Dedicated To The Tribal Aquaculture Program
 |
June 1994 - Volume 8 |
http://www.fws.gov/midwest/ashland/mtanhome.html
| Administrative
Coordinator:
Frank G. Stone
(715-682-6185) Ext.12 |
Edited By: Elizabeth W. Greiff
(715-349-2195) |
Topics Of Interest:
Application Process for an Environmental Protection Agency
Permit To Use MS-222
By: Mike Donofrio, Keweenaw Bay Tribal Biologist, Keweenaw Bay Indian
Community
Tricaine Methane Sulfonate (MS-222) is used as a fish anesthetic in hatcheries. We will use MS-222 to anesthetize our lake trout yearlings for the fin clipping and microtagging process. The Food and Drug Administration has approved MS-222 for use on food fishes. MS-222 is restricted to a level of 15 to 330 PPM and fish must not be used, for food purposes, within 21 days following the anesthetic treatment. However, you must obtain a permit from the Environmental Protection Agency (EPA) to discharge this chemical with your effluent. EPA permits are issued through the regional office in Chicago, IL. We sent a letter describing the product, its proposed use, and product label to: Claudia Johnson-Schultz, Indian Programs Coordinator, Water Division, EPA W-15J, 77 West Jackson Blvd. Chicago, IL 60604-3590. We also sent a copy of the request to Mary Ann Starus, Michigan Tribal Environmental Liaison. There is also a Tribal Environmental Liaison for Minnesota and Wisconsin. Ms. Starus was instrumental in making direct contact with a telephone call to the Regional office.
You shouldn't expect to receive a timely reply from EPA since they are understaffed. I recommend you apply for this permit a year in advance and use your Tribal Environmental Liaison to expedite the permit process.
Proper Storage and Handling of Fish
Feeds
By: Terrence Ott , U.S. Fish and Wildlife Service, La Crosse Fish Health
Center, La Crosse, WI 608-781-6238
One of the main ingredients in raising and producing healthy fish for the resource or as table fare is the quality of feeds you put into your fish. Quality fish feeds are just as important at your hatchery as the quality of water you use to raise your fish in.
Without providing your fish a properly balanced diet, growth will be abnormal, food conversion will be low, and health of your fish will be at stake. Malnourished fish cannot remain healthy and grow properly to produce the next generation of fish with an improper diet.
Feeding your fish on a daily basis can become and often does a routine practice at your hatchery. Routine experiences can lead to ignoring or skipping many common rules in the proper storage, handling, and feeding of your fish. Proper handling and storage procedures must be followed to protect the quality of the feed you purchase from the manufacturer. If proper storage conditions are not met, fish feed with their high levels of proteins, fats, and vitamins will rapidly spoil. These high levels must be maintained in order to meet the nutritional requirements your fish need to grow and reproduce.
The formulated fish feeds you use at your hatchery are soft and fragile and care must be expended in their handling to reduce breakage of the pellets. Broken pellets result in dust, and small particles which feed manufacturers describe as "fines" in your feeding operation. Fines that exceed 3.0 percent are not consumed by your fish and can increase feed conversions, result in poor water quality, create suitable habitat for fish pathogens, and cause environmental gill disease. To eliminate or keep to a minimum the fines in your feeding operation follow these four easy steps;
Don't walk or stand on bags of feed.
Don't throw or drop bags of feed onto hard surfaces.
Don't forcibly stack bags of feed.
Don't stack bags of feed over 10 high.
Proper storage of fish feeds at your hatchery depends if the feed you receive is dry, semi-moist, or moist. Dry feeds or semi-moist feeds do not require refrigeration. Ideal conditions for storing these types of feeds involve stacking the feed bags on wooden pellets so the bags are 3 to 4 inches off the floor. Keep the stacks in a cool, dry storage area where relative humidity is below 75 percent. Humid conditions can lead to insect infestations and moldy feed. It is also important to set feed stacks apart in order to provide air circulation and rodent control. Moist pellets should be stacked similar and stored in a freezer at temperatures below 0EF, until you are ready to feed your fish. Then thaw only the required amount to feed your fish at that time.
Never leave a pail of feed sitting out next to a raceway during warm days. Excessive heat can quickly turn that pail of quality feed into a costly supply of fertilizer for your garden by a process called, "oxidation". Oxidation will destroy the vitamins in the feed and turn the feed rancid. Rancid feed can be toxic to your fish, and will produce an undesirable flavor in fish eating the feed. Off-flavor of the feed can also occur due to the spoilage of the proteins and fats in the feed. Rancid feed can taste poorly to the fish, making them eat less and waste more. This can result in an accumulation of feed material on the bottom of the raceway, providing excellent habitat for fish pathogens to hide in and multiply.
Recommended maximum storage time for feeds is 90 days from the time of manufacturer, not from the time the feed arrives at your hatchery. Bags of feed which have exceeded the recommended shelf life or do not have a readable manufacturing date stamped on them should not be fed, or even purchased. Also any feed that appears abnormal (excessive fines, moldy, off-colors and odors) should not be given to your fish.
If these rules for the proper handling and storage of fish feeds are not met at your hatchery, you could be jeopardizing the health of your fish and your fishery program.
Great
Lakes Fish Disease Control Policy Special Publication 93-1
By: Great Lakes, Fishery Commission, 2100 Commonwealth Blvd. Suite 209,
Ann Arbor, MI 48105, 313-741-2077
The MTAN sincerely believes that every Tribal hatchery should adopt an aggressive fish disease prevention and monitoring program. This is especially important for all salmonid fish hatchery facilities. The MTAN also encourages you to become actively involved in the Great Lakes Fish Disease Control Committee. If you are not aware of the goals of the Committee, the following information may help you to better understand the management role they have in the Great Lakes fishery program.
The Great Lakes Fish Disease Control Committee (Committee) was established by the Great Lakes Fishery Commission (GLFC) in 1973 to recommend measures to protect the health of cultured and wild-fish populations in the Great Lakes basin. The Committee is comprised of representatives from state, provincial, and federal agencies involved with Great Lakes fishes and from private aquacultural interests. Decisions are made by a consensus of the membership. In 1985, the Committee developed a "Model Program" for controlling fish diseases in the basin. This Model Program was subsequently adopted as a policy of the GLFC.
Increasing national and international interest in importation of salmonid fishes for fisheries management and aquacultural purposes, and the damage caused by disease introductions indicated that expanded guidelines were needed to protect the health of salmonid fish stocks within the Great Lakes basin. Consequently, the Committee recommended at first a ban on importation's of salmonid fishes from regions where diseases were enzootic. For example, importation's were banned from the U.S. and Canada west of the Continental Divide, areas where infectious hematopoietic necrosis virus (IHN) and the parasite Ceratomyxa shasta, neither of which are known to occur in the Great Lakes basin, are found.
Fish disease control in the Great Lakes basin is the responsibility of those agencies that manage the fisheries resources. The Model Program as established by the Committee, sets forth the essential requirements for the prevention and control of serious infectious diseases, and includes a system for inspecting and classifying fish hatcheries as well as the technical procedures to be used during these evaluations.
The Committee does not seek fish-disease control authority.
The recommendations they propose are provided as an aid to the member agencies in the development of legislation and regulations. The Committee seeks the advice and counsel of these agencies in the continuing development of fish-disease control programs to assure that such programs are in the best interests of the fishery resources of the Great Lakes.
CONTROL POLICY
Efficient propagation of fish may be severely affected by the occurrence of fish diseases. Disease outbreaks have caused serious losses in fish hatcheries and potential exists for such losses in feral Great Lakes fish populations. Disease problems have resulted in reduced survival of stocked fish, production cost increases of 20%-30% or more, significant losses of fish to the public, and diminished economic returns to Great Lakes communities.
The policy of the Great Lakes FDCC encourages each agency to work toward the control of fish diseases in the Great Lakes basin by:
Developing legislative authority and regulations to allow control and possible eradication of fish diseases.
Preventing the release of seriously infected fish.
Discouraging the rearing of diseased fish.
Preventing the importation into the Great Lakes basin of disease infected fish.
Preventing the transfer within the Great Lakes basin of fish infected with restricted diseases.
Eradicating fish diseases, where practicable.
The Great Lakes FDCC will strive to coordinate the fish disease control program of the agencies. To this end, the FDCC endorses and supports the Great Lakes Fish Disease Control Policy and Model Program as a guide for agency program development.
An on Site Mini
Hatchery System for Walleye
By: Lee Newman, U.S. Fish and Wildlife Service, Ashland Fishery Resources
Office Ashland, WI 715-682-6185
The Grand Portage Natural Resources Department and the Ashland Fishery Resources Office identified a problem limiting the walleye fishery on the Pigeon River. That problem was that walleye recruitment was very erratic and limited, probably due to poor hatch and rearing characteristics of the river.
Stocking hatchery fish from outside sources was considered but rejected because of concerns about disease introductions through transported fish and because of concerns for damaging the genetic makeup of the native stock. Assuming that the ideal system would be to use eggs from native Pigeon River walleye and to rear them on site, we tested whether this could be done economically.
During assessment work on the spawning run in 1993, we collected and fertilized about three quarts of walleye eggs on the Pigeon River. We placed the eggs in a Big Redd in the basement of the Grand Portage Tribal Office for development. An estimated total of 100,000 fry were hatched. About 20,000 of the fry were immediately stocked in the Pigeon River and the remaining 80,000 were introduced into a small natural pond for rearing.
The fry were allowed to feed on natural food in the pond for about 5 weeks. At the end of that period the pond was allowed to empty directly into the Pigeon River. No attempt was made to count the number of fry produced in the pond, but many 1.0 to 1.4 inch fry were observed as the pond was emptied. We suspect that survival was good.
Total cost for materials for this project was very low, the Big Redd was borrowed and other materials needed cost about $50. Approximately 55 man-hours of labor were required. Operation of the Big Redd requires only a dependable supply of well water and 110 volt electrical power.
It appears at this point that the on site, mini-hatchery, might prove to be both an economical system for producing small numbers of walleye and, a system which could provide important benefits in terms of eliminating some of the problems associated with stocking transported fish.
In 1994, Grand Portage will construct a wetlands project that will incorporate a drainable natural pond for rearing walleye and an intensive culture pond. These ponds and the mini hatchery system should produce all of the walleye needed for stocking on reservation.
Common-Sense Rules
for Research Recordkeeping
By: Jean Lang, University-Industry Research Program, University of
Wisconsin-Madison - originally printed in Touchstone, August 1990
If one thing can be guaranteed about cool water fish culture, it's that the fish culture practices used during one season will not always guarantee the same results the very next year. Despite how hard you work at it, or how consistent you try to maintain your fish culture practices, the nature of the business will often give you varying degrees of success. One of the best solutions to this problem is to record everything you do during the rearing process. This would include the quantities, frequency, duration, time of day and dates, which you conduct any fish culture practice. This "Diary" of your stations operations is especially important for newly activated fish hatcheries, or even for well established facilities which have recently incorporated a new rearing system. If detailed records are consistently maintained, the result will be a much easier time for your employees to remember all the fine details which will be needed to repeat the successes from the previous year. If problems occurred during the rearing season, the station diary will also enable you to review last years activities and attempt to determine how to avoid repeating previous mistakes.
The MTAN thought that the following article (edited to conserve space) would help our readers to better appreciate how to best document their stations fish culture activities. Although some of the material may not be relevant to your program, it does help to illustrate how detailed your field notes should be. This article was distributed by Peggy Ross, University-Industry Research Program, 1215 WARF Bldg., 610 Walnut St., University of Wisconsin-Madison, Madison WI. The MTAN would also like to express our thanks to Chuck McCuddy (BIA-Ashland WI) who brought this article to our attention
Any researcher's success in obtaining a patent these days may depend as much on how carefully he or she has kept a research notebook as on how original and patentable the discovery is.
University researchers generally do not feel the need to follow the strict notebook-keeping practices of their industrial colleagues. They should, nevertheless, take a few common-sense steps to protect themselves and their work. These practices can not only back up patent claims, but also insure against loss of valuable data, provide proof of fulfillment of contracts, and protect against allegations of conflict of interest or research fraud.
Standard Practice
The basic mechanics of keeping a notebook are familiar but bear repeating:
Research should be recorded in a bound notebook.
Every page should be numbered and dated. It is best to begin each new day's work on a new page.
Use pen, not pencil, and if possible, use the same pen throughout the day. This helps support the case that an entry was made all at the same time and not altered later.
If there is need to correct an old data entry, you may return to a previous page, but the change must be easily identified as separate from the original and must be dated.
Notebooks represent months of work. If for no other reason, they should be locked up at the end of the day in a fireproof cabinet.
A Record of Thoughts and Data
A good research notebook is a diary, not a memoir. It reports things as they happen, not as they are recalled. It is also like a diary in that it reports what you are thinking in addition to what you are doing. This means writing out an easy-to-read summary of your ideas, why you are planning a particular experiment, what materials and processes you will use, and what you hope to find out.
Too many notebooks are simply static data repositories with no hint of the evolving thoughts that are driving the research. A researcher can spend months on experiments that have the makings of a new discovery, or that reduce a concept to practice. But if that researcher's notebook fails to indicate any awareness of the potential, that person may have a hard time later answering challenges to the ownership of the ultimate discovery. If you don't state your mental constructs as you go along, the supporting evidence may not be there.
The theoretical description can be as simple as writing, "I think factor X is controlling response of Y by blocking position Z on the enzyme. I will begin testing that today with the following series of experiments." A week later, an entry might read, "Today I've had second thoughts about this theory for these reasons," and list them. The point is to state your ideas clearly enough that someone can pick up your notebook years later and understand what you were thinking as well as doing on a particular date.
Scientists who learn to regularly summarize their findings in words as well as numbers generally find that their notebooks become more useful to them as references. An articulate notebook helps to maintain workplace continuity when a long-term project is passed from one laboratory assistant to another, and it also makes the process of witnessing and archiving data much more efficient.
Reviewing and Archiving
To verify the legitimacy of data, especially for patent purposes, notebooks should be witnessed on a regular basis by an impartial party. The ideal arrangement occurs when two colleagues who understand, but do not participate in, each other's work, can review each other's work in confidence. Together reviewers go through their respective notebooks and discuss the recent work. If your reviewer finds a section unclear, you make an entry to that effect on the current day's page and proceed to clarify it in writing to the reviewer's satisfaction. Then the reviewer writes that he or she has read and discussed in confidence the work of pages X to Y on Z date, and signs off.
Electronic Notebooks
Some scientists today keep their research records on computer. These records may be validly archived by copying them onto disc or tape that is then held in a safe location. If electronic records are to be the only record, then it is advisable to have a "time stamping" program that automatically dates an entry into the system and would detect any late entries into already complete work. Whether in notebooks or on computer, records should be kept for as long as the researcher wants to be able to verify the legitimacy of his or her work. If the scientist has applied for a patent, the records should be kept for the 17-year life of the patent plus an additional 10 years.
In summary, researchers should make their notebooks into real diaries of their thinking processes. This is the greatest failing of most notebooks. Many scientists don't realize the importance of recording their speculations and daydreams or of noting down where they got an idea. The other major failing is that notebooks are not understandable to others. To determine whether your current system of note-taking is okay, try to review one of your ten-year-old notebooks. If you have trouble reading your own material, then you'd better change your approach. Finally, it is very important to have a disinterested party review and archive sequential copies of data.
Influence of Stocking Densities on Walleye Fry Viability
in Experimental and Production Tanks
By: Alan Moore and Margaret A. Prange, Iowa Department of Natural
Resources, Rathbun State Fish Hatchery RR 2, Moravia, Iowa 52781 and Brian T. Bristow and
Robert C. Summerfelt Department of Animal Ecology
Iowa State University Ames, Iowa 50011
The main constraints to development of a production scale system for intensive culture of larval walleye have been low incidence of gas bladder inflation (GBI) and poor survival. Noninflation of the gas bladder (NGB) has been a critical factor in most attempts to rear larval walleye. Walleyes without inflated gas bladders are of little value to fishery managers because without an inflated swim bladder a fish has little chance for long term survival. Noninflation of the gas bladder in walleye is considered to be due to a surface tension-oil film barrier preventing small larva from penetrating the surface film for the initial intake of atmospheric gas. Noninflation of the gas bladder, however, has been a problem in live feed experiments and has been observed in pond-cultured fingerlings as well. An oil film is not the only obstacle to successful gas bladder inflation: larval walleye need a clean surface to inflate their gas bladders because, when air is gulped from the surface and forced into the swim bladder, feed and bacteria may also enter.
Surface sprays have been shown to increase gas bladder inflation in larval walleye reared intensively. In 1991, at both the Rathbun Hatchery (Moravia, IA) and at Iowa State University (Ames, IA), vertical (Rathbun) and horizontal (ISU) surface sprays were used to disrupt surface films and to create turbulence on the water surface of walleye rearing tanks. Walleye reared in these experiments had gas bladder inflation percentages ranging from 71.5 to 100 (mean of 89) at Rathbun, and from 18.3 to 82 (mean of 45) at Iowa State. After sprays were used at Iowa State University and Rathbun Hatchery, GBI averaged 83.3% in tanks with surface spray, but only 38.8% in experiments conducted without surface sprays.
Additional experiments were conducted at Rathbun Hatchery to determine if intensively reared larval walleye performances in deep cylindrical tanks with a circular flow and in a cuboidal tank with an upflow pattern were different. This experiment showed that walleye fry in the deep cylindrical tanks performed better than those in cuboidal tanks in two out of three trials. Fry viability ranged from 3.8 to 60.8%. Evidently, continuous recirculation of feed by the upwelling current in the cuboidal tanks had an adverse affect on gas bladder inflation and survival.
The next step in developing techniques for mass culture of larval walleye will require determining optimal fry densities. Experimentally, larval walleye have been cultured intensively at densities ranging from 1 to 1000 fry/L. It has been suggested that the optimal density for the intensive culture of larval walleye is somewhere between 10 and 1000 fry/L, but this range is far too broad to serve as a guideline for practical culture. Most studies have indicated that stocking densities of less than 20 fry/L result in higher survival. Li and Mathias (1982) stated that "increasing fish density above one fish per liter had a deleterious effect on survival." The decrease in survival at higher densities may be caused by successful cannibalism or by injury caused by cohort attacks. For intensive culture to be a practical alternative to pond or tandem culture, however, densities must be above 20 fry/L. Most studies published to date have concentrated on some variable other than stocking density, such as diet or culture environment, and the observed effects of stocking density on survival have been incidental.
In 1992, the goal of the research at Rathbun State Fish Hatchery was to replicate the excellent results of the 1991 production season, to evaluate fry stocking density, and to compare performance in large (679 L) cylindrical tanks, which simulate a full-scale hatchery production system, with smaller (278 L) cylindrical experimental tanks.
Methods and Procedures:
Experimental Design
Effect of stocking density on gas bladder inflation and survival of walleye fry was studied. Six 278-L cylindrical tanks were used in each of three trials (T-1, T-2, and T-3), between April 11 and June 15, 1992. Five different stocking densities (20, 30, 40, 50, and 60 fry/L) were evaluated with two replicates at each density. In trials one (T-1) and three (T-3), the densities were 20, 30, and 40 fry/L, and in trial two (T-2) the tanks were stocked at 20, 50, and 60 fry/L. In addition, three 679-L tanks stocked at 20 fry/L were used to evaluate fry viability in production-scale tanks. Fry performance in these large tanks was compared with that in each of the two 278-L tanks stocked at 20/L of each trial.
Fry Culture Tank Designs and Water Flow
The small cylindrical tanks were the same design used in 1991 experiments. The entire inside walls of the tank, standpipe, screen, and inlet pipe were painted black; the tank bottom was light gray. Flow was directed at a point midway between the wall of the tank and the edge of the screen, in a clockwise direction. The flow rate was regulated by a flow meter connected to the inlet pipe by plastic tubing. In all trials, the flow rate was set at 3.0 L/min for the first day after stocking and was maintained between 5.5 and 6.5 L/min thereafter.
Each 278 L cylindrical tank was provided with one surface spray directed into the tank with a 180 degree perimeter nozzle and a flow regulation valve. Spray was directed at an angle (80 degrees) so that it contacted the surface between the screen and the right side of the tank in the same direction as the water flow.
The large cylindrical tank rearing volume was approximately 679 liters. Flow was directed at a point midway between the wall of the tank and the edge of the screen, in a clockwise direction. The inside wall of the tank was painted flat black, and the bottom was light green. The screen was painted black, and the standpipe was white. Initially the flow rate was 8.0 L/min, but it was increased to as much as 11.0 L/min as the fry grew.
Each large tank was provided with two surface sprays connected to the water source by a 1.3-cm ID PVC pipe placed over the center of the tank, approximately 20 cm above the water surface. Spray was directed at an angle so that it contacted the surface between the screen and each side of the tank in the same direction as the flow. Spray volume/sprayer was the same as with the small tanks.
Lighting
All tanks were illuminated by overhead 150-watt projector flood lamps. The lights were on from 1500 hours to 1000 hours the next day. Black plastic covered frames surrounding the small tanks to prevent exposure to light entering from nearby windows and doors. The large tanks were not surrounded with black plastic and were exposed to light from nearby windows and doors.
Water supply
The water supply was from Rathbun Reservoir, and the temperature of the water supply was that of the lake, but three heaters were placed in the head tank to increase the initial water temperature to 12.2 C in T-1 and to 18.3 C in T-2. Heaters were not used in the T-3 trial because the lake temperature exceeded 18.3 C. Temperatures during T-1, T-2, and T-3 averaged 13.3, 18.7, and 20.3 C, respectively.
Source of Fry and Stocking Densities
T-1: April 11-May 2 - Upon arrival at Rathbun Hatchery, eyed eggs were tempered for one hour to reduce the temperature from 9.4 to 6.6 C before they were placed in incubation jars. Incubating eggs were warmed gradually to approximately 12.2 C. Eggs finished hatching nine days later. At 2d post-hatch, fry were enumerated for stocking with a mechanical fry counter (model FC2, Jensorter, Inc., Bend, Oregon). Densities were 20, 30, and 40 fry/L in the six small tanks (two tanks at each density), and 20 fry/L in each of the three large tanks.
T-2: May 5-May 26 - Eyed eggs were counted with a mechanical fry counter. Densities were 20, 50, and 60 fry/L in the six small tanks (two tanks at each level) and 20 fry/L in each of the three large tanks.
T-3: May 25-June 15 - One-day-old fry numbers were estimated volumetrically. The stocking densities were the same as in T-1, that is, 20, 30, and 40 fry/L in the small tanks, and 20 fry/L in the large tanks.
Feeding
Small tanks - Variable speed auger feeders were used to dispense the feed. Three marbles were placed in each feeder to reduce bridging of food over the auger. All feeders were connected to an automatic timer (Sweeney Enterprises, Inc., Boerne, Texas) which controlled duration and frequency of feedings. Fry were fed every 4 min for 16 h each day (1530-0730 hours). Duration of each feeding ranged from 3 to 14.1 seconds (s).
Large tanks - Each tank had two auger feeders placed 30 cm above the water on opposite sides of the tank. Feeding frequency was controlled with an automatic timer. Fry were fed every 5 min during T-1 and every 4 min during T-2 and T-3. Feeders were on for 22 h/d (1000 to 0800 hours). Duration of each feeding ranged from 0.1 to 0.5 s.
Proposed feeding rates (grams of food/1,000 fry) were based on trials two and three in 1991. Feeding rates were adjusted for fish age and for the estimated number of fry in the tank. Changes in food size and type were based on fry length. Fry were fed the Kyowa Fry Feed (Kyowa Hakko Kogyo Co. Ltd., Japan) throughout the experiments.
Tank Cleaning
Three to five h/d were spent siphoning and cleaning the nine tanks. The daily routine for tank cleaning included siphoning waste feed and dead fry, removing and spraying the drain screens with pressurized water until clean. Dead fry removed by the siphon were counted. A coarse sponge was used to wipe inlet pipes and all inside surfaces of tanks beginning 9 d posthatch in trial T-1 and 6 d posthatch in trials T-2 and T-3. Before the sidewalls were cleaned, the water level was decreased by 50%. When cannibalism became evident (11, 7, and 6 d posthatch during trials T-1, T-2, and T-3, respectively), up to 10 min/tank/d were spent siphoning out cannibals and recording their numbers.
Water Quality
Water samples were collected at the water source (head tank) to the culture tanks. Carbon dioxide, total hardness, and alkalinity were measured using a Hach Kit (Hach Company, Loveland, Colorado). A Hach DR/3000 spectrophotometer was used to determine nitrite and total ammonia nitrogen levels and turbidity. An Orion pH meter was used to measure pH. A dissolved gas meter (Common Sensing, Inc., Bainbridge Island, Washington) was used to measure DP, N2 mm Hg, % N2, total gas pressure (TGP) mm Hg, and % TGP.
Fry Sampling and Final Count
From 6 to 18 d posthatch, three to four random samples of 20 fry were removed from each tank and observed microscopically for presence of inflated gas bladders, food in gut, and oil globules. The percentages of fry with food in gut and with inflated gas bladders was recorded along with oil globule diameter, gas bladder length, and fry length. Final samples of 100 fry/tank were removed at the end of each trial. Presence of food in gut, lengths of gas bladders, and lengths of fry were recorded. The surviving fry were hand counted at the end of each trial.
Discussion
Density Effects
Fry survival was not significantly affected by stocking densities between 20 and 40 fry/L, but survival was significantly lower at densities of 50 and 60 fry/L than at a density of 20/L. The number of survivors (i.e. production per tank), however, was significantly greater in the tanks stocked at 50 and 60/L (216% greater at 60/L) than in the tanks stocked at 20/L. Considering the comparative value of a newly hatched fry to that of a 21 d old, feed-trained fingerling, it would be more economical to stock fry at densities of 50 than at 20 fry/L despite the reduced percentage of survival, because the output of trained fingerlings is greater at the higher densities.
Production Scale Tanks
With the exception of the third trial, when bacterial gill disease substantially reduced survival, fry viability was greater in the large than in the small tanks. Gas bladder inflation, however, was significantly lower in the large tanks in 2 of 3 comparisons. The differences in gas bladder inflation may have been due to the lower number of spray heads per unit surface area of tank in the large tanks; i.e. more spray heads may be needed in the large tanks. There was one spray per 4,778 cm2 of surface area in the small tanks and one spray per 5,941 cm2 of surface area in the large tanks. Small tanks had 24% more spray coverage than did the large tanks.
The lower incidence of cannibalism in the large tanks may be related to the difference in the length of the feeding period. The small tanks were fed for 16 hours/day, and the large tanks for 22 hours/day. The mean percentage of fry removed as cannibals was 4.9% (range 1.9 to 7.5%) in tanks fed 16 hours/day, but only 1.8% (range 0.3 to 3.4%) in tanks fed 22 hours/day.
Effects of Temperature
The best performance of fry in the three trials was in T-2 when the mean temperature was 18.7 C. The optimum temperature for raising walleye from 0-22 days posthatch seems to be between 17 and 18 C. Fry raised at 13.3 C (T-1) developed more slowly and were more likely to be killed or injured during cleaning and to succumb to cannibalism. In T-3, when mean temperature was 20.3 C, fry performance was less favorable because of bacterial gill disease. Temperature, crowding, and the general cleanliness of the inflowing water may have been a factor in the development of bacterial gill disease during the third week of this trial. In T-1, T-2, and T-3, respectively, survival and mean temperatures were 3.8% at 15.0 C, 34.2% at 15.8 C, and 68.2% at 19.2 C. In T-3, the mean temperature during the first nine days of culture was 18.2 C; close to the projected optimum.
Detecting
Swim Bladder Inflation in Fingerling Walleyes
By: Frederic T. Barrows, Greg A. Kindschi, and Ronald E. Zitzow, U.S. Fish
and Wildlife Service, Fish Technology Center, 4050 Bridger Canyon Road, Bozeman, Montana
59715
In the March '94 issue of the MTAN, we included two articles regarding the importance of surface water spray and swim bladder inflation to guarantee the successful rearing of walleye fry. As a follow up, we wanted to explain how you can detect the early signs of swim bladder inflation problems. The following information has been edited to conserve space.
The lack of swim bladder inflation is a major problem in the intensive culture of several species of larval fish. This phenomenon has been reported several times in intensively reared walleyes, striped bass, white seabass and madai. Recently, however, in pond-reared walleye fingerlings, 9,490 of 13,000 (73%) had uninflated swim bladders (personal observation). Although some populations of pond-reared walleyes had been observed to have low percentages of fish with uninflated swim bladders, such high percentages have not been previously documented.
Determining swim bladder inflation status of larval walleyes (< 40 mm total length) can be completed easily with the aid of a dissecting microscope. This method, however, is time-consuming, cumbersome, and not effective with large fish. Chapman (1988) developed a method of separating larval striped bass with inflated swim bladders from those with uninflated swim bladders. An anesthetic solution was used to separate the fish by their buoyancy. Henderson-Arzapalo (1992) used Tricaine (MS-222) in a saltwater solution to separate normal phase-I striped bass from those with uninflated gas bladders. Radiography (X-ray photography) has also been used to monitor development and to determine swim bladder inflation in salmonids and European bass.
The development of methods to easily detect swim bladder inflation status in fingerling walleyes (25-155 mm total length) would enable large groups of fingerlings to be monitored. Our objective was to compare four methods for determining swim bladder inflation, in terms of their accuracy, cost, and effect on walleye survival.
Methods
The light table (glow box, model 12-20E-3, Instruments for Research and Industry, Cheltenham, Pennsylvania), anesthesia, saltwater float, and radiographic methods were tested for detecting swim bladder inflation in walleyes that were 80-167 mm long. The light table method involved the use of a box that contained two fluorescent light bulbs and had a translucent top; such a box is normally used for viewing photographic slides. Fish were anesthetized in Tricaine (100 mg/L) and them passed over the light box. The swim bladder is easily identifiable as a light tubular area between the spinal column and the viscera.
In the anesthesia method, MS-222 was used to relax the fish so that an inflated swim bladder would have a buoyant effect on fish position. Preliminary trials indicated that the concentration of the anesthetic affected the buoyancy and relaxation time of fish with inflated swim bladders. Seven concentrations of MS 222 were tested (40, 60, 80, 100, 120, 140, and 160 mg/L), and the time required for the cessation of all fish movement, except gill movement, was recorded.
Salt water was used to separate fish with inflated swim bladders from those with uninflated swim bladders according to buoyancy, as in the anesthesia method. However, only dead fish were used in this trial. Saltwater concentrations of 1, 2, 3, and 4% by weight and plain water (no salt) were used to separate walleye fingerlings. Fish were placed in the bucket, stirred, and allowed to settle for about 30 s. Buoyant fingerlings were removed with an aquarium net and placed in a separate bucket of fresh water. Nonbuoyant fish (those lying flat on the bottom, unable to be raised into the water column with gentle stirring) were removed and then placed into another bucket containing fresh water. After the samples were separated in this manner, they were again removed from the water and placed on a light table so that swim bladder inflation could be verified visually and the fish of the different groups could be counted. Twelve frozen samples, each known to contain a mixture of walleye fingerlings with and without inflated swim bladders, were available for the trial. Three samples were tested for each salt concentration. Samples separated with plain water were mixed back together and separated again with 4% salt solution.
An additional experiment was conducted to determine if freezing and thawing had an effect on the detection of inflated swim bladders. Anesthetized fish were separated in 2% salt solution, and nonbuoyant fish were checked for inflated swim bladders under a dissecting scope.
The fourth method involved X-ray radiography of several walleyes lying flat and close together on a film plate. This procedure was conducted at a local veterinary clinic. In the present trial, no efforts were made to keep the X-rayed fish alive. This method could be used for determining the percentage of swim bladder inflation in a population of fish, whereas the other three methods could be used to sort live fish with inflated swim bladders from those with uninflated swim bladders.
Different triplicate lots, each with 30 fish of mixed swim bladder status, were examined with the light table, anesthesia, and X-ray methods. All fish were then sacrificed, dissected, and visually examined for the presence of an inflated swim bladder. The accuracy of each method was evaluated by comparing the ratio of inflated to uninflated swim bladders determined by the method being tested with the ratio determined by dissection. To evaluate the effect of the methods on survival, three lots of fish were sorted by the light table method and three lots were sorted by the anesthesia method with the high concentration of MS-222 (160 mg/L). They were then placed in 19-L rearing containers for 14 d to monitor survival.
Results and Discussion
The anesthesia and X-ray methods were 100% accurate in detecting swim bladder inflation status of the tested walleye fingerlings, which were longer than 75 mm and weighed about 18 g each. The anesthesia method with MS-222 at a concentration of 100 mg/L seems to be inexpensive and accurate, but fish condition must be closely monitored to prevent mortality during the treatment. At this MS-222 concentration, fish could be separated after 75 s. Walleyes with inflated swim bladders were upside down or floating at the surface, whereas those with uninflated swim bladders were lying flat on their sides on the bottom of the container. Shallow and wide containers, which provide more water surface area, were more useful than deep and narrow containers.
The light table method for detecting swim bladder inflation in walleye fingerlings is inexpensive, fast and did not adversely affect the fish after treatment. The accuracy, however, of the light table method was only 91.5%. In preliminary studies, the accuracy of this method was observed to be higher for fish shorter than 75 mm than for the larger fish used in this trial. This difference is attributable to the greater transparency of small walleyes, which allows an unobstructed view of the swim bladder. Neither the MS-222 method nor the X-ray method was tested on fish less than 75 mm long.
The saltwater float method was 99.7% accurate and simple to conduct. More time was required to separate and remove fingerlings in the 0, 1, and 2% salt solutions, because numerous buoyant fish remained throughout the water column and at the bottom. Only a few fish with inflated swim bladders were found among the nonbuoyant ones.
Separation of fingerlings according to swim bladder status worked well in 3 and 4% salt solutions. Very few buoyant fish remained in the water column or near the bottom. One fish with an uninflated swim bladder was found among buoyant fish of one sample separated in 3% salt solution. This fish may have stuck in some way to a buoyant fish. It appears that 4% salt solution may be too strong, as several fish with uninflated swim bladders separated with buoyant fish in all three samples. Based on these results, our recommendation is that 3% salt solution be used to separate 2.5-5-cm-long walleye fingerlings with inflated swim bladders from those with uninflated swim bladders after fish have been frozen and then thawed.
There was an effect on freezing and thawing on swim bladder inflation status. After fish had been frozen and thawed, a higher rate of noninflation was observed than before freezing. When expressed as a percentage of fish with uninflated swim bladders before freezing, the average increase was 7.9%. However, when expressed as a percentage of the total sample, the increase was only 0.3%. The effect of this method of storage and evaluation was considered minor.
The X-ray method is expensive but extremely accurate for testing walleyes of any size. Whether keeping fish alive for this method is practical is not known, but a good approach for this method may be to radiograph only a subsample of a population to determine relative percentages of swim bladder inflation and noninflation. This method also provides, however, a permanent record for future use.
Survival was not affected by evaluating the swim bladder status of fish with either the anesthesia (160 mg MS-222/L) or light table method. No mortality occurred during the 2 weeks after the use of these methods.
We recommend either the light table method or the MS-222 method in conjunction with a 3% salt solution for routine screening of populations of walleye fingerlings. These methods are simple, inexpensive, and fairly accurate. Training personnel to perform these evaluations could consist of first using one of these methods to sort the fish according to their swim bladder inflation status and then checking the results by either dissection or the X-ray method.
Hydrogen
Peroxide Controls Fungal Infections On Trout Eggs
By: Leif Marking, Jeffrey Rach, and Theresa Schreier, National Fisheries
Research Center, P.O. Box 818, La Crosse, WI 54601 (608-783-6451)
Malachite green, once the preferred fungicide for fishery use, is no longer permitted for treatment of fish or fish eggs for control of fungus because the Food and Drug Administration has restricted the use of malachite green on food fish. Formalin is an effective fungicide, but fishery managers are concerned about safety to the user and effluents in the environment. Other antifungal agents, therefore, are needed to maintain healthy fish and eggs in fish culture systems.
Hydrogen peroxide is active against a wide variety of organisms-bacteria, yeasts, fungi, viruses, and spores. Federal agencies list this chemical as "Generally Recognized As Safe" when used as a bleaching agent in manufacturing or feeding practices or as an antimicrobial agent in cheese production or drinking water treatment. Furthermore, hydrogen peroxide has been used as an antiseptic and a treatment for skin parasites, protozoans, and monogenetic trematodes on fish and is proposed as a treatment for sea lice on salmon. We designed experiments to evaluate the effectiveness of hydrogen peroxide for control of fungal infections on eggs of rainbow trout.
In Vivo Procedures
Green eggs of rainbow trout were placed in Heath incubation trays and maintained in well water with a flow rate of 1 L/min. Groups of 500 eggs were confined within 15-cm diameter acrylic rings that were fastened to the screen of each incubation tray. Eggs were inoculated with fungus (Saprolegnia parasitica) actively growing on hemp seeds suspended by tea balls in the upper tray of each duplicated treatment. Infection of eggs generally occurred within 7 days. The infection rate of about 10% in each tray was obtained by exchanging infected eggs between trays. Eggs infected at the 0 and 10% level were then exposed to hydrogen peroxide for 15, 30, or 60 min every other day for 2 weeks. Treatments ceased when the eggs began to hatch. A positive control group was inoculated with fungus but not treated with hydrogen peroxide. A negative control group was neither inoculated with fungus nor treated with hydrogen peroxide.
Efficacy of Hydrogen Peroxide
Artificial infection of eggs was successful - infection rates increased substantially in the positive control and the hatch rate for those eggs was only 2.1%. Hydrogen peroxide treatments of 500 ppm controlled fungal infections on eggs not infected (O% infection) at exposures of 15, 30, and 60 min; hatch rates were significantly higher than for control groups. This treatment concentration was effective for control of fungus at the 10% infection rate only in the 60-min exposure. The 1,000-ppm treatments, however, were effective for control of fungus and increasing the hatch rate at all three exposure periods. Both treatment concentrations controlled fungus and increased hatch rate of uninfected eggs (O% infection) at all exposures.
In reality, the 10% infection rate is perhaps a "worst case" situation in hatcheries where treatments are normally done when the fungus first appears or is suspected. The 1,000-ppm treatment rate may be excessive, especially for the longer exposures. It is premature to recommend a general treatment rate, therefore, a suitable treatment rate should be developed to meet the particular conditions in individual hatcheries. Hydrogen peroxide seems to be equally or more effective than formalin for control of fungus on trout eggs in similar in vivo experiments.
Availability for Use in Fisheries
Hydrogen peroxide is one of the most environmentally compatible chemicals because the primary decomposition products are oxygen and water. This versatile chemical already has widespread application and acceptance in pulp and paper, textile, waste treatment, mining, petroleum, food and chemical processing, cosmetic, and pharmaceutical industries. The U.S. Food and Drug Administration recently approved a petition from the National Fisheries Research Center-La Crosse that hydrogen peroxide be classified as a low regulatory priority when used to control fungi on all species and life stages of fish, including eggs. This ruling means that hydrogen peroxide can be used as a fungicide without an investigational new animal drug permit or a new animal drug application.
Over the last several months the MTAN has been accumulating additional information from product manufacturers which sell aquaculture related supplies. The following list will hopefully help you with locating different suppliers to insure your getting the best product and price available.
Little Titan: This pond aeration system offers an efficient and cost effective way to improve water quality and to provide fish with oxygen enriched water. The device is produced by Otterbine/Barebo, Inc., 3840 Main Rd. East, Emmaus, PA 18049 (215-965-6018).
AquaTornado Aerator: This large pond aeration system will mix and destratify, increase available dissolved oxygen, plus help to control algae and ice. The device is produced by Aeromix Systems, Inc., 2611 N. Second St., Minneapolis, MN 55411-1634 (800-879-3677).
PT4 - Hauling Oxygen Meter: Multichannel monitor for reading oxygen, temperature and pH. Serves to constantly monitor DO in incubators and fish transport tanks. Used at St. Croix Tribal hatchery during incubation periods with great success. This meter is made by Point Four Systems, Inc. and sold by Aquatic Ecosystems, Inc. 2056 Apopka Blvd., Apopka, FL 32703 (800-422-3939).
Sentry 3: This oxygen and temperature monitor is ideal for monitoring water for fish hatcheries or recreational sport fishing. The device is produced by Otterbine/Barebo, Inc., 3840 Main Rd. East, Emmaus, PA 18049 (215-965-6018).
Portable Dissolved Oxygen Meter: Features include a rugged waterproof design, automatic compensation of temperature, salinity, and partial pressure, plus an analog and digital outputs. The device is produced by Royce Instrument Corp., 13555 Gentilly Rd., New Orleans, LA 70129 (800-347-3505).
Fishing News Books-1994 Catalog: This reference book list several detailed descriptions of many other fisheries/aquaculture related books and journals which are available through Fishing News Books. The list is unique in its all-embracing coverage of the many technical and scientific aspects of fish farming, commercial and environmental management of aquatic resources and technical and scientific aspects of commercial fishing. An order form is also included with the catalog so you may purchase any of the books which are listed. The 1994 catalog can be obtained by writing to Philip Saugman, Fishing News Books, Blackwell Scientific Publications Ltd., Osney Mead, Oxford 0X2 0EL, UK (phone number is 0865-240201).
Fiberglass Rearing Tanks: Numerous sizes are available for round, half round, cone, rectangular and square tanks. Custom orders for specialized tanks are also taken. The standard features include; smooth molded gelcoated interior, exteriors are gelcoated over a mat finish, handlaid multiple laminates of mat and roving, flanges, drains, sumps, fittings, rounded corners and sidewall stiffening ribs. These tanks are produced from:
Hulls Unlimited East, Inc., PO Box 70, Deltaville, VA 23043 (804-776-9711).
Peterson Fiberglass Laminates, 300 Stariha Dr., Shell Lake, WI 54871 (715-468-2306).
Polytank, Inc., 62824 250th St., Litchfield, MN 55355 (612-693-8370).
J & S Tanks, 935 Highway 29 N., P.O. Box 144, Alexandria, MN 56308 (612-763-4191).
Will-O'-The-Wisp's: This device presents a totally new concept in commercially feeding your pond fish. The feeder is placed on a post overhanging the water. A circular ultra violet light attracts the insects and are pulled into the feeder by a high speed fan. The fan then blows the insects against a screen at the back of the feeder, dropping them into the water to feed your fish. The monthly electrical cost are reported to be less then $5.00. The device is distributed by Aquatic Biologist, Inc., N5174 Summit Ct., Fond du Lac, WI 54935 (414-921-6827).
Fish Farm and River Ecosystem: High schools and other public educational programs who want to get into aquaculture (in a small way) may want to look into these devices. The Fish Farm is a self-contained recirculating system (10' * 10') that provides "hands-on" fish rearing capabilities. The rearing system comes with a galvanized steel tank, liner, air pump, molded clarifier, PVC water lines and a rotating biological filter. The River Ecosystem is a small cross-section of a working river, complete with banks, shallows, pools, waterfalls and rapids. This system allows you to watch and monitor fish, frogs, lizards and plants in a "natural" setting. The system includes a tank, pump, light, heater, filtration cartridge and owners manual. To receive more information contact the Stoney Creek Equipment Company, 11073 Peach Ave., Grant, MI 49327 (800-448-3873).
Nets and Seines: Nylon gill nets, experimental gill nets, herring-pike-perch nets, hoop/fyke nets, seines, small mesh netting, survival and bait nets, landing and dip nets, mosquito nets, rope, leaded and float lines, nylon twine and netting needles, floats and leads, net pens, bird and animal capture nets and hydroelectric dam tail race nets. These products are available from:
InterNet Inc., 2730 Nevada Ave. North, Minneapolis, MN 55427-9949 (800-328-8456).
H. Christiansen Co., 22 North 2nd Ave. West, Duluth, MN 55802 (218-722-1142).
FNT Industries, Inc., 927 First St., PO Box 157, Menominee, MI 49858 (800-338-9860).
Minnow Traps: Cylinder, Cloverleaf, B, and Box type traps are available. All the traps are made of galvanized metal or treated wood frames, zinc coated screws, welded or woven galvanized hardware cloth and steel/aluminum rivets. They also have PVC coated wire. Easy-slide doors and smooth edges are part of every trap. Custom orders are also accepted. For more information contact Jerry Radermacher, Community Services Workshop, PO Box 787, Willmar, MN 56201 (612-235-4613).
Everything Else:
If you like one stop shopping for feeds, chemicals, equipment and books, be sure to subscribe to Aurum Aquaculture, 11818 115th Ave. N.E., Kirtland, WA 98034 (800-817-5808).
Two additional sources for a full range of aquaculture related materials are through:
Aquaculture Supply, 33418 Old Saint Joe Rd., Dade City, FL 33525 and Eagar Inc., P.O. Box 540476, North Salt Lake, UT 84054 (800-423-6249).
To find equipment for aeration, measuring, monitoring, pumps, filtration, sterilization, laboratory, chemicals, feed and feeders, nets, cages, predator control, tanks and liners, be sure to subscribe to Aquatic Eco-Systems Inc., 2056 Apopka Blvd., Apopka, FL 32703 (800-422-3939).
To find equipment for aeration systems, regenerative blowers, air and water pumps, air diffusers, vinyl tubing, RBC biofilters, biofiltration media, fish feeding devices, telephone alarms, plus hauling and rearing tanks, be sure to contact the Stoney Creek Equipment Company, 11073 Peach Ave., Grant, MI 49327 (800-448-3873).
If you need materials for water aeration (destratification of ponds, maximizing oxygen transfer, airlift systems, packed columns, diffusers, degassers and air curtains) or water heating/chilling systems, you may want to contact Aquaculture Research/Environmental Associates (AREA), P.O. Box 1303, Homestead, FL 33090 (305-248-4205).
Vertical tray incubation systems with or without isolation options, circular fish tanks, rearing troughs, egg shipping cases, fish measuring boards and "Heath" water/egg tray replacement components. For more information contact F.A.L/Heath, 4540 So. Adams St., P.O. Box 9037, Tacoma, WA 98409 (800-851-1510).
To develop a career in aquaculture you may want to contact the Alexandria Technical College, 1601 Jefferson St., Alexandria, MN 56308 (612-762-0221).
A new wallet size handout which describes how to identify the Eurasian Ruffe is now available through the Ashland FRO (715-682-6185).
Do you need transport containers for fish? Dynoplast has developed a series of versatile food containers which can withstand rough usage. These containers (manufactured from food approved materials) will ensure delivery of cool and fresh products in prime condition. For more information contact Canam Trading Corp., Elgin, IL(800-231-9721 - ask for Jim Quinn).
Product and company names mentioned in this publication are for informational purposes only. It does not imply endorsement by the MTAN or the U.S. Government.
