x°/EPA
United States
Environmental Protection
Agency
Environmental Research
Laboratory
Narragansett Rl 02882
Research and Development
EPA-600/S3-82-017 July 1982
Project Summary
Development of Techniques
and Methodology for the
Laboratory Culture of Striped
Bass, Morone saxatilis
(Walbaum)
Bruce A. Rogers, Deborah T. West in, and Saul B. Saila
This summary describes the research
undertaken to develop laboratory culture
techniques for striped bass (Morone
saxatilis) that could be used to provide
an adequate supply of various life stages
of this important fish species for water
quality and hazard evaluation testing.
For each of the four life stages defined
here (egg, larval, juvenile, and adult) the
upper and lower lethal levels where ap-
plicable and an approximation of optimum
conditions were defined with regard to
physical characteristics of the environ-
ment including temperature, salinity,
dissolved oxygen, light, and turbidity.
Satisfactory laboratory diets were de-
fined and verified for each life stage. A
comprehensive set of procedures was
developed and described in a step-by-
step manner for use by research person-
nel wishing to maintain laboratory pop-
ulations of striped bass for physiological
and toxicological use.
This Project Summary was developed
by EPA's Environmental Research Labo-
ratory, Narragansett, Rl, to announce
key findings of the research project that
is fully documented in a separate report
of the same title (see Project Report
ordering information at back}.
Introduction
Striped bass, Morone saxatilis, is an
important commercial and sport fish
species with a center of distribution be-
tween the Hudson River and the mouth
of Chesapeake Bay. Individuals of this
species ascend major rivers to spawn,
use coastal estuaries as nursery grounds,
and as adults make seasonal migrations
along the coast rarely straying more than
five miles from the shoreline. Because it
passes its entire life cycle in the waters
immediately adjacent to the Boston-
Washington, D.C., megalopolis, it is
subjected to the most intense effects of
man-made pollution and environmental
alteration. In spite of these abuses, the
Atlantic population of striped bass has
until recently enjoyed great abundance.
Although in the past a considerable
amount of research has been done on
the culture of the species for stocking
into southern reservoirs, no reliable cul-
ture methodology has been developed
for maintaining all of the life stages of
the striped bass in the laboratory where
the effects of various pollutants may be
determined in physiological studies and
bioassay experiments.
This study was undertaken to develop
a reliable culture protocol for all life
stages of the striped bass. Armed with
such a protocol, researchers will be in a
better position to examine the effects of
water borne pollutants on this resilient
but vulnerable species.
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Discussion
The striped bass is a desirable candi-
date for toxicological investigations in
the United States for the following
reasons. The species inhabits a wide
range and is distributed along all three
coasts. It is a commercially and recrea-
tionally important species throughout its
range. It is also an ecologically important
member of the community it inhabits,
not only along the coasts, but also within
the coastal plain rivers. Its life stages are
euryhaline and eurythermal, making them
extremely useful in studies to determine
sublethal differences in the physiology
of toxicants over broad salinity and/or
temperature ranges. In addition to being
easy to culture, a great deal of the back-
ground research on this species has been
reported.
To date, striped bass culture has been
undertaken primarily by federal or state
fish hatcheries to stock lakes, reservoirs
and impoundments for sport fishing and
shad (Dorosoma sp.) control. This work
is done almost exclusively in fresh water,
either in a hatchery or in ponds (Bonn et
al., 1976). The culture methodology
recommended in this report for the life
history stages of the striped bass, how-
ever, utilized sea water wherever possible.
This was the case not only because of its
availability and cost-effectiveness of
use, but primarily to keep disease prob-
lems to a minimum. Although some
fresh water (to reduce salinity) is needed
during spawning and early larval stages,
juveniles and adults feed and grow in sea
water.
Using the methods recommended in
this research and summarized below,
striped bass life history stages can be
cultured which are representative of the
species. Figure 1 and Table 1 describe
developmental stages of striped bass
larvae to metamorphosis.
Culture Methods Outlined
The outline of recommended methods
to secure batches of larvae or juveniles
for toxicological studies that follows is
based on the details presented in Sec-
tions 8-1 1 of the full report available from
NTIS. In general, if 20,000 prolarvae are
required for studies, then a minimum of
40,000 eggs are needed. This estimate
is based on 50% survival, although egg
survival varies from 10-20% for artifi-
cially spawned to 60-90% for naturally
spawned (see Table 2) and fertilized
eggs. This survival rate can be increased
to 40-50% by using antibiotics in the
rearing water. If 2,000 post-larvae are
required, then 2,500 prolarvae (80%
survival through initial feeding), or 5,000
eggs are required. These estimates are
provided as a guide and may vary with
broodstock, investigator, facilities and
other variables.
A. Fertilized eggs can be obtained from
natural or artificial spawnings.
1) Artificially spawned eggs re-
quire maintenance of broodfish
(mature adults) in culture system
equipped with temperature, sa-
(b)
(a)
Figure 1. Developmental stages (after Manuseti, 1958) of striped bass larvae to metamorphosis.
Refer to Table 1 for further description of stages.
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Table 1. Developmental Stages
Age
of Striped Bass, Reared at About 17°C, Unless Otherwise Stated, Through Transformation
Length mm TLa Characteristics
25.8 hours after fertilization (4)>> 3.25-4.06
36-48 hours after fertilization (2) 2.5-3.7
51.8 hours after fertilization (4) 3.25-4. 71
1st day after hatching (4) 3.58-5.12
4.23-5.20
2-5th day after hatching (1,2) 4.5-5.2
3rd day after hatching (3) 5.2
(41 4.71-6.23
5.O4-5.77
4th day after hatching (3) 5.8
(4) 5.5-7.5 (live)
5th day after hatching /I) 5.5-5.8
6th day after hatching 13) 6.0
6th- 7th day after hatching (4) 5.5- 7.5 (live)
8th day after hatching (1) 5.8-6.5
(3) 6-9
10-15th day after hatching (2) 7.5
10th day after hatching (3) 9.0
15th day after hatching (1) 10-12.5
18th day after hatching (3) 13.0
20-30th day after hatching (1,2) 10, 12-16
30 days after hatching (4) 13.1-15.4
30-40th day after hatching (2) 15 (stunted)
40 days after hatching (4) 11.9-20.4
40-50th day after hatching (1) 22-35
50- 70th day after hatching (1) 35-45
(2) 20
60-80th day after hatching (2) 25
Hatching completed for eggs at 24 °C. (aft
Hatching occurs, (a)
Hatching completed for eggs at 18°C. (a)
Eyes almost fully pigmented; pigmented ventrally; one-third yolk
reabsorbed at 24 °C.
Eyes only partially pigmented; yolk slightly reabsorbed at 18°C.
Yolk sac partly absorbed, eyes pigmented yellow, black & orange,
differentiation of jaws and digestive tract begun, pectoral buds
formed fan-like fin, 21 -23 myotomes. (b)
Eyes pigmented, jaws developing, pectoral fins become differentiated.
Eyes pigmented; mouth pans moving; pigmented ventrally jaw to oil;
yolk three-fourths reabsorbed; pectoral buds present at 24 °C.
Eyes pigmented; gut differentiated; ventrally pigmented; pectoral
buds visible at 18°C.
Small chromatophores along ventral edge of entire yolk sac.
Yolk absorbed at 24 °C.
One-third yolk reabsorbed, commencement of intestinal peristalsis,
23-24 myotomes. Swimming pelagically. (c)
Oil globule and yolk nearly absorbed, pigmentation ventrally. (c)
Yolk absorbed at 18°C. (d)
Teeth on jaws, orange pigment in caudal (heteroceral) area,
differentiation of stomach, three-fourths yolk reabsorbed, 25
myotomes. Transition to active pelagic feeding, (d)
Second dorsal and anal slightly differentiated, well-developed mouth
parts, (d)
Yolk sac fully absorbed and no oil globule visible, pectorals only fins
visible, teeth visible, generally pigmented on body, (e)
Pectorals only fins developed, ready for food.
Division of fin fold into three divisions, complete reabsorption of oil
globule, single-chamber gas bladder filled with air. Feeding on
plankton, (e)
Dorsal and anal fin rays well differentiated and rudimentary spines
observed, (f)
Differentiation of rays in caudal, anal and dorsal fins. First dorsal
elements and pelvic fins absent, myotomes correlated with number of
vertebrae, (g)
Metamorphosis at 24 °C.
Soft dorsal, anal and caudal (homocercal) fins well differentiated,
spinous and pelvic fins not well developed and well ossified, no
stripes visible yet. Initial formation of lateral-line scales (Murawski,
1958). (h)
Metamorphosis at 18°C.
Differentiation of rays in first dorsal and pectoral fins. Full
complement of lateral-line scales by 30 mm (Murawski, 1958).
Scales
Scales observed for first time, fins except larval pelvic in various
stages toward full meristic count, pigmentation stronger.
Covered with scales, 3 anal spines and full complement of meristic
characters, body covered with melanopores.
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Table 1.
(continued)
Age
Length mm TL"
Characteristics
80-90th day after hatching (1) 50-80
70-100th day after hatching (2) 30
3-4 weeks after hatching 13) 36
Appearance of longitudinal stripes.
Meristic counts complete except for fin rays, body pigmentated.
Fully developed fins and rays, pigmentation of black dots.
' Total length measured on preserved samples unless otherwise stated.
b Numbers in parenthesis refer to source, i.e., (1) Doroshev (1970); (2) Mansueti (1958); (3) Pearson (1938); and (4) Rogers et al.
(1977).
0 Letters in parenthesis refer to Figure 11.
Table 2. Percent Survival Through Hatching of Striped Bass Eggs from Artificial
and Natural Spawnings
Incubation
Salinity* (O/oo)
Artificially induced spawning
0
5
10
15
Incubation Temperature (°C)
16 18 20
58.5 64.3
(561) (280)
1.2 -
(249)
19.3 -
(165)
31.2
(160)
—
7.4
(244)
11.6
(215)
0
21
4.7
(536)
5.4
(185)
11.6
(205)
• 0
Natural matured spawning
0
10
15
77.9
(384)
90
(10)
90
(10)
90
(10)
71.0
(473)
71.5
(421)
90
(10)
90
(10)
80
(10)
* Percent survival at Q°/oo and 16°C (60°F), 18°C (65°F), and 21 °C (70°F)
reported by Shannon (1970). Survivals at the other salinity-temperature combi-
nations are results of this study.
+ ( ) = number of eggs per treatment.
Unity, and photoperiod control
with subsequent controlled
spawning.
2) Naturally spawned eggs may be
obtained easily by plankton or
neuston net fishing in spawning
rivers at the time of spawning
(February-May).
3) Collection of naturally spawned
eggs insures genetic diversity
not available among progeny of
a mating under controlled
spawning.
B. Handling of fertilized eggs to maxi-
mize survival and hatching.
1) Eggs collected from plankton
tows must be separated before
transporting them to rearing
containers.
2) Eggs secured from artificial
spawnings can be stocked
directly into rearing containers
at a rate of approximately 100
per liter.
3) Handle eggs only in water, i.e.,
dip or pipette or siphon. Do not
use dip nets.
4) When transferring shipped eggs
to rearing containers check that
water temperature of two are
within 1 °C of each other and
that rearing water quality is opti-
mum for egg survival (Table 3).
5) Water quality, especially tem-
perature, dissolved oxygen and
salinity, should be monitored
daily and maintained at optimum
levels (Table 3).
C. Handling of larvae to maximize sur-
vival and growth.
1) Easiest method of securing pro-
larvae is from eggs on hand that
hatch.
2) The recommended larval rearing
system is static prepared tank
system modified from Houde &
Ramsey (1971) and described
in greater detail in the full report
available from NTIS.
3) Larvae can be stocked at 100
per liter until actively feeding,
when densities should be re-
duced to approximately 50 per
liter.
4) Growing larvae should be graded
to nearly equal size fish to reduce
cannibalism.
5) The water quality in larval rearing
containers should be monitored
daily and maintained at optimum
level (Table 3).
6) Larvae should be fed 10-20% of
their dry body weight at least
twice daily beginning about 4-5
days after hatching an approved
strain of newly-hatched brine
shrimp nauplii. Table 4 lists bio-
chemical characteristics of some
life diets for larval bass.
7) Growth rates at various temper-
atures are detailed in Figure 2.
8) As larvae reach metamorphosis
other foods such as ground squid
or prepared diets (moist pellets)
can be added to adult brine
shrimp.
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Table 3. Summary of Optimal Rearing Conditions for the Various Striped Bass Life Stages
Eggs
Pro larva
Larvae
Postlarvae
Juveniles &
Subadults
Adults
ABIOTIC FACTORS
Temperature
Salinity
Dissolved Oxygen
Light
Turbidity
BIOTIC FACTORS
Diet
Density
16-20°C 16-21°C
2-IQO/oo 5-150/oo
air saturated
natural
photoperiod
500 mg/1"
not applicable not applicable
50- 75 per liter 50-25 per liter
air saturated
natural
photoperiod
18-22°C >10and<25°C
10-200/00 70-300/oo
air saturated
natural
photoperiod
<4 nig/ft
15-20% body weight 5-8% body weight
(dry) twice daily (wet) per day
30- 1 0 per liter
1 0-2 bass per 100
liters
>10and<24°C
70-300/oo
air saturated
natural
photoperiod
3-5% body weight
(wet) daily
2.4 g/l maximum
a Fine grained sediment.
b Bentonite.
25-
20-
6
OS
ID-
Humphries and Gumming.
1973
Mansueti, 795.
Rhodes and
Merriner,
1973
Rogers et al.,
1977
10
I
20
30 40
Days after hatching
50
60
70
Figure 2. A comparison of growth rates observed under fixed temperature
regimes (Rogers et al.. 1977) with those obtained in earlier studies under
conditions of increasing temperature.
D. Handling juveniles.
1) Juveniles, if needed for research,
can be reared from eggs or lar-
vae, or collected by seining in
spawning rivers.
2) Juveniles collected from the
field should be kept separate
from any reared or other col-
lected bass already in the cul-
ture system.
3) Water quality should be moni-
tored daily and maintained at
optimum conditions (Table 3).
4) Juveniles can be fed frozen brine
shrimp, ground squid, prepared
diets, or commercial trout feeds;
the first is generally preferred.
Conclusions
During the course of this study all of the
life stages of the striped bass from egg to
adult were successfully maintained under
laboratory conditions. The temperature,
salinity, dissolved oxygen, light and tur-
bidity requirements of all life stages were
either determined empirically, approxi-
mated from environmental data, or where
reported by other workers corroborated
in our laboratory. Optimum and survival
limits for each of these parameters were,
where appropriate, specified. By main-
taining conditions within these bounds,
striped bass eggs were repeatedly reared
through to the juvenile stage. A popula-
tion of striped bass adults was success-
fully maintained in captivity for five
years. Despite repeated attempts, we
were unable during the course of this
study to successfully induce spawning
in the laboratory. Sexually mature adults
of both sexes, however, did occur
among our captive population. A step-
by-step culture methodology has been
prepared for use by future workers.
References
Bonn, E.W., W.M. Bailey, J.D. Bayless,
K.E. Erickson, and R.E. Stevens(eds.).
1976. Guidelines for striped bass cul-
ture. Striped Bass Committee, South-
ern Division American Fisheries Society.
103 p.
Doroshev, S.I. 1970. Biological features
of the eggs, larvae and young of the
striped bass (Roccus saxatilis [Wal-
baum]) in connection with the problem
of its acclimation in the U.S.S.R. J.
Ichth. 10(2):235-278.
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Helf rich, P. 1973. The feasibility of brine
shrimp production on Christmas Island.
U.S. Sea Grant Technical Report UNIHI-
SEA-GRANT-TR-73-02. 1 73 p.
Humphries, E.T., and K.B. Gumming.
1973. An evaluation of striped bass
fingerling culture. Trans. Amer. Fish.
Soc., 102(1):13-20.
Laurence, G.C. 1977. Caloric value of
some North Atlantic calenoid cope-
pods. U.S. Fish Bull., 75:218-220.
Mansueti, R. 1958. Eggs, larvae, and
young of the striped bass, Roccussax-
atilis. Ches. Biol. Labs. Contr. No.
112:1-35.
Paffenhofer, G.A. 1967. Caloric content
of larvae of the brine shrimp Anemia
salina. Helg. Wiss. Meer., 16:130-
135.
Pearson, J.C. 1938. The life history of
the striped bass, or rockfish, Roccus
saxatilis (Walbaum). U.S. Bureau of
Fish. Bull., 28(49):825-851.
Raymont, J.E., J. Austin, and E. Linford.
1963, Biochemical studies on marine
zooplankton. I. The biochemical com-
position of Neomys/s integer. J. Cons.
Inter. Explor. Mer., 28(1):354-363.
Rhodes, W., and J. V. Merriner. 1973. A
preliminary report on closed system
rearing of striped bass sac fry to fin-
gerling size. Prog. Fish Cult., 35(4):
199-201.
Rogers, B.A., D.T. Westin, and S.B.
Saila. 1977. Life stage duration in
Hudson River striped bass. University
of Rhode Island Marine Technical
Report 31. 11 1 p.
Shannon, E.H. 1970. Effect of tempera-
ture changes upon developing striped
bass eggs and fry. Prog. 23rd Ann.
Conf. S.E. Assoc. Game & Fish Comm.
pp. 265-274.
Slobodkin, L.B., and S. Richman. 1961.
Calories/gm in species of animals.
Nature (London). 191:299.
Table 4. Caloric and Percent Composition of Some Live Larval Food Items
Food Item
Artemia salina
nauplii
adults
Acartia clausi
Acartia tonsa
Calanus finmarchicus
Calanus helgolandicus
Calories/gram
(ash-free, dry)
5800-6000(1)*
5454-5953(3)
5115-5854(3)
5664 ± 8612)
6835 ± 191(2)
5515 ± 277(5)
Percent of Dry Weight
Lipid Protein
15.04-27.24
6.51
5.8
10.5-47.0
11.5
42.5-50. 2(1 J
62.78(1)
82.6(4)
30-77(4)
75.2(4)
*Numbers in parentheses refer to source: (1) Helfrich et al. (1973); (2) Laurence
(1977); (3) Paffenhofer (1967); (4) Raymont et al. (1963); and (5) Slobodkin
and Richman (1961).
Bruce A. Rogers, Deborah T. Westin, and Saul B. Saila are with the Graduate
School of Oceanography, University of Rhode Island, Kingston, Rl 02881.
Allan D. Beck is the EPA Project Officer (see below).
The complete report, entitled "Development of Techniques and Methodology for
the Laboratory Culture of Striped Bass, Morone saxatilis/' (Order No. PB
82-217 795; Cost: $22.50, subject to change) will be available only from:
National Technical Information Service
5285 Port Royal Road
Springfield, V'A 22161
Telephone: 703-487-4650
The EPA Project Officer can be contacted at:
Environmental Research Laboratory
U.S. Environmental Protection Agency
South Ferry Road
Narragansett. Rl 02882
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