Nick C. Parker, Gerald T. Klar, Theodore I. J. Smith, and Jerome Howard Kerby
Striped bass (Morone saxatilis) and its hybrid striped bass produced in state and federal hatcheries are normally reared in static earthen or vinyl-lined ponds using techniques common for other warmwater species (Dupree and Huner 1984). However, increased demand for fish by both public and private sectors has stimulated interest in improving production efficiency through a variety of techniques, including semi-intensive production involving supplemental aeration. Efforts have also focused on techniques to reduce stress to improve survival of fish during all phases of rearing, as well as during and after transport. Research on techniques to produce progeny from controlled spawning of domesticated brood stock and through the use of cryopreserved sperm are on-going. We describe some special culture techniques now in use in some facilities involved in semi-intensive and intensive culture of the striped bass and its hybrids.
Aeration and Oxygenation
Dissolved oxygen (DO) concentrations in striped bass rearing ponds may decline below desirable levels for several reasons: dense phytoplankton blooms; high organic loading; high fish biomass; cloudy, still weather; and others. At these times, supplemental or emergency aeration may be necessary to insure adequate DO levels for fish survival.
There are two approaches to artificial pond aeration. Emergency aerators, which are relatively portable devices that can be moved from pond to pond as needed, are often used. Secondly, continuous aeration or pond circulation can be provided through the use of permanently installed devices such as axial flow pumps, submerged fans, paddle wheels, airlift pumps, air diffusers, oxygen generators, and liquid oxygen injection systems.
Circulation of pond water has proven beneficial, not only in striped bass ponds (Parker 1980, Parker et al. 1984), but also in catfish and shrimp ponds (Busch 1980; Busch and Goodman 1983; Rogers and Fast 1988). Mixing surface and bottom layers of water reduced thermal stratification and increased levels of DO at the pond bottom in all of these studies. Many of the supplemental aeration systems can be commercially purchased, so with the exception of airlift systems, are not described in detail here. We next discuss inherent advantages and disadvantages, and other information, on each system.
Parker (1980) reported that fingerling striped bass production was 2.4 times greater in ponds aerated continuously with airlift pumps than in 0.1-acre ponds without aeration. Continuously operated airlift pumps appear to maintain uniform temperatures and DO throughout the pond by continuously circulating the water, which results in mixing oxygen-rich surface water with oxygen-deficient bottom water (Figure 12.1). Air injected into water with an airlift pump adds very little oxygen to a pond compared with oxygen diffusion at the air-water interface. Oxygen produced by photosynthesis in the upper layer of the water column is uniformly distributed throughout the pond by airlift pumps, which serve to alleviate potentially dangerous levels of gas supersaturation common in pond surface water in the afternoon and low levels of DO in the morning (Parker et al. 1984). Emergency aeration may still be required during prolonged periods of cloudy weather or if pond vegetation is dense. Another advantage of this type of aeration is that it can be supplied in the culture of phase I fish (about 1-2.5 inches long) with little risk of damage to larvae and small fish. Water velocities in airlift pumps are low, and any fish that might be entrained are gently moved through the pumps. A negative concern is that operation of continuous aeration systems in ponds with large accumulations of
|Figure 12.1. Concentration of dissolved oxygen at the bottom of striped bass production ponds equipped with airlift pumps to circulate water (treatment) and in ponds without water circulation (control).|
organic matter on the pond bottom may actually reduce the average level of DO. This reduction in DO results from oxygenated surface water moving to the bottom of the pond, where oxygen is used to oxidize accumulated organic materials. To alleviate this situation, ponds should be drained, dried, and disked to oxidize organic deposits before they are filled, and frequent aeration procedures should later be used (see Chapters 8 and 9). Ideally, organic material added daily would be oxidized daily and no oxygen deficient accumulations would occur.
An airlift system consists of a blower, an air distribution line, and an airlift pump. Blowers deliver large volumes of low-pressure air and may be of the rotary vane or regenerative type (see Figure 6.2). A l-kW blower will supply enough air to operate 25 airlifts in a 2.5-acre pond at a cost of about $1.25 per day, based on US $0.07/kW-h. The blower may be installed on the pond bank or in a central location so it can supply several ponds. Protection of the blower from weather is desirable.
A wide variety of materials are suitable for air distribution lines since the operating pressure is usually about 1 psi. Rigid polyethylene or polyvinyl chloride (PVC) pipe may be permanently installed underground, or fiber-reinforced polyethylene pipe may be installed at the surface, to supply air to the airlift pumps. Air lines in excess of 100 feet should be at least 4 inches (and preferably 6 inches) in diameter to reduce friction and prevent excessive back pressure on the blower. Thin-walled, 6-inch diameter PVC pipe buried just beneath the surface makes an excellent main line for an air distribution system. A standard 6-inch glue-on coupling is placed at each airlift location. A hole is drilled into the side of the coupling and tapped to receive a 0.5-inch national pipe thread (NPT) fitting. A 0.5-inch PVC valve is attached to the tapped hole with a length of PVC pipe sufficient to place the valve above the surface of the ground. Air is transferred through 0.5-inch flexible polyethylene tubing from the valve to the airlift pump. The valve is adjusted (or a small plate with a 1/8- to 1 /4-inch orifice is fitted inside the 0.5-inch tubing) to maintain a minimum of 30-32 inches of water pressure (1.08-1.16 psi) in the main air distribution line; this insures even delivery of air to all pumps, even if they are not uniformly submerged in water.
Airlift pumps, which may be purchased commercially or constructed from readily available materials, consist of a vertical PVC pipe with a 90° elbow at the water surface (Figures 12.2 and 12.3). Air is injected into the vertical pipe 24 inches below the water surface. The air-water mixture becomes lighter than water, rises, and is discharged through the elbow surface. The elbow directs the water flow horizontally across the pond. The pivot point (Item 8, Figure 12.3), may be adjusted to match pond depth. The pump holder, a floating bracket (Item 6, Figure 12.3) then pivots to maintain discharge at the water surface when pond levels
rise or fall. Support for the airlift can be a post set in the pond bottom (Item 7, Figure 12.3) or in a concrete block (Item 4, Figure 12.2), that can be readily removed from the pond during harvest or moved from pond to pond. Any support system that maintains discharge at the surface would be suitable for airlift operations.
Size and pumping rates. Airlift pumps can economically mix oxygen-rich surface water with oxygen-deficient bottom water to improve environmental conditions in ponds (Parker,
(insert figure 12.2 here)
Figure 12.2. Airlift pump (1), with float (2), located on the swing arm assembly (3), and with the vertical support (4), cemented into a concrete block. Air supply to the pond is controlled by a valve. Photo Credit: Nick C. Parker.
|Figure 12.3. Construction detail and pond installation of a floating airlift pump. Detailed view of airlift pump with (1) hole for mounting airlift pump, (2) polyvinyl chloride (PVC) 90° elbow, (3) vertical riser constructed from PVC pipe, (4) 900 hose adaptor for air injection pipe, and (5) ballast with (a) depth of air injection dependent on air pressure available and (b) depth of airlift pump intake established by pond depth. Top view of (6) pump holder, (7) post anchor system, (8) pivot point, (9) flotation material, and (10) attachment of airlift pump to the pump holder. Side view of airlift pump installed with center of discharge at the pond surface (11), and with the intake just above the pond bottom (12). Direction of water (solid arrows) and air (open arrows) flow through the airlift pump (From Parker 1983).|
1979,1980). On the basis of material costs and operation, airlift pumps with a diameter of 3-4 inches seem to be the most efficient in circulating water within a pond. A detailed description of airlift pumping rates was given by Parker and Suttle (1987). When 3 cubic feet per minute (cfm) of air was injected 24 inches below the surface and the center line of the discharge was at water level (zero vertical lift), a 3-inch airlift pumped 43 gallons/minute and a 4-inch airlift pumped 75 gallons/minute. If the centerline of the discharge was raised 0.5 inches above the water level, flows decreased 16% for 3-inch and 38% for 4-inch diameter pumps, respectively. The proper adjustment of smaller diameter airlifts was easier to maintain than larger diameter airlifts. Regenerative blowers from 0.75-3 hp (0.5~2.24 kW) operated 27 3- or 4-inch airlift pumps per kW. The water pumping rate per kW of power was 300 gallons/minute for properly adjusted 3-inch airlifts, and 530 gallons/minute for 4-inch airlifts. Twenty-five airlift pumps, with a total operational cost of about $1.25/day at $0.07/kW-h, are recommended for 2.5-acre ponds with an average depth of 4 feet. This would circulate the entire pond volume in 2.1 days for 3-inch airlifts, and 1.2 days for 4-inch airlifts.
The point of air injection, not the total length of the vertical riser, is the critical measurement in airlift pump construction. Air injected into water at depths less than 24 inches decreases flow, whereas injection at depths greater than 24 inches requires higher air pressure and reduces total airflow available from regenerative blowers. Vertical risers 3 feet long work well in ponds that average 4 feet deep. Cutting the bottom of the vertical riser at a 45° angle allows continuous pumping if the water level drops and the riser touches bottom.
Regenerative blowers and airlift pump aeration systems have several advantages and disadvantages:
1. They can replace or greatly reduce the number of electrically operated surface agitators.
2. Personnel safety is increased when several electrically powered surface agitators are replaced by one blower.
3. Small ponds can be economically circulated and destratified with airlift pumps.
4. Emergency aerator requirements are reduced.
5. Airlifts can be used during early culture of phase I fish.
6. Fish production can be increased when ponds are continuously circulated and aerated.
1. Air delivery lines must be installed to all ponds with airlift pumps.
2. Pumps require adjustment at irregular intervals to keep them operating efficiently.
3. Airlift pumps can interfere with seining and may need to be removed when fish are harvested.
4. Airlift pumps may not provide adequate aeration in certain situations where DO declines rapidly.
Surface agitators (paddle wheels, pumps, floating aerators, etc.) are typically used to provide emergency aeration on an intermittent basis (Boyd and Ahmad 1987), but may be used continuously during summer in some locations. Small floating aerators are usually preferred over large paddle wheels for use in the small ponds typical of phase I culture. Emergency aeration with surface agitators is generally not recommended during the first half of phase I culture because of the risk of injury to small fish. They can, however, be used safely and effectively during late phase I and advanced culture. Depending on type, some surface agitators produce small zones of well aerated water with little circulation, or greater circulation with lower levels of DO.
Electric paddle wheels can be effective in rapidly restoring oxygen to ponds with low DO. Boyd and Ahmad (1987) found that in small ponds (from which all oxygen had been chemically removed by the addition of sodium sulfite and cobalt) this type of aerator produced oxygen at 1.1 pounds per hour and 10.6 pounds per hour at an operational cost of US 50.16/hour for 1.5-kW units and $0.79/hour for 8.5-kW units, respectively. They also determined that paddle wheel aerators were more efficient than diffused air systems, judged by the weight of oxygen added to deoxygenated ponds per unit of time. Rapid recovery rates make this type of aerator highly desirable for use in emergency situations, and improved designs now allow more cost-effective use of paddle wheels on a continuous basis.
One 1 /3-tap floating aerator will normally provide sufficient aeration on an emergency basis for a l-acre pond during production of phase I and phase II striped bass (about 3-10 inches long). Experience at the Southeastern Fish Cultural Laboratory has indicated that the availability of one 1 /3-tap floating aerator for each two ponds in production should be sufficient to handle most emergency situations.
Liquid Oxygen and Oxygen Generators
Aeration may be provided for fish culture by an external source of pure gaseous oxygen. Oxygen can be purchased in liquid form, in which a large volume of oxygen gas is compressed in a relatively small cylinder; however, the tanks must be refilled or replaced (with full tanks) at frequent intervals. Commercially available oxygen generators provide a continuous flow of relatively pure (95%) oxygen at rates up to 96 L/min. In most oxygen generators, compressed air is supplied to a material, such as zeolite, that absorbs and removes nitrogen.
Methods of dissolving pure oxygen into water include diffusers, packed columns, U-tube systems, and Aquatectors® (Speece 1969; Watten and Beck 1985). Diffusers consist of porous stones, rods, or plastic pipe, as well as serrated or slotted pipes, which are immersed directly in fish tanks or ponds. Oxygen is transferred more efficiently by small pore than by large pore diffusers or non-porous ones because the smaller bubbles they produce have larger surface area to volume ratios. However, small-pore diffusers require higher operating pressures and tend to clog easily. The efficiency of large pore diffusers can be increased by using an enclosed water column packed with plastic rings to increase bubble retention time.
Packed columns probably represent the minimum design to make the use of pure oxygen cost effective (Visscher and Godby 1987). A packed column usually consists of a 4- to 12-inch diameter sealed tube filled with plastic or other materials with high surface area to volume ratios. Water injected into the top of the column spreads across the surface of the packing media and absorbs the oxygen injected into the bottom of the column. Packed columns are normally operated under a vacuum of 20-50 mm Hg to effectively strip out nitrogen gas as oxygen is injected (Colt and Watten 1988).
In a U-tube system, water and oxygen are mixed and pumped down through a vertical pipe housed in a second, larger diameter pipe that contains the return flow. As the mixture of water and oxygen passes through the U-tube, hydrostatic pressure increases and oxygen rapidly diffuses into the water. Water exiting from the U-tube is fully saturated or supersaturated with oxygen. Commonly used U-tubes range from about 35-100 feet in depth. Essentially, they consist of a pipe placed inside a well casing with a plug at the bottom of the casing.
An Aquatector® (Zeigler Brothers, Inc., Gardners, Pennsylvania) is a device that mixes water and oxygen under pressure, shears the oxygen into micro-bubbles, and discharges the mixture into the rearing unit. The oxygen bubbles are so small and numerous that the discharge water appears milky white. Diffusion occurs across the tremendous surface area of the small bubbles.
Liquid oxygen, oxygen generators, U-tubes, and Aquatectors® are typically used in intensive culture situations where they are cost effective. The use of U-tube or Aquatector® use in ponds requires either circulating the pond water through the device or the continuous use of new water. Biological fouling may be a problem with recycling pond water if sufficient fresh water is not available.
Transportation of Striped Bass Fingerlings
Successful transportation of striped bass begins at harvest. In summer, fish should be harvested in the early morning when temperatures are lowest, and fish are least sensitive to stress. If possible, fish should not be harvested when pond temperatures exceed 85º F. We recommend that fingerlings be held in tanks for at least 24 hours after harvest to allow for recovery from stress. During this time, they should be treated in water containing 10 ppt salt (sea salt or NaCl) for 2-5 hours daily. Salt content should be reduced gradually through the slow addition of fresh water. Prophylactic treatments are commonly used for parasitic and bacterial infections before the fish are stocked (recommended therapeutants are discussed in Chapter 13). Antibiotics are commonly used as prophylactic treatments, before and during transportation, but their use as a standard practice is not recommended due to risk of inducing drug resistance in known pathogens (Brown 1989).
The hauling medium that we recommend for striped bass is water with a total hardness of 100 ppm (CaCO3) or greater, to which salt has been added to bring the concentration to 10 ppt. Synthetic sea salt is preferred, but solar salt or non-iodized table salt may be substituted. Transportation of striped bass in soft water (<20 ppm CaCO3) is not recommended; however, if only soft water is available, 50-1,000 ppm calcium chloride (CaCl2) should be added to the transport medium unless a source of calcium is provided in the initial mix (see Chapter 8 for additional comments). Use of tricaine methanesulfonate (MS-222) is recommended to reduce activity and stress during transport (see Chapters 4 and 5 for additional comments on MS-222). MS-222 is approved by FDA for use on food fish, but requires a 21-day withdrawal period before treated fish are safe for human consumption (Schnick 1988). However, this anesthetic may not be the most desirable sedative for long-term exposure because its method of action induces stress in fish and may reduce oxygen consumption by asphyxiation rather than by lowering metabolism. (See Chapter 8 for additional comments.)
Dissolved oxygen during transport should be at saturation for the entire time the animals are held in the hauling tank. These levels are maintained by bubbling compressed oxygen or the gas from liquid oxygen through air diffusers, such as air stones, perforated tubing, or micropore tubing. Smaller bubbles yield more efficient oxygen transfer. Oxygenation of transport water should be started about 15 minutes before loading fish to ensure that sufficient oxygen is available. Surface agitators should only be used for larger (phase II) fish. Antifoam compounds may be used if desired; they are particularly beneficial when agitators are used for aeration. A complete change of water may be needed on hauls in excess of 8-10 hours to prevent ammonia and CO2 build-up. When temperature of the transport and receiving water differs, the fish should be tempered at a rate no greater than 8°F per hour. Salt and any other additives must also be replaced if water in transport tanks is exchanged with fresh water.
Hauling temperatures ideally should be kept below 68°F on long hauls and below 75ºF on short hauls for phase I fish. For phase II fingerlings, temperature should be kept below 68°F. Ice may be used to lower temperature; however, non-chlorinated ice is difficult to obtain while traveling. To protect fish from chlorine, ice made from chlorinated water can be sealed in plastic bags and floated in the transport water. Alternatively, a dechlorinating agent (e.g., sodium thiosulfate) can be added to the water.
Hauling Density and Tanks
Striped bass 1-2 inches long (500-1,000 fish/pound) may be hauled at 0.5 pounds per gallon for 1-4 hours, 0.3 pounds per gallon for 4-8 hours, or at 0.25 pounds per gallon for over 8 hours (Geiger and Parker 1985). Fingerlings averaging 5 per pound have been transported at rates of 1.5 pounds per gallon for 10 hours and 0.75 pounds per gallon for 15 hours. Many excellent insulated fiberglass or aluminum hauling tanks are commercially available. These tanks usually come complete with plumbing for oxygen distribution and provisions for mounting agitators. Continuous temperature and oxygen monitors visible to the driver are desirable. In the absence of oxygen monitoring equipment, an ammeter visible to the driver should be installed to verify the operation of each agitator. When small fingerlings are being hauled, the agitator wells should be covered with fine-mesh screen to prevent entrainment of fish.
After arrival at the stocking site, fish must be acclimated to the temperature and quality of the receiving water before release. Receiving water should be pumped through the hauling tank for at least 1 hour (longer if the temperature difference is more than 2°F). Delivery of the fish through a quick-release valve is less stressful than removal with dip nets. If quick-release valves are used, the fish should be delivered through a tube submerged in the water, rather than allowed to free-fall to the surface. If fish must be netted, a soft, knotless, untreated net should be used; remove only a small number of fish at a time. Fish delivered to another hatchery or holding area should be slightly sedated with MS-222 before transfer by dipping, but stocking of anesthetized fish in the wild is not recommended due to the risk of predation on disoriented fish.
Stocking locations should be selected to allow fish the best chance for survival. Shoreline stocking into turbid water or cover of some type may reduce the risk of predation. Mid-reservoir stocking is less desirable if the fish must be handled an additional time. Also, the probable lack of adequate food and cover in mid-reservoir locations would require fish to move a considerable distance to locate food, thus increasing the possibility of predation.
Some agencies use nursery ponds constructed adjacent to, and connected with, a reservoir (see Chapter 8). Fry may be stocked directly into nursery ponds, reared to phase I and released. Phase I fish reared elsewhere may be stocked into a nursery pond and allowed time for recovery from handling before release.
Stress and Its Complications
Intensively cultured fish may be continuously or intermittently exposed to stressful conditions (Pickering 1981). Understanding the role of stress, its associated complications, and methods of avoiding stressful conditions, is imperative for successful culture.
The same basic physiological changes occur in men, mice, fish, and other vertebrates that are exposed to situations perceived as life-threatening. These changes may include a quickening of pulse, an increase in respiration rate, widening of the eyes, contraction of muscles, alertness for danger, and development of the general adaptation syndrome described by Selye (1976).
Two hormones released almost immediately when a life-threatening situation is perceived are the catecholamines, adrenaline and noradrenaline (also known as epinephrine and norepinephrine). These hormones act very quickly to prepare an animal to fight or flee. The release of a second set of hormones, the corticosteroids, helps to sustain the fight-or-flight reaction.
The corticosteroids include three hormones: cortisone, cortisol, and corticosterone. Cortisol seems to be the most common and important of the corticosteroids in fish, but cortisone and corticosterone may be important in some species (Chester Jones et al.1969). These hormones in fish are identical with those in humans and produce similar physiological changes.
Short-term elevations of corticosteroids may allow fish to flee from stressful life-threatening situations and thus may be beneficial. However, long-term elevation of cortisol and cortisone can be detrimental to the health of fish and result in reduced growth or actual weight loss (Davis et al. 1985), suppression of the immune system resulting in increased susceptibility to disease (Klinger et al.1983; Pickering 1984), and disruption of osmotic balance or the ability to regulate electrolyte and water balance within the body (Mazeaud et al. 1977; Nikinmaa et al. 1983; Tomasso et al. 1980).
General characteristics of the blood can be measured to indicate the level of stress in an animal. Hormone levels (catecholamines and corticosteroids), electrolytes (sodium, potassium, and chloride ions), and metabolic by-products (glucose and lactate) can be evaluated biochemically. The cellular components of blood (red and white blood cells) can be analyzed histologically. Any deviations from normal resting levels may indicate that fish have been stressed.
Water Qualify and Fish Density
Fish produced in hatcheries are commonly cultured at densities exceeding those found in the wild. As density increases, water quality usually decreases, and fish must adapt to less than optimum conditions. Crowding, low DO, high ammonia, and other factors can stress cultured fish. Environmental conditions suitable for survival of fish may not be adequate for growth, and even more exacting environmental conditions may be required to ensure successful reproduction (Parker and Davis 1981).
The stress response of fish is influenced by the rate at which water quality deteriorates. Tomasso et al. (1981) found that rapid deterioration of water quality was much more stressful than gradual changes, which allow fish more time to adapt.
Aquaculturists have long known that survival of fish handled and transported is typically higher in winter than in summer. These differences may be partly attributed to differences in oxygen saturation of cold and warm water; however, in some species, temperature (Figure 12.4) strongly affects the resting level and secretion rate of corticosteroids (Strange 1980; Davis et al.1984). The handling and transportation of fish at reduced water temperatures has been recommended (McCraren and Millard 1978), but this measure may reduce stress more effectively in some species than in others.
Handling, grading, and transporting are some of the most stressful procedures to which fish are exposed. Just as physicians and dentists treat patients with anesthetics to reduce pain in surgery and medicine, aquaculturists can use anesthetics to reduce stress, or the secondary effects of stress, in fish. The principal anesthetics used for fish are M~222, quinaldine, and etomidate. Corticosteroids increased significantly when striped bass were exposed to 25 ppm MS 222, 2.5 ppm quinaldine, or 10 ppt NaCl, but not when they were exposed to 0.1 ppm etomidate, either alone or in combination with 10 ppt salt (Davis et al. 1982). When striped bass were stressed by confinement in a net for 10 minutes, corticosteroid increases were reduced by quinaldine and MS-222, but not as effectively as by etomidate (Figure 12.5; also see Davis et al. 1982). Tomasso et al. (1980) found it beneficial to treat hybrid striped bass with 50 ppm MS-222 for 15 minutes before handling, and then to transport them in water containing 25 ppm M~222 and 10 ppt of salt. Etomidate is an experimental drug that is not available to aquaculturists in the United States. Other experimental anesthetics may be equally effective, but will require additional evaluation and clearance by regulatory authorities before they become available for fish culture.
Osmotic balance, the maintenance of salt (electrolytes) and water balance within the body of fish, is easily disrupted by stress. In general, fish blood has roughly one-third the salt content of sea water. In fresh water, fish must continuously expend energy to keep salt in the body and to excrete excess water. Conversely, fish in sea water must take water into the body and excrete excess salts.
Transportation of hybrid striped bass for 2 hours elevated corticosteroids almost immediately, and resulted in the delayed loss of chloride ions from blood plasma 72 hours after fish were released into fresh water (Tomasso et al. 1980). Transportation in anesthetic (25 ppm MS-222) and salt (10 ppt) did not prevent the delayed loss of electrolytes. However, when fish were placed in 10 ppt salt after transport and allowed to recover for 3 days, electrolytes remained within the normal range.
These data suggest that the detrimental effects of stress can be reduced by adjusting the salt concentration of the transport and recovery water to levels similar to that in fish blood. A reduction in the differential between the internal and external environment reduces the energy required by fish to maintain osmotic balance and helps to ameliorate osmotic disturbances caused by stress.
|Figure 12.4. Plasma-corticosteroid secretion profiles for channel catfish acclimated to one of several temperatures, then stressed by confinement. Each point represents a fish. (From Davis et al. 1984).|
|Figure 12.5. Plasma-corticosteroid concentrations (mean + SE) in undisturbed striped bass controls (C), fish exposed for 15 minutes to sedating concentrations of anesthetics and to salt (upper panel), and in fish exposed to anesthetics and salt and confined for 10 minutes in a dipnet (lower panel). F= freshwater controls; T= tricaine methanesulfonate; Q= quinaldine; E= etomidate; and S= salt. Number of fish is shown at the base of each column. Similar letters in each panel represent similar subsets at the 0.05 level by Duncans multiple range test. Data represented in the upper and lower panels were not compared; undisturbed controls for reference only. (From Davis et al. 1982).|
A single acute stressor may trigger a series of physiological changes that persist for several days or weeks after the initial stress. After the initial recovery phase, a secondary rise in corticosteroids has been reported in channel catfish (Tomasso et al. 1981), hybrid striped bass (Tomasso et al. 1980), and largemouth bass (Carmichael 1984). This secondary increase resulted from osmotic failure, as indicated by the loss of electrolytes that followed the initial stress. Depletion of energy reserves (Nikinmaa et al. 1983), reduction in numbers of lymphocytes (Klinger et al. 1983; Pickering 1984), and reduction in granulocyte and thrombocytes (Klinger et al. 1983) probably increased the susceptibility of fish to secondary bacterial, parasitic, or fungal infections (see review by Wedemeyer 1970). Fish may survive one stressful event only to die when exposed to a second stressor.
Aquaculturists may not be able to prevent fish from being stressed, but they can reduce the effects. In addition to the factors previously mentioned (species, temperature, water quality, handling procedures, etc.), other stress factors are sexual maturity, sex, and spawning (Mazeaud et al. 1977); time of year and time of day (Lamba et al. 1983; Pickering and Pottinger 1983); environmental contaminants (Passing 1984); and degree of domestication (Mazeaud et al. 1977). Factors that are stressful to warmwater fish may not be stressful to coldwater fish, and response to stressors in saltwater fish may differ from that in freshwater fish. Awareness of the stress response in fish and the factors influencing stress should allow culturists to reduce the effects. Fish loss due to stress can be reduced by several methods: (l) maintaining optimum environmental conditions; (2) handling and transporting fish at the lowest practicable temperature; (3) anesthetizing fish before and during handling; (4) handling and transporting freshwater fish at approximately isosmotic salinity (10 ppt salt); and (5) allowing fish to recover from one stressful situation before exposing them to another.
Controlled Spawning of Domesticated Brood Stock
Federal, state, and private hatcheries currently depend on collection and use of wild brood stock to produce striped bass and striped bass hybrids (Harrell 1984a). Such reliance results in some uncertainty regarding exact timing of natural spawning runs, and whether sufficient brood stock will be available to support desired production levels. Climatic conditions and other natural factors, as well as discharges from man-made water control structures, often determine the outcome from various hatcheries. Historically, use of wild brood fish has generally been satisfactory, but this may no longer be true as demand for fish for stocking purposes has increased substantially (Stevens 1984)as has demand by commercial culturists. Concomitantly, recreational fishing pressure for potential brood stock is high (IJ.S. Department of the Interior 1982). Dependence on wild brood stock to support hatchery operations places private aquaculturists and, to a lesser degree, state and federal personnel in the position of competing with the public sector for "their" fish. Indeed, this perception may limit commercial development (Smith 1988). In an examination of the potential of striped bass and striped bass hybrids for aquaculture, the Joint Subcommittee on Aquaculture of the Federal Coordinating Council on Science, Engineering and Technology (1983) stated: 'The major constraint to private U.S. striped bass aquaculture is nonavailability of seed stock." Although many state hatcheries produce striped bass and hybrids, such hatcheries are typically unwilling to provide fish for commercial aquaculture (American Fisheries Society 1983). However, recent work suggests that controlled spawning of domesticated brood stock will offer hatchery operators an alternate source of fry (Smith and Jenkins 1984,1988a; Smith 1987).
Striped bass can be reared from small phase I fingerlings to adults in indoor or outdoor tank systems. Cylindrical tanks, 20 feet in diameter and 5 feet deep, are preferred, although smaller tanks (12 feet in diameter x 3 feet deep) have been successfully used (Smith and Jenkins 1985). Fish can be reared in fresh or salt water, but water quality must be monitored regularly and maintained within normal culture limits. Recirculated water has been successfully used, although a flow-through water source is also acceptable and may be preferable.
Brood stock nutrition is important. Commercial trout diets have been used successfully as the main source of nutrition, supplemented with natural foods such as squid and fish. Striped bass should be fed several times daily when they are young, then feeding can be reduced to 1 or 2 times daily after they attain adult size. A pellet larger than the standard 0.25inch trout grower pellet may be required for large adult striped bass to ensure that they receive sufficient feed. During culture of brood stock, handling should be minimized to reduce stress. Fish can be safely handled using anesthetics (e.g., MS-222), and handling appears less stressful to fish held in salt water than to those kept in fresh water. Periodically, parasitic and bacterial infections will occur, but normal prophylactic treatments will control most problems encountered (see Chapter 13).
Cultured striped bass can grow to nearly 1.3 pounds at age 1, 5.7 pounds at age 3, 9 pounds at age 4, and 14.3 pounds at age 5 (Figure 12.6). After juveniles have adjusted to pelleted feed, mortality should be insignificant, provided water quality and disease problems are carefully controlled. Wild white bass (M. chrysops) can also be successfully reared under captive conditions (Figure 12.6).
Age and Size at Maturity
Smith and Jenkins (1987) found that under culture conditions, about 22% of the males matured by age 2 and most were mature by age 3 (Table 12.1). In contrast, females did not begin to mature until age 3, when only 16% were mature; at age 4, 59% were mature.
Cultured striped bass males and females have been artificially conditioned to spawn out-of-season by manipulating photoperiod and temperature (Smith and Jenkins 1987). Natural conditions were simulated by using double tube fluorescent lights (three cool-white 3tW tubes per tank) and water chillers. The annual spawning cycle was accelerated 2-3 months, and fish were spawned in January and February (Figure 12.7). Similarly, striped bass males and white bass males and females can readily be brought into spawning condition in outdoor culture tanks exposed to natural temperature and photoperiod conditions.
|Figure 12.6 Growth of cultured striped bass and wild white bass in tanks.|
|Figure 12.7 Natural cycles of photoperiod and temperature in South Carolina and the controlled cycle used to condition striped bass to spawn.|
Maturation characteristics of striped bass reared as
brood stock under tank culture conditions (Smith 1988).
aFish from which milt could be expressed.
bFish with eggs more than 700 µm in diameter.
cBased on number of males and females at 5 years of age; however, 19% of the fish were not mature at 5 years of age.
In general, procedures for identification of mature fish, use of hormones, spawning, and egg incubation are similar to those used with wild fish (see Chapters 5 and 6). Extra care is required during handling to reduce stress and physical damage because the brood fish will be reused over several years. Typically, fish should be anesthetized before handling and treated prophylactically during and after handling to control disease. Brood fish are individually tagged for long-term identification so that the background of each fish is known during each spawning season (Smith and Jenkins 1987).
Controlled spawning techniques have been demonstrated on a research scale. Work now underway to develop these techniques for use in state, federal, and private hatcheries. Control over the spawning cycle has several advantages: (1) reduced dependence on wild brood fish; (2) year-round production of fry; (3) increased fry production; (4) accelerated domestication of stocks; and (5) development of selective breeding programs. For the geneticist, utilization of controlled spawning techniques would provide increased opportunities for genetic research and development.
Cryopreservation of Sperm
Cryopreservation of spermatozoa has become an important animal husbandry technique for mammals and birds. Extensive work has been done with various fish species with varying degrees of success. Yet, the technology for cryopreserving fish gametes has lagged behind that in other animal husbandry fields. Currently, spermatozoa from a variety of fish species can be successfully cryopreserved, although fertilization percentages are normally less than when fresh semen is used. Methods are sufficiently advanced for some species to be used successfully for experimental breeding programs, but are not yet adequate for normal use in production facilities (Kerby 19&3; Kerby and Bodolus 1988). Recent progress, particularly in the area of extender development and freezing techniques, promises more practical use in the future.
Striped bass brood stock are usually collected on or near the spawning grounds; still, hatcheries sometimes have difficulty in obtaining an adequate supply of ripe males (Bayless 1972; Bonn et al. 1976). This paucity of males is particularly evident late in the spawning season when ripe females may be abundant. Additionally, because the spawning of white bass typically peaks earlier than that of striped bass, collection of male white bass for use in production of hybrids is often difficult.
These difficulties provided the impetus for efforts to cryopreserve striped bass spermatozoa (Figure 12.8) (Kerby 1983,1984; Kerby et al. 1985; Kerby and Bodolus 1988).
(insert Figure 12.8 here)
Figure 12.8. Scanning electron micrograph of striped bass spermatozoan shoveling the relative positions and dimensions of the head (H), mitochondrial collar (MC), and flagellum (F). Bar = 2 µm (from Kerby and Bodolus 1988).
Although efforts were successful, and several million larvae were produced over a period of 4 years, methods are not yet sufficiently refined to be used reliably in production hatcheries. The techniques can, however, support experimental work where consistently large numbers of progeny are not required. The best results have been obtained using controlled-rate freezing systems (Linde BF4/6 or CRYO-MED Model 900 systems) utilizing liquid nitrogen (LN2) for the freezing process. We next summarize previous results and preferred methods for those interested in applying the methods in their work.
Initial efforts (Kerby 1983) were designed to identify suitable extenders and cryoprotectants for striped bass by evaluating various extenders and cryoprotectants developed for other species. An extender is defined as a solution of salts, sometimes including organic compounds, which helps maintain the viability of cells during cryopreservation, whereas a cryoprotectant is defined as an organic compound which protects the viability of cells during the freezing and thawing process (Ott 1975). These trials normally involved small amounts (1 mL) of extended semen (0.2 mL of semen and 0.8 mL of freezing mediumextender plus cryoprotectant). Different protocols, including various freezing rates, were also evaluated. Freshly ovulated striped bass eggs (about 300 per trial) were challenged with thawed cryopreserved sperm to determine fertilization capacity.
Results demonstrated that striped bass sperm could be successfully cryopreserved by using several different extenders combined with dimethyl sulfoxide (DMSO) as the cryoprotectant. The best overall results were obtained with an extender (OH-189) developed by Ott (1975) for salmonid sperm (Table 12.2). Dimethylsulfoxide, added at an optimum concentration of 5%, was the only cryoprotectant tested that was successful. The freezing medium was mixed in a 1:4 sperm:medium (volume:volume) ratio and frozen in 1 mL aliquots in 2 mL A/S NUNC polypropylene cryotubes obtained from Union Carbide Incorporated. The best mean fertilization percentage obtained with cryopreserved sperm was 23.6 (range, 0-87%). Other extenders producing good results were OH-134, OH-235, and OH-275 (Table 12.2). Mean freezing rates between 5 and 20°C per minute were more effective than slower rates (Kerby 1983).
Although some fertilization could be consistently obtained with cryopreserved sperm, and fertilization capacity could be retained for at least 4 years, fertilization percentages of individual tests varied considerably from one trial to the next. Inconsistency was partially due to variation in gamete quality among females and perhaps males. Other important variables that may affect fertilization success include small variations in technique and repeatability from one trial to the next in both the freeze-thaw and the fertilization processes. Fertilization percentages from cryopreserved sperm were seldom comparable with those of fresh sperm. Results of 48 individual tests with extender OH-189 (Table 12.3) showed the variability obtained at different freezing rates, with different sample lots, and with different females.
Female striped bass normally produce from 1-4 quarts (1-4 L) of eggs, depending on the size of the fish (1 quart per 10 pounds or 1 L per 4.5 kg of body weight as a rule-of-thumb), and
|Table 12.2. Chemical composition of extenders used to cryopreserve striped bass sperm (grams of solute/liter total solution). Add 5% dimethyl sulfoxide (volume/volume) as the cryoprotectant.|
|Table 12.3. Percent fertilization (mean :l SE; ranges in parentheses) of striped bass ova challenged with sperm frozen and stored in liquid nitrogen (-196°C). Extender was OH-189, containing 5% DMS ,. Sperm:medium ratio was 1:4. SAC.|
hatchery managers usually use semen from at least two males, with a total volume of semen ranging from 10-30 mL per batch of eggs. Kerby (1983) noted that cryopreserved sperm cannot be expected to have a fertilization capacity equivalent to fresh sperm. Semen is diluted 1:4 with the protective extender, so large quantities of extended semen must be preserved and stored.
Storage containers for freezing large volumes of semen are not commercially available, so for production studies, polyethylene, polypropylene, stainless steel, and teflon containers were fabricated and evaluated (Kerby and Bodolus 1988). Because acceptable results were obtained in initial experiments with 2-mL and 5-mL polypropylene cryotubes (Kerby 1983), the custom-fabricated tubes had inside diameters and a configuration similar to the original vials. Wall thickness and heat transfer properties varied among tubes constructed of various materials. The tubing was cut in 11.8 inch lengths and sealed at one end by tightly inserting a rubber stopper. The volume of the fabricated containers was approximately 20 mL.
Semen was extended in OH-189 containing 5% DMSO at temperatures (15-18°C) corresponding to hatchery water temperatures. Each sample tube was filled with 18 mL of extended semen and immediately frozen in a CRYO-MED controlled-rate freezer at rates ranging from 5-15°C per minute, or the samples were placed directly in the freezing chamber pre-chilled to temperatures of -60 or -100°C. All samples were lowered to a temperature of -60 to -80°C in the freezer before being transferred to LN2 for storage.
Samples of cryopreserved sperm were rapidly thawed in a 50-60°C water bath and tested with fresh eggs to determine fertilization rates. To simulate production conditions, 200 mL egg aliquots (ca. 200,000 eggs) were fertilized by adding nearly thawed semen from three or four tubes (54 or 72 mL), fresh hatchery water, and stirring gently. Eggs were then incubated in modified MacDonald hatching jars (Kerby and Bodolus 1988).
Fertilization percentages for sperm preserved in "production" quantities ranged from 0- 61.5%. In general, a freezing rate of 5°C per minute provided the best overall results; mean fertilization percentages were 16.2 for the polyethylene tubes, and 14.2% for the polypropylene tubes. Results often ranged between 10 and 40%. Trials in which semen was rapidly frozen by placing it in the chamber pre-chilled to -60 or -100°C usually yielded fertilization percentages between 9 and 35%. Trials with stainless steel and teflon produced generally poor fertilization percentages.
Growth and Survival Experiments
Striped bass eggs fertilized with frozen-thawed sperm resulted in apparently healthy larvae. However, it was desirable to determine if growth and survival of "cryogenic" larvae were comparable with those of "natural" larvae obtained from eggs fertilized with fresh sperm.
Kerby et al. (1985) reported that in two pond experiments designed to assess relative growth and survival between natural and cryogenic larvae to the phase I fingerling stage, there were no significant differences between harvest numbers, weights, or survival. Although larger numbers of natural fish were produced, cryogenic fishprobably because of their lower relative densities in the pondswere significantly bigger than natural fish.
Due to the variability inherent in pond culture experiments, it is difficult to adequately assess the lack of significant differences unless many replicates are used. Between-pond variability was so large that it could have masked significant differences between treatments. Based on previous experience, Kerby et al. (1985) concluded that production and size differences were probably caused by environmental differences in culture conditions between ponds, and not to the experimental treatments. Results from the study clearly indicated that striped bass produced by cryopreservation techniques were normal, and they appeared as viable as fish produced with fresh sperm (Kerby et al. 1985).
There are a number of technique "tricks" in the cryopreservation process that can be learned only by trial-and-error or from experience. We here attempt to provide some of those techniques.
Use only freshly-collected males that produce free-flowing milt when slight pressure is applied to the abdomen. Wipe the abdomen with a damp towel to avoid contamination of semen with water or mucus from the body. Strip into a cool, dry container. Discard samples contaminated with blood, contain large quantities of urine, or are highly viscous and do not mix easily with water (forms clumps).
Before freezing, test for sperm motility by placing a very small drop of semen on a glass slide. Activate the spermatozoa with a drop of hatchery water and observe them under oil immersion with a compound microscope (magnification 1,000X). This procedure must occur rapidly, as motility lasts only 30 to 60 seconds after activation in fresh water. Use only semen containing highly motile spermatozoa. Keep the collected semen cool (15-18°C), process it, and initiate the freezing procedure immediately. It is best to process the semen at a room temperature of 15°C or less.
Processing semen. Extenders should be refrigerated (38°F). Prepare a freezing medium by adding 5% DMSO to the extender. Add the semen to the freezing medium (1 part semen:4 parts medium). Extender OH-189 is preferred (Table 12.2).
The semen-medium mixture can be prepared several ways. In one, the cap (or needle) from an appropriately-sized syringe is removed and the semen is drawn into the syringe. Then the appropriate amount of semen is added to a pre-measured volume of medium. Alternatively, remove the plunger from the syringe (with the hub capped) and pour the desired amount of semen into the barrel of the syringe. Replace the plunger, remove the cap, and adjust the volume of semen to coincide with the prepared volume of medium. Add to the freezing medium and mix well. After mixing, an automatic pipes system, such as a Repipet Jr. ® (available from scientific supply companies) is an excellent way to rapidly and accurately dispense the semen-medium mixture to the freezing containers. At present, probably the best freezing containers are the 5-mL NUNC cryotubes. If larger containers are desired, they can be fabricated from polyethylene or polypropylene tubing as previously discussed.
Immediately after mixing, distribute the extended semen to the freezing containers, place them in the freezing chamber, and initiate the freezing process. If 5-mL cryotubes are used, they should be attached to aluminum holding canes and placed in a freezing rack. If longer containers are used, they can be placed in the rack individually. A freezing rate between 9 and 18°F (5 and 10°C) per minute is recommended. Individual containers should be labeled to identify each frozen batch. If later experience indicates that a particular batch is of poor quality, it can be discarded.
As an alternative to controlled-rate freezing equipment, which is very expensive, containers of extended semen can be immersed in an ice chest containing crushed dry ice (frozen CO2) or suspended in the vapor above liquid nitrogen in a LN2 refrigerator. Based on experience, ambient temperature in the freezing chamber should be from -60 to -100°C.
The temperature inside one of the containers of semen should be monitored by means of a thermocouple placed within the container. Temperature of the frozen sperm should be at least -60ºC before removal from the freezing chamber. It is very important that the containers of frozen sperm be transferred as rapidly as possible to the storage container containing LN2. Even short-term contact with ambient room temperatures can cause a rapid elevation of sample temperatures. The frozen sperm should be stored in LN2 until ready for use.
Thawing frozen semen. Preparations for use of stored sperm should be made before eggs are taken from the female. A water bath containing about 55°C water is necessary for thawing frozen semen. The containers of sperm to be used should be isolated in the LN2 storage container for quick access. About 40 to 50 mL of extended semen should be used per 100 mL of freshly ovulated eggs. Thawing should begin shortly before (or at the same time) the female is spawned. The containers of stored sperm should be agitated vigorously in the water bath until almost thawed (slush phase). At this stage, pour the semi-thawed semen into a beaker, keeping it separate from the eggs. Be sure not to completely thaw the semen before removing it from the water bath as over-heating can easily occur and kill the spermatozoa. When the semen has been thawed and concentrated in the beaker, add it to the eggs simultaneously with hatchery water. Gently stir the mixture of eggs and semen for at least 1 minute. Leave the eggs in the semen-water mixture for 3 to 5 minutes, decant the water, and incubate the eggs according to normal practice.
The whole process of thawing and fertilization should be done as rapidly as possible. Ideally, thawed sperm should reach the "slush" phase just as stripping of the female is completed.
Accurate fertilization percentages can easily be determined 3 to 4 hours following fertilization. It is recommended that at least one group of eggs be fertilized with fresh sperm as a control if possible to provide some assessment of the quality of each batch of stored sperm.
Reprinted from R.M. Harrell, J.H. Kerby, and R.V. Minton (eds.) Culture and propagation of striped bass and its hybrids. American Fisheries Society, Bethesda, Maryland, 1990.
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