• TABLE OF CONTENTS
HIDE
 Historic note
 Front Cover
 Front Matter
 Table of Contents
 Introduction
 Hatchery facility
 Brood fish conditioning
 Brood fish capture and handlin...
 Determination of gender
 Controlled maturation in indoor...
 Sampling the eggs in the ovary
 Hormone injections
 Spawning
 Triploid grass carp
 Verification of triploidy
 Care of spawn in incubators
 Developmetn of the larvae
 Transportation of fry
 Pond production to fingerling...
 Intensive production to fingerling...
 Conclusion
 Additional reading














Group Title: Bulletin - University of Florida. Cooperative Extension Service ; no. 244
Title: Hatchery manual for grass carp and other riverine cyprinids
CITATION THUMBNAILS PAGE IMAGE ZOOMABLE
Full Citation
STANDARD VIEW MARC VIEW
Permanent Link: http://ufdc.ufl.edu/UF00025532/00001
 Material Information
Title: Hatchery manual for grass carp and other riverine cyprinids
Series Title: Bulletin
Physical Description: 28 p. : ill. ; 28 cm.
Language: English
Creator: Rottmann, R. W
Shireman, J. V
Florida Cooperative Extension Service
Publisher: Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida
Place of Publication: Gainesville Fla
Publication Date: [1992]
 Subjects
Subject: Fish hatcheries -- Florida   ( lcsh )
Ctenopharyngodon idella -- Florida   ( lcsh )
Cyprinidae -- Florida   ( lcsh )
Genre: government publication (state, provincial, terriorial, dependent)   ( marcgt )
bibliography   ( marcgt )
non-fiction   ( marcgt )
 Notes
Bibliography: Includes bibliographical references (p. 27).
Statement of Responsibility: R.W. Rottmann and J.V. Shireman.
General Note: Title from cover.
General Note: "Printed 4/92"--P. 28.
Funding: Bulletin (Florida Cooperative Extension Service)
 Record Information
Bibliographic ID: UF00025532
Volume ID: VID00001
Source Institution: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: aleph - 001753225
oclc - 26898559
notis - AJG6188

Table of Contents
    Historic note
        Unnumbered ( 1 )
    Front Cover
        Page i
    Front Matter
        Page ii
    Table of Contents
        Page iii
    Introduction
        Page 1
    Hatchery facility
        Page 2
        Page 3
        Page 4
        Page 5
        Page 6
        Page 7
        Page 8
    Brood fish conditioning
        Page 9
    Brood fish capture and handling
        Page 10
    Determination of gender
        Page 11
    Controlled maturation in indoor tanks
        Page 11
    Sampling the eggs in the ovary
        Page 11
    Hormone injections
        Page 12
        Page 13
        Page 14
    Spawning
        Page 15
        Page 16
    Triploid grass carp
        Page 17
    Verification of triploidy
        Page 18
    Care of spawn in incubators
        Page 18
        Page 19
    Developmetn of the larvae
        Page 20
    Transportation of fry
        Page 20
    Pond production to fingerling size
        Page 20
        Page 21
    Intensive production to fingerling size
        Page 22
        Page 23
        Page 24
        Page 25
    Conclusion
        Page 26
    Additional reading
        Page 27
        Page 28
Full Text





HISTORIC NOTE


The publications in this collection do
not reflect current scientific knowledge
or recommendations. These texts
represent the historic publishing
record of the Institute for Food and
Agricultural Sciences and should be
used only to trace the historic work of
the Institute and its staff. Current IFAS
research may be found on the
Electronic Data Information Source
(EDIS)

site maintained by the Florida
Cooperative Extension Service.






Copyright 2005, Board of Trustees, University
of Florida







Bulletin 244


Hatchery Manual for Grass Carp and

Other Riverine Cyprinids


R.W. Rottmann and J.V. Shireman


V..


Florida Cooperative Extension Service / Institute of Food and Agricultural Sciences /


University of Florida / John T. Woeste, Dean


7 a '"-
.... : -^ .' 'l.^..,:.^ -^y ii6u S



;.~5~~ikl


s~s~-





















SCIEPWC

































R. W Rottmann is a Biologist III and J. V. Shireman is Chairman, Department of Fisheries and Aquacul-
ture, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, Florida, 32611.








Table of Contents


Introduction . . . . . . ..

Hatchery Facility .......................
W ater supply . . . . . . .
Hatchery ponds ............
Hatchery buildings .. ... ... .. .. ... .. .
Brood fish holding tanks ......... .........
Incubating apparatus .. ...................
Semi-intensive and intensive fry rearing tanks and cages .

Brood Fish Conditioning ..................

Brood Fish Capture and Handling ............


1.. . . . .


. . . . . . 9

. . . . . . 10


Determination of Gender ................................

Controlled Maturation in Indoor Tanks ........................
Early spaw n . . . . . . . . . .
Late spawn .... ...............................
Normal-season spawn . .. .. .. .. .. .. .. .. .. ... ... .


Sampling the Eggs in the Ovary ........

Hormone Injections ................
Carp pituitary . . . . .
Human chorionic gonadotropin and carp pituitary .
Synthetic LH-RH analogue .............
Synthetic LH-RH analogue and dopamine blocker


Spawning ................
Hand stripping/dry fertilization . .
Tank spawning .............

Triploid Grass Carp ........ ..
Production techniques for triploid fish .

Verification of Triploidy ........

Care of Spawn in Incubators ...
Estimation of percent viable embryos .
Duration of incubation .........
Hatching of eggs ............

Development of the Larvae .....


. . . . . . . 11


. . . .. . . . . . 15
. . . . . . . . . 15
. .. . . . . . . 16

... .. .. .. . ... .. ....... 17
. . . . . . . . . 17

. . . . . . . . 18


Transportation of Fry ......................

Pond Production to Fingerling Size . . . .
Fry pond preparation .................. ....
Stocking of fry . . .. . .. . ...
Feeding of the fry ........ ... ... ..
Harvesting fingerlings from ponds ..... . . ...

Intensive Production to Fingerling Size . . .
Semi-Intensive fry rearing . . . . .
Intensive fry rearing .......................
Live food culture . . . . . .
Pond production to stocking size . . . ..
Intensive production to stocking size . . . .


Conclusions .................


. . 21

. . 21
. .. 21
. . 22
. . 22
. . 22

. 23
. . 23
. . 23
. . 23
. 26
. . 26


. . .. .. .. . 2 7


Additional Reading


. . . 27











Hatchery Manual for Grass Carp


and Other Riverine Cyprinids


Introduction
The Cyprinidae (carp) family has more species
than any other family of fish, and many are of consid-
erable economic importance. The Chinese carps, i.e.,
grass carp (Ctenopharyngodon idella), bighead carp
(Hypophthalmichthys nobilis), and silver carp (H.
molitrix), have been cultured for food in China for
many centuries. These fish were introduced in
Europe during the end of the eighteenth century, and
more recently, in 1963, grass carp were stocked into
U.S. waters. The bighead and silver carp were
brought to this country somewhat later. The primary
interest in the grass carp in Florida is for aquatic
weed control. The grass carp is an effective and
economical control of submersed aquatic weeds, such
as Hydrilla. In Florida, only triploid grass carp are
legal for stocking, because they are reported to be
sexually sterile.
Grass carp in their natural habitat normally spawn
in large, swift-flowing rivers. The adult fish feed in
the shallow backwater areas of the lower river
where vegetation is abundant; they then migrate
upstream in large schools. When the river starts to
swell with spring rains and environmental condi-
tions are suitable, they spawn in the swift current.
Spawning in the flowing water of a flooded tribu-
tary has definite advantages for a fish species. The
continuous current of the water provides oxygen
and keeps the semi-buoyant eggs and larvae in
suspension, thereby keeping them off the bottom
where they could be silted over and die. The turbid-
ity of the swift-flowing water also conceals the eggs
and fry from predators. The current carries the fry
downstream to inundated areas, which are rich in
food organisms required for development of the fry.
The gonads of grass carp confined in lakes and
ponds develop only to a certain stage and then
remain dormant until resorption takes place. This
process can be repeated year after year, without
spawning ever occurring. Hormone-induced spawn-
ing is the only technique presently available that
will initiate hatchery reproduction of these fish.
The hormone-induced spawning technique has
been around for about 30 years. Surprisingly,
hormone-induced spawning of fish appears to be
almost universally successful, with only minor
modifications. Hormones have been used to spawn


the entire range of fish from the primitive sturgeon
(Family Acipenseridae), to salmon (Family Sal-
monidae), sea bass (Family Serranidae), and mullet
(Family Mugilidae). Commercial producers of
striped bass (Morone saxatilis) as well as grass
carp, use hormones routinely to spawn their fish.
The interaction of hormones on the reproductive
system is a complex mechanism, and additional
information is being added yearly to the volumes of
scientific knowledge. However, it is unnecessary to
understand the precise action of the hormones on
the fish in order to use the technique of hormone-
induced spawning. Our experience has taught us
that at least 95% of induced spawning involves
maintaining suitable water quality, having
adequate facilities, providing proper nutrition,
minimizing stress to the fish during capture and
handling, preventing parasite and disease out-
breaks, and simulating natural conditions in the
spawning container.
Induced hatchery spawning and rearing of fish to
marketable size requires a continuous series of de-
cisions, any of which can diminish or completely
obliterate the success of the project. There are
many ways to fail at each step. The process starts
long before the spawning season, with the prepara-
tion of facilities and the conditioning of brood fish,
and continues until the fish produced arrive at
their destination. Consistent performance requires
strict attention to detail.
Hatchery spawning and rearing of fish can either
be consistently smooth, productive, and gratifying,
or frustrating, aggravating, and a waste of time,
money, and fish. Success depends primarily on the
level of practical experience of the hatchery man-
ager and staff, as well as their technical expertise.
This publication is a manual for the hatchery pro-
duction of grass carp and other Chinese carp, and
is directly applicable to other species of fish. It is
the result of 14 years of experience in hormone-
induced spawning of grass carp as well as bighead
carp, silver carp, goldfish (Carassius auratus), red-
tailed black shark (Labeo bicolor), rainbow shark
(L. erythrurus), striped bass, white bass (Morone
chrysops), walleye (Stizostedion vitreum), paddlefish
(Polydon spathula), and sturgeon (Scaphirhynchus
platorynchus).









Hatchery Facility
The hatchery facility is important to the success
of induced spawning and fish rearing. Makeshift
construction of the water supply, brood fish holding,
egg hatching, and rearing facilities usually results
in system failure. The system need not be expen-
sive and elaborate, but it should be properly de-
signed, uncomplicated, and well-made to minimize
problems.
The prerequisites of a fish hatchery are: (1) an
adequate quantity of good quality water; (2) suffi-
cient land of suitable soil type for ponds; (3)
adequate buildings with hatching setups, tanks,
and storage; (4) appropriate equipment; (5) skilled
hatchery workers; and (6) operating capital.


HATCHERY


SPRING A


It is imperative that all appropriate government
agencies be contacted before building or acquiring a
hatchery to ensure that all regulations are followed
and that all necessary permits are obtained.
Water supply
A dependable source and sufficient volume of
high quality water is essential for a successful
hatchery. Good water quality is especially critical
for maintaining brood fish, hatching eggs, and rear-
ing fry. The water supply should be thoroughly
tested prior to hatchery construction. Ground water
sources, such as springs (Figure la) and wells
(Figures lb and Ic), are usually preferred to reser-
voirs and streams for hatchery water supply.
Ground water normally does not contain parasite


WEL
PUM





7!


PRESSURE TANK


L
P


HATCHERY


B


WELL RESERVOIR
PUMP POND


SCREEN
INTAKE


HATCHERY IMPOUNDMENT
A OF A STREAM


HATCHERY
PUMP
LAKE OR SCREEN
RIVER INTAKE


Figure 1. WATER SOURCES.

A. A Spring.
B. A well and pressure tank.
C. A well and reservoir combination.
D. Gravity flow from a surface water source.
E. Pumping from a surface water source.


SCREEN
INTAKE


DAM


HATCHERY


r










Table 1. Optimum water quality parameters


PARAMETER VALUE

Temperature 70 77" F
pH 7.0 8.5
Alkalinity greater than 25 ppm
Dissolved Oxygen 4 10 ppm
Carbon Dioxide less than 5 ppm
Hydrogen Sulfide less than 0.3 ppm
Un-Ionized Ammonia less than 0.02 ppm
Nitrite less than 0.1 ppm
Copper less than 0.1 ppm
Iron less than 0.5 ppm
Phosphate less than 6.0 ppm
Potassium less than 2.0 ppm
Sodium less than 5.0 ppm
Zinc less than 0.1 ppm

and disease organisms, predacious insects, or wild
fish, which may be present in surface water. Aera-
tion is usually the only treatment necessary to con-
dition ground water for fish culture. Ground water,
however, may contain iron, nitrogen, carbon dioxide,
hydrogen sulfide, methane, and pesticide residues
in toxic levels. Surface water quality may be
superior if ground water is high in toxic products.
Optimum water quality parameters for the hatch-
ery production of grass carp fry are presented in
Table 1.
The water system is best kept simple to minimize
failure. When possible, water should be delivered to
the hatchery by gravity flow, thereby eliminating
the high cost of pumping. Springs, artesian wells,
reservoirs, and diverted streams may be ideally
suited for gravity flow systems (Figures la and Id).
If water is pumped (Figures Ib, Ic, and le), the sys-
tem should be designed so that only one lift is re-
quired. When pumps or aerators are used to main-
tain water quality for fish, back-up systems and an
auxiliary power supply are essential to prevent
mass mortality due to pump or power failures.
Flow-through water systems (i.e., where water is
used in the incubating aparatus or fish tanks, then
discharged) are simpler to construct than recirculat-
ing systems, and fish in these systems are less
prone to disease outbreaks and water quality prob-
lems.
In some situations, recirculating systems may be
the best option. Recirculation systems use less
water, and water temperature is easier to control.
However, greater management skill and attention
to fish health is required when recirculated water
is used.
PVC pipe is by far the easiest and simplest to as-
semble and is one of the least expensive materials


for construction of water distribution and drainage
lines. Leaching of heavy metals from water lines is
also avoided when PVC pipe is used.

Hatchery ponds
The most important considerations in pond de-
sign and construction, particularly the fry and
fingerling rearing ponds, are that they should be:
(1) reasonably watertight; (2) easily supplied with
fresh water; (3) relatively small (0.1 to 1.0 acres);
(4) rectangular in shape for ease of seining; (5) sur-
rounded with levees wide enough for vehicle access;
(6) completely drainable or easily pumped dry for
complete harvest; and (7) equipped with drain
structures and catch basins, if possible, to simplify
harvest. A typical drainable hatchery pond is shown
in Figure 2. Before starting pond construction, a
soil survey should be performed to determine if the
soil type will retain water. Ponds are usually con-


Figure 2. A typical drainable hatchery pond.


Figure 3. A. Concrete catch basin, water inlet and drain
structure ("monk").











Floor Slope
1/8'/Foot
S 12-9


S-4-6


ei
PI


Figure 3. B. Diagram of concrete catch basin and "monk".
structed with a bulldozer, dragline, loader, trackhoe
or backhoe. Construction is greatly simplified if the
site has been previously cleared, and there are no
trees, stumps, roots, or brush to contend with. Fol-
lowing construction, the pond banks or levees
should be immediately sodded and/or seeded with
grass to reduce erosion.
The concrete catch basin and drain structure
("monk") illustrated in Figure 3 is used in many pri-
vate, state, and federal fish hatcheries. Water level
is controlled by a standpipe, and the channels in
the monk accommodate drain screens and baffles.

Hatchery buildings
The hatchery buildings should be designed to be
simple, functional, and durable, with adequate
lighting, ventilation, and temperature control. A
steel-reinforced concrete slab with brick or concrete
block walls is the preferred construction, because of
the dampness and splashing experienced in a fish
hatchery. Water is heavy, so floors must be designed
to withstand the load. Floor drains should be over-
designed to handle the inevitable leaks and spills,
and to carry discharge water from the hatching
apparatus and fish tanks.
The hatchery building should be insulated to
minimize temperature fluctuation and to reduce
the cost of heating and cooling. Adequate work area
and storage space are a necessity. Electrical outlets
should be numerous and easily accessible, but
placed high enough to avoid becoming wet. Lighting
is best provided by standard incandescent bulbs;
they are less troublesome in the damp environment
of a hatchery and do not emit ultraviolet light,
which may be harmful to fish eggs, sperm, embryos,
and fry.

Brood fish holding tanks
Circular tanks at least 6 feet in diameter are
recommended for holding grass carp brood fish.
This tank design is self-cleaning, minimizes injury


to brood fish, and provides a swift water current,
simulating natural spawning conditions.
Fish tanks can be either commercial fiberglass or
plastic units, or constructed from galvanized metal
cattle watering troughs available at most livestock
supply stores. The troughs are strong, lightweight,
relatively inexpensive, and are available in several
sizes ranging from 4 to 10 feet in diameter; depth is
usually 2 feet. These troughs have a small side
drain that is inadequate for fish rearing purposes;
a suitable drain can be constructed from a PVC
closet flange.
Before the drain is attached, the galvanized sur-
e face on the inside of the tank must be treated and
e painted to prevent leaching of toxic zinc into the
water. The galvanized surface is first etched with
acetic acid or copper sulfate solution, then
thoroughly washed with fresh water, and air dried
before epoxy paint is applied. Check with the man-
ufacturer to ensure that the paint is non-toxic. The
paint should be applied in several thin coats, rather
than one thick layer. A paint sprayer is useful, be-
cause it saves time and it applies a more even coat
than is possible with a brush.
Next, a hole is cut in the center of the tank bot-
tom. The closet flange is placed base down on the
tank bottom and the inside circumference is drawn.
A 1/4-inch hole is drilled on the inside of the drawn
circle to accommodate the metal-cutting blade of a
saber saw, and the circle is carefully cut.


Tank bottom


2" PVC standpipe



L 4" PVC pipe


-4"x3" PVC coupler


Slots


PVC closet flange
- 3"x2" PVC bushing

2" PVC drain pipe


Figure 4 Drain connection, center standpipe, and venturi
pipe for large brood fish holding tank.










Figure 5. An external standpipe controls
the water level in the tank. A net basket for
collecting the eggs for tank spawning.


Figure 6. A heavy knotted nylon
tank cover, attached to a hinged,
wood frame prevents fish from
jumping out and dying.


The drain can now be attached to the tank bot-
tom (Figure 4). The closet flange is secured to the
outside of the bottom of the tank with 3/16-inch
round head bolts. During assembly, silicone sealer
is applied under the bolt heads and between the
flange and the outside of the tank bottom.
It is advantageous to control water level by an ex-
ternal standpipe, especially for tanks 6 feet or
greater in diameter and for tank spawning (Figure
5). The drain opening in the tank need not be cov-
ered when holding large fish; however, a short
standpipe is useful to prevent complete draining
when water level is lowered prior to injecting or
handling the fish.
If an internal standpipe with a venturi pipe is
preferred, the internal stop of a 3-inch by 2-inch
PVC bushing is removed with a rotary rasp on an
electric hand drill, so a 2-inch pipe can extend
through the bushing (Figure 4). The bushing is
glued into the bottom of the closet flange with PVC
cement. The 2-inch drain line that extends below
the tank bottom is measured so that it occupies
only half of the length of the bushing area. The
upper half of the bushing will accommodate the 2-
inch diameter standpipe, which is held in place by
friction fit. The venturi pipe is constructed from a
3-inch by 4-inch coupler and a length of 4-inch
diameter PVC pipe. Slots are cut in the 3-inch
diameter end of the coupler which makes a tight
friction fit in the top of the closet flange.
Water is sprayed into the tanks through a par-
tially open gate valve at a 450 angle to the surface
to provide aeration and circular flow. The recom-
mended exchange rate of water is once per hour.
Tanks must be covered to prevent fish from jump-
ing to their deaths. Tank covers constructed of
heavy nylon netting attached to hinged wooden or
plastic frames are functional and inexpensive
(Figure 6). These covers must be securely fastened
to the tank.


Provisions must be made for photoperiod and
temperature control if the tanks are to be used to
condition brood fish for out-of-season spawning.
Standard incandescent lights controlled by electric
timers are recommended for photoperiod manipula-
tion. Water heaters and/or chillers may be neces-
sary to adjust water temperature, and tanks should
be insulated.

Incubating apparatus

Several types of incubators have been used to in-
cubate grass carp eggs in hatcheries: (1) hatching
jars (Figure 7); (2) conical baskets (Figure 8); (3)
conical vats (Figure 9); (4) screened conical vats (Fi-
gure 10); and (5) circular tanks (Figure 11).
The hatching jars, conical baskets, conical vats,
and screened conical vats all have rounded or coni-
cal bottoms, and all function in a similar manner
but differ in size and construction. Water is deliv-
ered through a tube to the bottom of the container.
The upwelling current keeps the eggs oxygenated,
suspended, and in constant motion. High water
pressure can cause a strong current or supersatu-
rated gases can cause tiny air bubbles that may
carry the eggs out of the incubators; water should
be thoroughly aerated and delivered by a low-pres-
sure manifold system (Figure 12).
Hatching jars have less than 2 gallons of volume
and can accommodate up to 50,000 eggs. Conical
baskets are constructed of fine-mesh netting. The
baskets are suspended in a tank of water and hold
approximately 70,000 eggs. Conical vats and
screened conical vats are much larger (50 gallons)
and can hold 800,000 eggs. These vats are usually
constructed of stainless steel, plastic, or fiberglass.
The larvae swim vertically after hatching and are
carried by water current from the hatching jar and
conical vat into screened tanks or aquaria (Figure
13). The drain standpipe in the tanks or aquaria


Brood fi:
holding













Water


Figure 7. Hatching jar for incubation.








Overflow



















Tripod stand


Water level









0.5mm mesh
netting


Funnel--



Figure 8. A conical basket for incubation.




Air supply 0.5mm me



Water discharge .





Ti f il 'hng '.
IPerforated air tung



Perforated air tuDlng g -


Figure 10. A screened conical vat for incubation.


Figure 9. A conical vat for incubation.








Air suppy


I i


supply


Gate valve

50 Ell


Figure 11. A circular tank for incubation.


Water discharge


/2" Gate valve

V2" Galvanized nipple
1/2" Galvanized nie 1/2" Threaded PVC Street Ell

1/2" Polypipe nipple
I" PVC Pipe
Clear flexible hose or tubing


Figure 12. A low-pressure water delivery manifold system for jars, baskets or vats.


Center


PVC pipe


pipe


cutouts


Reservoir tank






















Figure 13. A screened tank for retaining larvae.

are surrounded by a screen cylinder with a perfo-
rated airline attached around the base of the
screen. The screen pipe is constructed from a PVC
pipe, cut as illustrated, covered with polyester cloth
(50 mesh/inch or 0.5 millimiter), secured to the pipe
with silicone or hot glue and attached to the tank
bottom by tight friction fit or silicone cement.
The larvae in conical baskets and screened coni-
cal vats are retained after they hatch and are trans-
ferred by hand. Screens used for retaining grass
carp larvae should be 50 mesh/inch (0.5 millime-
ter).
Fiberglass or plastic circular tanks (4 feet in
diameter) have a much greater volume than the
other types of incubators (150 to 200 gallons) and
have flat bottoms with center drains. A circular
tank can accommodate over 1 million grass carp
eggs. The center screen pipe is constructed from
PVC pipe cut as illustrated in Figure 11 and cov-
ered with polyester cloth. The screen must be com-
pletely sealed to the pipe with hot glue or silicone
cement. Some circular tanks have a recessed area
in which the center screen pipe fits snugly; if not, it
must be securely glued with silicone cement to the
tank bottom. Fry can be lost through any small
holes or cracks. A perforated air hose is attached
around the base of the screen.
Although water is added to the tank (2 gallons
per minute), compressed air gently bubbled around
the center screen generates the water current re-
quired to keep eggs in suspension. In addition, the
air bubbles help to keep the center screen from
clogging and help provide oxygen for the developing
embryos. Vigorous aeration of eggs or larvae, how-
ever, can be damaging and potentially
lethal. Rough internal surfaces of the tank may
also result in damage to the eggs. Larvae are held
in the same tank in which they were hatched until
they utilize their yolk sac and are ready to start
feeding.
Round, metal cattle troughs, 4 feet in diameter,
may also be used for egg incubation. The trough is
painted with epoxy, as described in the previous
section ("Brood fish holding tanks"). A smaller
drain assembly, based on the sink basket-strainer


assembly, is used. A circle is drawn on the tank bot-
tom using the inside of the rubber gasket as a
template. A 1/4-inch hole is drilled on the inside of
the drawn circle to accommodate the blade of a
saber saw, and the circle is carefully cut in the
tank. A liberal amount of silicone sealer is applied
around the hole on the inside of the tank. The
drain basket is inserted from the inside of the tank
and the rubber gasket, paper gasket, and threaded
ring are attached from below. The threaded ring is
secured as tightly as possible and the excess
silicone sealer is smoothed around the flange to en-
sure a watertight seal. The external drain connec-
tion is made using a 1 1/2-inch diameter threaded
female PVC adaptor (Figure 14). The internal
standpipe is a 1 1/4-inch diameter PVC pipe with
several wraps of duct tape around the bottom to
make a tight friction fit in the sink drain basket.
The screened pipe is constructed from a 3 1/2-inch
long section of 3-inch diameter PVC sewer pipe, a 3-


1 1/4" PVC
standpipe


Duct tape


4" PVC sewer
pipe


S'Smm
mesh




*- 4"x3" coupler




3" PVC sewer pipe


1 1/2 PVC
- J threaded
female adaptor


Figure 14. A drainpipe connection for small tanks.









inch by 4-inch PVC sewer coupler, and a section of
4-inch diameter sewer pipe. The 3-inch diameter
sewer pipe makes a tight friction fit into the sink
drain basket; the opposite end slips into the coupler
which connects to the 4-inch diameter pipe. The 4-
inch center pipe is cut as illustrated and covered
with nylon screen.

Semi-intensive and intensive fry rearing
tanks and cages
The semi-intensive technique of fry rearing
utilizes tanks or cages as culture containers. The
same fiberglass or plastic tanks or galvanized metal
cattle troughs used for incubating eggs and holding
larvae can be used for fry rearing. The cages are
made of 42 mesh/inch (0.6 millimeter) polyester fab-
ric (Figure 15). Tanks and cages have cylindrical
center drain screens of 42 mesh/inch (0.6 millime-
ter). Perforated tubing around the bottom of the
screen delivers compressed air for aeration and re-
duces clogging of the screen.

COMPRESSED AIR


Figure 15. A cage for semi-intensive fry rearing in ponds.

Tanks are supplied with zooplankton-ladened
water pumped from a fertilized pond. The intake of
the pump is covered nylon screen to prevent intro-
duction of predators. Cages are suspended directly
in the pond and water is moved through the cage
with an airlift. Water turnover time is approxi-
mately 10 to 15 minutes.
Intensive fry rearing is usually conducted in
either glass aquaria or fiberglass, metal, concrete,
or plastic tanks. Although not absolutely necessary,
transparent containers are preferred for fry rear-
ing, because of the ease of observing the fry and
their food.
Water quality is maintained with airlift sponge
filters (approximately 5 by 5 inches square and the
height of the water depth minus 1 inch). A 1 1/4-
inch diameter hole is cut through nearly the entire
length of the sponge with a piece of metal tube
sharpened and notched as illustrated in Figure 16.
A glass, ceramic, plexiglass, or aluminum plate
glued to the bottom of the sponge with silicone ce-


AIR TUBING



SPONGE -


AIRLIFT CYLINDER -


LEAD WEIGHT
BASE


SERATIONS CUT WITH HACKSAW AND
EDGE SHARPENED WITH GRINDER


METAL CONDUIT->


Figure 16. Airlift sponge filters maintain water quality for inten-
sive fry rearing.
ment provides ballast for the filter, and an air tube
is inserted. Water is pumped through the sponge by
bubble airlift. One filter (5 by 5 by 7 inches) is used
for approximately every 20 gallons of water. A
gradual water exchange (one turnover every 48
hours) will further improve water quality without
flushing too much of the food from the tank. Drip ir-
rigation emitters will maintain this constant, slow
water flow. Fry rearing containers should be placed
in an area that is protected from direct sunlight
and is convenient for feeding and observation.

Brood Fish Conditioning
Grass carp must attain a certain degree of matur-
ity and ripeness before induced spawning can pro-
ceed successfully. In the southern United States,
grass carp become sexually mature in 2 to 3 years;
4 to 6 years are required in northern areas. In
Florida, the spawning season for grass carp is dur-
ing the months of April and May. In more northern
latitudes of the United States, the spawning season
is in May and June.
Although grass carp normally spawn in the
spring, the preceding summer and fall is the time
during which egg and sperm development occurs in
preparation for spawning. This point is of utmost
importance. If the brood fish are not properly con-
ditioned during this period, induced spawning will
be unsuccessful. The hormone-induced spawning
technique does not make eggs and sperm, it only
triggers the release of eggs and sperm that are al-
ready in an advanced stage of development.
Proper nutrition and suitable water quality play
a major role in conditioning. Grass carp to be used
for spawning should be reared in vegetated ponds
and fed floating catfish pellets during summer and
fall to ensure good condition. When aquatic plants
are limited, brood fish should be fed fresh terres-
trial vegetation and floating catfish pellets to attain
prime spawning condition.










Studies have shown that plankton is the primary
natural food of silver carp and bighead carp. Brood
fish ponds should be fertilized to stimulate
plankton production. The fish's diet should also be
supplemented with minnow-meal feed.

Brood Fish Capture and Handling
Grass carp, as well as the other Chinese carps,
are extremely excitable and ae spectacular jump-
ers; they must be handled carefully to minimize
physical injury and stress. The importance of this
point cannot be overemphasized. Cumulative physi-
cal stress to the fish can have a greater detrimental
effect on spawning success than almost any other
factor. Speed and gentleness of handling are of ut-
most importance. Conditions of low oxygen and
high temperatures must be prevented. Female
brood fish ready for spawning are in a particularly
delicate condition and should be treated accord-
ingly.
Grass carp for spawning can be captured by sein-
ing the holding pond. Always check dissolved oxy-
gen before seining; if levels are below 4 ppm, the
pond should be aerated, flushed with fresh water,
and the capture of the brood fish should be delayed
until conditions improve. Bag seines with extra
lead weight are more effective for grass carp cap-
ture than straight seines. Brood fish should be cap-
tured and transported only during the cool of early
morning hours. Partial draining of the pond can
simplify capture by seining; however, during warm
months, drain only at night. The brood fish pond
level should not be lowered during the heat of the
day; shallow water heats up rapidly, stressing the
fish. Dissolved oxygen should be checked frequently
during draining. If dissolved oxygen drops below 4
ppm, stop draining, refill the pond, and aerate.
Handling of brood fish should be kept to an abso-
lute minimum. Knitted 1/2-inch mesh dip nets
(Figure 17) are recommended for handling grass
carp to minimize scale loss. When transferring


1/4 INCH X 1 1/2 INCH ALUMINUM
BENT TO SHAPE


Figure 17. Knitted mesh dip nets for handling brood fish.


brood fish from bag seine to transport to holding
tanks, work as quickly as possible. A few seconds of
time spent with the fish out of the water in the dip
net can mean the difference between a perfect
spawn, partial spawn, or no spawn. This situation
is understandable when you consider that fish are
basically in a zero gravity environment, cushioned
on all sides by water. When fish are lifted from the
water, their full weight is now resting on their
gonads and damage can easily result. Ideally, brood
fish should be placed in net bags in containers of
water during transfer.
Transport tanks should be large enough to allow
complete freedom of movement to the brood fish
and have no sharp corners or edges to injure fish.
Transport units should be equipped with 12-volt
agitators and bottled oxygen with diffusers. Fish
must be protected from rapid temperature changes;
ice may be added to the water during hauling to
prevent an increase in water temperature; one-half
pound of ice will reduce the temperature of one gal-
lon of water by about 10F. Do not transport brood
fish long distances during spawning season; plan to
obtain the fish during the preceding year. A
prophylactic treatment of one part per thousand
salt and Nitrofurazone at 5 to 10 ppm active ingre-
dient should be used in the transport water to
minimize bacterial infection. Federal law prohibits
the use of Nitrofurazone on foodfish. A dip treat-
ment of copper sulfate (500 ppm) for up to 2
minutes has also been shown to be an effective
bacterial control for brood fish. Successful spawns
are impossible if fish are dying from infection.
Water temperature must exceed 68F for success-
ful induced spawning. Temperatures above 80OF,
however, may have a detrimental effect on spawning
success. Holding brood fish in ponds at spawning
temperature for an extended period results in re-
sorption of the eggs in the ovary and failure to
spawn; therefore, induced spawning should be con-
ducted in the shortest period of time possible.
Brood female grass carp captured from the pond
when temperature exceeds 68F should be injected
within 1 day; further delay may result in resorption
of eggs and failure to spawn. All suitable brood
females captured in the seine should be brought in
for injection and spawning. The stress of capture
may result in resorption of eggs, even if the fish is
released back into the pond. If additional brood fish
are still in the pond after seining, the pond should
be flushed with fresh water to counteract the ef-
fects of disturbing the bottom sediments on reduced
dissolved oxygen and release of toxic hydrogen sul-
fide to the water.
To minimize stress, do not weigh brood females
using a platform or hanging scale before injections.
It is better to estimate weight when determining
hormone dosage than inflict damage to the fish.
However, if neccesary, fish should be weighed in a
net bag in a container of water on the scale.








Determination of Gender
The gender of the brood fish is determined by ex-
ternal characteristics that manifest themselves just
prior to and during the spawning season. Male
grass carp can be distinguished by the sandpaper-
like texture of the dorsal surface of the pectoral
fins. The male silver carp and bighead carp have
numerous sharp spines along the back side of the
thickened first ray of the pectoral fins. Milt can usu-
ally be stripped (gently squeezing the area in front
of the vent) from the male fish a month or more be-
fore the females are ready for spawning.
The pectoral fins of female fish are slightly short-
er, less robust, and are smooth to the touch. Female
fish ready for spawning have soft, full abdomens;
fat deposits in the body cavity of females may, how-
ever, give the appearance of full, ripe ovaries. There-
fore, it is difficult, if not impossible, for the hatch-
ery worker to determine by external appearance the
suitability of brood females for induced spawning.
Once the sex of the fish has been determined, the
fish should be marked by fin clips, tags or some
other means to indicate gender during periods
when fish are not exhibiting external sex charac-
teristics.

Controlled Maturation in Indoor
Tanks
The reproductive cycles of fish can be altered by
artificial manipulation of environmental conditions.
Temperature and photoperiod are important factors
in gonadal maturation and spawning. Studies at
our laboratory indicate that grass carp captured
during the winter when the gonads are in the rest-
ing stage, and held in indoor tanks under condi-
tions of artificial photoperiod and temperature, un-
dergo the final stages of development leading up to
hormone-induced spawning. By manipulating these
variables, spawning can be accomplished during
months other than April and May. The hatchery
manager then has the opportunity to schedule
spawning to optimize use of the hatchery facility
and staff, production costs, food supply, and time of
stocking for weed control.
Grass carp brood fish brought indoors in good
condition during the winter can be starved several
months prior to induced spawning and still develop
mature ova and sperm. The fish utilizes the fat de-
posited in the body cavity as an energy source dur-
ing periods of limited food intake.

Early spawn
To produce grass carp fry early in the calendar
year, brood fish should be transferred from ponds to
the indoor, environmentally controlled circular
tanks during early winter when pond water tem-
perature is below 55F. At this temperature, the fish


are easier to capture and less likely to injure them-
selves. Initial water temperature in the tanks
should be identical to outdoor pond temperature.
The cold temperature acts as a biological check-
point; final ripening of the eggs and sperm proceeds
when temperature increases. Photoperiod should be
set for long-day regime (16 hours of light), and the
water warmed gradually (2F/day) until tempera-
ture is between 70'F to 75F Brood fish are held
under these conditions for approximately 45 days
prior to initiating the induced-spawning procedure.
Fish can be held for an additional 75 days and still
be successfully spawned. Temperatures greater
than 800F will cause female fish to resorb ova, re-
sulting in reduced spawning success.
Late spawn
Delayed spawning of grass carp can be accom-
plished by artificially perpetuating winter tempera-
ture (less than 50F) and short-day photoperiod
regime (8 hours of light). Brood fish should be cap-
tured in late winter or early spring (temperature
less than 550F) and transferred to the environmen-
tally controlled circular tanks. Approximately 60
days prior to scheduled spawning, photoperiod
should be increased to 16 hours of light, and tem-
perature should be elevated 2F/day to 70F to 75F
and held constant for 45 to 120 days until hormone-
induced spawning.

Normal-season spawn
In addition to out-of-season spawns, indoor tanks
can also be used for final ripening of brood fish
for hormone-induced spawning during the normal
season. Brood fish are captured from the ponds in
late winter or early spring when water temperature
is less than 550 Temperature and photoperiod are
allowed to increase with ambient conditions. In-
duced spawning can begin 45 days after the temper-
ature increases to 70F or above. Bringing fish in-
doors at this time separates the stress of capture
from the stress of spawning, allowing fish the op-
portunity to heal and recover from injuries sus-
tained during capture. The result is more consis-
tent spawns due to reduced stress, as compared to
the traditional method of capture from ponds just
prior to induced spawning.


Sampling the Eggs in the Ovary
Female brood fish whose ovaries have not yet
reached the pre-ovulation stage following the com-
pletion of vitellogenesis will not ovulate success-
fully. During the process of pre-ovulation, the ger-
minal vesicle or nucleus migrates from the center
toward the micropyle (opening through which the
sperm enters the egg), and the egg absorbs fluids
(hydrates) and swells in size.








Determination of Gender
The gender of the brood fish is determined by ex-
ternal characteristics that manifest themselves just
prior to and during the spawning season. Male
grass carp can be distinguished by the sandpaper-
like texture of the dorsal surface of the pectoral
fins. The male silver carp and bighead carp have
numerous sharp spines along the back side of the
thickened first ray of the pectoral fins. Milt can usu-
ally be stripped (gently squeezing the area in front
of the vent) from the male fish a month or more be-
fore the females are ready for spawning.
The pectoral fins of female fish are slightly short-
er, less robust, and are smooth to the touch. Female
fish ready for spawning have soft, full abdomens;
fat deposits in the body cavity of females may, how-
ever, give the appearance of full, ripe ovaries. There-
fore, it is difficult, if not impossible, for the hatch-
ery worker to determine by external appearance the
suitability of brood females for induced spawning.
Once the sex of the fish has been determined, the
fish should be marked by fin clips, tags or some
other means to indicate gender during periods
when fish are not exhibiting external sex charac-
teristics.

Controlled Maturation in Indoor
Tanks
The reproductive cycles of fish can be altered by
artificial manipulation of environmental conditions.
Temperature and photoperiod are important factors
in gonadal maturation and spawning. Studies at
our laboratory indicate that grass carp captured
during the winter when the gonads are in the rest-
ing stage, and held in indoor tanks under condi-
tions of artificial photoperiod and temperature, un-
dergo the final stages of development leading up to
hormone-induced spawning. By manipulating these
variables, spawning can be accomplished during
months other than April and May. The hatchery
manager then has the opportunity to schedule
spawning to optimize use of the hatchery facility
and staff, production costs, food supply, and time of
stocking for weed control.
Grass carp brood fish brought indoors in good
condition during the winter can be starved several
months prior to induced spawning and still develop
mature ova and sperm. The fish utilizes the fat de-
posited in the body cavity as an energy source dur-
ing periods of limited food intake.

Early spawn
To produce grass carp fry early in the calendar
year, brood fish should be transferred from ponds to
the indoor, environmentally controlled circular
tanks during early winter when pond water tem-
perature is below 55F. At this temperature, the fish


are easier to capture and less likely to injure them-
selves. Initial water temperature in the tanks
should be identical to outdoor pond temperature.
The cold temperature acts as a biological check-
point; final ripening of the eggs and sperm proceeds
when temperature increases. Photoperiod should be
set for long-day regime (16 hours of light), and the
water warmed gradually (2F/day) until tempera-
ture is between 70'F to 75F Brood fish are held
under these conditions for approximately 45 days
prior to initiating the induced-spawning procedure.
Fish can be held for an additional 75 days and still
be successfully spawned. Temperatures greater
than 800F will cause female fish to resorb ova, re-
sulting in reduced spawning success.
Late spawn
Delayed spawning of grass carp can be accom-
plished by artificially perpetuating winter tempera-
ture (less than 50F) and short-day photoperiod
regime (8 hours of light). Brood fish should be cap-
tured in late winter or early spring (temperature
less than 550F) and transferred to the environmen-
tally controlled circular tanks. Approximately 60
days prior to scheduled spawning, photoperiod
should be increased to 16 hours of light, and tem-
perature should be elevated 2F/day to 70F to 75F
and held constant for 45 to 120 days until hormone-
induced spawning.

Normal-season spawn
In addition to out-of-season spawns, indoor tanks
can also be used for final ripening of brood fish
for hormone-induced spawning during the normal
season. Brood fish are captured from the ponds in
late winter or early spring when water temperature
is less than 550 Temperature and photoperiod are
allowed to increase with ambient conditions. In-
duced spawning can begin 45 days after the temper-
ature increases to 70F or above. Bringing fish in-
doors at this time separates the stress of capture
from the stress of spawning, allowing fish the op-
portunity to heal and recover from injuries sus-
tained during capture. The result is more consis-
tent spawns due to reduced stress, as compared to
the traditional method of capture from ponds just
prior to induced spawning.


Sampling the Eggs in the Ovary
Female brood fish whose ovaries have not yet
reached the pre-ovulation stage following the com-
pletion of vitellogenesis will not ovulate success-
fully. During the process of pre-ovulation, the ger-
minal vesicle or nucleus migrates from the center
toward the micropyle (opening through which the
sperm enters the egg), and the egg absorbs fluids
(hydrates) and swells in size.








Determination of Gender
The gender of the brood fish is determined by ex-
ternal characteristics that manifest themselves just
prior to and during the spawning season. Male
grass carp can be distinguished by the sandpaper-
like texture of the dorsal surface of the pectoral
fins. The male silver carp and bighead carp have
numerous sharp spines along the back side of the
thickened first ray of the pectoral fins. Milt can usu-
ally be stripped (gently squeezing the area in front
of the vent) from the male fish a month or more be-
fore the females are ready for spawning.
The pectoral fins of female fish are slightly short-
er, less robust, and are smooth to the touch. Female
fish ready for spawning have soft, full abdomens;
fat deposits in the body cavity of females may, how-
ever, give the appearance of full, ripe ovaries. There-
fore, it is difficult, if not impossible, for the hatch-
ery worker to determine by external appearance the
suitability of brood females for induced spawning.
Once the sex of the fish has been determined, the
fish should be marked by fin clips, tags or some
other means to indicate gender during periods
when fish are not exhibiting external sex charac-
teristics.

Controlled Maturation in Indoor
Tanks
The reproductive cycles of fish can be altered by
artificial manipulation of environmental conditions.
Temperature and photoperiod are important factors
in gonadal maturation and spawning. Studies at
our laboratory indicate that grass carp captured
during the winter when the gonads are in the rest-
ing stage, and held in indoor tanks under condi-
tions of artificial photoperiod and temperature, un-
dergo the final stages of development leading up to
hormone-induced spawning. By manipulating these
variables, spawning can be accomplished during
months other than April and May. The hatchery
manager then has the opportunity to schedule
spawning to optimize use of the hatchery facility
and staff, production costs, food supply, and time of
stocking for weed control.
Grass carp brood fish brought indoors in good
condition during the winter can be starved several
months prior to induced spawning and still develop
mature ova and sperm. The fish utilizes the fat de-
posited in the body cavity as an energy source dur-
ing periods of limited food intake.

Early spawn
To produce grass carp fry early in the calendar
year, brood fish should be transferred from ponds to
the indoor, environmentally controlled circular
tanks during early winter when pond water tem-
perature is below 55F. At this temperature, the fish


are easier to capture and less likely to injure them-
selves. Initial water temperature in the tanks
should be identical to outdoor pond temperature.
The cold temperature acts as a biological check-
point; final ripening of the eggs and sperm proceeds
when temperature increases. Photoperiod should be
set for long-day regime (16 hours of light), and the
water warmed gradually (2F/day) until tempera-
ture is between 70'F to 75F Brood fish are held
under these conditions for approximately 45 days
prior to initiating the induced-spawning procedure.
Fish can be held for an additional 75 days and still
be successfully spawned. Temperatures greater
than 800F will cause female fish to resorb ova, re-
sulting in reduced spawning success.
Late spawn
Delayed spawning of grass carp can be accom-
plished by artificially perpetuating winter tempera-
ture (less than 50F) and short-day photoperiod
regime (8 hours of light). Brood fish should be cap-
tured in late winter or early spring (temperature
less than 550F) and transferred to the environmen-
tally controlled circular tanks. Approximately 60
days prior to scheduled spawning, photoperiod
should be increased to 16 hours of light, and tem-
perature should be elevated 2F/day to 70F to 75F
and held constant for 45 to 120 days until hormone-
induced spawning.

Normal-season spawn
In addition to out-of-season spawns, indoor tanks
can also be used for final ripening of brood fish
for hormone-induced spawning during the normal
season. Brood fish are captured from the ponds in
late winter or early spring when water temperature
is less than 550 Temperature and photoperiod are
allowed to increase with ambient conditions. In-
duced spawning can begin 45 days after the temper-
ature increases to 70F or above. Bringing fish in-
doors at this time separates the stress of capture
from the stress of spawning, allowing fish the op-
portunity to heal and recover from injuries sus-
tained during capture. The result is more consis-
tent spawns due to reduced stress, as compared to
the traditional method of capture from ponds just
prior to induced spawning.


Sampling the Eggs in the Ovary
Female brood fish whose ovaries have not yet
reached the pre-ovulation stage following the com-
pletion of vitellogenesis will not ovulate success-
fully. During the process of pre-ovulation, the ger-
minal vesicle or nucleus migrates from the center
toward the micropyle (opening through which the
sperm enters the egg), and the egg absorbs fluids
(hydrates) and swells in size.








An egg sample may be taken from a small
number of the female fish to determine the stage of
maturity. A piece of Intramedic polyethylene tubing
of very small thickness (1.57 mm outer diameter),
slipped over the end of a 18-gauge needle, attached
to a syringe with a hole drilled in the barrel, is
used to obtain a sample of the eggs (Figure 18). Be-
fore sampling, the female fish can be quieted with
MS-222 if neccesary. The tube is gently inserted
into the oviduct, a transverse slit in the front of the
urogenital opening of the female fish (Figure 19).
The more developed a female fish is, the easier it is
to insert the tube and obtain an egg sample. Once
the tube is inserted into the ovary, the hole in the
barrel is covered with a finger, and a small amount
of suction is applied with the syringe to draw the
eggs into the tubing. The tube containing the egg
sample is then slowly withdrawn; the finger over
the hole in the barrel of the syringe is released be-
fore the end of the tube is removed from the
oviduct.


NEEDLE


SYRINGE


DRILLED HOLE


-. POLYETHYLENE TUBING


Figure 18. The device used for egg sampling.

Carp eggs that are ripe and ready for hormone-
induced spawning, are various shades of brown or
green; under the microscope, they appear uniformly
grey. Ripe eggs are approximately one millimeter or
larger in diameter; immature eggs are smaller. In
addition, the nucleus (germinal vesicle) of the ripe
egg is nearer one end rather then in the center; the
nucleus appears as an almost clear circle in the
egg. A cover slip is placed over the eggs on a glass
slide; the weight of the cover slip slightly compres-
ses the eggs. When illuminated from below, the pos-
ition of the nuclei can be observed with a 10X hand
lens or microscope (Figure 20). Once the eggs have
been determined to be ripe, the fish can be injected
with the hormones for spawning. Eggs that have
begun to resorb or break-down in the ovary appear
whitish in color; under the microscope, the egg con-
tents are irregular in composition and appear to
have pulled away from the egg shell. Once resorp-
tion of the ova begins, induced spawning will no
longer be successful.
Males should be checked for milt flow before
being selected for hormone-induced spawning.


VENT AREA
"Inv


ANUS


GENITAL OPENING




- URINARY OPENING


Figure 19. Sampling the eggs in the ovary of the fish is done
by inserting a tube into the genital opening.



Hormone Injections
Several hormone preparations have been used to
induce spawning of grass carp and other Chinese
carps. Regardless of the injections used, the gonads
must be in an advance stage of development and
the fish must be physically healthy in order for
spawning to be successful.
Every effort must be made to minimize stress to
the brood fish while administering the injections.
Injections usually require two hatchery workers;
however, an experienced technician can inject the
fish alone. The water depth in the holding tank is
lowered to reduce the chance of fish jumping out of
the tank during netting and injuring themselves.
The fish are captured in a dip net but are not re-
moved from the water to administer the injection.
Do not grab the fish by the caudal peduncle, be-
cause scales can be easily dislodged, opening a site
for infection.
During the preparation and use of hormones,
sterile disposable syringes and needles should be
used. Bacteriostatic or sterile water should be used
to reconstitute hormones. A tissue grinder, or
mortar and pestle, is used to mix carp pituitary
hormone. Reconstituted hormones are labeled and
stored in a freezer in 1- or 2-cc ampules. These
items should be boiled in water for 10 minutes be-
fore use, to minimize bacterial contamination.
The volume of the injection should be kept to a
minimum; 2 cc or less is optimum for grass carp.
Proportionally smaller injections are recommended
for smaller species of fish. It is more convenient to



























Figure 20. A. Sample of immature eggs from the ovary.


C. Egg that has undergone migration of the germinal vesicle
(nucleus).

prepare the hormone for several brood fish all
together and then administer by volume.
All four injection schedules presented below have
been successfully used to induce spawning of grass
carp at our laboratory.

Carp pituitary
The traditional hormone preparation used for in-
duced spawning of grass carp has been either fresh
or acetone-dried common carp (Cyprinus carpio)
pituitary. The pituitary gland acts as an inter-
mediary between the brain and the gonads. In
nature, spawning is regulated and initiated by the
gonadotropic hormone produced and stored in the
pituitary. The stored hormone is released into the
blood when conditions are favorable for spawning.
The injection of pituitary material for induced ovu-
lation and spawning "overrides" the fish's own
hormonal system.
Carp pituitary extract is the dried, powdered
pituitary glands of common carp. Dried carp pitui-
tary is normally purchased in 1-gram vials from
commercial suppliers. A hand tissue grinder or


B. Egg prior to migration of the germinal vesicle (nucleus).


D. Egg that has begun to breakdown in the ovary. The contents
seem to have pulled away from the shell.

mortar and pestle is needed to prepare the hor-
mone for injection. Weigh out 0.5 grams of dried
carp pituitary material, and pulverize with a dry
mortar and pestle or tissue grinder. Then add 10 cc
of bacteriostatic water a little bit at a time, mixing
thoroughly to produce a suspension. The hormone
of the carp pituitary goes into solution after 20 to
30 minutes, and the tissue residue should be sepa-
rated by centrifuge or by simply drawing off the
supernatant from above the settled solids. Only the
liquid above the settled solids is used for injections.
Divide the hormone preparation into 1 or 2-cc am-
pules, label, and store in the freezer or use im-
mediately to inject the fish. Carp pituitary is in-
jected intramuscularly (Figure 21) at the base of
the last ray of the dorsal fin with a 20-gauge nee-
dle.
Female grass carp are administered two injec-
tions of carp pituitary 18 to 24 hours apart. Total
dosage of carp pituitary administered is 2 to 4
milligrams/pound female body weight (0.4 to 0.8 cc
of the mixture described above for each 10 pounds
fish weight). The first injection is 1/10 and the


L~ "B ,-.,
"'
F, k 'C
luln
"i~i-
.I
.

.*.


-.

-.


Tf~5~, ~Lul'-
~
r f;
;i:
't ~e









DORSAL FIN SYRINGE



.. .. :. ,: i..;'


Figure 21. The hormones are administered by an intramuscu-
lar injection at the base of the last ray of the dorsal fin.

second, or resolving dose is the remaining 9/10 of
the total dosage.
Male grass carp are given one injection of carp
pituitary at the same time the female receives the
second injection. The dosage of carp pituitary for
male fish is 1 to 2 milligrams/pound body weight
(0.2 to 0.4 cc for each 10 pounds of fish weight).
The potency of carp pituitary is variable, depend-
ing upon its processing and the age and condition
of the donor fish. Variation in potency has made it
difficult to accurately quantify the dosages required
for induced spawning. Most authors suggest that
hormone should be given in excess to ensure an
adequate quantity of active ingredient to attain the
threshold level for spawning.
Grass carp usually spawn 7 to 12 hours after the
second dose of carp pituitary, depending on temper-
ature. Higher temperatures usually shortens the re-
sponse interval.

Human chorionic gonadotropin and carp
pituitary
Human chorionic gonadotropin (HCG) has been
successfully used to spawn some species of fish;
however, it will not induce ovulation in grass carp
when used alone. Hormone treatment with HCG is
useful for advancing the preovulation phase.
Female grass carp do respond favorably to two injec-
tions of HCG, followed by a resolving dose of carp
pituitary. Injections are administered 12 to 24
hours apart. HCG is measured in International
Units (IU), and usually comes in 10,000 IU vials
available from commercial sources.
Mix the contents of a 10,000 IU vial of HCG with
1 cc of sterile or bacteriostatic water (10,000 IU/cc).
The dose of the initial injection is 100 to 200 Inter-
national Units (IU)/pound body weight (0.1 to 0.2 cc
of the mixture described above for each 10 pounds
of fish weight), with 800 to 900 IU/pound body
weight administered in the second injection (0.8 to
0.9 cc for each 10 pounds of fish weight). HCG is
injected intramuscularly at the base of the dorsal
fin with a 20-gauge or smaller needle (Figure 21).
The third injection, carp pituitary, is prepared
and administered as described in the previous
section ("Carp Pituitary") at a rate of 2 to 4 milli-
grams/ pound body weight (0.4 to 0.8 cc of the


described mixture for each 10 pounds of fish
weight). Male grass carp are given 1 to 2 milli-
grams carp pituitary/ pound body weight at this
time (0.2 to 0.4 cc for each 10 pounds of fish
weight). Grass carp usually spawn 9 to 12 hours
after the carp pituitary injection.

Synthetic LH-RH analogue
Luteninizing hormone-releasing hormone (LH-
RH) is produced in the portion of the brain called
the hypothalmus. It acts on the pituitary to trigger
the release of gonadotropic hormone into the blood.
Mammalian LH-RH has been used to induce ovula-
tion of fish, but a comparatively large dose and
frequent injections were required. The synthetic
LH-RH analogue, des-GLY10 [D-Ala6]LH-RH ethyl-
amide (LH-RHa), has proven to be an even more
effective releasing hormone for fish. LH-RHa stimu-
lates the fish's own pituitary to produce the
gonadotropic hormone necessary for ovulation or
spermiation, rather than circumventing the endo-
crine system with crude pituitary materials from
donor fish or gonadotropic hormones derived from
other species.
Studies conducted at the University of Florida
concluded that a single injection of LH-RHa is effec-
tive, reliable, and safe for induced spawning of
grass carp. In the study, 92% of female grass carp
three years and older spawned after a single injec-
tion of LH-RHa. The percentage of viable embryos
was good (90% for hand spawning), suggesting good
egg quality. The use of LH-RHa for induced spawn-
ing of grass carp is strongly recommended. This
material is effective, requires only a single injec-
tion, and is easier to use and less costly than other

hormone preparations.
LH-RHa is available in 1-milligram or 5-millig-
ram vials from commercial sources. The unopened
vial of hormone should be stored in a refrigerator or
freezer. The contents of the 5-milligram vial should
be mixed with 10 cc of sterile or bacteriostatic
water (1-milligram in 2cc) and divided into small
volumes to be either used immediately or kept in a
freezer, otherwise its level of activity may be re-
duced. Store the reconstituted hormone in 1 or 2 cc
ampules so that only the required amount need be
defrosted, saving the potency of the remaining
quantity. Caution should be used while reconstitut-
ing, using, and storing the material to minimize
the possibility of microbial contamination.
The recommended dosage is 5 micrograms/pound
of female fish (0.2 cc of the mixture described above
for each 15 to 20 pounds of fish body weight).
Dosages of 0.5 to 50 micrograms/pound have been
successful, so there is a wide safety margin. Males
are given one-half the dose administered to the
females. The hormone is injected intramuscularly
at the base of the dorsal fin with a 20-gauge or
smaller needle.









Response interval (time between injection and
spawning) for female grass carp receiving a single
dose of LH-RHa varies from 13 to 24 hours.
Response time is negatively correlated with water
temperature; a shorter response interval is ob-
served with warmer water.
Some cyprinids, such as the bighead carp, do not
respond to a single injection of LH-RHa. Research
has shown that two injections 6 to 10 hours apart,
each injection made up of one-half the total dosage,
results in ovulation. Two injections produce a
greater amplitude of gonadotropic hormone in the
blood of the fish than a single injection of the same
total dosage.

Synthetic LH-RH analogue and dopamine
blocker
Dopamine is also produced by the brain and like
LH-RH, it controls the release of gonadotropic
hormone from the pituitary. Unlike LH-RH,
dopamine inhibits the release of hormone from the
pituitary, functioning as a feed back mechanism,
shutting down the pituitary. There is a family of
drugs that act as dopamine blockers, either by
preventing the release or by inhibiting the absorp-
tion of dopamine. Research has shown that the
combination of a dopamine blocker with LH-RHa
greatly improves the release of gonadotrophic hor-
mone from the pituitary. Some species of cyprinids,
such as redtailed black shark and rainbow shark,
do not respond at all to injections of LH-RHa. In-
duced spawning has been successful, however, when
a dopamine blocker is used with LH-RHa.
Some of the dopamine blockers that have been
used for fish are reserpine, pimozide, domperidone,
and haloperidol. We initially used reserpine (23 mg/
pound body weight of fish), but we have switched to
haloperidol because of its greater potency. Halo-
peridol powder, can be put into solution by mixing
50 milligrams with 10 cc bacteriostatic water and
acidifying with lactic acid to a pH of 3.0 to 3.6. Use
pH paper to determine the end point. Haloperidol
is injected intramuscularly at a rate of 0.23 mg/
pound of fish (0.45 cc of the above mixture for each
10 pounds body weight). The dopamine blocker is
administered with either a single injection of LH-
RHa or with the first of two LH-RHa injections. We
have used haloperidol with LH-RHa to spawn grass
carp, bighead carp, redtailed black shark, rainbow
shark, and shovel nose sturgeon.

Spawning
Hand stripping/dry fertilization
Ovulation is a complicated process lasting several
hours; the exact duration is controlled primarily by
temperature. Ovulation starts with the disappear-
ance of the nuclear membrane and the appearance
of the chromosomes, leading to the first meiotic di-


vision. At this time, the follicle, which attaches the
egg to the ovary, splits and partially dissolves, re-
leasing the egg. The eggs may then flow from the
ovary through the genital opening.
The traditional method of taking the spawn of
grass carp has been the hand stripping/dry fertili-
zation procedure. Although the procedure for tank
spawning of grass carp, developed more recently, is
much simpler and less strenuous on brood fish and
hatchery workers, hand stripping techniques must
still be used to produce triploid grass carp. Three
hatchery workers are usually required to take the
spawn.
Brood fish are separated by sex prior to hormone
injection to prevent spawning in the holding tank,
when hand stripping is desired. Individual fish may
be identified by plastic tags, fin clips, or distin-
guishing scars or marks.
One hour before earliest anticipated spawn,
female fish are checked for readiness to spawn.
Female fish are checked every hour until ovulation
is verified. During ovulation, the eggs detach from
the ovarian tissue, thereby severing the oxygen
source supplied by the blood of the female. If the
eggs remain in. the female, they become anoxic and
egg quality deteriorates. The maximum grace
period between ovulation and taking the eggs is
approximately 30 to 40 minutes.
Handling should be kept to an absolute
minimum. Frequent or rough handling of female
brood fish retards ovulation, reduces success, and
increases mortality. Checking the fish does not
require that it be removed from the water. The
hatchery workers capture the fish in a dip net, turn
it belly up, and restrain it by holding the pectoral
fins. The abdomen of the female fish is gently
stroked with the fingertips (Figure 22). Do not try
to squeeze or force the eggs from the fish; this will
only injure the female and reduce success. If no
eggs are obtained, the fish is released to the tank
and examined again after another hour. If only a
few eggs flow from the vent when slight pressure is


Figure 22. Verifying ovulation is done by gently stroking the
abdomen and observing if eggs flow freely from the vent.








applied, partial ovulation has occurred and the fish
should be released and checked again 20 to 30
minutes later. Attempting to hand strip a female
fish that has only partially ovulated will result in
few eggs and will usually damage the ovaries so
that a complete spawn will not occur.
When the female grass carp is ready to release
her eggs, the muscles of the fish tighten and begin
to quiver; the vent appears to enlarge. Within sec-
onds, a stream of eggs nearly 1/2 inch in diameter
is ejected from the vent by the contraction. The first
hatchery worker immediately plugs the flow of eggs
by placing a thumb over the vent. The second
worker lifts the fish in the dip net while the first
worker holds the fish by the caudal peduncle and
continuously holds their thumb over the vent until
the eggs are taken from the fish. The female fish in
the dip net is immediately placed in a plastic trash
can filled with water and MS-222, until she no
longer struggles (three tablespoons of MS-222 are
used for 30 gallons of water). The first worker sits
on a chair next to the container of anesthetic, hold-
ing the vent closed. The anesthetized fish is then
lifted onto a dry towel spread on the lap of the first
worker and then thoroughly dried with the towel.
The third worker wipes the water and slime from
the vent and tail area and then holds a clean, dry
pan to catch the eggs. The fish is held on her side,
slightly tail down, and the stream of eggs is di-
rected into the pan (Figure 23). The pan is held so
that water from the fish does not drip onto the eggs
in the pan.


Figure 23. The stream of ovulated eggs from the female is
directed into a dry pan or other container.
Ripe spawn flows readily from the genital open-
ing of the female. The eggs are not forcibly
squeezed from the fish; slight pressure is applied to
the anterior portion of the abdomen to assist in
emptying the ovaries. Good spawn has little ovarian
fluid and is grayish-green to brownish-orange. Over-
ripe spawn is watery and many eggs are cloudy
white.
Female grass carp can produce in excess of 1 mil-
lion eggs. One milliliter of unfertilized grass carp
spawn contains 800 to 1,000 eggs. Approximately


the same number of eggs are contained in 1 gram
of unfertilized spawn. Estimation of the quantity of
spawn obtained from each female is determined by
measuring the volume or weight.
Speed is not essential when fertilizing the eggs;
dry eggs can be fertilized within a 30-minute period
without affecting percent fertilization. However, the
spawning team must ensure that absolutely no
water comes in contact with the eggs before adding
the milt. If water accidentally comes in contact
with the eggs before the milt has been added, the
egg begins to swell and the micropyle closes, pre-
venting the sperm from penetrating the egg. The
male is captured, wiped off, and held belly down
over the pan containing the eggs. The portion of the
abdomen posterior to the pelvic fins is gently mas-
saged to extrude the milt on the eggs. Good quality
milt is white in color and creamy in texture.
Milt can be collected from the males and stored
prior to stripping the eggs. Male fish are captured
in a dip net and anesthetized. The fish is then
turned belly up, and the vent area dried by blotting
with a towel. The area just behind the pelvic fins is
gently massaged toward the vent to extrude the
milt. The first few drops of milt are not used. The
milt is collected by inserting a plastic tube attached
to a syringe into the urogenital opening, and suc-
tion is applied while stripping to draw the milt into
the syringe. Care must be taken to ensure that
water, urine, intestinal contents, slime, or blood are
not mixed with the milt. It is best to collect and
store milt separately from each male to avoid con-
tamination. The milt is expelled into a plastic bag;
50 micrograms of dry streptomycin sulfate per mil-
liliter of milt is added to control bacteria. The bag
is filled with oxygen and sealed with a rubber band
and the milt is then stored on ice in a cooler or
refrigerator. Do not freeze milt, as this will kill the
sperm. Milt of some species of fish can be held in
this manner for approximately 3 weeks.
The milt of two or more males should be used to
ensure fertilization of the spawn. The fresh milt is
spread over the eggs and thoroughly mixed; only
then is water added to activate the sperm. Only
enough water is added to completely cover the eggs.
Stored milt should be mixed with water first and
then shaken for 5 seconds before being added to the
pan with the eggs. The sperm remains alive and
active in water for less than 1 minute. The pan con-
taining the eggs should be gently rocked during
this period to ensure uniform distribution of the
sperm.
The fertilized eggs are transferred to the hatch-
ing apparatus after a minute or two. Care should
be taken not to allow the fertilized eggs to be ex-
posed to direct sunlight. Subdued lighting should
be used in the spawning and incubation area. Dur-
ing development, the eggs water-harden (take up
water and expand). The diameter of the unfertilized
egg is 1 to 1.2 millimeters; during incubation, they
swell to 5 millimeters or more.









Tank spawning
The procedure for tank spawning or natural fer-
tilization of hormone-induced grass carp has been
developed at the University of Florida. Brood fish
are injected with the hormone as in the hand
stripping/dry fertilization method, but the eggs and
milt are not manually stripped. Fish of both sexes
are placed in a 6-feet diameter or larger circular
tank with a freshwater flow of one exchange per
hour spraying into the tank to produce water cur-
rent and aeration. The male fish stimulate the
female by rubbing their roughened pectoral fins
along her abdomen and by butting her with their
heads. Eggs are released in the water when the
female fish is physiologically ready, and the male
fish fertilize the spawn. A 6-feet diameter tank
with two or three males is used for each female.
Group spawning can be accomplished if the tank is
of sufficient size to accommodate many individuals.
Too many breeders in a small tank, however, may
disturb each other.
Better fertilization can be expected if male fish
are accustomed to the tank, uninjured, and have
been injected with a preparatory dose of hormone
the previous week. This is in addition to the injec-
tion they receive at the same time as the female.
The same males can be used for spawning time
after time. Brood fish should not be disturbed fol-
lowing injectionss, and subdued lighting is recom-
mended. Foam is observed on the surface of the
water immediately after the spawning act.
Water flow from the spawning tank carries the
semi-buoyant fertilized eggs through the drain pipe
and into a fine-mesh net basket, where they are
dipped out for transfer to the hatching apparatus.
Gentle bubble-aeration is recommended around the
net basket to keep the eggs in suspension. A small
mesh aquarium dip net is used to transfer the eggs
from the net basket to a pan or bucket, and the
eggs are immediately transported to the hatching
apparatus. Always avoid bright light when transfer-
ring and incubating eggs.
Estimating the number of eggs in tank spawning
is accomplished by taking samples during the pro-
cess of egg transfer to determine the mean volume
per 1,000 eggs and recording the total volume of
water-hardened eggs placed in the hatching ap-
paratus. Egg number can also be estimated by tak-
ing several volumetric samples from the hatching
apparatus, counting the number of eggs per sam-
ple, and extrapolating to the volume of the hatch-
ing apparatus.
The tank spawning procedure results in a signifi-
cant reduction in labor requirements and injury to
brood fish. Three hatchery workers are required for
hand stripping, while a single worker can handle
many tank spawns without difficulty. Brood fish
that have been tank spawned are injured far less
than those that are manually stripped. In addition,


the tank-spawning method does not require hatch-
ery personnel skilled in predicting time of spawn-
ing, and the danger of overripening of eggs in the
ovary is avoided. Fertilization percentage may be
slightly lower for tank spawns, as compared to
hand spawns by an experienced hatchery team;
however, the advantages of tank spawn far out-
weigh the diminished fertilization. Hatcheries pro-
ducing diploid grass carp are advised to try the
tank spawning technique.

Triploid Grass Carp
Production techniques for triploid fish
Fear of uncontrolled reproduction of grass carp
has prompted research into production of triploid
fish for aquatic weed control. The triploid has a
chromosome count of 72, as compared to 48 for the
parent species. It is thought that the sterility of
induced triploid fish is a result from the failure of
synapses during meiosis as a result of the third set
of chromosomes.
Production techniques for triploid grass carp use
the hand stripping/dry fertilization procedures.
Pressure shock, heat shock, cold shock, or chemical
shock applied to the embryo shortly after fertiliza-
tion promotes retention of the second polar body,
resulting in the additional set of chromosomes in
triploid progeny. Pressure shock and heat shock
have been the most successful of the techniques re-
ported for the production of triploid grass carp, as
well as salmonids and other fish species.
Hydrostatic pressure appears to be one of the
most consistent methods for producing triploid
grass carp. Pressure shocks are administered using
a stainless steel cylindrical vessel closed by a brass
piston fitted with an O-ring. The piston is also fit-
ted with a pressure gauge and relief valve. An
external hydraulic press is used to apply pressure
to the piston (Figure 24). The optimum parameters
for this technique are reported to be 7,000 to 8,000
pounds per square inch for a duration of 1 to 2
minutes, starting 4 minutes after water is added to







Figure 24. Hydrostatic pressure
chamber and press used to treat
fertilized eggs to produce
triploid grass carp.








the eggs and sperm in the temperature range of
74F to 760F. Decompression is instantaneous at the
end of the treatment, and the eggs are then trans-
ferred to the incubation apparatus.Treatments
using these parameters have resulted in near 100%
triploid fish and a hatch relative to controls of
nearly 70%. Success in the production of triploid
fish may be adversely affected by poor egg quality,
temperature variation, and sloppy spawning
technique. When the percent of viable embryos of
the untreated group is less than 60%, due to poor
egg quality, few if any triploid fish are produced by
pressure treatment.
Temperature shocks are administered via water
bath in insulated tanks or large ice chests. Therm-
ostatically controlled submersible heaters or chill-
ers are used to adjust water temperature to the
desired level. Fertilized eggs are placed in small-
mesh dip nets or net baskets, and immersed at the
proper time in the aerated treatment bath. Nets
containing the eggs are swirled to ensure uniform-
ity of treatment to all the eggs. At the end of the
treatment, eggs are immediately placed in the incu-
bation apparatus with ambient water temperature.
The reported heat shock parameters vary from
1080F for 1 minute, starting 4 minutes after water
was added to the eggs and sperm, to 1010F for 3.5
minutes, starting 1 minute after the addition of
water. Although these treatments are somewhat
successful, there appears to be a great deal of vari-
ability and inconsistency in the production of trip-
loid grass carp by heat shock.

Verification of Triploidy
Each individual grass carp must be determined
to be a triploid by analyzing their red blood cell
erythrocytee) nuclear volumes, using an electronic
particle analyzer, before they can be stocked in
Florida waters. The size range for blood cells is
determined by calibration with standard particles.
Diploid grass carp are reported to have a nuclear


j.- -.

Figure 25. Obtaining a blood sample from each grass carp
to be analyzed for triploidity before stocking in Florida's waters.


volume of 10.06 cubic micrometers, while mean
triploid nuclear volume is 14.82 cubic micrometers.
Blood samples are taken from grass carp greater
than 2 inches long. Fish may be anesthetized if
necessary. The head of the fish is tilted back and
the branchial artery on the isthmus is punctured
with a blood lancet rinsed in EDTA between sam-
ples. Approximately 1 microliter of blood is with-
drawn, using a positive displacement micropipette
with disposable polypropylene tips (Figure 25). The
blood sample is immediately dispersed into an
Accuvette vial filled with 10 milliliters of Isoton II
electrolyte and one drop of Zapoglobin lysing agent.
The sample is processed immediately while the fish
is held in a net container suspended in water.
Diploid individuals are discarded.

Care of Spawn in Incubators
Good water quality is a necessity for incubating
eggs and holding larvae. Water used for incubation
must be filtered if particulate matter or predatory
animals are present. During the initial stages of
development, oxygen consumption is negligible; as
development progresses, the oxygen demand in-
creases considerably. Also during their develop-
ment, the eggs excrete carbon dioxide and am-
monia. If these products are allowed to accumulate,
they can kill the embryos.
The developing embryos needs a continuous sup-
ply of water, nearly saturated with oxygen. A con-
tinuous flow of water through the incubator will
also remove the carbon dioxide and ammonia. We
strongly advise the use of well water for incubating
eggs. Thorough aeration of the water before it is de-
livered to the incubators is required to provide suffi-
cient oxygen. The use of aerated well water in a
single use, flow-through incubation system elimi-
nates many problems. Recirculated water is not re-
commended for egg incubation.
Normal development of the embryos requires
water of the appropriate temperature. Incubation
temperatures below 68F may result in reduced via-
bility and problems with fungus (Saprolegnia sp.)
infestation of the embryos and larvae. Excessively
high water temperature (850F or greater) may also
adversely affect the development of the embryos.
Care should be taken to maintain incubation water
temperature within the optimum range.
Unscreened McDonald jars and conical vats must
be monitored during the incubation period. Water
flow must be sufficient to keep the eggs in suspen-
sion without flushing the eggs from the jar or vat.
Egg buoyancy changes during incubation; therefore,
water flow adjustments must be made. Super-
saturated gasses may come out of solution and the
bubbles may become attached to the eggs, carrying
them out of the hatching apparatus. This may be a
problem if the water is heated, from a well, or
pumped under high pressure. Strong aeration in
the reservoir tank will alleviate this problem.








the eggs and sperm in the temperature range of
74F to 760F. Decompression is instantaneous at the
end of the treatment, and the eggs are then trans-
ferred to the incubation apparatus.Treatments
using these parameters have resulted in near 100%
triploid fish and a hatch relative to controls of
nearly 70%. Success in the production of triploid
fish may be adversely affected by poor egg quality,
temperature variation, and sloppy spawning
technique. When the percent of viable embryos of
the untreated group is less than 60%, due to poor
egg quality, few if any triploid fish are produced by
pressure treatment.
Temperature shocks are administered via water
bath in insulated tanks or large ice chests. Therm-
ostatically controlled submersible heaters or chill-
ers are used to adjust water temperature to the
desired level. Fertilized eggs are placed in small-
mesh dip nets or net baskets, and immersed at the
proper time in the aerated treatment bath. Nets
containing the eggs are swirled to ensure uniform-
ity of treatment to all the eggs. At the end of the
treatment, eggs are immediately placed in the incu-
bation apparatus with ambient water temperature.
The reported heat shock parameters vary from
1080F for 1 minute, starting 4 minutes after water
was added to the eggs and sperm, to 1010F for 3.5
minutes, starting 1 minute after the addition of
water. Although these treatments are somewhat
successful, there appears to be a great deal of vari-
ability and inconsistency in the production of trip-
loid grass carp by heat shock.

Verification of Triploidy
Each individual grass carp must be determined
to be a triploid by analyzing their red blood cell
erythrocytee) nuclear volumes, using an electronic
particle analyzer, before they can be stocked in
Florida waters. The size range for blood cells is
determined by calibration with standard particles.
Diploid grass carp are reported to have a nuclear


j.- -.

Figure 25. Obtaining a blood sample from each grass carp
to be analyzed for triploidity before stocking in Florida's waters.


volume of 10.06 cubic micrometers, while mean
triploid nuclear volume is 14.82 cubic micrometers.
Blood samples are taken from grass carp greater
than 2 inches long. Fish may be anesthetized if
necessary. The head of the fish is tilted back and
the branchial artery on the isthmus is punctured
with a blood lancet rinsed in EDTA between sam-
ples. Approximately 1 microliter of blood is with-
drawn, using a positive displacement micropipette
with disposable polypropylene tips (Figure 25). The
blood sample is immediately dispersed into an
Accuvette vial filled with 10 milliliters of Isoton II
electrolyte and one drop of Zapoglobin lysing agent.
The sample is processed immediately while the fish
is held in a net container suspended in water.
Diploid individuals are discarded.

Care of Spawn in Incubators
Good water quality is a necessity for incubating
eggs and holding larvae. Water used for incubation
must be filtered if particulate matter or predatory
animals are present. During the initial stages of
development, oxygen consumption is negligible; as
development progresses, the oxygen demand in-
creases considerably. Also during their develop-
ment, the eggs excrete carbon dioxide and am-
monia. If these products are allowed to accumulate,
they can kill the embryos.
The developing embryos needs a continuous sup-
ply of water, nearly saturated with oxygen. A con-
tinuous flow of water through the incubator will
also remove the carbon dioxide and ammonia. We
strongly advise the use of well water for incubating
eggs. Thorough aeration of the water before it is de-
livered to the incubators is required to provide suffi-
cient oxygen. The use of aerated well water in a
single use, flow-through incubation system elimi-
nates many problems. Recirculated water is not re-
commended for egg incubation.
Normal development of the embryos requires
water of the appropriate temperature. Incubation
temperatures below 68F may result in reduced via-
bility and problems with fungus (Saprolegnia sp.)
infestation of the embryos and larvae. Excessively
high water temperature (850F or greater) may also
adversely affect the development of the embryos.
Care should be taken to maintain incubation water
temperature within the optimum range.
Unscreened McDonald jars and conical vats must
be monitored during the incubation period. Water
flow must be sufficient to keep the eggs in suspen-
sion without flushing the eggs from the jar or vat.
Egg buoyancy changes during incubation; therefore,
water flow adjustments must be made. Super-
saturated gasses may come out of solution and the
bubbles may become attached to the eggs, carrying
them out of the hatching apparatus. This may be a
problem if the water is heated, from a well, or
pumped under high pressure. Strong aeration in
the reservoir tank will alleviate this problem.









Conical baskets, screened vats, and circular
tanks retain the eggs regardless of the water flow,
but care must be taken to ensure that water flow is
not too great to impinge eggs and larvae on the
screen. Air bubbled gently along the screen can al-
leviate this problem, and it provides additional aer-
ation and water current to keep eggs in suspension.
Aeration cannot be too vigorous, because the deli-
cate embryos and larvae can be damaged.
When the quality of the spawn is good (70% or
greater), the dead eggs need not be removed from
the incubator. However, if there is a high percen-
tage of dead eggs, they should be siphoned off.
Dead eggs are more buoyant and form a layer above
the live ones in the incubator, and are easily
removed. In circular tanks, screened vats, and
hatching baskets, dead eggs do not stratify and can-
not be removed; the dead eggs, however, break
down prior to hatching and are flushed out of the
tank. Air bubbling in the incubation apparatus ap-
pears to facilitate this process.
Estimation of percent of viable embryos
Nonviable eggs can be distinguished 6 to 10
hours after spawning by their cloudy appearance

^ i ^c BI


and incomplete development. A sample of eggs
is withdrawn from the incubator, examined under a
10X hand lens or dissecting microscope, and enum-
erated to determine percent viable embryos. Suc-
cess is dependent not only on the quality of the
eggs and sperm released by the brood fish, but also
the skill of the spawning team and the quality of
the hatchery facility.

Duration of incubation
Temperature is the primary factor influencing
duration of incubation. At the optimum tempera-
ture (70F to 770F), hatching occurs after about 24
to 28 hours. As temperature increases to 80 to 82F,
incubation time decreases to approximately 18
hours. Figure 26 is a pictorial account of the de-
velopment of grass carp embryos from fertilization
through hatching.
Hatching
The embryos develop into larva and hatch by
breaking out of the eggs. Breaking of the egg shell
by the larva is a mechanical process, aided by
enzymatic weakening of the egg shell.


i
.. ... .t -


Figure 26 1) Embryonic stages of grass carp embryos. Eggs immediately after fertilization and water hardened. 2) Egg in
two cell division stage. 3) Early (large cell) morula stage approximately 2.5 hours after fertilization. 4) Late stage (small
cell) morula stage. 5) Late gastrulation stage, approximately, approximately 12 hours after fertilization, 6) Embryo immediately
before hatching (24-28) hours.








Grass carp larvae are separated from the dead
eggs and shells by swimming up with the water
current. The newly hatched larvae swim vertically
toward the surface of the incubation apparatus. If
an overflow outlet is present, the larvae are carried
out of the incubator with the current and are re-
tained by a screen in the larvae-holding aquarium
or tank. The larvae in screened vats and circular
tanks are retained in the apparatus; the dead eggs
and shells are quickly broken down by the bubble
aeration and flushed from the incubator.

Development of the Larvae
The newly hatched grass carp larvae are very
different from the adult fish. They do not have a
gut, mouth, vent, swim bladder, or gills. The yolk
sac provides the nutrients for growth and develop-
ment. The larvae alternately lie on the bottom,
swim up vertically in the water column, and then
fall back to the bottom. Grass carp larvae are held
in aerated aquaria or circular tanks until they
utilize their yolk sacs (1 to 3 days) and are ready to
start feeding. During this period, stocking densities
can be quite high (50,000 larvae/gallon), providing
water current is sufficient to keep them in suspen-
sion. Water exchange rate should be once every
half-hour when stocking rates are high (50,000 lar-
vae/gallon). The drain standpipe is surrounded by a
screen cylinder of 50 mesh/inch (0.5 millimeters).
Air is gently bubbled around the screen to provide
additional aeration and water current; this also
helps prevent clogging of the screen and impinge-
ment of the larvae. If circular tanks or screened
vats are used for incubation, larvae may remain in
the same container until time of stocking for fry
rearing.
The number of larvae in a container can be esti-
mated volumetrically. At 1 to 2 days after hatch, the
water is turned off and each aquarium or tank is
siphoned from inside the screen cylinder to a pre-
determined volume marked on the side. The air is
then shut off and the larvae are gently stirred to
achieve a uniform distribution within the tank. The
larvae are sampled with a tube or small container of
known volume inserted vertically to the bottom, and
the sample is withdrawn for enumeration. A
minimum of five samples should be collected from
each tank or aquaria. The number of larvae is calcu-
lated by multiplying the volume of the water in the
tank by the average number of larvae per volume of
sample. This technique is most accurate if there are
a minimum of 3 to 5 larvae/milliliter. Immediately
turn on the water and air after the samples have
been collected.
The inflation of the swim bladder, development of
the mouth, presence of pigmentation, and initiation
of horizontal swimming behavior marks the transi-
tion from larvae to fry and the need for an external
food source (Figure 27). It is difficult, if not impossi-


Figure 27. A grass carp fry 72 hours after spawning.

ble, to accurately estimate the number of fry once
they are swimming horizontal and are ready to start
feeding, because they can avoid the sampling con-
tainer.

Transportation of Fry
Fry are transported in oxygen-filled plastic bags.
It is best to use two bags, one placed inside the other,
to prevent leakage of water and oxygen, ensuring
survival of fry. These are packed in styrofoam-lined
cardboard boxes. The inlet to the fry tank water is
turned off and the fry are concentrated to a density
of 50,000 /gallon by siphoning water from inside the
cylindrical screen. Do not pull the standpipe to con-
centrate fry, because they may become injured by
impingement. Netting of the delicate fry is not recom-
mended. Water and fry are dipped, poured, or drained
carefully from the holding tank into the plastic bags
until approximately 2 gallons are contained in each
bag. Air should be squeezed from the bag and re-
placed with bottled oxygen to fill the remaining vol-
ume of the box. A small block of ice or an artificial
ice pack is sometimes added between the two bags
to maintain temperature within acceptable limits for
the fry. The bags are sealed with a heat bagsealer,
or two castration bands or rubber bands; the top of
the box is securely taped. Always avoid direct sunlight
when packing and transporting fry. Boxes of fry
should not be allowed to remain stationary for pro-
longed periods to prevent fry suffocation. Fry can be
transported in these containers for 24 hours with
little mortality at concentrations up to 50,000/gallon
(100,000/box).


Pond Production to Fingerling Size
Fry pond preparation
The traditional method of rearing fry to fingerling
size is direct stocking into prepared ponds at 3 to 5
days of age. Ponds suitable for the culture of grass
carp are the same as those used for fry of channel


P,








Grass carp larvae are separated from the dead
eggs and shells by swimming up with the water
current. The newly hatched larvae swim vertically
toward the surface of the incubation apparatus. If
an overflow outlet is present, the larvae are carried
out of the incubator with the current and are re-
tained by a screen in the larvae-holding aquarium
or tank. The larvae in screened vats and circular
tanks are retained in the apparatus; the dead eggs
and shells are quickly broken down by the bubble
aeration and flushed from the incubator.

Development of the Larvae
The newly hatched grass carp larvae are very
different from the adult fish. They do not have a
gut, mouth, vent, swim bladder, or gills. The yolk
sac provides the nutrients for growth and develop-
ment. The larvae alternately lie on the bottom,
swim up vertically in the water column, and then
fall back to the bottom. Grass carp larvae are held
in aerated aquaria or circular tanks until they
utilize their yolk sacs (1 to 3 days) and are ready to
start feeding. During this period, stocking densities
can be quite high (50,000 larvae/gallon), providing
water current is sufficient to keep them in suspen-
sion. Water exchange rate should be once every
half-hour when stocking rates are high (50,000 lar-
vae/gallon). The drain standpipe is surrounded by a
screen cylinder of 50 mesh/inch (0.5 millimeters).
Air is gently bubbled around the screen to provide
additional aeration and water current; this also
helps prevent clogging of the screen and impinge-
ment of the larvae. If circular tanks or screened
vats are used for incubation, larvae may remain in
the same container until time of stocking for fry
rearing.
The number of larvae in a container can be esti-
mated volumetrically. At 1 to 2 days after hatch, the
water is turned off and each aquarium or tank is
siphoned from inside the screen cylinder to a pre-
determined volume marked on the side. The air is
then shut off and the larvae are gently stirred to
achieve a uniform distribution within the tank. The
larvae are sampled with a tube or small container of
known volume inserted vertically to the bottom, and
the sample is withdrawn for enumeration. A
minimum of five samples should be collected from
each tank or aquaria. The number of larvae is calcu-
lated by multiplying the volume of the water in the
tank by the average number of larvae per volume of
sample. This technique is most accurate if there are
a minimum of 3 to 5 larvae/milliliter. Immediately
turn on the water and air after the samples have
been collected.
The inflation of the swim bladder, development of
the mouth, presence of pigmentation, and initiation
of horizontal swimming behavior marks the transi-
tion from larvae to fry and the need for an external
food source (Figure 27). It is difficult, if not impossi-


Figure 27. A grass carp fry 72 hours after spawning.

ble, to accurately estimate the number of fry once
they are swimming horizontal and are ready to start
feeding, because they can avoid the sampling con-
tainer.

Transportation of Fry
Fry are transported in oxygen-filled plastic bags.
It is best to use two bags, one placed inside the other,
to prevent leakage of water and oxygen, ensuring
survival of fry. These are packed in styrofoam-lined
cardboard boxes. The inlet to the fry tank water is
turned off and the fry are concentrated to a density
of 50,000 /gallon by siphoning water from inside the
cylindrical screen. Do not pull the standpipe to con-
centrate fry, because they may become injured by
impingement. Netting of the delicate fry is not recom-
mended. Water and fry are dipped, poured, or drained
carefully from the holding tank into the plastic bags
until approximately 2 gallons are contained in each
bag. Air should be squeezed from the bag and re-
placed with bottled oxygen to fill the remaining vol-
ume of the box. A small block of ice or an artificial
ice pack is sometimes added between the two bags
to maintain temperature within acceptable limits for
the fry. The bags are sealed with a heat bagsealer,
or two castration bands or rubber bands; the top of
the box is securely taped. Always avoid direct sunlight
when packing and transporting fry. Boxes of fry
should not be allowed to remain stationary for pro-
longed periods to prevent fry suffocation. Fry can be
transported in these containers for 24 hours with
little mortality at concentrations up to 50,000/gallon
(100,000/box).


Pond Production to Fingerling Size
Fry pond preparation
The traditional method of rearing fry to fingerling
size is direct stocking into prepared ponds at 3 to 5
days of age. Ponds suitable for the culture of grass
carp are the same as those used for fry of channel


P,








Grass carp larvae are separated from the dead
eggs and shells by swimming up with the water
current. The newly hatched larvae swim vertically
toward the surface of the incubation apparatus. If
an overflow outlet is present, the larvae are carried
out of the incubator with the current and are re-
tained by a screen in the larvae-holding aquarium
or tank. The larvae in screened vats and circular
tanks are retained in the apparatus; the dead eggs
and shells are quickly broken down by the bubble
aeration and flushed from the incubator.

Development of the Larvae
The newly hatched grass carp larvae are very
different from the adult fish. They do not have a
gut, mouth, vent, swim bladder, or gills. The yolk
sac provides the nutrients for growth and develop-
ment. The larvae alternately lie on the bottom,
swim up vertically in the water column, and then
fall back to the bottom. Grass carp larvae are held
in aerated aquaria or circular tanks until they
utilize their yolk sacs (1 to 3 days) and are ready to
start feeding. During this period, stocking densities
can be quite high (50,000 larvae/gallon), providing
water current is sufficient to keep them in suspen-
sion. Water exchange rate should be once every
half-hour when stocking rates are high (50,000 lar-
vae/gallon). The drain standpipe is surrounded by a
screen cylinder of 50 mesh/inch (0.5 millimeters).
Air is gently bubbled around the screen to provide
additional aeration and water current; this also
helps prevent clogging of the screen and impinge-
ment of the larvae. If circular tanks or screened
vats are used for incubation, larvae may remain in
the same container until time of stocking for fry
rearing.
The number of larvae in a container can be esti-
mated volumetrically. At 1 to 2 days after hatch, the
water is turned off and each aquarium or tank is
siphoned from inside the screen cylinder to a pre-
determined volume marked on the side. The air is
then shut off and the larvae are gently stirred to
achieve a uniform distribution within the tank. The
larvae are sampled with a tube or small container of
known volume inserted vertically to the bottom, and
the sample is withdrawn for enumeration. A
minimum of five samples should be collected from
each tank or aquaria. The number of larvae is calcu-
lated by multiplying the volume of the water in the
tank by the average number of larvae per volume of
sample. This technique is most accurate if there are
a minimum of 3 to 5 larvae/milliliter. Immediately
turn on the water and air after the samples have
been collected.
The inflation of the swim bladder, development of
the mouth, presence of pigmentation, and initiation
of horizontal swimming behavior marks the transi-
tion from larvae to fry and the need for an external
food source (Figure 27). It is difficult, if not impossi-


Figure 27. A grass carp fry 72 hours after spawning.

ble, to accurately estimate the number of fry once
they are swimming horizontal and are ready to start
feeding, because they can avoid the sampling con-
tainer.

Transportation of Fry
Fry are transported in oxygen-filled plastic bags.
It is best to use two bags, one placed inside the other,
to prevent leakage of water and oxygen, ensuring
survival of fry. These are packed in styrofoam-lined
cardboard boxes. The inlet to the fry tank water is
turned off and the fry are concentrated to a density
of 50,000 /gallon by siphoning water from inside the
cylindrical screen. Do not pull the standpipe to con-
centrate fry, because they may become injured by
impingement. Netting of the delicate fry is not recom-
mended. Water and fry are dipped, poured, or drained
carefully from the holding tank into the plastic bags
until approximately 2 gallons are contained in each
bag. Air should be squeezed from the bag and re-
placed with bottled oxygen to fill the remaining vol-
ume of the box. A small block of ice or an artificial
ice pack is sometimes added between the two bags
to maintain temperature within acceptable limits for
the fry. The bags are sealed with a heat bagsealer,
or two castration bands or rubber bands; the top of
the box is securely taped. Always avoid direct sunlight
when packing and transporting fry. Boxes of fry
should not be allowed to remain stationary for pro-
longed periods to prevent fry suffocation. Fry can be
transported in these containers for 24 hours with
little mortality at concentrations up to 50,000/gallon
(100,000/box).


Pond Production to Fingerling Size
Fry pond preparation
The traditional method of rearing fry to fingerling
size is direct stocking into prepared ponds at 3 to 5
days of age. Ponds suitable for the culture of grass
carp are the same as those used for fry of channel


P,








catfish (Ictalurus punctatus), bait minnows,
largemouth bass (Micropterus salmoides), or striped
bass. Drainable ponds are the easiest to manage and
harvest.
It is advisable to dry and disk fry rearing ponds
during the winter. This interrupts the life cycles of
disease, parasitic, and predatory organisms. Drying
and disking the pond also promotes aerobic decom-
position of the muck on the pond bottom. The ponds
should be filled with well water just prior (3 to 7
days) to stocking fry, when possible. This reduces
the threat of predacious insects (i.e. diving beetles,
dragonfly nymphs, etc.) and ensures a population of
small zooplankton of suitable size to be ingested by
the fry. When surface water sources are used to fill
ponds, inlets should be covered with a screen filter
sock to prevent introduction of fish or predacious
invertebrates.
Ground water ponds that cannot be drained
should be pumped with a 3-inch or larger trash
pump. The organic muck should be washed from
the bottoms and sides of the ponds to the trash
pump inlet with a smaller pump equipped with a
nozzle on the discharge hose. The ponds should
then be treated with hydrated lime (2,000 pounds/
acre), spread evenly over the entire surface of the
pond sides and bottom, and allowed to refill. The in-
itial dose of fertilizer is applied just prior to or
while the pond is filling (See "Feeding the fry").
Innoculating ponds with desired zooplankton has
been shown to be beneficial. Rotifers (Brachionus
sp) and cladocerans (Daphnia sp and Moina sp) are
preferred organisms. These zooplankton can either
be cultured (see "Live Food Culture") or collected
from hatchery ponds or other bodies of water. Do
not inadvertently introduce predaters with the
innoculum.
Diesel fuel sprayed at a rate of 4 gallons per sur-
face acre kills airbreathing insects in the pond and
can be used just prior to and at 3-day intervals
after stocking until fry are large enough to avoid
predation. The treatment is most effective when
applied during calm periods. The diesel fuel should
have air bubbled through it for 24 hours prior to
application to release volatile hydrocarbons that
can be toxic to zooplankton and fish.

Stocking of fry
Grass carp and other Chinese carp fry are
stocked at a rate of 100,000-500,000 per acre. Al-
ways check water quality parameters of the pond,
especially dissolved oxygen, before stocking the
fish. Be sure that there is at least 4 ppm dissolved
oxygen in the pond before stocking. The fry should
be stocked at night or during early morning hours
to reduce temperature shock. Water temperature in
the shipping bag should be adjusted to pond tem-
perature by floating the bags in the pond prior to
release. A tempering rate of 40F per hour is
suggested. Direct sunlight should be avoided during


the tempering process. When the water tempera-
ture in the bag is equal to the pond water, the bag
is opened, and approximately one-half gallon of
pond water is added every 30 seconds for 2 minutes
to aid in acclimating the fish to the new water con-
ditions. The fry are then immediately released. It is
important not to open the plastic bags until fish are
ready to be stocked, because of high carbon dioxide
levels in the water in the bag.

Feeding of the fry
The first food of the fish is primarily zoo-
plankton. Pond zooplankton abundance can be in-
creased through fertilization. Organic fertilizers are
usually preferred for producing zooplankton
blooms. Inorganic fertilizers tend to stimulate
phytoplankton blooms rather than zooplankton. A
combination of organic and inorganic fertilizers fre-
quently yields good results. A wide variety of mate-
rials have been used successfully for pond fertiliza-
tion; the choice depends primarily on availability
and cost. The fish farmer should experiment to see
what combination, rates, and schedule work best
for that particular farm. Fertilization rates may
vary from pond to pond. The application rate
should be determined for each individual situation.
Animal manure (chicken and swine) is an effec-
tive organic fertilizers for rearing fry to fingerlings.
Weekly applications of 300 to 400 pounds/acre of
manure in combination with inorganic fertilizer has
been used to produce fingerlings. However, in-
creased fish disease and parisitic problems have
been experienced when manure is used as a fer-
tilizer. Liquid inorganic fertilizer (ammonium
polyphosphate) is more efficient to use than granu-
lar forms. It should be applied at 1 1/2 to 2 gallons/
acre-foot, with the rate adjusted for your particular
conditions. You may need to apply fertilizer every
week to maintain the bloom.
Hay has also been a favored source of organic ma-
terial for fry pond fertilization. Rates of 200 to
1,000 pounds of hay/acre have been used. The main
advantage of hay is its long-term production of
zooplankton. Hay does not usually produce a quick
bloom of zooplankton; this may be overcome by
grinding a portion of the hay or by the use of 100 to
500 pounds/acre of quick-acting organic fertilizers
such as soybean meal, cottonseed meal, or animal
manure. A relatively small initial dose followed by
daily applications of small amounts of finely
ground organic fertilizer appears to be a more logi-
cal way to sustain a zooplankton bloom without
causing devastating water quality changes. The
more traditional method of applying a large quan-
tity of fertilizer at infrequent intervals can result in
a boom or bust situation. If minnow meal is the or-
ganic fertilizer applied daily, this direct method of
feeding the zooplankton would also speed the trans-
ition by the fish from live food to artificial diet. Fre-
quently, minnow meal is less expensive than some








of the high protein organic fertilizers such as soy-
bean meal, cotton seed meal, and alfalfa meal.
Regardless of the fertilization program used, sup-
plemental feed should be offered, especially when
stocking density is high. Minnow meal, used in bait
fish farming, is adequate until the fish are 3 to 4 in-
ches long. Fish are fed two to six times daily de-
pending on available manpower. Feed is spread by
hand from the windward side of the pond. It is im-
portant to get an even distribution of the meal over
the surface, rather than dumping it all in one spot.
Feeding activity should be observed to anticipate
water quality or disease problems, or to adjust feed-
ing rate. Water quality monitoring is especially im-
portant when fish receive supplemental feed. Cau-
tion should be observed when more than 30 pounds
per acre is fed.

Harvesting fingerlings from ponds
When grass carp and other Chinese carp are
stocked at high density in the ponds, they should be
harvested at approximately 3 to 4 inches in length
and redistributed at a density of 3,000 to 5,000 fish
per acre for growth to stocking size (10 inches). It is
generally advisable to harvest at dawn when tem-
perature is most favorable. Diseased or poorly con-
ditioned fish should not be harvested until the
problem is corrected.
Ponds with catch basins that can be drained by
gravity flow are the simplest for harvesting finger-
ling fish. Ponds are usually drained slowly during
the night to avoid stranding the fish. Check dis-
solved oxygen frequently during draining. If dis-
solved oxygen is below 4 ppm or the fish exhibit
symptoms of stress or disease, immediately refill
the pond and diagnose and treat the problem before
attempting harvest. The kettle should have a fresh-
water source to reduce losses due to low-dissolved
oxygen, elevated water temperatures, or damage to
gills from suspended solids. As the fingerlings con-
centrate in the concrete kettle, they are dip-netted,
placed into a container with water, and transferred
to the hauling tank.
Ponds that cannot be drained can be harvested
with a small-mesh, woven, bag seine. Extra lead
weights may be needed in some situations. Be sure
that the fish are large enough so that they will not
be gilled in the mesh of the net. Do not allow the fish
to be covered by the bottom sediments brought in by
the seine. Minimize the time that the fish are in the
seine and dip nets. Transfer fish to hauling tanks in
containers with water. Ponds may be
partially pumped before seining to facilitate harvest.
The pump inlet must be screened to prevent loss of
fish. Fingerlings can also be concentrated in one area
by spraying fresh water or by feeding.
Survival of fry to fingerling-size fish in ponds is
variable. A return of 40 to 60% is considered good.
Total mortality of fry in ponds can occur.


Intensive Production to Fingerling
Size
Studies at the University of Florida have shown
that the fry of grass carp and other carps can be
raised in intensive culture systems. Previously,
problems had occurred with the culture of fry, be-
cause they are relatively sensitive to environmental
stress, susceptible to predation by many aquatic
organisms, and selective in feeding habits.
Generally, intensive culture refers to the rearing
of a large number of organisms in a limited space.
Advantages of intensive culture are (1) reduced
land area required and construction cost of
facilities, as compared to ponds; (2) improved sur-
vival due to exclusion of predators; (3) food is
supplied as needed rather than relying on the fluc-
tuating zooplankton populations in the ponds; and
(4) water quality for the fry can be controlled. The
two major problems of intensive fry rearing are
providing adequate and appropriate food and main-
taining water quality. Intensive culture does, how-
ever, require greater supervision, skill, time and
technical knowledge on the part of the hatchery
manager and staff.
Two culture systems have been successfully
tested at the University of Florida for fry rearing:
(1) semi-intensive and (2) intensive. Both
techniques have resulted in nearly 100% survival of
fry to a 1-inch size.

Semil-ntensive fry rearing
Cages and tanks (see "Hatchery Facility") used
for semi-intensive fry rearing are stocked at 100 fry/
gallon. Zooplankton laden water is pumped to fry
rearing tanks from ponds fertilized and treated as
outlined in the section titled "Fry production To
Fingerling Size". The inlet to the pump is screened
to prevent introduction of predators to the fry
tanks. Cages are suspended directly in the ponds
and water is drawn through the tanks by an airlift
pump. The water in the tanks and cages is ex-
changed every 10-15 minutes. Dry diet is added to
the culture container after the first week. Approxi-
mately five times the production of grass carp per
surface acre can be expected using this technique
as compared to direct pond stocking. A major disad-
vantage is that water quality is largely determined
by conditions in the heavily fertilized pond.

Intensive fry rearing
The intensive technique of fry rearing utilizes
aquaria or tanks as culture containers. Fry are
stocked at 100 individuals per gallon. Water quality
is maintained with simple airlift sponge filters (Fi-
gure 16). During 3-week feeding trials, no fresh
water was added; however, a gradual freshwater
drip, with an exchange time of approximately 2
days, would be beneficial for improving water qual-









ity and fry growth. If a freshwater flush is used, the
outlets from the tanks or aquaria must be screened
(42 mesh/inch or 0.6 millimeter polyester cloth) to
prevent escape by the fry.
Live food is considered important for raising fish
fry, especially during the first weeks of feeding.
Some fry require a food item that shows indepen-
dent movement. This live food must also be appro-
priate for the mouth size of the fish and must pro-
vide a nutritionally complete diet. Cultured live
foods suitable for fry include freshwater rotifers
(Brachionus rubens), daphniids (Monia), brine
shrimp (Artemia), and nematodes (Panagrellus).
Commercially available fry diets are more conve-
nient to feed, and by introducing the fry to an artifi-
cial ration early in life, the transition from live food
is less traumatic. Live foods are fed at least twice
per day, and dry foods are fed more frequently
using automatic feeders.
Any of these foods can be used alone; however, a
fry feeding program utilizing a combination of live
food items and a dry diet is superior, because it pro-
vides for a backup food supply in case of production
failures, and is better able to meet the nutritional
requirements of the fish fry.

Live Food Culture
Rotifers Most of the culture procedures used in
this country for rotifers are similar to those de-
veloped for the marine rotifer, Brachionus plicitilis,
in Japan starting in the early 1960's. This proce-
dure utilizes jars, aquariums, tanks, or vats for in-
door or greenhouse culture of algae and rotifers.
Specially formulated chemical fertilizers are used
for phytoplankton culture. Living rotifers are trans-
ferred to the culture media, when the maximum
phytoplankton density has been reached. Rotifers
can either be batch-cultured in this manner, or a
continuous culture can be maintained by excluded
rotifers from separate phytoplankton cultures. The
algae media is transferred to the rotifer tanks as
needed. A yeast-based diet is fed to the rotifers to
supplement the algae. When the rotifer population
has increased to 50 individuals/milliliter, it should
be harvested daily with a small-mesh net (300
mesh/inch or smaller) to maintain the productivity
of the culture. Meticulous cleaning of tanks and
equipment is required to prevent contamination of
the separate algae cultures by rotifers. Rotifer den-
sities of 50 to 100 individuals/milliliter can be
maintained. However, at higher density (500 indi-
viduals/milliliter) rotifer populations can peak and
decline very quickly.
A simplified procedure for the production of fresh-
water rotifers (B. rubens) (Figure 28) has been used
to provide live food for fry rearing studies at our
laboratory. The culture procedure utilizes shallow
(12 to 18 inches deep) outdoor pools. The cultures
are fertilized with liquid swine manure. The


Figure 28. Rotifer.

maximum amount of this organic fertilizer should
be used, while ensuring that the free ammonia con-
centration remains below 15 parts per million. The
rotifers consume phytoplankton and bacteria. The
pools are inoculated with phytoplankton (Chlorella)
and later with rotifers (B. rubens). To ensure a con-
stant supply of rotifers, a minimum of two or three
pools should be inoculated in this manner. Rotifer
production and decline periods normally last about
one week each during warm months. High densities
(50 to 500 rotifers/millimeter) were present in at
least one of the two pools through out the three
month study period with no additional inputs of fer-
tilizer.
The relative health of the rotifer culture can be
evaluated by not only the number/volume, but also
by the number of females with eggs and the rela-
tive swimming speed of the rotifers. These paramet-
ers are excellent indicators of water quality prob-
lems or food deficiencies in the rotifer culture.
The freshwater rotifer (B. rubens) will frequently
congregate at the surface of rotifer pools in very
high concentrations (1,500 rotifers/milliliter). This
is usually observed early in the morning as a result
of oxygen depletion and appears as an orangish-
brown scum at the water surface. When this occurs,
large numbers of rotifers can be quickly harvested
by simply skimming off this surface concentration.









The rotifers should be fed to the fry in a concen-
trated form with very little of the culture water,
otherwise the ammonia and organic matter present
in the rotifer culture may cause water quality prob-
lems in the fry tanks. Freshwater rotifers are one of
the best live cultured food for Chinese carp fry.
Studies at our laboratory have shown that growth
of grass carp and bighead carp fry fed rotifers is
superior to those fed brine shrimp, microworms, or
commercial dry diet.
Daphnia Many species of the family
Daphniidae occur throughout the world and are
collectively known as daphnia. The genera Daphnia
and Moina are the more common. Many fishery
workers have emphasized the importance of these
zooplankton as food for both young and adult fish.


Figure 29. Moina.
Moina (Figure 29) have several advantages over
Daphnia as a live food for fish. The maximum body
length of Moina is less than half the length of
Daphnia magna and D. pule. Adult Moina (0.7-1.0
millimeter) are larger and young (<0.4 millimeter)
Moina are smaller than newly hatched brine
shrimp (0.5 millimeter). Young Moina are slightly
longer than adult Rotifers (0.3 millimeter). Moina
are therefore suitable as the initial food for the very
small fry of the Chinese carps. In Singapore, Moina
are grown in ponds fertilized with mostly chicken
manure or, less frequently, with pig manure. Moina
are used as the sole food of fry of many ornamental
tropical fish; a 95 to 99% survival to 1/2 inch length
is quite common. The optimum temperature for
Moina (75F to 88F) is more suitable for rearing
Chinese carp fry than Daphnia (64F to 720F). In
addition, high population densities of Daphnia can
result in a dramatic decrease in their reproduction,
but this is not apparently the case with Moina .
The egg output of Daphnia magna drops sharply at
a density as low as 95 to 115 mature individuals/
gallon. Moina cultures routinely reach densities of
19,000 individuals/gallon. A comparison of the pro-
duction of Moina macrocopa and Daphnia magna
cultures fertilized with yeast and ammonium ni-


trate, showed that the average daily yield of Moina
was 3-4 times the daily production of Daphnia.
Moina can be cultured in almost any sized con-
tainer. Earthen or concrete ponds, and metal, plas-
tic, glass and fiberglass vats or tanks can be used.
Water depth should be no greater than 3 feet. A
depth of 16 to 20 inches is probably optimum, allow-
ing light penetration through the water column for
photosynthesis. Gentle bubble aeration provided by
one or two outlets of aquarium air tubing for each
400 gallons of culture volume is recommended. A
small trickle of water into the culture container
may also improve water quality and production. Dif-
fuse light or shade over approximately 1/3 to 1/2 of
the surface of the culture container is recom-
mended. If Moina cultures are protected from rain,
production is more stable.
Moina can be cultured in combination with their
food, or separate cultures of phytoplankton can be
used to feed them. Combined culture is by far the
easiest. The separate culture of Moina and phyto-
plankton can only be maintained in closed, tightly
controlled systems. Production using this system
had been reported to be approximately 1/4 to 1/3
higher. However, since this method requires addi-
tional space for the cultivation of the phyto-
plankton, it follows that production per unit vol-
ume, especially in large scale cultures, is equiva-
lent or even higher when the animals are grown
together with their food.
Moina have been cultured on a variety of organic
and mineral fertilizers. Organic fertilizers are
usually preferred to mineral fertilizers because they
provide bacterial and fungal cells and detritus as
well as phytoplankton as food for Moina. This vari-
ety of food items more completely meets their nutri-
tional needs resulting in maximum production.
Mineral fertilizers may be used alone; however, they
work better in earthen ponds than in tanks or vats.
Mineral fertilizers are also used in the separate cul-
ture of phytoplankton to be fed to Moina. The com-
bined application of organic and mineral fertilizers
can result in good production of Moina, by creating
favorable conditions for the growth of a variety of
foods.
Fresh organic fertilizers are preferred to old or
moldy materials because they are richer in organic
matter and microbes. Manure is usually dried be-
fore use. Coarse organic materials such as manure,
hay, bran, and oil seed meals are usually soaked in
water and then poured into a bag that is suspended
in the water column. Nylon stockings work well for
this purpose; they are inexpensive, readily avail-
able, and nylon also holds up much better in water
than cotton or burlap. Although not necessary, the
use of a bag prevents large particles from being a
problem when the Moina are harvested.
Moina have been cultured on a wide variety of
materials such as livestock manures, hay or other
vegetation, cereal grains, oil seed meals, bran,








yeast, mineral fertilizers, and garden soil. The fol-
lowing are some of the simpler and more successful
methods used at our laboratory.

1) Cow manure, cotton seed meal, and yeast: Use
5 ounces of commercially available dehydrated
cow manure, 1 ounce of cotton seed meal, and
0.5 ounces of baker's yeast, for 100 gallons of
water.
2) Alfalfa, and yeast: Use 2.5 ounces of alfalfa pel-
lets or meal and 0.5 ounces of baker's yeast, for
100 gallons of water.
3) Cow manure, bran, and yeast: Use 5 ounces of
commercially available dehydrated cow manure,
2.5 ounces of bran, and 2 ounces of baker's
yeast for 100 gallons of water.

Do not over-fertilize the cultures. Regardless of
the fertilizer used, start with small amounts added
at more frequent intervals; slowly increase the
amount used as experience is gained. If fungus
occurs in large quantities in the culture due to an
excessive application of organic fertilizer (usually
observed 2 to 3 days after fertilization), the culture
should be discarded.
Cultures are inoculated with 100 Moina per gal-
lon after two hours of aeration, assuming good
water quality. If fewer are used, the population in
the culture will increase more slowly. Therefore, the
amount of fertilizer used initially should be 1/4 to
1/2 the recommended dosage; the remaining por-
tion is added as the population increases. A greater
number used for inoculation reduced the time to
harvesting, but does not result in proportionally
higher production. Occasionally the mortality of the
initial inoculation is high and an additional inocu-
lation is required.
The cultures should be fertilized with 1/4 to 1/2
the initial amount whenever the transparency is
more than 16 inches. This can be determined with a
secchi disk or a pie plate on the end of a yard stick.
Depth of transparency is where the disk is just
barely visible.
A partial harvest of no more than 1/5 to 1/4 the
population at regular daily intervals is necessary to
maintain the steady productivity of the culture.
Moina can be harvested by simply dipping out the
required number with a baby brine shrimp net or
by siphoning or draining the required volume of
culture water through a net. Cultures can be main-
tained in this manner for several months before a
new culture needs to be started.
Brine Shrimp Fish culturists have tradition-
ally used newly hatched brine shrimp, Artemia,
(Figure 30) almost exclusively as the initial food for
fish fry. Brine shrimp are incubated in transparent
containers with vigorous aeration at a salinity of
twelve parts per thousand and a temperature of
80F. Hatching occurs in 24 to 48 hours. Brine
shrimp can be separated from the salt water and


Figure 30. Brine shrimp nanplii (Artemia).


shells by turning off the aeration, allowing the
nauplii to settle to the bottom, and siphoning the
nauplii into a fine-mesh net. Avoid feeding un-
hatched eggs of egg shells to fry, because these can
obstruct the digestive tract of the fish. This problem
can be totally eliminated by decapsulating the
brine shrimp eggs, there by chemically removing
the shell before hatching (See "Additional Read-
ing"). A disadvantage of brine shrimp is that they
die of osmotic stress after several hours in freshwa-
ter. Over feeding can quickly foul the water in the
fry rearing container.
Nematodes -Nematodes (microworms) are one of
the simplest and most dependable live foods to cul-
ture (Figure 31). Almost any shallow, flat, water-
tight container can be used to culture microworms;
8 by 12 inch plastic refrigerator boxes with approxi-
mately 10 holes (1/16 inch) drilled in the lid are
especially convenient. Experience has shown that a
media of steam-rolled oats, available in 50-pound


Figure 31. Microworms (nematodes.).








bags from livestock feed stores, is one of the most
productive and economical foods for microworms.
Use slightly less than 1/2 quart of rolled oats with
one quart water for each 8 by 12 inch container. The
oats are cooked for 3 to 5 minutes, covered, and
allowed to cool. The media is placed in the culture
container and spread to a thickness of 1/2 to 3/4
inch. Any media on the sides of the container
should be removed with a damp cloth. A teaspoon
or more of yeast is sprinkled over the oatmeal, and
the starter culture containing the microworms is
spread on the surface. The microworms feed on the
yeast and bacteria. Keep the culture container at a
temperature of approximately 800F; cooler tempera-
tures result in less daily production, but the culture
lasts longer. After 3 days to 1 week, the worms
begin to crawl up the sides of the container, where
they can be easily scraped off. A wooden slat laid
diagonally across the media works well for provid-
ing additional surface from which the nematodes
can be harvested. Once established, a microworm
culture of this size will provide a continuous har-
vest of approximately 1 1/2 teaspoons of worms
daily for 3 weeks or more. Studies at our laboratory
have shown that growth of grass carp and big head
carp fed microworms is not significantly different
from those fed brine shrimp.

Pond production to stocking size
Small grass carp are extremely vulnerable to
largemouth bass predation. Therefore, when grass
carp are stocked into waters with existing bass
populations for weed control, they should be at
least 10 inches long to minimize bass predation.
Ponds have traditionally been used for production
of stocking-size grass carp. Fingerlings are usually
stocked at 3,000 to 5,000 per acre for growout. It is
advantageous to use ponds with existing sub-
mersed aquatic vegetation for rearing grass carp;
however, the fish can be fed floating catfish pellets
and green terrestrial grasses such as golf course
grass, millet, or Bermuda grass. Fresh-cut vegeta-
tion is fed at a daily rate of approximately 200
pounds per acre. Floating catfish feed is scattered
on the upwind side of the pond at a rate of 2% to
4% of the weight of the standing crop. Never feed
more than what the fish will eat in 15 to 30
minutes. When small fish are being fed, it may be
necessary to continue feeding minnow meal until
they are large enough to take pellet rations. Produc-
tion of grass carp can exceed 2,000 to 4,000 pounds
per acre.
Seining, handling, and transporting Chinese
carps, such as the grass carp, are more difficult
than for other warm-water fish. Bag seines with
additional bottom weights are more effective for
capture than are straight seines. The top of the
seine is usually held above the water surface to
minimize escape by jumping. This technique is
labor intensive and the workers may be injured by


the jumping fish. Injury and stress to the fish
during handling should be minimized, to avoid sec-
ondary bacterial infection, especially when water
temperature exceeds 80F If fish are seined during
the summer, seining should be conducted during
the morning when the water is cooler. The seine
should be brought in near the inlet pipe so that
cool, fresh water can be supplied.
The fish are usually graded by size, and any
foreign fish, vegetation, or animals, such as tad-
poles, are removed. Prophylactic treatment for
bacterial infection and removal of parasites is rec-
ommended before shipping. Holding tanks or vats
must be covered to prevent fish from jumping out.
All grass carp stocked in Florida waters or sold
from Florida fish farms must be verified as triploid
(see "Verification of triploidy").
Hauling tanks should be equipped with agitators
and bottled oxygen. Most of the better transport
units are equipped with a large-diameter discharge
that enables rapid transfer of the fish into holding
vats or waters which are to be stocked. Ice may be
added during hauling to prevent rapid increases in
water temperature; one-half pound of ice reduces
the temperature of 1 gallon of water by about 10F.
Water in the transport tank should be tempered
with pond or lake water in which the fish are to be
stocked; approximately 20 to 60 minutes of temper-
ing are required for each 10F change in water tem-
perature. A 12-volt bilge pump is useful for this
purpose.

Intensive production to stocking size
Grass carp are well suited for high density cul-
ture in flow-through circular tanks. Circular tanks
should be equipped with venturi drains for self-
cleaning. Water is jetted into the tank at a 45
angle to induce a counter-clockwise circular cur-
rent. Turnover time should be approximately 1
hour. Also, compressed air should be provided for
additional aeration and as a backup in case of
water pump failure. Fish should be fed at least
twice daily with a nutritionally complete channel
catfish ration in combination with vegetation.
Duckweed (Lemna) is an excellent food for finger-
ling grass carp, but the high water content makes
it cumbersome to feed. Stocking densities in excess
of 2 fish/gallon have been successful.

Conclusions
Induced spawning and rearing of fish to market-
able size is a complicated process requiring
facilities, capital, hard work, and strict attention to
detail. Before initiating an aquaculture venture,
learn as much as possible from books, magazines,
articles, established fish farmers, and hatchery
workers. Lessons learned in this way are much less
painful than those punctuated by dead fish and lost
revenues.









Technical information is important, but cannot
replace experience. Start small and build as you
learn from your mistakes; small mistakes are easier
to correct than large ones. Always keep the welfare
of the fish foremost in any consideration or deci-
sion. When in doubt, contact the fisheries spe-
cialists in your area or state for additional informa-
tion.

Additional Reading


Introduction to Fish Disease, Parasites and Treat-
ment. IFAS Extension Circular 716.
Hormone Spawning of Tropical Aquarium Fish.
IFAS Extension Circular 246.
Microworms: An Alternative Live Food for Fish Fry.
IFAS Extension FA 9.
A Simple Procedure for Decapsulating and Hatch-
ing Brine Shrimp. Campton, D. E., 1989. Prog.
Fish-Cult. 51:176-179


Management of Water Quality for Fish. IFAS Exten-
sion Circular 715.

SUPPLIERS OF PRODUCTS

AMPULES (For Freezing and Storing Hormones)


Curtin Matheson Scientific, Inc.
P.O. Box 1546
Houston, TX 77251


CARP PITUITARY
Argent Chemical Laboratories
8702 152nd Ave. N.E.
Redmond, WA 98052
206-842-3777
Stewart Fish Supply, Inc.
P.O. Box 15061
St. Louis, MO 63110
314-865-2000


HALOPERIDOL AND RESERPINE
Sigma Chemical Company
P.O. Box 14508
St. Louis, MO 63178
1-800-325-3010

HUMAN CHORIONIC GONADOTROPIN (HCG)
Argent Chemical Laboratories
8702 152nd Ave. N.E.
Redmond, WA 98052
206-885-3777
Fritz Aquaculture
Division of Fritz Chemical Co.
P.O. Drawer 17040
Dallas, TX 75217
1-800-527-1323

INTRAMETIC POLYETHYLENE TUBING
Curtin Matheson Scientific, Inc.
P.O. Box 1546
Houston, TX 77251


Fisher Scientific Co.
711 Forbes Ave.
Pittsburg, PA 15219
412-562-8300


Crecent Research Chemical, Inc.
5301 N. 37th PI.
Paradise Valley, AZ 85253
602-945-4733
Stoller Fisheries
Box B
1308 Memphis
Spirit Lake, IA 51360
712-336-1750


Crecent Research Chemical, Inc.
5301 N. 37th PI.
Paradise Valley, AZ 85253
602-945-4733
Stewart Fish Supply, Inc.
P.O. Box 15061
St. Louis, MO 63110
314-865-2000


Fisher Scientific Co.
711 Forbes Ave.
Pittsburg, PA 15219
412-562-8300









POLYESTER SCREEN (Filter Cloth)
Fritz Aquaculture
Division of Fritz Chemical Co.
P.O. Drawer 17040
Dallas, TX 75217
1-800-527-1323

EQUIPMENT AND SOLUTIONS FOR VERIFICATION OF TRIPLOID


Curtin Matheson Scientific, Inc.
P.O. Box 1546
Houston, TX 77251


Fisher Scientific Co.
711 Forbes Ave.
Pittsburg, PA 15219
412-562-8300


STERILE DISPOSABLE NEEDLES AND SYRINGES


Curtin Matheson Scientific, Inc.
P.O. Box 1546
Houston, TX 77251


Wholesale Veterinary Supply, Inc.
P.O. Box 2256
Rockford, IL 61131
1-800-435-6940

SYNTHETIC LH-RH ANALOGUE (LH-RHa)
Argent Chemical Laboratories
8702 152nd Ave. N.E.
Redmond, WA 98052
1-800-426-6258


Fisher Scientific Co.
711 Forbes Ave.
Pittsburg, PA 15219
412-562-8300


Sigma Chemical Company
P.O. Box 14508
St. Louis, MO 63178
1-800-325-3010


TISSUE GRINDER OR MORTAR AND PESTLE


Curtin Matheson Scientific, Inc.
P.O. Box 1546
Houston, TX 77251


Fisher Scientific Co.
711 Forbes Ave.
Pittsburg, PA 15219
412-562-8300


The information given herein is provided for your convenience with the understanding that no discrimination is
intended and no endorsement by the University of Florida is implied. The listing of specific trade names and
suppliers does not constitute an endorsement of these products or vendors in preference to others containing the
same active ingredients or providing similar items.












COOPERATIVE EXTENSION SERVICE, UNIVERSITY OF FLORIDA, INSTITUTE OF FOODANDAGRICULTURAL SCIENCES, JohnT. Woeste,
Director, in cooperation with the United States Department of Agriculture, publishes this information to further the purpose of the May 8 and June
30,1914 Acts of Congress; and is authorized to provide research, educational information and other services only to individuals and institutions that
function without regard to race, color, sex, age, handicap or national origin. Single copies of extension publications (excluding 4-H and youth
publications) are available free to Florida residents from county extension offices. Information on bulk rates or copies for out-of-state purchasers
is available from C.M. Hinton, Publications Distribution Center, IFAS Building 664, University of Florida, Gainesville, Florida32611. Before publicizing
this publication, editors should contact this address to determine availability. Printed 4/92.




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