Settlement of larvae of the giant scallop, Placopecten magellanicus (Gmelin), on various artificial and natural substrata under hatchery-type conditions

Aquaculture Aquaculture 141 (1996) 201-221

Settlement of larvae of the giant scallop, Placopecten magellanicus ( Gmelin) , on various artificial and natural substrata under hatchery-type conditions ’ Christopher

M. Pearce *, Edwin Bourget

GIROQ, DGporremenr de Biologic, Uniuersire Luvul, Suintr-Fey, Quebec GIK 7P4, Cunudu

Accepted 7 October 1995

Abstract Multi-choice experiments on larval settlement of the giant scallop, Pfacopecten mugellanicus (Gmelin), in response to various artificial and natural substrata were conducted in a 1000-I plastic tank to determine what conditions optimize larval and spat collection under a normal, hatchery-type environment. The first experiment tested a variety of substrata (many of which have been previously used in bivalve spat collection studies) including the following: (I) nylon monofilament of diameters 0.50, 0.75, 1.10, 1SO, and 2.00 mm (N.B. the OSO-mm diameter monofilament was tested with and without a marine microbial film at the beginning of the experiment (designated as ‘filmed’ and ‘non-filmed’, respectively) whereas all other monofilament treatments and the artificial substrata to follow lacked a microbial film at the start of the experiment), (2) polyethylene onion bag material, (3) polyethylene Astroturf, (4) smooth and roughened clear acrylic plastic, (5) polyester aquarium filter-wool, and (6) filmed and non-filmed adult giant scallop shells. Standardized for surface area, filter-wool collected significantly more larvae and spat than any other non-filmed substratum (Student-Newman-Keuls (SNK) test, P I 0.05). There was no significant difference among all other non-filmed substrata in numbers of larvae or spat collected (SNK test, P 2 0.05). Filmed 0.50-mm diameter monofilament and filmed adult shell collected significantly more spat than their respective non-filmed counterparts (SNK test, P I 0.05). A second experiment examined settlement on various types of polyethylene meshes (Vexar) including the following mesh sizes (mm): 3.0, 3.0 X 11.0, 3.8, 7.0, 11.0, and 19.0. All substrata

* Corresponding author at: Department of Biology, Dalhousie University, Halifax, Nova Scotia B3H 45 I, Canada. Tel.: (902) 494 6697; Fax.: (902) 494 3736; e-mail: [email protected]. ’ Contribution lo the programs of OPEN (Ocean Production Enhancement Network, one of the 15 Networks of Centres of Excellence supported by the Government of Canada) and GIROQ (Groupe Intenmiversitaire de Recherches OcCanographiques du Quebec). 00448486/96/$15.00 0 I996 Elsevier Science B.V. All rights reserved SSDl 0044-8486(95)01210-9

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were black except the 3.0.mm mesh which was green. Standardized for surface area, the 3.0-mm mesh collected significantly more larvae and spat than any of the other mesh sizes (SNK test, P 5 0.05). There was only one other significant pairwise comparison of numbers of spat collected; 3.8-mm mesh collected significantly more spat per unit surface area than 1 I.O-mm mesh (SNK test, P I 0.05). Polyester filter-wool, which was by far the best substratum for spat collection, is inexpensive and re-usable and could make an excellent settling material for use in hatcheries. Keyrvor&: Artificial substrata; Hatchery conditions; Larva; Spat collection

Plucoppecten mugel1unicu.s;

Scallop; Settlement;

1. Introduction The giant scallop, Placopecten mageflanicus (Gmelin), has long formed the basis for a lucrative natural fishery in the Atlantic provinces of Canada and the northeastern United States (Boume, 1964; Serchuk et al., 1979; Sinclair et al., 19851, but only recently has it received some interest as a potential aquaculture species (Couturier, 1990; Dadswell and Parsons, 1991). Despite the increasing interest in the development of an aquaculture industry for the giant scallop in this region, very little scientific information is available on the reproduction and larval development of this species. Although much work has been carried out on various biological, chemical, hydrodynamical, and physical factors influencing larval settlement and metamorphosis in other benthic marine invertebrates (see reviews by Thorson, 1966; Meadows and Campbell, 1972; Crisp, 1974; Crisp, 1976; Scheltema, 1974; Burke, 1983; Butman, 1987; Bourget, 1988; Morse, 1990; Pawlik, 1990; Pawlik, 1992; Rodtiguez et al., 1993; Pearce, 1996), very little has been done with giant scallop larvae. This is despite, or perhaps because of, the fact that larval settlement is a particularly critical event in the life cycle of this species in which high mortality often occurs (Tremblay, 1988). No published studies have compared larval settlement of giant scallops on various potential collecting substrata under controlled hatchery-type conditions. A variety of artificial and natural materials have been used to collect settling pectinid larvae in the laboratory and the field including: shell material, meshes, monofilament, Astroturf, polyethylene films, plastic plates, and ropes (Appendix A). Most studies on pectinid settlement/recruitment have been conducted in the field and have made use of some sort of mesh bag filled with fishing line, gill netting, or monofilament as collectors, technology which was first developed in Japan (Ventilla, 1982). Although field collection of pectinid juveniles with ‘spat bags’ is relatively cheap for the aquaculture industry, it is often unpredictable, both spatially and temporally, and is usually only successful at certain times of the year (Brand et al., 1980; Hortle and Cropp, 1987; Sause et al., 1987; Fraser, 1991; Ruiz-Verdugo and Cgceres-Martinez, 1991; Tripp-Quezada, 1991; Young et al., 1992; Robins-Troeger and Dredge, 1993). In some localities natural spatfall may be minimal, thus precluding field collection via spat bags (Cropp, 1993a). Hatcheries, while more expensive and labour intensive, can provide a reasonably steady supply of spat or seed for a greater portion of the year. Several hatcheries producing giant scallop larvae have already gone into production in the Atlantic Canada region. In order to maximize giant scallop spat production in the hatchery, a greater understanding of broodstock development, fertilization success, larval

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rearing, settlement, and metamorphosis is necessary. This study examines larval settlement on a variety of potential spat collection materials under hatchery-type conditions in order to determine what substrata are most efficient at collecting giant scallop larvae and spat.

2. Materials

and methods

2.1. Larval culture Larvae for the experiments were kindly supplied by the Fisheries Resource Development Ltd. (FRDL, a subsidiary company of National Sea Products Ltd.) hatchery at Sandy Cove, Nova Scotia, Canada (44”28’06” N, 63”33’33” W>. Larvae were transported from the hatchery to the laboratory in an 11.4-l plastic carboy within 0.5 h and then placed in a 1000-l insulated polyethylene tank (Xactics Ltd.) that had been previously filled with 0.2-pm cartridge filtered seawater (FSW) at 15°C. Larvae were 28 days old and had a shell length of 208.6 + 6.8 pm (mean + s.d., n = 50) when added to the tank (N.B. shell length is defined as the longest straight line distance between the anterior and posterior margins of the shell running parallel to the hinge). Generally, giant scallop larvae do not become competent to metamorphose until a shell length of 230-260 pm is 260 000 attained (Culliney, 1974; Naidu et al., 1989; Couturier, 1990). Approximately larvae were introduced to the tank, producing a concentration of 0.26 larva ml-’ at the beginning of the experiment. Approximately every second day (i.e. every Monday, Wednesday, and Friday) the tank was drained to - 20 cm from the bottom, leaving the collecting substrata submerged (see following section). The water that was drained off was passed through a 44-pm Nitex screen to capture any larvae; the screen was partially submerged in FSW so that larvae were never exposed to the air for any period of time. These larvae were then re-introduced to the tank after having refilled it with fresh FSW at - 15°C. Larvae were then fed a mixture of phytoplankton consisting of 5000 cells ml-’ of Isochtysis galbana (Tahitian strain: clone designation TISO), 1000-2500 cells ml-’ of Thalassiosira pseudonana, and 1000-2500 cells ml-’ of Chaetoceros muelleri. Concentrations of T. pseudonana and C. muelleri varied depending on the quantity of culture available, but were usually 2500 cells ml- ’ . All cell counts were carried out on a model ZB Coulter counter. The tank was covered tightly for the duration of the experiment (light entered the tank only during times of water changes and feeding) and the water was gently aerated with a cylindrical plastic air stone (length: 2.0 cm, diameter: 0.7 cm>. Water temperature during the experiment ranged from 13.3 to 16.5”C and salinity ranged from 29.9 to 3 1.4%0. The experiments ran for 23 days from Sept. 30 to Oct. 23, 1992. 2.2. Experiment

1: various artificial and natural substrata

In this experiment a number of artificial and natural substrata were tested as potential spat collecting materials. Three replicates of each of the following substrata were placed separately in miniature collectors constructed out of 15 X 12-cm pouches of 3.8-mm mesh size black low-density polyethylene mesh (Vexar, made by DuPont Canada Inc.) and tested in the experiment:

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(1) Tangles of green nylon monofilament (Perlon, made by Mylon Slonek KG, Saal/Donau, Germany) of various diameters including: 0.50, 0.75, 1.10, 1.50, and 2.00 mm. The strength in lb/kg of these monofilaments were 25/ 11.4, 50/22.7, 100/45.5, 180/81.8, 300/136.4, respectively. The total lengths of monofilament used for each replicate collector were 15.00, 10.00, 6.82, 5.00, and 3.75 m, respectively. The different lengths for each diameter were chosen to give an equivalent surface area of 23 560 mm* for each replicate. These monofilaments all lacked a marine microbial film at the beginning of the experiment. From hereafter, treatments that had no film at the start of the experiment are referred to as ‘non-filmed’ substrata while those that were filmed in seawater prior to the experiment are referred to as ‘filmed’ substrata. In addition, a sixth treatment, where a 0.50-mm diameter monofilament had been filmed in running unfiltered seawater for 16 days prior to the experiment, was established. (2) Orange onion bag material (polyethylene mesh, unknown manufacturer) with a mesh size of 3 X 5 mm that was non-filmed. The total surface area of each replicate was 23 420 mm*. (3) Brown polyethylene Astroturf (PNS-1, made by Monsanto Canada Inc.) that was non-filmed. This Astroturf was made up of repeated units separated by 4-5 mm. Each unit consisted of a circle of eight finger-like projections, each projection being 2 mm wide and 20 mm high. The total surface area of each replicate was 47650 mm*. (4) Smooth and roughened plates (length X width X height: 106 X 106 X 3 mm) of clear acrylic plastic (Acrylite FF, made by Cyro Canada Inc.). The roughened panels were created by sanding both sides with #36 grit sandpaper using an automated belt-sander. Both treatments were non-filmed. The total surface area of each replicate was in the range of 23 610-23 890 mm*. (5) Aquarium filter-wool made of 100% polyester (‘Poly’ Filter Wool, made by Rolf C. Hagen Inc.) that was non-filmed (mean + s.d. filament diameter: 29.6 f 4.1 pm, n = 50). The total surface area of each replicate was in the range of 50 430-63 560 mm*. (6) Adult giant scallop valves that either had a natural marine microbial film developed over the lifetime of the organism (filmed) or were air-dried, scrubbed under hot freshwater, and rinsed with distilled water to remove the film (non-filmed). The total surface area of each replicate was in the range of 19 440-23 750 mm*. (7) Control. No substratum in collector. All surface areas were calculated by direct measurements except for scallop shells and filter-wool. Areas of shells were measured using the computer image-analysis system BioScan OPTIMAS 4.02. Images of shells were projected onto a video monitor (SONY Trinitron PVM-1342Q 13-inch colour video monitor) with the use of a black and white, high resolution charge-coupled device camera (Pulnix TM-7EX). Images were then frozen with an Imaging Technology Inc. frame grabber. OPTIMAS was then used to take area measurements of the insides of the shells. The total surface area was then calculated as twice this amount. For filter-wool, a regression equation was calculated correlating volume (as measured by water displacement) to weight. This produced the following equation with an r* of 0.991: volume = (weight + 0.020) / 1.404

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where volume is measured in ml and weight in g. Knowing the weight of a quantity of filter-wool, thus, allowed simple conversion to a volume and this allowed calculation of surface area, since the average filament diameter was known. The Vexar mesh pouches containing the afore-mentioned substrata were attached to a rack made from 18-mm (internal diameter) PVC tubing. The three replicates of each treatment were randomly assigned to three separate areas of the rack in a completely randomized block design. The collectors were hung vertically lengthwise from the rack with about 2-4 cm between each collector. The whole rack was hung horizontally a few centimetres off the bottom of the tank so that collectors were not actually touching the bottom. The substrata were put into the 1000-l tank 2 days after larvae had been added and were left for 23 days. At this time there were no planktonic larvae evident. At the end of the experiment the rack was gently pulled up to the surface of the tank and the collectors (i.e. Vexar mesh bag with substratum) were individually placed in separate specimen bags and preserved with 70% ethanol for later counting. Prior to counting, the collectors were soaked in a 1% bleach solution in FSW (i.e. - 0.05% sodium hypochlorite) for at least 10 min to dislodge any byssally-attached spat (Boume et al., 1989; Heasman et al., 1994), rinsed with a jet of 500 ml of FSW, and then rinsed with a jet of 5-7 1 of hot freshwater to remove all larvae and spat. The ethanol that the collectors were originally preserved in, the bleach that they were soaked in, and the seawater and freshwater rinsings were all passed through a 75pm Nitex mesh to collect the larvae and spat. These were then rinsed into a small crystallizing dish for counting. Collectors were generally not examined directly, but previous tests, where similar substrata had been examined after similar bleaching and rinsing procedures, showed that few individuals (i.e. < 0.1%) remained on the substrata. Counts of larvae and spat were made with the use of a dissecting microscope set up for polarized-light viewing (Gallager et al., 1989) since shells of larvae and spat of the giant scallop are birefringent and luminesce under polarized light. Quick scans were made under normal light after the polarized light procedure, but few individuals were ever found. Counts of larvae and spat were standardized among treatments to numbers per 100 cm2 of surface area. Counts of individuals include those on the settlement substratum itself as well as the mesh bag which contained the substratum. Individuals were classified as spat only if new dissoconch (i.e. post-metamorphic) shell growth was evident, otherwise they were counted as larvae. This may have lead to a conservative estimate of the number of spat present, since very recently metamorphosed individuals may not have had time to deposit a visually detectable layer of dissoconch. The criterion of new shell growth, however, is a definitive indicator that an individual has settled and survived through metamorphosis and is more reliable than other potential criteria such as the presence of developed gill bars or the absence of a velum, both of which can be difficult to determine under normal light in preserved samples and impossible to ascertain under polarized light. 2.3. Experiment

2: various arti’cial

mesh substrata

This experiment examined larval settlement (DuPont Canada, Inc.) of varying mesh sizes and 3.0-mm low-density polyethylene green (length 3.0 X 1 l.O-mm low-density polyethylene black

response to panels of Vexar mesh colours including: X width: 15.7 X 12.0 cm> (length X width: 15.5 X 13.8 cm)

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3.8-min low-density polyethylene black (length X width: 15.2 X 11.6 cm) 7.0-mm low-density polyethylene black (length X width: 15.1 X 1 1.9 cm) 11 .O-mm high-density polyethylene black (length X width: 15.0 X 12.3 cm) 19.0-mm low-density polyethylene black (length X width: 15.0 X 13.2 cm) Mesh sizes given are from side to side of the opening, not diagonally. DuPont product numbers for the different meshes are (in order as they appear above): L-9, L-3, L-36, L-30, L-37, and L-38. These meshes were tested in the same tank and at the same time as the previous experiment (i.e. Experiment l), so that all experimental conditions (including larval batch, experimental duration, water temperature, etc.) were as described previously. The meshes were hung vertically lengthwise from the PVC rack mentioned above; the meshes were a few centimetres off the bottom of the tank and not actually touching it. Five replicates of each mesh size were randomly placed in five separate regions of the rack in a completely randomized block design. The experiment was run for 23 days after which the rack was pulled up and the meshes placed in separate specimen bags and preserved with 70% ethanol for later counting. Prior to counting, the meshes were soaked in a 1% bleach solution, rinsed with a jet of 500 ml of FSW, and then rinsed with a jet of 3-5 1 of unfiltered seawater to remove all larvae and spat. The ethanol that the meshes were originally preserved in, the bleach that they were soaked in, and the seawater rinsings were all passed through a 75pm Nitex mesh to collect the larvae and spat. These were then rinsed into a small crystallizing dish for counting. The Vexar meshes were examined directly to see if any individuals were still left on the surface following these procedures, but no larvae or spat were ever found on any of the replicates. Counts of larvae and spat were made as in Experiment 1 and standardized for different surface areas of meshes. Surface areas were calculated by direct measurements of filaments making up the meshes, compensating for overlap of filaments. 2.4. Statistical methods Completely randomized block design analyses of variance (ANOVAS) were carried out on log-transformed numbers of larvae and spat per unit surface area in both experiments. Untransformed data from the first experiment suffered both from non-normality and heterogeneity of variances (as judged by probability plots and Cochran’s test, respectively), but the log-transformation solved both problems. Data from the second experiment were neither non-normal nor significantly heteroscedastic, but the log-transformation was used anyway since it increased normality and made the variances even more homogeneous. Post-hoc comparisons were made with Student-Newman-Keuls (SNK) multiple comparisons tests at CI= 0.05. Data analysis was performed with the computer packages SYSTAT 5.2.1 (Wilkinson, 1992) and SuperANOVA 1.l 1 (Abacus Concepts, 1989). Fig. 1. Number of scallop larvae (a) and spat (b) per unit surface area collected by various potential settlement substrata. Bars are the means of three replicates. Error bars are the standard errors of the means. Letters by the error bars indicate the result of a Student-Newman-Keuls post-hoc comparisons test; treatments with the same letters do not differ significantly (P > 0.05). Different bar shadings are as follows: white bars = monofilament, stippled bars = all other artificial substrata, wavy bars = shells, and vertical striped bar = control (i.e. mesh collector with no settlement substratum in it).

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(a) Filmed 0.50 mm Monofilament 0.50 mm Monofilament 0.75 mm Monofilament 1.10 mm Monofilament 1.50 mm Monofilament 2.00 mm Monofilament Onion Bag Astroturf

d

Filter-Wool Smooth Acrylic Plastic Rough Acrylic Plastic Filmed Shell Non-filmed

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of Spat/100

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3. Results 3.1. Experiment I The number of larvae and spat collected per unit surface area varied significantly with position on rack or block (larvae: F = 9.99, df = 2, 26, P I 0.001; spat: F = 4.55, df = 2, 26, P I 0.05) and also with substratum (larvae: F = 9.81, df = 13, 26, P 5 0.0001; spat: F = 9.87, df = 13, 26, PI 0.0001) (Fig. I(a), (b)). The five different

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# of Spat/100 cm2 Fig. 2. Number of scallop larvae (a) and spat (b) per unit surface area collected by various sizes of artificial polyethylene meshes (Vexar). Bars are the means of five replicates. Error bars are the standard errors of the means. Letters by the error bars indicate the result of a Student-Newman-Keuls post-hoc comparisons test; treatments with the same letters do not differ significantly (P > 0.05).

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diameters of non-filmed monofilament did not differ significantly from one another in numbers of larvae or spat collected per unit surface area and did not collect significantly more larvae or spat per unit surface area than the smooth and roughened acrylic plastic plates. Roughened acrylic plastic did not differ significantly from smooth acrylic plastic in numbers of larvae or spat collected per unit surface area. Astroturf, onion bag material, and non-filmed shell did not collect significantly more larvae or spat per unit surface area than any of the non-filmed monofilament treatments. The 0.05mm diameter monofilament that had been filmed in running seawater prior to the experiment had significantly more larvae and spat per unit surface area than non-filmed monofilament of the same diameter. Similar results were shown with spat for the filmed and non-filmed giant scallop shells. Although the filmed shell had more larvae per unit surface area than the non-filmed shell, the difference was not significantly different at P I 0.05. Interestingly, only three treatments collected significantly more larvae and spat per unit surface area than a control collector consisting of a mesh pouch with no settlement substratum in it. These were filmed OSO-mm diameter monofilament, filmed shell, and filter-wool (which was not filmed at the beginning of the experiment). There was no significant difference among these three substrata in numbers of larvae or spat collected per unit surface area. Filter-wool collected at least twice as many larvae and three times more spat per unit surface area than any other non-filmed settlement substratum. 3.2. Experiment 2 The number of larvae and spat collected per unit surface area varied significantly with position on rack or block (larvae: F = 9.56, df = 4, 20, P I 0.0005; spat: F = 7.06, df = 4, 20, P I 0.001) and also with mesh size (larvae: F = 25.06, df = 5, 20, P I 0.0001; spat: F = 8.92, df = 5, 20, P I 0.0001) (Fig. 2(a), (b)). The smallest mesh size of 3.0 mm collected significantly more larvae and spat per unit surface area than any other mesh size. The next three smallest meshes (i.e. 3.0 X 11.0, 3.8, and 7.0 mm) collected significantly more larvae per unit surface area than the two largest meshes (i.e. 11 .O and 19.0 mm). For spat, only one other pairwise comparison was significant: 3.8-mm mesh collected significantly more spat per unit surface area than 11 .O-mm mesh.

4. Discussion The significant block effect in both experiments indicates that larval and spat collection by the substrata was not equivalent among different positions on the rack. In general, significantly more larvae and spat were collected in the centre and at one end of the rack than at the other end. This is most probably due to subtle differences in water movements that may have affected the transport of larvae to different areas of the tank. Although the larvae were reared under near-static conditions, the airstone, which was kept at one end of the tank throughout the experiment (i.e. the end with lower larval and spat numbers), may have induced a characteristic flow pattern within the tank which could have ‘delivered’ larvae to certain areas of the rack. Also, flow was induced during

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water changes. Although larvae that were screened off were not re-introduced into the tank until after it had been refilled, many larvae could have been left in the water that remained in the bottom of the tank covering the collecting substrata. These larvae may have been carried to particular areas of the rack by the inflowing water. The block effect is interesting in that it shows how even subtle differences in water flow can affect larval settlement rates over small spatial scales. Hatcheries settling out scallop spat in large containers should be aware of this possibility and place settlement substrata throughout the tanks in order to maximize spat collection. The standard scallop spat collector used in most field studies of pectinid settlement has consisted of some sort of mesh bag stuffed with a filamentous substratum such as fishing line or gill netting (see Appendix A). This type of collector was first developed and used on a commercial basis in Japan (Ventilla, 1982) and the basic design has since been adopted by a number of countries for their own scallop industries. Despite the large amount of pectinid research that has made use of this type of collector and its prevalence in the aquaculture industry, surprisingly little work has examined the effect of filament diameter on spat collection efficiency. Wallace (1982) reported that spat of the Iceland scallop, Chlunzys islundicu, were always found attached to 0.15mm diameter nylon monofilament but never to thicker 0.8-mm diameter monofilament. The total surface areas of the two monofilament treatments were not given in that study, however. Similarly, Thorarinsdottir (1991) reported that smaller diameter monofilament (0.2 mm) collected more spat of C. islandica than larger diameter monofilament (0.4 mm). However, it is difficult to say that collection efficiency was directly related to monofilament size in that study since the two treatments were contained in different types of collecting bags with dissimilar mesh sizes and different quantities of the two monofilaments were used. Pouliot et al. (1995) compared spat collection of the giant scallop, Placopecten mageflanicus, by various diameters of monofilament (0.17, 037, 0.55, 0.77, and 0.90 mm, all sizes standardized for surface area) in standardized collectors in the field and found no significant difference among the five sizes. Miron et al. (1995) found that the same five diameters of monofilament did not vary significantly in the numbers of inert particles they passively caught under flume conditions, suggesting that settlement of giant scallop larvae on collectors in the field may be a passive process. The results of the present experiment agree with those of Pouliot et al. (1995) and Miron et al. (1995) in that there was no evidence for larval settlement choice among five different diameters of monofilament (0.50,0.75, 1.10, 1.50, and 2.00 mm>. The present study was conducted under near-static conditions in a relatively confined space and should facilitate the detection of larval choice among treatments (i.e. the study was more powerful than field or flume experiments). It is apparent that, within the size range tested, there does not appear to be any direct selection for certain monofilament diameters, although certain diameters could interact with external bag meshing size to enhance or reduce attachment (see Pouliot et al., 1995; Miron et al., 1995). Marine microbial films are known to enhance larval settlement and metamorphosis in a wide variety of benthic marine invertebrates including hydrozoans, scyphozoans, polychaetes, gastropods, bivalves, cirripedes, bryozoans, asteroids, and echinoids (see reviews by Meadows and Campbell, 1972; Crisp, 1974; Burke, 1983; Pawlik, 1992; Rodriguez et al., 1993; Pearce, 1996). Work with pectinids on the effect of microbial

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films on larval settlement has given various results. Hodgson and Boume (1988) found that, under laboratory conditions, significantly more larvae of the spiny scallop, Chfarnys hastutu, metamorphosed in response to filmed artificial surfaces (i.e. pieces of glass slides and paraffin plastic) than to similar non-filmed surfaces. Parsons et al. (1993) greater giant scallop reported similar results from the field showin g significantly (Placopecten mugellunicus) recruitment occurring on artificial substrata with a high biofilm coverage than on those with less coverage. Xu et al. (1991) showed how larval settlement rates of the bay scallop, Argopecten irrudiuns, could vary based on the bacterial strain of the film. Some strains induced 76% more larvae to settle than a control while others failed to initiate or even inhibited settlement. Our own laboratory work has shown how filmed filter-wool collects significantly more spat of A. irrudiuns than non-filmed filter-wool (Pearce and Bourget, unpublished results, 1993). Bacterial films appear to be less important in the settling response of larvae of Pecten maximus, however (Tritar et al., 1992). In the present experiment, filmed monofilament and filmed adult shell collected significantly more spat per unit surface area than their respective non-filmed counterparts. Filmed monofilament also collected significantly more larvae per unit surface area than non-filmed monofilament. These results, obtained under near-static conditions in the laboratory at a scale where substratum choice was possible, confirm the conclusions of Parsons et al. (1993) that giant scallop larvae actively select filmed substrata. The reason(s) for pectinid larvae to actively choose a filmed surface at settlement is/are unknown, but may be related to a transitional form of feeding occurring between the ciliated-velum phase of the planktonic larva and the gill filament phase of the benthic juvenile (Boume and Hodgson, 1991). In most species of bivalves the foot persists after metamorphosis and, in a number of species, has been implicated in direct deposit feeding of young juveniles (Caddy, 1969; Bayne, 197 1; Aabel, 1983). Bivalve larvae that settle on appropriately filmed substrata may be able to bridge the transitional period between velar and gill feeding by depositional feeding with the foot (i.e. pedal-palp feeding). 6 Foighil et al. (1990) found that significantly higher numbers of Japanese scallop (Putinopecten yessoensis) spat re-attached to lightly fouled cultch than to clean cultch and that the presence of this microbial film significantly enhanced early post-metamorphic growth. 0 Foighil et al. (1990) pointed out, however, that the nutritional importance of feeding on benthic films was much less than that of suspended microalgae and suggested that this post-metamorphic foot-feeding may be a vestigial behaviour in this species. Settlement of pectinid larvae on filmed surfaces could also be related to an indication that the substratum is not actively grazed or browsed and, hence, is a ‘safe’ area to settle into (an indication akin to the detection of congeners in sessile species). Surface heterogeneity is another factor known to influence settlement of benthic marine invertebrate larvae (see reviews by Crisp, 1974; Crisp, 1976; Burke, 1983), but very little research has examined directly the effect of surface complexity on larval settlement in pectinids. From the scant information available it appears that roughened surfaces are not a necessary prerequisite for settlement of larval scallops; a number of laboratory studies have shown settlement on smooth glass slides or on plastic/glass containers or tanks @astry, 1965; Comely, 1972; Castagna, 1975; Dix, 1976; Hodgson

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and Bourne, 1988; Tremblay, 1988; Kingzett et al., 1990; Chevolot et al., 1991). Culliney (1974) found no difference in larval settlement rate of giant scallop larvae in response to small pebbles, fragments of adult shell, and glass shards. In the present study there was no significant difference in numbers of larvae or spat collected by roughened or smooth plates of acrylic plastic and non-filmed shell did not differ significantly from either of these two treatments. These results suggest that surface roughness, at the scale tested, did not affect settlement rates of giant scallop larvae. Ambrose and Lin (1991) found that settlement preferences of bay scallop ( Argopecfen irrudians) larvae for artificial meshes in controlled multi-choice laboratory experiments were dependent on what other substrata were present for settlement. Larvae settled in significantly greater densities on 3-mm clear unoriented polypropylene mesh than on 4-mm black oriented polypropylene mesh when the two meshes were presented without any other substrata in two-choice experiments. When presented with three other substrata (5-mm black unoriented polyethylene mesh, Astroturf, and white burlap cloth), however, there was no significant difference in initial settlement densities between the two meshes; Astroturf collected the most spat over time, however. In field experiments, comparing recruitment on collectors filled with either the 3-mm or 4-mm mesh, more spat were found in collectors with the smaller mesh (Ambrose et al., 1992). In both of those studies, however, the different substrata were not standardized for surface area, leading the authors to conclude that this was the predominant factor controlling larval settlement rate (Ambrose and Lin, 1991; Ambrose et al., 1992). Our multi-choice experiment revealed that the smallest polyethylene mesh size of 3.0-mm collected significantly more larvae and spat per unit surface area than any other mesh size. Since results were standardized for surface area, it is not simply a question of substratum surface availability. Although all meshes were composed primarily of polyethylene (92-96% for the low-density polyethylene meshes and 98% for the high-density polyethylene mesh), they may have also contained other chemicals in variable and low concentrations that were added for colour (e.g. pigments), softening (e.g. ethyl-vinyl acetate), and slipperiness. Since pigments differ between the green and black meshes and concentration of agents added for softening and slipperiness can vary with lot, larvae may have been responding to differences in chemical cues and/or surface texture. One could argue that colour, in itself, was the important factor controlling settlement rate in this experiment since the 3.0-mm mesh was green whereas all other meshes were black, but Naidu (1980) reported that colour per se was not a critical factor in the settlement of giant scallop larvae. By far the best substratum tested for settlement of larval giant scallops under the hatchery-like conditions in these experiments was the filter-wool. It collected significantly greater numbers of larvae and spat per unit surface area than any other non-filmed substratum. Although not comparable statistically, it also collected more larvae and spat per unit surface area than the 3.0-mm green Vexar mesh (165.1 f 47.5 vs 129.8 f 10.9 larvae per 100 cm* respectively, and 7.3 + 2.3 vs 3.4 f 0.3 spat per 100 cm*, respectively: mean + s.e.m.). Filter-wool is most likely an even more efficient settlement substratum than these results suggest, since a large portion of its surface area would have been unavailable for larval settlement due to overlapping of compacted filaments. Thus, the surface area available for larval settlement would have been much smaller than

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the actual total surface area. All other settlement substrata, except for the monofilaments, would not have experienced this compacting problem. Conditioning filter-wool in running seawater prior to the experiment would have increased larval settlement even further. This has been shown in our laboratory with bay scallop larvae (Pearce and Bourget, unpublished results, 1993). On a per surface area basis, filter-wool is a relatively cheap substratum (- $0.32 m- 2, Canadian currency) in comparison with monofilament (- $.5.00-$6.00 mm2), the standard cultch material used by industry at present. It is inert in seawater (made from polyethylene), would be easy to clean and re-use, and would last a relatively long time. All these properties point to filter-wool being an excellent cultch material for giant scallop larvae under hatchery-type conditions, but further research is warranted on a commercial scale to determine its practicality. For large-scale spat production, greater quantities of filter-wool would be required and a collector that prevented compacting of the fibres would be necessary, since the filter-wool filaments tend to pack together quite tightly when immersed in water. Stretching the filter-wool out in a sheet-like fashion in a horizontally placed collector may be all that would be required to prevent compacting. Also, further work would be needed to determine the difficulty of removing live spat from the filter-wool fibres. Samples of filter-wool were often examined in the present study after the spat removal process and no individuals were ever found, but this was after the spat had been preserved in alcohol, soaked in a sodium hypochlorite solution, and rinsed vigorously with a jet of water. Live spat may be more difficult to remove from the filaments. Commercial-scale experiments examining the relative difficulty of spat removal from a number of standard cultch materials in relation to filter-wool would be necessary.

Acknowledgements

Much appreciation is extended to R.K. O’Dor and R.E. Scheibling for the use of dry and wet lab facilities. D.L. Krailo is thanked for rearing the phytoplankton. Special gratitude is extended to Charlie Thompson for help with making the collector units. Alan Jeffs, of DuPont Canada Inc., was very helpful in providing product information on Vexar. The suggested changes of three anonymous reviewers were greatly appreciated. The work was carried out with funding to E. Bourget from OPEN (Ocean Production Enhancement Network, one of the 15 Networks of Centres of Excellence supported by the Government of Canada). C.M. Pearce has been funded by NSERC (Natural Sciences and Engineering Research Council), OPEN, FCAR (Fonds pour la Formation de Chercheurs et 1’Aide ‘a la Recherche), GIROQ (Groupe Interuniversitaire de Recherches OcCanographiques du Quebec), IFRP (Interim Funding Research Programme), the O’Brien Foundation, and Universite Laval.

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Appendix settlement

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141 (1996) 201-221

A. Review of types of artificial substrata that have been used as surfaces for pectinid spat collection in both laboratory and field studies

Species Laboratory studies Amusium halbti Argopectm irrdians

Argopecten purpurutus Chlomys austrulis Chlamys firrreri Chlrmys hastutu

Chlamys islandicu Chlamys nohilis ( = Mimachlamys n0hili.s) Equichlamys hifrons Patinopecten yrssoensis ( = Mizuhopecten yess0en.si.s)

Pecten maximus

Plucopectm mogellunicus

Field studies Amusium japonicum halbti Argopecten circularis ( = A. uentricosus)

Substratum

Reference

Old monofilament shark netting Plastic container a plastic Panels of wood d, Mylar, fibreglass, container a,d 3-mm transparent polypropylene mesh, 4-mm black polypropylene mesh, 5-mm black polyethylene mesh, polypropylene artificial turf, burlap cloth Sieves Plastic web nets, plastic film, plastic plates, coir rope, coir film Netron net Old monofilament shark netting Plastic web nets, plastic film, plastic plates, coir rope, coir film Frayed polypropylene line, scallop shell, filmed d/non-filmed glass slides and paraffin plastic Monofilament (0.3 mm) b Plastic web nets, plastic film, plastic plates, coir rope, coir film

Cropp, 1993b Sastry, 1965 Castagna, I975

Polyethylene containers a Hemp palm bark Oyster and scallop shell, monofilament line, polypropylene rope, Vexar ‘, artificial turf, jute, sisal, Kinran dz 24well tissue culture plates a Conditioned Kinran ’

Dix, 1976 Tanaka et al., 1987 Boume and Hodgson, Boume et al.. 1989

Sieves (125 pm) f Plastic web nets, plastic film, plastic plates, coir rope, coir film Slate, tile, bottom of glass containers hd Bottom of tanks a Polyester nets Pebbles, fragments of scallop shell, glass shards Shell material, fine mesh, gill netting, Nitex s, Kinran d.e, plastic container a.d

Polypropylene

onion bags (5 mm)

f

Shrubs, polypropylene nets, shells, polyfilm Plastic filament (0.4-0.5 mm) b Monofilament

string d, shrubbery,

shells

Ambrose

and Lin, 1991

Kamey, 1991 Lou, 1991 Piquimil et al., 1991 Cropp, l993a Lou, 1991 Hodgson

and Boume.

1988

Harvey et al., 1993 Lou. 1991

1991;

Kingzett et al., 1990 6 Foighil et al., I990 &chard et al., 1991 Lou, 1991 Comely, I972 Roman, 1991 Tritar et al., 1992 Culliney, 1974 Tremblay, I988

Robins-Troeger I993 Felix-Pica, I99 Ruiz-Verdugo Martinez, 1991 Tripp-Quezada,

and Dredge, I and Caceres1991

C.M. Peurce, E. Bourget/Ayurrculture Argopecten

3-mm transparent polypropylene mesh, 4-mm black polypropylene mesh, 3-, 5, 15mm black polyethylene meshes 3-mm transparent polyethylene mesh, 4-mm black polyethylene mesh Plastic netting, nylon threads Onion and potato mesh sacks (2 X IO mm) f Netlon d.h, gill netting, acetate film

irradiuns

Ch1amy.s jkeri Chlamys glahru Chlumys

hastata

Chlumys

ruhidu

Chlumys

islandicu

Chlumys

opercularis

( = Aequipecten

and

opercu-

lurk)

Chlamys

tehuelchus

( = Aeyuipecten tehuelchus) Chlcrmys uariu Drcatoprcten

Flexopecten Hinnites

d. strangei

jlexuosus

multirugosus

Leptopecten

k

luticrurutus

Nylon monofilament (0.15 d or 0.8 mm) b Nylon gill netting (0.2 d or 0.4 mm) b Netlon greenhouse shading net ‘, Hairlok d,‘, onion bags d, teased polypropylene rope, orange bags Netlon greenhouse shading material (3 mm) f, plastic mesh sacks (6 mm) Q Monofilament nylon mesh Monofilament garden netting Monofilament nylon netting (20 mm) f Monofilament Teased polyethylene monofilament Polyethylene monofilament, hemp fibre, shrub branches d Monofilament Weathered nylon gill netting, polypropylene onion bags (5 mm) f Polypropylene onion bags (5 mm) f Monofilament Gill netting (twine size #I41 d, gill netting partially coated with cement d, scallop shells, dry chaparral sticks Hanging culture oyster strings Gill netting (twine size #I41 d, gill netting partially coated with cement d, scallop shells, dry chaparral sticks Weathered nylon gill netting, polypropylene onion bags (5 mm) f Polypropylene

Patinopecten ( = Mizuhoprcten yessoensis)

Pecten albu Pecten jiuncrtus

yessoensis

215

141 (1996) 201-221

onion bags (5 mm) f

Cedar leaves, Japanese cypress leaves, pine needles, Hizex ‘, scallop shells, bark of hemp palm, used gill netting d, Netlon net Cedar twigs, scallop shells, nylon gill netting d, Netlon d.h, Hizex d.’ Japanese cedar twigs, plastic sheets, plastic plates, gill netting d Moulded polyethylene cones Old nylon gill netting Monofilament shark net Monofilament shark net Old nylon gill netting

Ambrose

et al., 1932

Peterson and Summerson, 1992 Lou, 1991 Lykakis and Kalathakis, 1991 Rhee (unpublished data cited in Rhee, 1991) Wallace, 1982 Thorarinsdottir, 199 I Brand et al., 1980

Paul et al., 1981 Fraser, 1983 Fraser and Mason, 1987 Fraser, 199 I Margus, 1991 Ruzzante and Zaixso, 1985 Zaixso, 1980 (referenced in Orensanz et al., 1991) Margus, 199 I Sumpton et al., 1990 Robins-Troeger and Dredge, 1993 Magus, I99 I Phleger and Gary, 1983

Boume, 1991 Phleger and Gary, 1983

Sumpton

et al., 1990

Robins-Troeger 1993 Motoda, I977

Ventilla, Sekino,

and Dredge,

1982 1983

Kalashnikov, 1991 Sause et al., 1987 Hortle and Cropp, 1987 Cropp and Hortle, 1992 Young et al., 1992

216

C.M. Pearce, E. Bourget /Aquaculture Polypropylene

Pecten jacobaeus Pecten murimus

Pecten novuezelundiae Pecten vogdesi Placopecten mugellunicus

141 (1996) 201-221

onion bags (5 mm) f

Monotilament Netlon ’ Net bags of nylon or polyethylene Netlon greenhouse shading material (3 mm) f, plastic mesh sacks (6 mm) d.j Monofilament nylon mesh Monofilament garden netting Monofilament mesh (6 mm) f Monofilament nylon netting (20 mm) f Netlon netting (5 mm) f Polypropylene rope, monofilament netting, Japanese lantern nets m Synthetic rope, polyethylene mesh, used monofilament gill netting Plastic filament (0.4-0.5 mm) b Frayed rope ends, wooden buoys d, scallop shells d Navigation buoy Hizex ’ Monofilament gill netting (twine size #14) d, Hizex ’ Monofilament gill netting d, Hizex ’ Monofilament gill netting 9/ 13 monofilament gill netting Monofilament gill netting Nylon monofilament gill netting (twine size #14: 0.6 mm b, Monofilament (0.17, 0.37, 0.55, 0.77, 0.90 mm) b

Robins-Troeger and Dredge, 1993 Margus, 1991 Mason, 1969 Pickett, 1978 Paul et al., 1981 Fraser, 1983 Fraser and Mason, 1987 Wilson, 1987 Fraser, 1991 Thouzeau, 1991a, Thouzeau, 1991b Minchin, 1992 Bull, 1991 Ruiz-Verdugo and Martinez, 1991 Naidu, 1970

Cbceres-

Merrill and Edwards, 1975 Naidu and Scaplen, 1979 Naidu et al., 1981 Naidu and Cahill, 1986 Tremblay, 1988 Dadswell and Parsons, 1991 Robinson et al., 1992 Parsons et al., 1993 Pouliot et al., 1995

a Larvae metamorphosed on container that they were reared in. b Diameter of filaments. ’ Low density polyethylene mesh. d Best substratum tested for collecting spat. e Artificial fibrous material manufactured in Japan. f Diameter of mesh. g Nylon monotilament screen. h Mesh. ’ Rubberized fibre. j Diameter of mesh (stretched diagonal). ’ Now known as Crussuddma gigantea. ’ Polyethylene film. m Three best substrata, but many others were reported on (see paper).

References Aabel, J.P., 1983. Morphology and function in postmetamorphal Abra alba (Bivalvia: 213-219. Abacus Concepts, 1989. SuperANOVA. Abacus Concepts Inc., Berkeley, 322 pp.

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Ambrose, W.G., Jr. and Lin, J., 1991. Settlement preference of Argopecren irradiuns (Lamar& 1819) larvae for artificial substrata. In: S.E. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. 1. The World Aquaculture Society, Baton Rouge, pp. 16-20. Ambrose, W.G., Jr., Peterson, C.H., Summerson, H.C. and Lin, J., 1992. Experimental tests of factors affecting recruitment of bay scallops (Argopecfen irradians) to spat collectors. Aquaculture, 108: 67-86. Bayne, B.L., 1971. Some morphological changes that occur at the metamorphosis of the larvae of Myti!us edulis. In: D.J. Crisp (Editor), Proceedings of the 4th European Marine Biology Symposium. Cambridge University Press, Cambridge, pp. 259-280. Bourget, E., 1988. Barnacle larval settlement: the perception of cues at different spatial scales. In: M. Vannini, M. Chelazzi and G. Chelazzi (Editors), Behavioural Adaptations to the Intertidal Life. Plenum Press, New York, pp. 153-172. Boume, N., 1964. Scallops and the offshore fishery of the Maritimrs. Fish. Res. Board Can. Bull., No. 145.60 PP. Boume, N., 1991. Fisheries and aquaculture: west coast of North America. In: S.E. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 925-942. Boume, N. and Hodgson, C.A., 1991. Development of a viable nursery system for scallop culture. In: S.E. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. 1. The World Aquacuhure Society, Baton Rouge, pp. 273-280. Boume, N., Hodgson, C.A. and Whyte, J.N.C., 1989. A manual for scallop culture in British Columbia. Can. Tech. Rep. Fish. Aquat. Sci., No. 1694, 215 pp. Brand, A.R., Paul, J.D. and Hoogesteger, J.N., 1980. Spat settlement of the scallops Chlamys opperculuris CL.1 and Pecten muximus CL.1 on artificial collectors. J. Mar. Biol. Assoc. UK, 60: 379-390. Bull, M.F., 1991. Fisheries and aquaculture: New Zealand. In: SE. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 853-859. Burke, R.D., 1983. The induction of metamorphosis of marine invertebrate larvae: stimulus and response. Can. J. Zool., 61: 1701-1719. Butman, C.A., 1987. Larval settlement of soft-sediment invertebrates: the spatial scales of pattern explained by active habitat selection and the emerging role of hydrodynamical processes. Oceanogr. Mar. Biol. Annu. Rev., 25: 113-165. Caddy, J.F., 1969. Development of mantle organs, feeding, and locomotion in postlarval Macomu balthicu (L.) (Lamellibrancbiata). Can. J. Zool., 47: 609-617. Castagna, M., 1975. Culture of the bay scallop, Argopecten irrudiuns, in Virginia. Mar. Fish. Rev., 37: 19-24. Chevolot, L., Cochard, J.-C. and Yvin, J.-C., 1991. Chemical induction of larval metamorphosis of Pecten maximus with a note on the nature of naturally occurring triggering substances. Mar. Ecol. Prog. Ser., 74: 83-89. &chard, J.C., Comps, M., Buestel, D. and Paquotte, P., 1991. Introduction of the Japanese scallop Patinc+wcf~n yasoensis (Jay) into France and the hatchery production of the Fl generation. In: S.E. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. 1. The World Aquaculture Society, Baton Rouge, pp. 281-286. Comely, CA., 1972. Larval culture of the scallop Pccrcn muximus CL). J. Cons. Perm. Int. Explor. Mer. 34: 365-378. Couturier, C., 1990. Scallop aquaculture in Canada: fact or fantasy? World Aquaculture, 21: 54-62. Crisp, D.J., 1974. Factors influencing the settlement of marine invertebrate larvae. In: P.T. Grant and A.M. Mackie (Editors), Chemoreception in Marine Organisms. Academic Press, New York, pp. 177-265. Crisp, D.J., 1976. Settlement responses in marine organisms. In: R.C. Newell (Editor), Adaptation to Environment: Essays on the Physiology of Marine Animals. Butterworths, London, pp. 83- 124. Cropp, D.A., 1993a. Hatchery culture potential of the scallop Chlumys ousrralis in Western Australia. Aquaculture, 1 15: 3 I-40. Cropp, D.A., 1993b. Development of large-scale hatchery production techniques for Amusium balloti (Bemardi 18611 in Western Australia. Aquaculture, I 15: 285-296.

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Cropp, D.A. and Hortle, M.E., 1992. Midwater cage culture of the commercial scallop Pecren @murus Reeve 1852 in Tasmania. Aquaculture, 102: 55-64. Culliney, J.L., 1974. Larval development of the giant scallop Plucoppecren magellunicus (Gmelin). Biol. Bull., 147: 321-332. Dadswell, M.J. and Parsons, G.J., 1991. Potential for aquaculture of sea scallop, Plucopecren mugeUunicua (Gmelin, 1791) in the Canadian Maritimes using naturally produced spat. In: SE. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. I. The World Aquaculture Society, Baton Rouge, pp. 300-307. Dix, T.G., 1976. Larval development of the queen scallop, Equichlumys hifrons. Aust. J. Mar. Freshwat. Res., 27: 3999403. Felix-Pica, E.F., 1991. Fisheries and aquaculture: Mexico. In: S.E. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 943-980. Fraser, D.I., 1983. Observations on the settlement of pectinid spat off the west coast of Scotland in 1982. Int. Count. Explor. Sea Comm. Meet. 1983/K:40, 12 pp. Fraser, D.I., 1991. Settlement and recruitment in Pecren maximus (Linnaeus, 1758) and Chlumys (Aequipec1758). In: S.E. Shumway and P.A. Sandifer (Editors), An International ten) operculuris (Linnaeus, Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. 1. The World Aquaculture Society, Baton Rouge, pp. 28-36. Fraser, D.I. and Mason, J., 1987. Pectinid spat studies in selected areas of the west coast of Scotland, 1982-1986. Int. Count. Explor. Sea Comm. Meet. 1987/K:4, 9 pp. Gallager, S.M., Bidwell, J.P. and Kuzirian, A.M., 1989. Strontium is required in artificial seawater for embryonic shell formation in two species of bivalve molluscs. In: R.E. Crick @ditor), Origin, Evolution, and Modem Aspects of Biomineralization in Plants and Animals. Proceedings of the Fifth International Symposium on Biomineralization, Arlington, Texas. Plenum Press, New York, pp. 349-366. Harvey, M., Bourget, E. and Miron, G., 1993. Settlement of Iceland scallop Chkumys iskundicu spat in response to hydroids and lilamentous red algae: field observations and laboratory experiments. Mar. Ecol. Prog. Ser., 99: 283-292. Heasman, M.P., O’Connor, W.A. and Frazer, A.W., 1994. Detachment of commercial scallop Pectenfimatus, spat from settlement substrates. Aquaculture, 123: 40-407. Hodgson, C.A. and Boume, N., 1988. Effect of temperature on larval development of the spiny scallop, Chkumys hastutu Sowerby, with a note on metamorphosis. J. Shellfish Res., 7: 349-357. Hortle, M.E. and Cropp, D.A., 1987. Settlement of the commercial scallop, Pecfenfitmu~us (Reeve) 1855, on artificial collectors in eastern Tasmania. Aquaculture, 66: 79-95. Kalashnikov, V.Z., 1991. Fisheries and aquaculture: Soviet Union. In: S.E. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 1057-1082. Kamey, R.C., 1991. Ten years of scallop culture on Martha’s Vineyard. In: S.E. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. I. The World Aquaculture Society, Baton Rouge, pp. 308-3 12. Kingzett, B.C., Boume, N. and Leask, K., 1990. Induction of metamorphosis of the Japanese scallop Purinopecten yessoensis Jay. J. Shellfish Res., 9: 119- 125. Lou, Y., 1991. Fisheries and aquaculture: China. In: S.E. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 809-824. Lykakis, J.J. and Kalathakis, M., 1991. Fisheries and aquaculture: Greece. In: SE. Shumway (Editor), Scallops: Biology, Ecology and Aquaculture. Developments in Aquaculture and Fisheries Science, Vol. 21. Elsevier, New York, pp. 795-808. Margus, D., 1991. Settlement of pectinid larvae in the Krka River estuary of Yugoslavia. In: S.E. Shumway and P.A. Sandifer (Editors), An International Compendium of Scallop Biology and Culture. World Aquaculture Workshops, No. 1. The World Aquaculture Society, Baton Rouge, pp. 37-42. Mason, J., 1969. The growth of spat of Pecten mu_ximu.\ CL.). Int. Count. Explor. Sea Comm. Meet. 1969,‘K:32, 5 pp.

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