolve the problem before having to take serious corrective measures. This type of approach would widen the gap between the intensity of the treatment required to irradiate the biofoulers, and that which minimizes the consequences on oysters, not only in terms of mortality, but also with respect to growth performance.
In the studies they performed on controlling marine biofouling in power plant cooling water, Rajagopal et al. (1995) noted that oysters had a greater resistance to high water temperatures than other fouling species. Oyster mortality reached 100% after approximately 40 minutes in water at 42°C, while it took only a few minutes to irradiate the mussels. As part of their studies on the same research subject, Nel et al. (1996) observed that exposure to water at 70°C for 40 seconds did not raise the meat temerature of the Cupped Oyster (Crassostrea gigas) to over 30°C, while, according to Arakawa (1980), cupped oysters die when their meat temperature reaches between 44 and 48°C.
In France, 3 to 5-second treatments in a water bath at 80 to 85°C are recommended for large oysters, while 2-second treatments are recommended for small oysters (CREAA, 2004). Arakawa (1980) and Park et al. (1988) recommend 15-second treatments at 60°C to control marine biofouling on oyster culture structures in Japan and Korea. Although they are cupped oysters (Crassostrea gigas) and may have thicker shells than American oysters grown in suspension, scalding principles should be the same. The results of this study indicate that 20 seconds in a water bath at 60°C appears to be the maximum treatment duration for all size groups. With the exception of small oysters (under 40 mm), a 15-second treatment in a water bath at 60°C would be the most appropriate duration to minimize the impact on the oysters, which is in keeping with the approaches recommended by Arakawa (1980) and Park et al. (1988).
The results of this study are also consistent with the assumption put forward by Arakawa (1980), to the effect that oysters die once their meat reaches a critical temperature. Based on this premise, small oysters should be affected by a hot water bath more quickly because their shells are thin. These types of trends were observed in this study, taking into account the apparent correlation between the survival rates of the various size groups and treatment intensity. Therefore, given the mortality rates observed for 25-35 mm oysters after 15-second treatments, the maximum treatment duration would be 12 seconds for this size group.
It was impossible to evaluate the impact of scalding treatments on oyster over-collection because of the lack of oysters collected from culture structures and other oysters. However, signs of over-collection were observed on bags and oysters in Saint Simon Bay South in 2007, which suggests that these types of infestations can and are expected to occur in the area, given the increase in the volume of oysters currently being grown in Saint Simon Bay and the anticipated volume in coming years. Lanteigne (2008) used a 15-second treatment at 60°C to irradiate 5-15 mm oyster spat at the end of August. Although no tests could be performed for oyster over-collection, trends related to the fluctuation of the various size groups' mortality rates based on treatment intensity suggest that a 15-second treatment at 60°C would irradiate the over-collection as long as it is administered after spatfall. Spat measures less than one mm at this time and is very fragile. According to Mallet (pers. comm.), oyster spat cannot survive for more than two hours out of water, while mussels can survive for more than one day during the same period. A 10-second treatment might even be sufficient.
Oyster farmers met as part of a study led by the French company indicated that they only treat their oysters once per season: at the end of winter and the beginning of spring, when an over-collection of oysters is confirmed. However, they have less reason to worry about mussel infestations on their culture sites than we do on our production sites, where mussels can accumulate until mid-September, which gives them more leeway. Spat is relatively large at this time of year, and it emerges from bags for 3-4 seconds in a water bath at 80-85°C. For some, the preventive approach recommended by Vienneau Aquaculture Inc. would be very beneficial and would enable them to reduce treatment intensity, given that it is not truly necessary to wait so long to confirm the presence of an over-collection. They could then operate at a lower temperature, which could reduce stress on the oysters. They said that they would be interested in testing this approach. For others, the intensty of the treatment they currently use minimizes the loss of oysters; however, oyster meat may reach a temperature that could lead to stressful conditions and, consequently, have a negative impact on growth performance. French farmers feel that this justifies testing out the new approach.
Although the number of mortalities was low, as previously reported for 55-65 mm oysters grown in Dark Sea trays in September with shorter treatment durations (5 and 10 seconds), and for the control group, this suggests that oysters from this size group are more fragile. The results associated with the levels of siltation in trays and poor meat yields suggest that oysters' physiological condition could impact their tolerance threshold. Mussels, in addition to acting as sediment traps, produce more biodeposition than oysters (Tenore and Dunstan, 1973) and can thereby increase the effects of the siltation process on oyster culture structures.
The results of this study have led to the conclusion that infestations of biofoulers and of mussels, specifically, can have cumulative consequences on the survival of oysters treated in hot water baths. They can worsen the physiological condition of the oysters, which consequently become more fragile when it comes time to administer treatment to remove the biofoulers. These conclusions are consistent with studies performed by Willemsen (2005) and Arakawa (1980), who pointed out that siltation on culture structures caused by marine biofouling tends to impact the physiological condition of oysters and, in extreme cases, leads to oyster mortality.
Preventive treatment would therefore minimize the risk of siltation, which would not only improve the effectiveness of the treatment process, but could also increase growth rates. Italian oyster farmers who use Ostriga trays for suspension culture, a technique similar to Dark Sea trays, clean cultivation structures and wash oysters up to three times in an 18-month production cycle. They do not seem to worry about the impact of mussel infestations on oysters. They use a manual technique to control marine biofouling and do not seem to consider use of the scalding technique as a control measure. It must be stressed that, as mentioned previously, their production cycle is only 18 months and they therefore only have to deal with one or two infestations of mussels. However, they do wash the oysters in the culture structures to reduce water volume and, if necessary, transfer them to clean structures.
According to Rajagopal et al. (1994), juvenile mussels can easily be killed in hot water, and there is a correlation between mussel resistance and mussel size. This was confirmed by Graham et al. (1975), McNair (Fisheries, Aquaculture and Rural Development, P.E.I., unpublished data) and studies by Vienneau Aquaculture Inc. in 2007 (Lanteigne, 2008). According to Arakawa (1980), mussels exposed to air die when their meat reaches a temperature of 38-40°C. Mussel spat have much thinner shells than adults. Their meat could therefore reach this temperature more quickly and would withstand a preventive approach, in which treatment is administered when mussels are small. This analysis was confirmed by the differences in survival rates observed in this study and the variations in mussel size.
The strategy involving irradiation of mussels on culture structures capitalizes on mussels' poor resistance to heat (compared to that of oysters) and their difference in size, which can further emphasize this difference. From a preventive standpoint, mussels generally measure no more than 15 mm in mid-summer. However, the intensity of the mussel spat collection and the accumulation of mud associated with these infestations can affect treatment effectiveness if there is a delay in the treatment of culture structures. Fewer mussels were observed in June than in mid-August and late September. These mussels were also larger, which suggests they had been accumulating for a year.
The first trials performed by the company in 2007 (Lanteigne, 2007) revealed that an 18-second treatment at 60°C removed all mussels and oyster spat younger than one year (and under 20 mm) with no apparent mortalities of oysters below market size. These results are consistent with those of studies performed by the CREAA and the CRAB in Europe, and by Arakawa (1980) and Park et al. (1988) in Japan and Korea, as well as those reported in P.E.I. as part of McNair's preliminary trials (Fisheries, Aquaculture and Rural Development, P.E.I., unpublished data). However, this treatment appears too aggressive for the smallest size of oysters (25-35 mm). The findings of Lanteigne (2008) showed that treatments at 50 and 55°C controlled mussel infestations on the smallest sizes of oysters grown in suspended culture structures. However, from an operational standpoint, it is more difficult to vary water temperature than treatment duration. The results of this study indicate that a shorter treatment (10 seconds) at 60°C would be sufficient to treat bags of 25-35 mm oysters infested with 0-8 mm mussel spat. This type of approach could involve administering treatment at the beginning of July and immediately after oyster spat has settled, depending on the amount that has accumulated. However, Mallet et al. (2009) expressed reservations about handling oysters at this time of year because this could cause mortality problems. Nevertheless, the risk is greater for the largest sizes of oysters because of the spawning period, and consequently, should not impact juvenile oysters.
The death of a significant proportion of mussels could lead to further mussel loss because they will attach themselves to any substrate. Mussels with high densities and increased growth will gradually attach themselves to one another, rather than to the cage, weakening their attachment to the culture structure. Mortality rates of 40-50% for commercial-size mussels on socks can lead to significant losses, and farmers must double-sock to prevent mussels from falling off. This is especially clear with lines of stacked cages, and may result from a significant accumulation of mussels on them, compared to that observed on bags. This situation may explain variations in mussel survival rates on the exterior and inside culture structures. Depending on the condition and size of the mussels, byssal threads may hold the shells in place, which can increase the effects of siltation on oyster culture structures. This is particularly relevant to mussel shells in Dark Sea trays.
The intensity of mussel byssal thread production varies according to oceanographic conditions and sometimes collection density, water temperature and movements of the water mass caused by waves or currents. Mussels in more vibrant sectors will produce more byssal threads than those in less vibrant sectors with higher water temperatures, which is true in the case of floating bags compared to Dark Sea trays. They have greater densities, which, in theory, would weaken their attachment to the collection substrates; however, the impact of dynamic factors on floating bags could reduce the number of mussels that fall off, despite high mortality levels.
Mud accumulation on culture structures can also act as an insulator and reduce efficiency of the scalding process, which may explain the differences between mussel survival rates inside them and on their exterior. According to Arakawa (1980), in the case of desiccation, mussels die when the temperature of their meat reaches between 38 and 40°C. The greater the mussel accumulation, the longer they remain on and inside structures, and the more difficult it will be to remove them due to the production of feces and pseudo-feces, as well as its accumulation, particularly with respect to oysters in culture structures. Arakawa (1980) also states that clusters of mussels are more resistant to desiccation. The greater the infestation, the easier it is to irradiate mussels on the exterior of structures, but the harder it is to remove them from inside the structures.
Treatment must be sufficiently intense to irradiate mussels, as even survival rates as low as 10% can indicate that a significant residual population remains. Although survival rates were as low as 10-12% following a 10-second treatment of Dark Sea trays in August, approximately 100 to 200 mussels were still alive in the culture structures. This density of mussels can have adverse effects on growth performance and oyster quality, and can increase the field team's workload when they sort and harvest.
Oysters, like barnacles, are better adapted to periods of exposure to air than mussels, and should theoretically be more resistant. Studies performed by Graham et al. (1975) on controlling marine biofouling in power plant cooling water indicate, howevr, that barnacles are more quickly eliminated in hot water baths than mussels. They pointed out that hydrozoans were the first species affected by hot water treatments, followed by barnacles and mussels. These results are consistent with those of studies performed by Arakawa (1980) and Park et al. (1988), who estimated that a 10-second treatment at 60°C was sufficient to control barnacle infestations on oyster culture structures. The results were similar for tests performed in Neguac, in which barnacles were small. However, it is possible for difference species of barnacles to colonize on culture structures and to show different levels of resistance. According to Foster (1969), barnacles that colonize in intertidal environments are more resistant to heat. According to Brinkhurst et al. (1973), different species of barnacles will colonize different habitats, making them relatively resistant to desiccation. For example, the Balanus balanoides colonizes rocks in the intertidal zone and is more resistant to desiccation. The Balanus balanus does not colonize beyond the lower intertidal zone, while the Balanus crenatus cannot withstand intertidal zone conditions. Different species can therefore be found between Dark Sea trays in deeper waters and floating bags. However, we did not identify any species of barnacles on culture structures.
Observations of barnacle survival rates reveal that they are consistent with those of mussels in terms of resistance in a hot water bath. The largest barnacles had the highest resistance, while spat was very fragile to submersion in a hot water bath at 60°C. It would therefore be more effective to treat infestations of barnacles soon after they are recruited. Depending on the number of barnacles accumulated and the timeframe in which they are treated, it may be difficult to irradiate them in culture structures if they are not treated right away. As is the case with mussels, the principle of the technique is based on the increase in meat temperature of the targeted organisms. Given the size of juvenile barnacles, it would therefore be easier to increase the temperature inside their shells. In the case of serious infestations, barnacles tend to attach themselves to one another as they grow, which enables them to develop a certain form of insulation.
Low barnacle mortality rates were observed following 10- and 15-second treatments in June. The barnacles were relatively large compared to those observed in mid-August in the Neguac area. We must evaluate the possibility of administering treatment immediately after barnacle spat collection, which, in theory, occurs after oyster spat collection. According to Mallet (2009), treatment administered during this period would also help to minimize the impact on oysters. Barnacles are generally collected at the beginning of August, at approximately the same time as oyster spat is collected. Treatment at this time of year would not only irradiate barnacle spat, but also the over-collection of oysters.
Washing culture structures prior to hot water bath treatment increased the effectiveness of mussel control treatment significantly. In August, when water temperatures are relatively high, this approach reduced mussel density by 50% or more prior to hot water bath treatment, which led to even greater mussel mortality rates after the treatment was administered. It is therefore possible to effectively treat culture structures for 5 seconds. However, given that there is time before oyster mortality begins, it is recommended that culture structures be submersed for 10 seconds to optimize the irradiation of mussels and other biofoulers. As previously mentioned, even relatively few mussels can impact oyster quality and, potentially, growth rates. However, it may be to the company's advantage to monitor the conditions of the various species' colonies in order to review the terms and conditions of their treatment strategies and consider washing culture structures on a monthly basis to minimize siltation and biofouler infestations on these structures. This type of approach may improve oyster growth and reduce the amount of effort required during the scalding process.
A structure that mechanizes this process must be developed and integrated into a marine biofouling control strategy. These results are consistent with the findings of Mallet et al. (2009), who indicated that efforts should be made to develop new approaches to maintaining culture structures.
However, we must consider the impact of this type of approach on biofouler settlement rates and, specifically, the settlement rates of oysters and barnacles, which prefer a cleaner substrate. The fact that oysters and barnacles settled on floating bags in 2007, while none of these species settled on Dark Sea trays, may be due to a cleaner substrate.
Controlling marine biofouling in a hot water bath is a common practice in cupped oyster cultivation in Asia and Europe. Results obtained in this study have led to the development of a procedure for controlling marine biofouling. A new machine that is better adapted to the reality of oyster culture activities in New Brunswick has been developed, along with local expertise.
Preventive treatment. It is most effective to treat biofoulers at the beginning of their infestation, while they are small. This approach reduces the need for intensive treatment, which could minimize the impact on oysters.
Washing prior to treatment. Washing reduces mussel density prior to a hot water bath treatment and significantly increases treatment effectiveness. This approach also removes mud from culture structures, which can cause death or, at least, negatively impact growth. This is more obvious with Dark Sea trays that are submersed deeper in the water column and are more stable as a result. Their position in the column tends to foster siltation in the lower trays. It may be advantageous to wash culture structures regularly in an attempt to foster better growth rates; however, it is important to take into consideration the impact of this approach on the settlement rates of biofoulers, as well as those of oysters and barnacles, specifically.
Treatment frequency. It is possible to be strategic and reduce the frequency of treatments to two periods. The first treatment may be administered at the beginning of August, immediately after the oysters have settled. Barnacles settle at approximately the same time, although this should be confirmed. Mussels are also still small. The mussels, the over-collection of oysters and the barnacles can all be irradiated in one step. The second treatment may be administered at the end of September, after the mussels have settled. It should not be administered too late into the fall due to the mussels' increase in growth and resistance.
Treatment duration. Washing structures and administering a preventive treatment should reduce the intensity of the treatment required to irradiate the biofoulers. It is possible to administer a treatment for as little as 5 seconds, but this is not recommended. A low mussel survival rate can nevertheless impact oyster quality and growth. Given that there is time before the oysters begin to die (20 seconds), it is recommended that 10-second treatments be administered to structures at the beginning of August, following the recruitment of biofoulers, and 15-second treatments in the fall to ensure that the mussels are completely irradiated.
Although the study validated the technique for controlling biofouling on culture structures for oysters grown in suspension, and allowed for the development of an approach to implement this technique, some questions remain. We must learn more about the specific effects of stressful conditions at maximum treatment durations on oyster physiology and growth, specifically. We must evaluate the impact of hydrozoan infestations on oyster growth performance and, if necessary, review treatment frequency. We must verify the settlement periods for the various fouling species, specifically, barnacles, oysters and mussels, to adjust treatment periods. Could washing culture structures on a monthly basis increase growth rates and reduce infestations of biofoulers? If necessary, how would this impact oyster spat and barnacle settlement?
With the exception of oysters, which prefer a clean substrate, the majority of biofoulers colonize surfaces in succession, with the development of biofilm as the first step. Is it possible that by irradiating biofoulers, we are creating conditions conducive to the development of this biofilm, which consequently accelerates the re-colonization process? Mussel and hydrozoan infestations seem to persist on lines of stacked cages. A sea grape infestation was observed on Dark Sea trays after structures were treated in August. Is it possible that scalding fosters biofouler re-colonization? If so, would it not be advantageous to integrate a step that involves washing the structures after the scalding treatment?
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