c/EPA
United States
Environmental Protection
Agency
EPA 600 8-87 004
January 1987
Research and Development
Methods for Spawning,
Culturing and Conducting
Toxicity-Tests with Early
Life Stages of Four
Atherinid Fishes:
The Inland Silverside,
Menidiaberyllina, Atlantic
silverside, M. menidia,
Tidewater Silverside,
M. peninsulae and
California grunion,
Leuresthes tenuis
-------
Methods for Spawning,
Toxicity-Tests with Ea
Atherinid Fishes: The
beryllina, Atlantic silvers
silverside, M. peninsulae
EPA/600/8-87/004
January 1987
(pulturing and Conducting
ly Life Stages of Four
inland silverside, Menidia
de, M. menidia, tidewater
and California grunion,
Leurestftes tenuis
Douglas P. Middaugh, Michael J. Hemmer and
Larry R. Goodman
Environmental Research Laboratory
Gulf Breeze, Florida 32561
Office of Research and Development
U.S. Environmental Protection Agency
Washington, DC 20460
-------
DISCLAIMER
This document has been subjected to the Agency's peer and
administrative review, and has been approved for publication as an
EPA document.
The mention of trade names in this document does not imply
endorsement by the U.S. Environmental Protection Agency.
11
-------
FOREWORD
Timely assessment of the environmental risks of pesticides and toxic
substances to fish requires that test species be readily available.
Methods for the acquisition, spawning, culture and testing of the
early life stages of marine and estuarine fishes should be formulated
and presented in a format that will enable the experienced aquatic
biologist to conduct tests with minimal difficulty. Moreover, a
compilation of methods should provide for utilization of fishes from
all coastal regions of the United States.
This manual presents methods for field and/or laboratory spawning
of four species of atherinid fishes including:
• Inland silverside, Menidia beryllina
Estuarine populations - Cape Cod, Massachusetts to Texas.
Freshwater populations - States adjacent to Mississippi River
Basin; and Texas, Oklahoma, California.
• Atlantic silverside, Menidia menidia
Estuarine populations - Maine to N.E. Florida.
• Tidewater silverside, Menidia peninsulae
Estuarine populations - N.E. Florida to Texas.
• California grunion, Leuresth.es tenuis
Coastal populations - San Diego to Los Angeles, California.
Procedures are also presented for culturing and conducting acute
and early-life stage toxicity tests with each of the species. All of the
methods have been used extensively by investigators at the Gulf
Breeze Environmental Research Laboratory. Guidelines provided in
this manual are based upon a compilation of studies that have been
published in the peer-reviewed literature.
111
-------
ABSTRACT
Procedures are presented for spawning, culturing and conducting
acute and chronic toxicity tests with four atherinid fishes: the
inland silverside, Menidia beryllina, Atlantic silverside, M.
menidia, tidewater silverside, M. peninsulas, and California
grunion, Leuresthes tennis. Guidelines also are provided for growing
of food organisms (Chlorella sp., Brachionus plicatilis, and Artemia
sp.) that are required for successful culture and testing of the
atherinid fishes.
IV
-------
ACKNOWLEDGEMENTS
The authors wish to thank D.A. Bengtson, Department of Food
Sciences and Technology, Nutrition and Dietetics, University of
Rhode Island; W.S. Hall, Aquatic Ecology Section, Applied Physics
Laboratory, Johns Hopkins University; and D.J. Klemm, Newtown
Fish Toxicology Laboratory, J.R. Clark, Gulf Breeze Environmental
Research Laboratory, and W.A. Rabert, Environmental Review
Divison, Office of Toxic Substances, U.S. Environmental Protection
Agency for providing a critical review of an early draft of this
document. Steven Foss drafted the figures and Val Caston typed
several early drafts of this document.
-------
CONTENTS
Foreword iii
Abstract iv
Acknowledgements v
I. Introduction 1
II. Biology of the Atherinids - distribution,
ecology, reproduction, identification, collection,
handling, spawning, larval culture 2
A. Inland silverside, M. beryllina 2
B. Atlantic silverside, M. menidia 12
C. Tidewater silverside, M. peninsulae 18
D. California grunion, L. tennis 23
III. Acute toxicity tests, M. beryllina, M. menidia,
M. peninsulae, and L. tenuis 29
A. Static tests (96 hours) 29
B. Flow-through tests (96 hours) 30
IV. Early life-stage toxicity tests, M. beryllina, M. menidia,
M. peninsulae 31
A. Seawater 31
B. Exposure system 31
V. Early life-stage toxicity tests, L. tenuis 33
A. Seawater 33
B. Exposure system 33
VI. Batch culture of the alga, Chlorella sp. and
mixohaline rotifer, Brachionus plicatilis 35
A. Apparatus used 35
B. Media preparation and Chlorella sp. culture 35
C. Culture of B. plicatilis 38
VII. Hatching of brine shrimp, Artemia sp 40
A. Strains used 40
B. Apparatus required 40
VI
-------
C. Hatching procedure 40
D. Enumeration ofArtemia nauplii 41
E. Nutritional quality ofArtemia sp 41
III. References 42
IX. Appendices 49
A. Selected biogeographical data for occurrence
of the inland silverside, M. beryllina 49
B. Recommended environmental variables and feeding
regimes for laboratory spawning of three
atherinid fishes 50
C. Recommended environmental variables and feeding
regimes for laboratory incubation and larval rearing
of four atherinid fishes 51
D. Selected biogeographical data for occurrence
of the Atlantic silverside, M. menidia 52
E. Selected biogeographical data for occurrence
of the tidewater silverside, M. peninsulae 53
F. Selected biogeographical data for occurrence
of the California grunion, L. tenuis 54
G. Recommended test parameters and feeding regimes
for conducting static or flow-through 96 hr acute
toxicity tests with 14 day-old atherinid fishes 55
H. Enrichment media for seawater used to grow
Chlorella sp 56
vn
-------
I. INTRODUCTION
This methods manual provides guidelines for conducting toxicity
tests with four atherinid fishes: the inland silverside, Menidia
beryllina; Atlantic silverside, Menidia menidia; tidewater
silverside, Menidia peninsulae; and California grunion, Leuresthes
tenuis.
We have conducted research to determine optimal conditions for
collecting, handling and transport of the three species of silversides
and for field stripping, fertilization and shipment of embryos of the
California grunion. Methods have been developed for laboratory
spawning of silversides and for incubation of embryos and culture of
larvae of all four atherinids. Toxicity test methods also were
developed. The methods described include acute static and flow-
through procedures as well as early life-stage (ELS) test methods for
each species.
An important aspect of fish culture is the availability of adequate
food resources. This manual provides information required for
growth of the mixohaline rotifer, Brachionus plicatilis, and Artemia
sp. nauplii. These two food items are essential for successful culture
and testing of the larval atherinid fishes.
The manual has been formatted to provide a complete synopsis of
culture and testing procedures for individual species. To avoid
redundancy and save space, sections describing procedures for M.
menidia and M. peninsulae at times refer the reader to methods
previously described for M. beryllina.
Other procedures for fish transport, culture of embryos and larvae,
and exposure to toxicants may work equally well. Our recommended
methods have been used repeatedly with success at the Gulf Breeze
Environmental Research Laboratory.
-------
II. Biology of the Atherinids
A. The inland silverside, Menidia berylllna, and the Mississippi
silverside, Menidia audens, are now considered conspecific
(Johnson, 1975; Chernoff et al., 1981).
1. Geographical distribution — M. berylima is a ubiquitous
resident of estuaries, coastal rivers and numerous lakes from
Massachusetts to Texas and is also found in the Mississippi
River Basin (Sisk and Stephens, 1964; Gomez and Lindsay,
1972; Clay, 1975; Johnson, 1975; Chernoff et al., 1981).
Moreover, it occurs in Clear Lake, California where it was
introduced in 1967 (Cook and Moore, 1970) and has since
moved to the Sacramento-San Joaquin River System (Moyle
et al., 1974), and the Lexington Reservoir (Fisher, 1973). The
general biogeographic distribution of M. berylllna is shown in
Figure 1 and a compilation of selected biogeographical data
are provided in Appendix A. These data, along with the
literature citation for each entry, provide a detailed listing of
potential collection locations for each of the species described
in this manual.
2. Ecology and reproduction — M. beryllina is euryhaline, living
in freshwater lakes, rivers and reservoirs, and in coastal
areas at salinities from 0 to 35 %o (Robbins, 1969; Hubbs et al.,
1971; Echelle and Mosier, 1982). Although estuarine forms
seem to prefer salinities of 19 %« or less (Johnson, 1975;
Middaugh et al., 1986); in the Laguna Madre, Texas, M.
beryllina has been found at a salinity of 75 %o and reported as
abundant at 45 %o (Simmons, 1957). The duration of
reproductive activity varies according to geographic location
and, apparently, water temperature. Sexually mature M.
beryllina are found in June and July at Woods Hole,
Massachusetts (Rubinoff and Shaw, 1960). Although the
spawning period is also brief in Rhode Island, lasting for only
several weeks (Bengtson, 1984) it has been noted that if M.
beryllina from these latitudes are maintained in the
laboratory, they will continue to spawn throughout much of
the year (pers. comm., Bengtson, Dept. of Food Science and
Nutrition, Univ. of Rhode Island, Kingston, 02881). In the
Chesapeake Bay, spawning occurs from early April to late
September (Hildebrand and Schroeder, 1928), at Beaufort,
North Carolina from March to September (Hildebrand, 1922)
and in Tampa Bay, Florida, throughout the year with the
-------
-------
exception of January and August (Springer and Woodburn,
1960). In coastal Texas, Gunter (1950) reported gravid
females in February and March at respective water
temperatures of 20 and 25°C. In Lake Texoma, Oklahoma,
reproduction generally occurs from late March through mid-
July at water temperatures of approximately 15 to 30°C
(Mense, 1967; Hubbs et al., 1971; Hubbs, 1982). Populations
in Lexington Reservoir, California, spawn from early May
until mid-September (Fisher, 1973). Clear Lake populations
spawn from about late March through early July (Cook and
Moore, 1970).
3. Identification - The inland silverside, Menidia beryllina, and
the Mississippi silverside, Menidia audens, are now
considered conspecific (Johnson, 1975; Chernoff et al., 1981).
The largest inland silverside, Menidia beryllina, examined by
Robbins (1969) was a female 90.7 mm standard length (SL).
Females attain a larger size than males. Scales are large,
usually well imbricated, and with well developed circulii and
radii. Usually there are not more than 37-50 scales in the
lateral series (Robbins, 1969; Chernoff et al., 1981). The first
dorsal fin has 2-7, usually 4 or 5 spines, with the origin well
in advance of the anal fin origin and lying over a point above
the anterior edge of the anus. The second dorsal fin has one
spine and 7-11, usually 9 or 10, rays. The anal fin has one
spine and 13-20, usually 15-17, rays (Robbins, 1969). The gas
bladder in M. beryllina is long and translucent (Echelle and
Mosier, 1982), and extends to a position approximately above
the fourth anal fin ray (Robbins, 1969). It is bluntly rounded
posteriorly and is a good characteristic for use in the field to
quickly identify M. beryllina and separate this species from
Menidia menidia and M. peninsulae which have a truncated
opaque gas bladder. Another diagnostic characteristic for
separating M. beryllina and M. peninsulae is measurement of
the horizontal distance between the origins of the first
spinous dorsal fin and anal fin (Chernoff et al., 1981). This
measurement is s7% of SL in M. beryllina and zl% of SL in
M. peninsulae. Moreover, mature and hydrated M. beryllina
eggs, 0.9 to 1.1 mm diameter, possess 1 or 2 long-thick
filaments (length usually is equal to 15 to 30 egg diameters)
and 1 to 15 short-thin filaments. In contrast, fully hydrated
Menidia menidia and M. peninsulae eggs possess 15 to 50
short-thin filaments and no long-thick filaments. M. beryllina
also may be taken in freshwater areas where the brook
silverside, Labidesthes sicculus, is present. M. beryllina is
more robust than L. sicculus. Viewed from above, the
premaxillary of L. sicculus is pointed, forming a cone shape
while that of M. beryllina is crescent shaped, not forming a
-------
pointed beak. L. sicculus scales are also smaller than those of
M. beryllina (Blair et al.f 1968).
4. Collection, handling, spawning and transport.
a. Where — Sexually mature fish generally will be available
in lakes, rivers and estuarine habitats from March or April
through August at water temperatures of 15 to '30°C
(Hubbs, 1982; Hubbs, 1976; Rubinoff and Shaw, 1960;
Hildebrand and Schroeder, 1928; Springer and Woodburn,
1960; Gunter, 1945). Certain locales such as the Upper
Laguna Madre, Texas have populations that spawn
throughout the year (Simmons, 1957) while at northern
latitudes, such as in Rhode Island estuaries, the breeding
season may be as short as two or three weeks in late June
and early July (Bengtson, 1982). M. beryllina frequents
shallow waters along shorelines where sandy to partially
vegetated substrates occur. Beaches, bordering open but
protected waters, are preferable for collecting. In Lake
Texoma, M. beryllina is found in areas with a sandy bottom
(Mense, 1967; Hubbs, 1982). They are found in a similar
habitat in Clear Lake, California (Cook and Moore, 1970).
Elston and Bachen (1976) made collections in the
Lexington Reservoir, California in a shallow sandy area
with sparse growth of rooted aquatic plants and a border of
tule beds, Scirpus spp.
b. When — The optimal time to collect M. beryllina is during
early to mid-morning between 0800 and 1200 hrs. This
time is recommended because of the diel reproductive
pattern noted for M. beryllina in Lake Texoma where most
mature fish were ripe between 0800 and 1200 hrs (Hubbs,
1976). Similar reproductive timing was noted in Lexington
Reservoir, California where spawning occurred during
mid-morning over a vegetated gentle slope at water depths
of 2.5 to 60 cm (Fisher, 1973). This same reproductive
timing was noted by Robbins (1969) in Lake Kustis,
Florida. Moreover, Middaugh et al., (1986) were able to
determine the sex of individual M. beryllina collected
during mid-morning from Blackwater Bay, Florida in mid-
April. Extrusion of ripe eggs and sperm was possible,
suggesting that estuarine populations also may be ready- to
spawn during early to mid-morning.
c. How - A 1 x 10-m bag seine with knotless 5-mm mesh is
ideal for collecting. Since M. beryllina typically resides in
shallow water (<1.5 m deep), they are easily captured by
seining close to shore. It is important to avoid total
beaching of the bag seine when collecting M. beryllina.
These fish will quickly die if removed from water and ripe
females often abort their eggs if stranded. Ideally, the bag
-------
portion of the seine containing captured adults should
remain in water 5 to 15 cm deep (Middaugh et al., 1986).
d. Spawning in the field.
1. Refer to Figure 2 for a diagrammatic explanation of the
procedure outlined below.
2. Immediately after seining (while still on the beach) --
Three to five ripe females should be dipped into a bucket
of water from the collection site to remove sand and
detritus.
3. Eggs — Females with hydrated eggs are stripped into a
20 cm diameter glass culture dish containing ambient
temperature water (Middaugh and Lempesis, 1976) or
directly onto a nylon screen (0.60 to 1.0 mm mesh) which
is then gently lowered into the culture dish of ambient
temperature water with the eggs on the upper surface of
the screen (Barkman and Beck, 1976). If excessive
pressure on the abdomen is required to strip eggs, the
female should be discarded. Mature eggs are clear with
an amber-green hue.
4. Milt - Several males should then be stripped into a
seperate culture dish containing ambient temperature
water. Eggs are then fertilized by pouring water from
the dish containing sperm into the dish containing eggs.
Upon contact with water, adhesive threads on mature
eggs uncoil, making enumeration and separation
difficult. If eggs are stripped directly into the culture
dish, one end of a nylon string may be dipped into the
dish and gently rolled so the embryos adhere (Middaugh
and Lempesis, 1976). The Barkman and Beck (1976)
technique for attaching the eggs to nylon screening
minimizes the natural clumping tendency due to
entanglement of the filaments on Menidia eggs and is
recommended.
5. Strings of embryos or embryos on screens - These are
transported to the laboratory by placing in an insulated
glass or stainless steel container half-filled with water
from the collection site.
6. In the laboratory - Embryos should be suspended in a 10
to 20 1 all-glass container. The temperature and salinity
of the water at the collection site should be used as
guidelines for holding conditions in the laboratory. If
convenient, water from the collection site should be used
for holding embryos in the lab. Since M. beryllina
embryos are euryhaline and eurythermal, slight
changes in salinity and incubation temperatures (to a
new constant salinity and temperature) are acceptable,
-------
FIELD SEINING
HAND STRIP EGGS
HAND STRIP SPERM
INTO A GLASS CULTURE
DISH- THEN ADO TO E06S
06-1 Omm
NYLON SCREEN
ADULTS DIPPED
TO REMOVE SAND
AND DETRITUS
HOLD EMBRYOS IN IO8
AQUARIUM FOR 6- 10
DAYS, 22-25'C
WIDE-MOUTH
VACUUM BOTTLE
WAIT 10-15 MINS
RNSE 3 TIMES
TRANSFER SCREEN
WITH EMBRYOS
~X PARTIALLY FILLED
WITH WATER
SCREEN WITH
s ATTACHEl
EMBRYOS
GLASS LINED
OR
STAINLESS STEEL
TRANSFER LARVAE TO
ALL GLASS AQUARIUM
WHEN 4-6 DAYS OLD
USE SYPHON TUBE
GLASS TUBE OR BEAKERS
DO NOT USE DIPNETS
ACUTE TOXICITY
TESTS
SCREENED
DRAIN
DAYS 1-5 FEED 10,000 ROTIFERS/J! 2X DAILY
DAYS 6- 14 FEED 5,000 ARTEMIA NAUPLII/* 2X DAILY
Figure 2. Diagrammatic explanation of procedure for collection,
fertilization, transport, culture and testing of
embryonic and larval stages ofMenidia.
-------
provided the constant laboratory regime is not likely to
result in salinity or thermal stress. The optimal
temperature for survival of embryos and growth of
young larvae is 25°C (Hubbs et al., 1971; Middaugh et
al., 1986). A salinity of 15 %0 produced the best survival
and growth of larval M. beryllina from parental stock
taken from a brackish water (1-5 %o) habitat (Middaugh
etal.,1986).
e. Transport of adults to the laboratory.
1. Handling — Adults should be removed from the bag end
of the seine by hand and placed immediately in a bucket
containing 12 to 15 1 of ambient temperature water from
the collection site. Not more than 25 adults should be
placed in a bucket at any one time and these should be
quickly and gently transferred to the transport tank.
2. Transport tank — A container of 100 to 350 1 volume
with smooth sides (fiberglass, rigid styrofoam or
stainless steel) should be used to transport adults to the
laboratory. A loading capacity of 5.Og I-1 (2adultsof ~2.5
g each) should not be exceeded. The transport tanks
should be partially filled with water before addition of
adult fish. Vigorous aeration from 2 to 3 airstones
should be provided continuously using a battery-
operated portable aerator.
3. Before transport - Care should be taken to prevent the
introduction of sand, mud, clay or any other abrasive
materials to the transport tank while loading of ambient
water or fish.
4. At the laboratory - Adult fish should be dipnetted with
fine mesh nets and transferred quickly from the
transport tank to buckets containing 12 to 15 1 of water,
then carried to the laboratory spawning system and
divided among the brood tanks as described in the next
section.
f. Laboratory spawning.
1. Spawning system - Inland silversides, M. beryllina may
be spawned in the laboratory on a year-round basis.
Procedures described by Middaugh et al., (1986) provide
for maintenance of a brood stock of 50 individuals, sex
ratio 1:1 (i.e., 25 males + 25 females) in each of 2
circular fiberglass tanks, diameter 1.3 m, water depth 38
cm. A filter reservoir system is employed to maintain
water quality and a March TE5C MD pump (March
Manufacturing Co., Glenview, Illinois) circulates water
from the filter-reservoir system to each spawning tank
(Figure 3). A once-through system without the filter-
-------
FigureS. Laboratory spawning system utilized with Menidia
beryllina, M. peninsulae and M. menidia, A. 1.3 m
diameter holding tanks; B. seawater circulation
pump; C. filter-reservoir tank; D. seawater discharge
lines from circulating pump; E. shunt-return line to
filter-reservoir; F. drain-return line from holding
tanks routed into reservoir tray; G. reservoir filter-
tray. The tray contains several strata of filter
material including (from top) aquarium filter fiber (2-
3 cm deep), activated charcoal (1-2 cm deep), coarse
stone gravel (2-3 cm deep) and crushed oyster shell (1-
2 cm deep).
-------
reservoir can also be used if the incoming water is of
sufficiently good quality. Flow rates are adjusted to
provide a surface current velocity of 4 to 8 cm sec-1 in
each holding tank. Water flows out of the bottom of each
tank through an inverted standpipe. A perforated air
tube positioned at the outer bottom edge of each inverted
standpipe maintains dissolved oxygen at > 5.0 mg I-1
and produces a gentle upwelling current.
2. Feeding schedule -- Fish in each holding tank are fed 8.0
g of Tetramin (Standard Mix-large flake) food each
morning, 0800 to 0900, 4.0 g at 1000 to 1100, 4.0 g at
1400 to 1500 and a final 8.0 g feeding at 1500 to 1700
hrs. Daily feedings may be supplemented between 1100
and 1200 hrs with -150,000 Artemia nauplii. At least
once each week, excess food is siphoned from the bottom
of each tank, and aquarium filter fiber and activated
charcoal in the filter-reservoir system changed.
Approximately 20% of the water in each system is
removed weekly and replaced with temperature and
salinity adjusted water.
3. Water quality — The spawning system should have
water quality similar to that at the collection site. M.
beryllina adults from freshwater habitats such as Lake
Texoma, Oklahoma; Clear Lake, California; or Lake
Eustis, Florida should be maintained in water with a
hardness, pH, alkalinity, and total organic carbon,
similar to the water from which they were collected. In
contrast, fish collected from estuarine locales should be
maintained at salinities similar to those measured at
the collection location.
4. Water temperature and salinity - These environmental
variables should resemble values -encountered at the
collection location. It appears that reproduction in M.
beryllina occurs over a temperature range of ~ 15 to 30°C,
with 25°C considered optimal for embryo development
and survival. Thus adult brood stock should be held at
25°C after an appropriate acclimation period if
collections are made early in the reproductive season
(i.e., when water temperatures may be low). M. beryllina
taken from Blackwater Bay, Florida, temperature 23.7
to 26.3°C, salinity 3 to 5 %o are routinely held at 25°C
and 5 %o while in brood tanks (Middaugh et al., 1986).
5. Photoperiod and light intensity — A photoperiod of
13L:11D is recommended for Menidia brood stock with a
light intensity of 300 lux provided by two banks of 16 cm
long 40 watt "cool white" fluorescent tubes mounted 1.5
m above holding tanks. Timer controlled lights are
10
-------
turned on at 0600 and off at 1900 hrs. Pairs of spawning
tanks must be isolated from outside ambient light and
general disturbances by light-tight curtains. These
curtains remain partially open to facilitate easy feeding
of adult fish during daytime, but are tightly closed at
night. A small exhaust fan is mounted on the plywood
ceiling of our enclosure to prevent a buildup of excess
heat (from the water circulation pump) and humidity
from the holding tanks.
6. Combinations of spawning signals - We recommend
running the circulation pump continuously.
Introduction of a 13L:11D photoperiod (lights on 0600
and off 1900 hrs) is used to mimic the natural
photoperiod during spring-early summer when
reproductive activity is evident throughout the
geographical range of M. beryllina. M. beryllina tend to
spawn throughout the day and night. However, in our
studies most egg release appears to occur between 0800-
1200 hrs each day for populations collected from
freshwater lakes and reservoirs, and 1800-2300 hrs for
estuarine populations. The laboratory spawning time for
freshwater populations is similar to times of spawning
in nature (Hubbs, 1976; Robbins, 1969). Daily egg
production from a tank containing 25 females and 25
males (from an estuarine locale), during 13 days when
eggs were enumerated, ranged from 659 to 4,649
(x = 2,316)(Middaugh et al., 1985). Egg production was
similar for a population of fish taken from Lake Chicot,
Arkansas (Middaugh and Hemmer, unpublished).
7. Spawning substrates — Polyester aquarium filter-fiber
substrates, size —15 cm long x 10 cm wide x 10 cm thick
are suspended just below the water's surface and in
contact with the side of each holding tank. A synopsis of
recommended environmental variables for spawning M.
beryllina and other silversides in the laboratory is
provided in Appendix B.
g. Culture of laboratory spawned Menidia beryllina.
1. Fertilized eggs - They should be removed from the
surface of the spawning substrate, generally between
1300 and 1400 hrs daily. No effort should be made to
tease individual eggs from the substrate, rather,
concentrations of eggs and accompanying substrate are
removed from the main body of the spawning substrate
with forceps. It should be noted, however, that
minimization of the amount of spawning substrate
removed with the embryos is desirable.
11
-------
2. Embryos -- Developing embryos that are attached to
polyester substrates are suspended in a 10 to 20 1 glass
aquarium containing 8 to 18 1 of water adjusted to the
temperature (25°C) and salinity (freshwater to ~20 %o) at
which adult brood fish were held and eggs spawned and
fertilized in the laboratory.
3. Water temperature and salinity -- Utilization of a single
temperature and salinity regime for spawning adults,
embryo incubation, and larval culture and testing
eliminates questions of acclimation and is often helpful
because the same source of water can be used in the
various production and testing systems.
4. Transfer -- Newly hatched larval M. beryllina should be
maintained and fed in the 10 to 20 1 glass culture
container for 4 to 6 days after hatching, with water
temperature and salinity appropriately adjusted. A
density of 5 to 10 larvae I-1 of water is desirable, thus a
maximum of 180 larvae should be placed in a tank
containing 18 1 of water. A 1.5 cm inside diameter (I.D.)
glass tube, approximately 45 cm in length, equipped
with a rubber squeeze bulb is used to make transfers of 4
to 6 day old Menidia. Alternatively a small siphon tube,
3 cm I.D. x 1.5 m in length may be used to siphon fish
from the hatching aquarium to the grow out tank. We
prefer to transfer larvae to 200 ml beakers for
enumeration and then pour the contents into a 20 to 40 1
tank. Under no circumstances should a dipnet be used to
transfer larval M. beryllina,. Mass mortalities will occur
if a dipnet is used.
5. Feeding — The mixohaline rotifer, Brachionus plicatilis,
must be provided on the day that M. beryllina hatch. Two
feedings, one between 0800 and 0900 hrs and a second at
1400 to 1500 hrs are required. At each feeding, rotifers
are added at a rate of 10,000 I-1 of water in the holding
aquarium. Thus an 18 1 volume would require the
addition of 180,000 rotifers, twice daily. This regime is
continued through the fifth day after larval M. beryllina
hatch. On days six through fourteen, 5,000 newly
hatched (<8 hr old) Artemia nauplii I-1 are added each
morning and afternoon. A synopsis of recommended
environmental variables for laboratory incubation of
embryos and larval culture is provided in Appendix C.
B. Atlantic silverside, Menidia menidia.
1. Geographical distribution -- M. menidia ranges as far north
as the Magdalen Islands, Province of Quebec, Canada (Cox,
1921). The southern range is limited to the Atlantic coast of
12
-------
northern Florida (Gosline, 1948; Robbins, 1969). Johnson
(1975) collected a few M. menidia as far south as New Smyrna
Beach, Florida where it seems to intergrade with the
tidewater silverside, M. peninsulae, which is generally found
from Daytona Beach, southward (Chernoff et al., 1981). M.
menidia has been taken from Scarborough Harbor and off
Todd Point in southern Maine (Robbins, 1969). It is a resident
of Essex Bay, Massachusetts (Conover and Kynard, 1981),
southern Rhode Island estuaries (Bengtson et al., 1986), and
is common at Woods Hole, Massachusetts (Kendall, 1902). It
is also found in southern Connecticut estuaries opening into
Long Island Sound (Cadigan and Fell, 1985). Moreover, M.
menidia occurs in the lower regions of Chesapeake Bay
tributaries (Bayliff 1950; Robbins, 1969). Massman (1954)
collected adults approximately 54 km upstream from
brackish water in the James, Rappahannock and Pamunkey
Rivers, Virginia. An offshore migration of M. menidia north
of Cape Hatteras occurs in fall and winter as the estuarine
and nearshore water temperatures drop to about 6 to 8°C
(Conover and Murawski, 1982). South of Cape Hatteras, M.
menidia was extremely abundant and the only fish occurring
in shallow water throughout the winter (Hildebrand, 1922).
In South Carolina, M. menidia is a year-round resident of
intertidal creeks (Cain and Dean, 1976; Shenker and Dean,
1979) and is found in the surf zone of barrier beaches
throughout the winter (Anderson et al., 1977). The general
distribution of M. menidia is shown in Figure 4 and a list of
selected biogeographical data, that should aid in selection of
collection sites, is provided in Appendix D.
2. Ecology and reproduction -- M. menidia is euryhaline and
eurythermal, living in the upper reaches of rivers in the
Chesapeake Bay (Massman, 1954) and migrating offshore
some 50 km during the colder months at northern latitudes
(Conover and Murawski, 1982). De Sylva et al., (1962)
collected M. menidia at salinities ranging from 2 to 35 %o in
the Delaware Bay and found the greatest abundance at water
temperatures from 12 to 30°C. Bayliff (1950) observed that
adults were less numerous in shallow waters of the
Chesapeake Bay during fall as water temperatures declined
to less than 12°C; at 6°C or lower, few M. menidia were to be
found in the shallows. Hildebrand and Schroeder (1928) were
able to collect specimens at a depth of approximately 50 m
during the winter in Chesapeake Bay. A decline in
abundance was noted at temperatures below 10°C in the
North Edisto River estuary of South Carolina (Middaugh,
unpublished). Moreover, Dahlberg (1972) reported that M.
menidia occurred in a Georgia estuary at water temperatures
from 7 to 31.5°C but observed that adults became scarce when
water temperatures dropped below 12°C. M, menidia is an
13
-------
Atlantic silverside, Men id I a menidio
Figure 4. Biogeographica! distribution of the Atlantic silverside, Menidia menidia.
-------
opportunistic omnivore, feeding on mysids, copepods,
molluscan larvae, annelid worms, amphipods, young
gastropods, crab larvae, diatoms and other fishes including
its own young (Kendall, 1902; Bayliff, 1950; De Sylva et al.,
1962; Mulkana, 1966). Sexually mature adults spawn from
March through August depending upon the latitude. Ripe fish
have been found in June at Prince Edward Island, Canada
(Leim and Scott, 1966), and in June and July at Woods Hole,
Massachusetts (Kuntz and Radcliffe, 1918; Rubinoff and
Shaw, 1960; Kendall, 1902). In Salem Harbor,
Massachusetts, Conover and Kynard (1984) noted that M.
menidia had a spawning periodicity that coincided with new
and full moons. Intensity and frequency of spawning was
correlated with the height of daytime high tides. Breeding
occurred from late-April through June at water temperatues
of 9 to 21°C. Daytime spawners deposited eggs on mats of
intertidal, filamentous algae. At the Pataguanset River
estuary in eastern Connecticut, Cadigan and Fell (1985)
reported reproductive activity in M. menidia from early-May
until late-July and observed that water temperature was an
important factor influencing reproductive activity. An
apparently critical lower temperature for reproduction, 16°C,
was reached in mid-late May. Interestingly, Middaugh (1981)
reported that 16°C was required for spawning to occur in M.
menidia from the North Edisto River estuary in South
Carolina. Spawning generally began in early to mid-March
when the critical minimum water temperature was reached
and ended in June or July as estuarine water temperatures
exceeded 30°C. In South Carolina estuaries, M. menidia
primarily spawns during a 3 to 4 day period on daytime high
tides following new and full moons (Middaugh, 1981;
Middaugh et al., 1981). Eggs are deposited on a variety of
upper intertidal substrates including, Spartina alterniflora,
detrital mats and in abandoned crab burrows along erosional
scarps (Middaugh et al., 1981).
3. Identification -- The Atlantic silverside, Menidia menidia is
the largest species in the genus. The largest specimen
examined by Robbins (1969) was a 117 mm SL female. Males
are generally smaller than females. Scales are small to
moderate in size, well inbricated, and usually with well
developed circulii. Branchial lateral line scales are usually
41-47, post pectoral lateral line scales usually 42-46;
predorsal scales 18-22. The first dorsal fin has 3-7, usually 4
or 5 spines; origin is over the posterior edge of the anus or
anal fin origin. The second dorsal fin has one spine and 7-11,
most frequently 8 or 9, rays. The anal fin is long with one
spine and 19-29, usually 21 to 26, rays. The air bladder
scarcely reaches a point above the anal fin origin and is
opaque and abruptly truncate in shape posteriorly. The snout
15
-------
is moderately blunt or sub-conic to angular and is usually
greater than eye length. Jaws are equal. The distal tip of
mandible does not project beyond tip of premaxillary, and the
premaxillary is rounded anteriorly (Robbins, 1969). Mature
and hydrated M. menidia eggs are 1.0 to 1.2 mm in diameter
and bear a cluster of 15 to 50 thin filaments and no thick
filaments.
4. Collection, handling, spawning and transport.
a. Where - North of Cape Cod sexually mature fish will
generally be available in estuaries during late April
through June or July, at water temperatures of 2 to 21°C
(Kendall, 1902; Kuntz and Radcliff, 1918; Rubinoff and
Shaw, 1960; Bengtson, 1985). At Salem Harbor,
Massachusetts sexually mature adults were taken from
late April through June at water temperatures of 9 to 21°C
(Conover and Kynard, 1981). From Cape Cod southward to
Florida, it appears that water temperatures >16°C are
required for spawning in spring (Cadigan and Fell, 1985)
and that reproduction ceases as water temperatures
approach 30°C (Middaugh, 1981).
b. When — Menidia menidia should be collected just prior to
natural spawning runs. These runs at all latitudes appear
to occur 25 during daytime and are timed at, or just after,
high tides (Middaugh, 1981; Conover and Kynard, 1984).
Generally, the 1 to 4 day period after new or full moons is
best for collecting. High tides will occur between —0800 and
1200 hrs on these days.
c. How ~ Refer to section II. A. 4. c. for Menidia beryllina.
d. Spawning in the field.
1. Immediately after seining -- Refer to sections II. A. 4. d.
1-6. for Af. beryllina.
e. Transport of adults to the laboratory.
1. Adults ~ Refer to section II. A. 4. e. 1-4. for M. beryllina.
f. Laboratory spawning.
1. Spawning system — Atlantic silversides, Menidia
menidia, may be spawned in the laboratory during the
natural reproductive season. Procedures described by
Middaugh and Takita (1983) should be utilized. The
system described by Middaugh and Hemmer (1984) and
Middaugh et al., (1986) for spawning M. peninsulae and
M. beryllina (Fig. 3) is also suitable for laboratory
spawning of Menidia menidia (Middaugh and Hemmer,
unpublished).
16
-------
2. Feeding schedule - Refer to section II. A. 4. f. 2. for M.
beryllina; and Appendix B.
3. Water quality - The pH, total organic carbon, and
ammonia in the spawning system should be maintained
at levels similar to the collection location.
4. Water temperature and salinity — M. menldia from
northern latitudes, north of Cape Cod, should be held at
18 to 24°C (Conover and Kynard, 1981) while those from
south of Cape Cod should be held at 20 to 25°C (Cadigan
and Fell, 1985; Middaugh, 1981). Reproductively active
M. menidia occur in freshwater (Massman, 1954) and in
nearly full strength seawater (Anderson et al., 1977;
Middaugh 1981). The salinity in spawning tanks should
be similar to that at the collection site, provided there is
evidence that M. menidia is reproductively active where
collected. Middaugh and Takita (1983) utilized
temperatures ranging from 16 to 25°C and a salinity of
30 ± 2 %o in laboratory spawning studies with M.
menidia. In contrast, Conover and Kynard (1984)
reported laboratory spawning at 13 to 24°C during May
through July for adults collected from Essex Bay,
Massachusetts.
5. Photoperiod and light intensity -- Refer to section II. A.
4. f. 5. for M. beryllina.
6. Combinations of spawning signals — "Tidal" signals,
interruptions of the current velocity, are accomplished
by using an electrical timer to turn off the water
circulation pump for 1 hr at the specified times of 1200 to
1300 and 2400 to 0100 hrs. M. menidia is sensitive to
interruptions in current velocity that mimic conditions
during slack high tides in nature. They will spawn in
the labortory during daytime 1200 to 1300 hrs in
response to the interruption in current velocity, but not
at night, between 2400 and 0100 hrs when the
circulation pump is also turned off (Middaugh and
Takita, 1983; Middaugh, 1981). A synopsis of
recommended environmental variables for spawning M.
menidia in the laboratory is provided in Appendix B.
7. Spawning substrates -- Refer to section II. A. 4. f. 7.
g. Culture of young Menidia menidia.
1. Fertilized eggs and embryos - Refer to section II. A. 4. g.
1-2. for M. beryllina.
2. Water temperature and salinity -- Utilization of a single
temperature and salinity regime for spawning adult M.
menidia; and for embryo incubation, larval culture and
17
-------
testing is advisable. This procedure minimizes questions
of acclimation and is often helpful because the same
source of water can be used in the various production
and testing systems. For northern latitudes we
recommend a culture temperature of 22°C and salinities
of 25 to 30 %o, unless environmental variables vary
widely from these parameters at the locale where fish
are collected. For southern latitudes (south of Cape Cod)
we recommend a temperature of 25°C and salinities of 25
to 30 %o, again with the caveat for environmental
variables.
3. Transfer — Refer to section II. A. 4. g. 4. for M. beryllina.
4. Feeding -- Refer to section II. A. 4. g. 5. for M. beryllina.
However, Middaugh and Lempesis (1976) were able to
obtain acceptable (70%) survival of M. menldia larvae by
feeding them on the day-of-hatch and daily thereafter
with very young (<8 hr old) Artemia nauplii. Similar
results were reported by Barkman and Beck (1976) and
Conover and Kynard (1981). The reader is cautioned: It
is absolutely essential to begin feeding M. menidia
larvae with young Artemia nauplii on the day that
Menidia hatch. A one day delay in adding Artemia
nauplii will result in markedly reduced survival of
larval M. menidia (Middaugh and Lempesis, 1976). If
the use of Artemia nauplii alone (without initial feeding
of Brachionus sp.) is unsuccessful then procedures
outlined in section II. A. 4. g. 5. should be utilized. A
synopsis of recommended environmental variables for
laboratory incubation of embryos and larval culture is
provided in Appendix C.
C. Tidewater silverside, Menidia peninsulae, recently recognized
as a distinct species (Johnson, 1975; Chernoff et al., 1981) was
once considered conspecific with Menidia beryllina (Gosline,
1948; Robbins, 1969). These two atherinids are often found in
close proximity in estuaries along the southeastern coast of
Florida and throughout the Gulf of Mexico (Echelle and Mosier,
1982; Lucas, 1982). The general distribution of M. peninsulae is
shown in Figure 5 and a list of selected biogeographical data are
provided in Appendix E. This list should aid in the identification
of appropriate collection sites.
1. Geographical distribution - M. peninsulae has a disjunct
distribution extending from Daytona Beach, Florida to Horn
Island, Mississippi and Galveston Bay, Texas to Tamiahua,
northern Veracruz, Mexico (Johnson 1975; Chernoff et al.,
1981).
2. Ecology and reproduction - Menidia peninsulae along the
northern part of its range intergrades with M. menidia. At
18
-------
Tidewater silverside, Menidia peninsulae
<£>
FigureS. Biogeographical distribution of the tidewater silverside, Menidia peninsulae.
-------
Fort George Inlet, Duvall Co., Florida, collections yielded 99%
Af. menidia. However, at Flagler Beach, Flagler Co., Florida,
the ratio was 63% M. menidia, 20% hybrids and 19% M.
peninsulae. In contrast, a more southerly location,
Melbourne, Brevard Co., yielded no M. menidia or hybrids
and 100% Af. peninsulae (Johnson, 1975). Moreover, Af.
peninsulae and Af. beryllina are often found in close proximity
in estuaries along the southeastern coast of Florida and in the
Gulf of Mexico (Echelle and Mosier, 1982; Lucas, 1982).
Despite sympatric occurrence and collection of Af. beryllina
and Af. peninsulae in the same seine haul at four localities;
two in Florida and two in Texas, Johnson (1975) observed that
the species were discrete in these areas. Collections yielded a
very low frequency of hybridization. Although M. peninsulae
has been collected at salinities of less than 5 %o to greater
than 35 %<>; Johnson (1975), Echelle and Mosier (1982) and
Middaugh et al., (1986) observed that M. peninsulae typically
resided at salinities of 15 %o or greater while Af. beryllina
generally occurred at salinities of 19 %o or less. In the Crystal
River, Florida locale, Lucas (1982) observed that M.
peninsulae was present throughout much of the year, but
disappeared during December and January, temperature 6.0
to 17.0°C, salinity 22 to 24 %0 and in July, water temperature
30.7°C, salinity 28 %o. In contrast M. peninsulae from the
Pensacola, Florida locale were present in most months but
disappeared from the shoreline in December and January
when water temperatures were below 12°C (Middaugh and
Hemmer, 1986a). Lucas (1982) described three feeding stages
for M. peninsulae from Crystal River, Florida. In early spring,
young-of-the-year fed on tychoplankton and detritus. During
late spring through winter, calanoid copepods and cypris
larvae were selectively eaten. Reproductively active M.
peninsulae fed primarily on amphipods and larval silversides
during February and March. A reproductive peak was
observed only in the spring, no such peak was apparent
during fall. Middaugh and Hemmer (1986a) observed
spawning during March at low tide on a red alga, Ceramium
byssoideum, which was growing in the cracks and crevices of
a rocky substrate just below the low tide line. The annual
reproductive cycle of Menidia peninsulae from Santa Rosa
Island, Florida extends from February through July or
August with the greatest spawning activity during March
through June at water temperatures of 16.7 to 30.8°C.
3. Identification — While differences in M. peninsulae and M.
beryllina are diagnostically clear, these differences are
difficult to express in terms of individuals (Johnson, 1975).
One of the best diagnostic characters for M. peninsulae is the
horizontal distance between the origins of the first spinous
dorsal and anal fins. In Af. peninsulae, this distance is >7%
20
-------
of SL while in M. beryllina the distance is <7% of SL
(Chernoff et al., 1981; Middaugh et al., 1986). Chernoff et al.
(1981) reported measurements of this distance in M.
peninsulae primary type material of 9.3% and 10.7% SL; in
M. beryllina 5.7% and in M. audens (now considered M.
beryllina) 4.4% SL. The least destructive way to identify live
M. peninsulae and M. beryllina is by the relative posterior
extension of the gas bladder. In M. peninsulae the opaque
gas bladder is truncated and extends to the first to third soft
anal ray (Johnson, 1975; Echelle and Mosier, 1982). In
contrast, the translucent gas bladder of M. beryllina is
bluntly rounded and extends to a position approximately
above the fourth or fifth soft anal fin ray. The fully hydrated
eggs of M. peninsulae are 0.9 to 1.1 mm in diameter and
possess 15 to 50 short-thin filaments and no long-thick
filaments.
4. Collection, handling, spawning and transport.
a. Where — Sexually mature fish will generally be available
from early March through late June or early July
(Middaugh et al., 1986). M. peninsulae frequents shallow
waters along shorelines where sandy to partially vegetated
substrates occur and prefers locales where salinities are
S15 %o (Johnson, 1975).
b. When -- On several occassions, M. peninsulae has been
observed spawning on morning (0730 to 1130 hrs) slack low
tides.
c. How — Refer to section II. A. 4. c. for M. beryllina.
d. Spawning in the field.
1. Immediatedly after seining -- Refer to section II. A. 4. d.
1-6. for M. beryllina.
e. Transport of adults to the laboratory.
1. Adults - Refer to section II. A. 4. e. 1-4. for M. beryllina.
f. Laboratory spawning.
1. Spawning system -- Tidewater silversides, Menidia
peninsulae, may be spawned in the laboratory
throughout the year. The system described in Figure 3
and section II. A. 4. f. 1. should be utilized.
2. Feeding schedule - Refer to section II. A. 4. f. 2. for M.
beryllina.
3. Water quality - Refer to section II. A. 4. f. 3. for M.
beryllina.
21
-------
4. Water temperature and salinity — These variables
should be similar to that from the collection site. M.
peninsulae from the the Santa Rosa Island locale,
collected at 22°C and 23 %o were successfully maintained
and spawned in the laboratory at 24°C (22.1 to 25.4°C)
and 26 %o (23 to 28 %o) Middaugh et al., (1986).
5. Photoperiod and light intensity - Refer to section II. A.
4. f. 5. for M. beryllina.
6. Combinations of spawning signals - "Tidal" signals,
interruption of the 8 cm sec-1 current velocity in holding
tanks should occur at 1200 to 1300 and 2400 to 0100 hrs.
These interruptions in current velocity are accomplished
by using an electrical timer to turn off the circulation
pump for 1 hr at the specified times of 1200 to 1300 and
2400 to 0100 hrs. Menidia peninsulae apparently is
sensitive to interruptions in current velocity that mimic
conditions during slack low tides in nature (Middaugh et
al., 1986). They will spawn in the laboratory during
nightime (2400 to 0100 hrs) in response to the
interruption in current velocity, but not during daytime
from 1200 to 1300 hrs when the circulation pump is also
turned off (Middaugh and Hemmer, 1984; Middaugh et
al., 1986).
7. Spawning substrates - Refer to section II. A. 4. f. 7. for
M. beryllina. A synopsis of recommended environmental
variables for spawning M. peninsulae in the laboratory
is provided in Appendix B.
g. Culture of young Menidia peninsulae.
1. Fertilized eggs - Refer to section II. A. 4. g. 1. for M.
beryllina.
2. Embryos -- McMullen and Middaugh (1985) learned that
M. peninsulae embryos from Santa Rosa Island, Florida,
incubated at 20°C, showed poor hatchability and post-
hatch survival and growth. They concluded that optimal
conditions for embryo incubation and subsequent larval
growth and survival are 25°C and 30 %o.
3. Water temperature and salinity - Utilization of a single
temperature and salinity regime for spawning adult M.
peninsulae', and for embryo incubation, larval culture
and testing is advisable. This procedure minimizes
questions of acclimation and is often helpful because the
same source of water can be used in the various
production and testing systems. For M. peninsulae we
recommend a culture temperature of 25°C and salinity of
30 %o. This recommendation is based upon the work of
McMullen and Middaugh (1985) who found this
22
-------
combination to be optimal for larval survival and
growth, and upon the successful spawning of M.
peninsulae at a similar temperature and salinity
(Middaugh and Hemmer, 1984; Middaugh et al., 1986).
4. Transfer -- Refer to section II. A. 4. g. 4. for M. beryllina.
5. Feeding — Refer to section II. A. 4. g. 5. A synopsis of
recommended environmental variables for laboratory
incubation of embryos and larval culture is provided in
Appendix C.
D. California grunion, Leuresthes tenuis.
1. Geographical distribution - L. tenuis ranges from Monterey
Bay, California to Bahia Magdalena on the outer coast of Baja
California Sur (Moffatt and Thomson, 1975). The general
distribution of L. tenuis is shown in Figure 6 and a list of
selected biogeographical data are provided in Appendix F.
This list identifies locations where L. tenuis has been
collected.
2. Ecology and reproduction — L. tenuis is perhaps the best
known of the four atherinid fishes described in this manual
because of its unique reproductive behavior. Adult L. tenuis
reside in near-shore waters of southern California where
annual water temperatures range from 12 to 28°C (Moffatt
;>nd Thomson, 1975). They are surface-dwelling fishes
(Reynolds et al., 1977) that attain a maximum length of 150
to 180 mm (Walker, 1952). Spawning generally occurs from
late February through August. L. tenuis spawns in a sand
substrate at the approximate time of new and full moons
(Middaugh et al., 1983). Spawning runs take place at night
and are timed just after the highest high tides during each
semilunar period, subsequent high tides and wave action
result in depositon of sand over the incubating embryos
(Moffatt and Thomson, 1978; Middaugh et al., 1983).
Approximately 2 weeks after deposition, developed embryos
are washed out of the sand by the next series of high tides of
the same or greater height (Shepard and LaFond, 1940). The
buried embryos are protected from thermal stress and remain
relatively moist even though they usually are not inundated
for a week or more during incubation (Walker, 1949;
Middaugh etal., 1983).
3. Identification - The California grunion, Leuresthes tenuis,
grows to 143 mm SL or larger (Moffatt and Thomson, 1975;
Clark, 1925). It is greenish above and silver on the sides with
a dark lateral stripe. Scales along the mid lateral band are
highly and irregularly crenulate. Lateral scale range is 69-
80, x = 75 (Moffatt and Thomson, 1975) with 7-9 scales
between the first and second dorsal fins (Miller and Lea,
23
-------
California grunion, Leuresthes tenuis
Figure 6. Biogeographical distribution of the California grunion, Leuresthes tenuis.
-------
1972). The premaxillary extends over the mandible. The first
dorsal fin has 5-7 spines, second dorsal fin 1 spine and 9-10
rays, and anal fin 1 spine plus 21-24 rays.
4. Collection handling, spawning and transport of embryos.
a. Where - California grunion eggs are collected along
beaches of southern California during natural spawning
runs which occur from February through August, with a
peak in spawning activity during April and May (Walker,
1949).
b. When - Runs occur at night and are timed just after the
highest high tides during each semilunar cycle. Generally,
the 1 to 4 day period just after new and full moons is best
for collecting. Runs usually occur as the high tide is
receding (Walker, 1952).
c. How - California grunion are collected by hand as they are
stranded on the beach during natural runs. Use of nets is
forbidden by the California Department of Conservation. A
scientific collection permit is required and is available
from the Department of Conservation, Sacramento, CA.
d. Spawning in the field.
1. Refer to Figure 7 for a diagrammatic explanation of the
following procedure.
2. Eggs - Twenty to 25 females, captured just prior to
natural spawning, should be stripped into a 20 cm
diameter glass culture dish containing clean seawater
(depth 1 to 2 cm) from the spawning locale.
3. Milt - Five to 8 males are then stripped into a separate
dish containing clean seawater. Water containing sperm
is then poured into the dish containing eggs and mixed
by gently stirring. After 10 minutes, the fertilization
water is carefully decanted from the culture dish which
is then refilled with clean seawater. This refill-rinse
procedure should be repeated at least 2 additional times.
4. Fertilized embryos — Grunion embryos are carefully
pipetted, using a 1.5 cm I.D. x 45 cm glass tube equipped
with a rubber squeeze bulb, into all-glass or stainless
steel vacuum bottles. Several thousand embryos may be
transported in a vacuum bottle containing
approximately 500 ml of seawater from the collection
site. Water temperatures along the California coast will
usually range from 16 to 20°C during the height of the
reproductive season and salinities are generally ^32 %<>
(Middaugh et al., 1983). Note: It is absolutely essential
25
-------
9
COLLECTION
DURING
SPAWNING RUN
CLEAN
WATER7"-
ADULTS DIPPED
TO
REMOVE SAND
INTO 20cm dram.
GLASS CULTURE DISH
HAND STRP SPERM
INTO A GLASS CULTURE
DISH- THEN ADD TO EGGS
CLEAN -
WATER
EMBRYOS ARE
BURIED IN CLEAN
BEACH SAND
^^
• ••*• " •]
WATT 10-15 MM THEN
RINSE THREE TMES
TRANSPORT
TO LAB
- CLUTCHES OF
150-3OO EGGS
WOE MOUTHED
VACUUM BOTTLE
TRANSFER EMBRYOS
GLASS
LINED OR
STAINLESS STEEL
AFTER 12-14 DAYS AT 25'C
EMBRYOS ARE REMOVED
FROM SAND TRANSFER TO
CULTURE DISH OF
SEAWATER
LARVAE POURED
INTO 80S GLASS
AQUARIUM
PARTIALLY FILLED
WITH WATER
ACUTE TOXICITY
TESTS
WATER IS AGITATED
BY STIRRING
DAYS I-14 FEED
5000 ARTEMIA
NAUPLII /;
TWCE DAILY
Figure?. Diagrammatic explanation of procedure for
collecting, fertilizing, incubating and testing of
California grunion, Leuresthes tenuis, embryos.
26
-------
that sand not be pipetted into the vacuum bottles with
fertilized embryos.
5. Sealed vacuum bottles -- Embryos from the collection
site should be transported to the laboratory in vacuum
bottles. Borthwick et al. (1985) and Goodman et al.
(1985a) were able to successfully ship fertilized eggs
from southern California to Florida in vacuum bottles.
The transport time ranged from 24 to 36 hrs.
e. Embryo incubation in the laboratory.
1. Upon arrival at the laboratory -- Embryos are pipetted
into clean, dry beach sand (2.0 cm deep) in a 20 cm
diameter glass culture dish. Small depressions (1.0 cm
deep) are formed at 3 or 4 equally spaced locations
within the sand and 150 to 300 embryos are then
pipetted into each of these shallow depressions.
2. Embryos — Dry sand is used to cover the embryos so that
they are buried 0.5 to 1.0 cm deep. Thirty ml of sea water
(from the vacuum bottle) is then poured into the sand at
each location where embryos were buried. The culture
dish is then covered to prevent evaporation and placed in
an incubator at 25°C. This incubation temperature is
based upon field measurements taken at locations where
grunion embryos incubate naturally, 19 to 32°C, x —
25°C (Middaugh et al., 1983). Moreover, Hubbs (1965)
reported that hatching only occurred between 14.8 and
26.8°C. Similarly, Ehrlich and Karris (1971) observed
that grunion embryos hatched when maintained at 14.0
to 28.5°C. Optimal hatching, close to 100%, occurred
between 16 and 27°C. Hubbs (1965) pointed out that
embryos incubated at temperatures of 19°C or above
would be able to hatch on the next series of highest tides
in nature (approximately 10 to 14 days after they were
fertilized).
3. After 12 to 14 days — Embryos are carefully removed
from the sand substrate with a stainless steel spatula or
spoon.
4. To initiate hatching -- Approximately 500 to 1000
embryos and surrounding sand are placed in a 20 cm
glass culture dish containing = 1.5 1 of 25°C, 28 to 35 %o
salinity seawater. A spatula or a glass rod moved around
the inner edge of the culture dish will create a circular
current. This agitation of embryos and suspension of
sand grains induces hatching.
5. Hatched larvae - Should be transferred to 160 1 (or
larger) glass aquaria by pouring them from the 20 cm
diameter hatching dish, or by using a 1.5 cm I.D. x 45 cm
27
-------
glass pipette equipped with a rubber squeeze bulb. Care
should be used to minimize the amount of sand and
nonviable embryos transferred to the grow-out aquaria.
6. Density - The number of larval grunion in the grow-out
aquaria should not exceed 10 I-1. A water temperature of
25°C and salinities of 28 to 35 %o should be maintained
(Borth wick etal., 1985).
a. In a flow-through (dynamic) system - The holding
aquarium (160 1) should receive 20 1 hr-1 of
temperature and salinity adjusted seawater.
b. In a recirculating (static) system — The holding
aquarium (160 1) should be equipped with a Dyna-Flo
Model 425 Aquarium Filter (or equivalent). This
filtration system should be charged with spun
polyester aquarium filter-fiber and activated
charcoal. It is essential to cover the siphon intake
with 300-400 mm mesh plankton netting so that
grunion larvae are not impinged on the syphons or
pulled into the filtration unit. Units should be turned
off between 0700-1600 hrs (during the twice daily
feedings). Filter media should be changed at least
twice weekly and the syphon screens checked and
cleaned daily if necessary.
7. Feeding - On the day of hatching and until used in
toxicity test, grunion should be fed 5,000 Artemia
nauplii 1-1, twice daily (i.e., 0800 to 0900 and 1500 to
1600 hrs). For a 160 1 holding aquarium, 800,000
Artemia nauplii should be added at each feeding.
28
-------
III. Acute toxicity tests, M. beryllina, M. menidia,
M. peninsulae and L. tenuis
A. Static tests (96 hours).
1. Conducted - In 4 1 wide mouth glass jars containing 3 1 of
filtered (5 um) seawater maintained at 25 ± 1°C and 0 to 35 %o
salinity (depending upon the species used). It is convenient to
utilize a salinity similar to that in the grow-out aquaria.
Refer to Appendix G for recommended test conditions.
2. Age of fish -- Fourteen day-old atherinids are recommended
for testing. Because hatching of larvae is synchronous, all
individuals should be of identical age.
3. Transfer of larvae — Under no circumstances should larvae be
dip-netted. Mass mortalities will occur. Transfer from the
grow-out aquaria is accomplished by siphoning atherinids
(use a 3.0 cm I.D. x 1.5 m long plastic tube) into a 12 to 15 1
glass or plastic container. Individuals are then pipetted (1.5
cm x 45 cm glass pipette equipped with a rubber squeeze bulb)
or dipped into a 200 ml beaker. It is easy to enumerate 5
larvae in each beaker. Volume is then reduced to 10 ml in the
beakers and the contents are poured into test jars. A squeeze
bottle of uncontaminated test media should be available to
wash larvae from beakers if they are stranded.
4. General guidelines — Procedures provided by ASTM (1980)
should serve as a guide in conducting acute toxicity tests with
the atherinids, however, it is essential to feed all species with
Artemia nauplii during 96 hr tests. Live Artemia (20 to 30
nauplii per fish) are provided 2 to 3 times daily during
exposure. This amount of food should provide minimal
nutrition without confounding results due to dissolved
oxygen or loading problems (Borthwick et al., 1985).
5. The toxicant - If the chemical being tested results in rapid
mortality of Artemia nauplii, it may be necessary to remove
the dead nauplii from the bottom of test containers prior to
each subsequent feeding with live Artemia. It is also
important not to over-feed if fish will not eat available
Artemia.
6. Test containers - No aeration should be required. Dissolved
oxygen concentrations should be >60% of saturation. A
29
-------
photoperiod of 14L:10D with a light intensity of 11,000 lux is
recommended.
7. Temperatures and salinities utilized — These variables
should be similar to conditions in adult spawning and larval
rearing tanks. The values summarized in Appendix G are
recommended.
B. Flow-through tests (96 hours).
1. Conducted in 601 glass aquaria supplied with =<20 to 60 1 hr1
of 20 um filtered seawater.
a. Toxicant metering system - Should be selected on the basis
of the amount of toxicant to be metered, the flow rate of
dilution water and the series of toxicant concentrations
under consideration (ASTM, 1980; Clark et al., 1985).
b. Toxicant carrier concentrations -- These concentrations
should be constant for all exposures within a test. A
seawater control and carrier control should be maintained
along with the 4 to 7 exposure concentrations of toxicant.
c. To facilitate observations and enumerate mortalities —
Fish may be confined to 4 1 wide-mouth glass jars with two
screened (100 -140 mm mesh nylon screen) openings, 3 cm
diameter, 1 cm above the base.
1. Each 60 1 aquaria may contain 2 to 4 of the 4 1 wide
mouth glass jars with 10 fish jar -l.
2. A self-starting syphon (6 cm I.D. ) in the 60 1 aquarium
provides a 10-15 cm draw-down of water approximately
once every 15 min., thus ensuring adequate water
exchange between the 4 1 holding jars and larger 60 1
aquaria.
3. Other operational parameters are identical to those
provided for static tests. Refer to section III. A. 2-7.
30
-------
IV. Early Life-stage toxicity tests, silversides, Menidia
beryllina, M. menidia, and M. peninsulas.
A. Seawater — Dilution water for this 28 day test should be passed
through a sand filter and 20 um polypropylene filter. Water
temperature should be continuously maintained at 25 ± 1°C for
all three species. Salinity should be maintained at
approximately 4 to 6 %o for M. beryllina (or at the collection site-
laboratory spawning salinity if higher or lower) and 18 to 30 %o
for M. menidia and M. peninsulae (Goodman et al., 1985b).
B. Exposure system - A glass proportional diluter similar to that
described by Schimmel et al., (1974) is used in each test. The
diluter delivers 1 1 of water to each control and treatment at
each cycle (approximately every 15 min). Glass splitter boxes
equipped with two 2 mm I.D. glass capillary tubes slowly deliver
the seawater from the diluter to each of the two exposure
aquaria for each treatment. A 4 mm I.D. self-starting siphon
varies the water depth in each glass aquarium (21 cm long by 22
cm wide by 10 cm high) between approximately 4 and 7 cm. The
draw-down time is 1.5 to 3.0 min and helps to ensure adequate
exchange of toxicant between the exposure cups and aquarium.
Exposure aquaria are partially immersed in a freshwater bath
that maintains the respective tests at 25 ± 1°C. The photoperiod
during all testing is 14L:10D and light intensity is
approximately 1400 lux.
1. Embryos —The embryos of each species have numerous
chorionic fibrils that must be gently teased apart and their
fibrils clipped before selecting individuals to be used in the
toxicity test. Embryos are examined microscopically; groups
of four or five viable and synchronous individuals are then
randomly placed in each incubation cup until the cup
contains 16 embryos. Two incubation cups are then placed in
each of the two duplicate aquaria for each of the seven
treatments.
2. Incubation cups ~ These containers should be constructed of
glass petri dish bottoms (9 cm I.D.) with 10-cm-high cylinders
of 363 pm nylon mesh attached to the inside walls with
silicone adhesive. These cups are used in tests with all three
species. At test initiation, embryos should be approximately
32 to 36 hrs post-fertilization. If held at 25°C, these embryos
31
-------
should be in stages 18 to 20 of development (Lagler et al.,
1977).
3. Observations - Daily observations are made for deaths and
signs of poisoning. Dead animals are removed and recorded
when observed. Larvae are usually first observed on day 4 in
tests with M. menidia and M. peninsulae and on day 5 in the
M. beryllina test. Silverside larvae are fragile and susceptible
to injury or stranding on the mesh of incubation cups.
Therefore, during intervals after hatching begins, the
incubation cups should not be removed from the exposure
aquaria. Larvae are counted in place. Intervals are two days
for M. menidia, five days for M. peninsulae, and ten days for
M. beryllina. Before incubation cups containing hatched fish
are removed from the aquaria, the water level should be
lowered slowly to below the top of the petri dish using a 4 mm
(I.D.) glass siphon.
4. Feeding — Rotifers, Brachionus plicatilis', and Artemia sp.
nauplii (<8 hours old) are used as food for Menidia spp. Each
M. menidia incubation cup is provided a 1:1 mixture of
rotifers (2,500) and Artemia (2,500) twice daily (0800 to 0900
and 1400 to 1500 hrs) for the first eight days after hatching
begins, and thereafter, only Artemia (5,000) twice daily until
test termination. Menidia peninsulae and M. beryllina are fed
rotifers three times (0800 to 0900, 1200 to 1300 and 1400 to
1500 hrs) daily for the first 8 days after hatching begins at a
rate of 5,000 feeding-1 followed by a three-day transition
period in which both rotifers (2,500) and Artemia (2,500) are
provided in 3 feedings day1 and then Artemia (5,000) 3 times
day1 until test termination. In each test, equal portions of
food are dispensed to all cups containing hatched fish except
those with few survivors, which should received
approximately one half as much if considerable food remains
from previous feedings (Goodman et al., 1985b). Fish are not
fed on the last day of the test.
5. Test completion — At termination of the 28-day experiments,
fish are killed by immersion in ice water, drained on paper
towels, weighed individually, and may be combined within
exposure concentrations and frozen for subsequent chemical
residue analysis.
6. Statistical treatment — Analysis of variance and Duncan's
multiple range test may be used to analyze survival and
weight data. Individual fish weights are used in weight
analyses. Percentages should be arcsine transformed; a
minimum significance level of a — 0.05 is used for all
statistical purposes.
32
-------
V. Early life-stage toxicity tests, L. tenuis
A. Seawater — Dilution water for this 35-day test should be passed
through a sand filter and 20 um polypropylene filter. Water
temperature should be continuously recorded and maintained at
25 ± 1°C. Salinity should be maintained at 25 to 30 %o (Goodman
etal.,1985a)
B. Exposure system — A glass proportional diluter similar to that
described by Schimmel et al., (1974) delivers 1.0 1 of water to
each treatment at each cycle (approximately every 15 min).
Glass splitter boxes equipped with two 2 mm I.D. glass capillary
drains delivered the seawater from the diluter to the two glass
exposure aquaria per treatment. Exposure aquaria (I.D. = 21
cm long x 22 cm wide x 10 cm high) should be partially
immersed in a freshwater bath that will maintain test water
temperature at 25 ± 1°C. During embryo exposure, the aquaria
are equipped with a 5-mm I.D. self starting siphon that
fluctuates the water depth between 2 and 7 cm. The only carrier-
solvent used successfully in California grunion ELS tests is
triethylene glycol at a concentration of 0.4 pi I-1. Attempted ELS
tests at higher triethylene glycol concentrations failed because
embryonic development in treatments receiving the carrier-
solvent was slower than in the seawater-control treatment
(Goodman et al., 1985a).
1. Embryos — During the test, embryos should be immersed in
seawater and kept in darkness prior to testing. The test
should be started within 48 hrs after embryos are fertilized.
Embryos should be in developmental stages 22 to 24 (Lagler
et al., 1977) or younger. The embryos are examined
microscopically and groups of four viable specimens placed
randomly in each of 28 or more incubation cups. This process
is repeated until all cups contain 16 embryos. Treatment and
duplicate numbers are assigned by using a random number
table. Two incubation cups are then placed in each of the two
duplicate aquaria per treatment except that at least two
additional cups should be placed in the carrier-control
treatment for hatching trials.
2. Incubation cups - Embryos are maintained in containers
constructed by replacing the bottom of a 250-ml glass beaker
with 450-mm nylon mesh secured with silicone adhesive. Two
7.5 cm long glass tubes (8-mm I.D.) attached horizontally on
33
-------
opposite edges support the cup above the aquarium bottom.
Embryos are incubated in darkness until hatching is
stimulated. During this pre-hatch interval, a flashlight is
used when taking water samples or checking viability of
embryos. During the later stages of embryonic development
care must be taken to prevent jostling the embryos because
this could stimulate hatching.
3. On exposure day 9 -- (embryos -12 days old) Embryos from one
of the extra cups in the carrier-control treatment should be
gently rinsed into a larval incubation cup, constructed by
attaching 9-cm-high cylinders of 450 pm nylon mesh to the
inside walls of 100 x 15 mm glass Petri dish bottoms with
silicone adhesive. The embryos are placed on a laboratory
shaker and then alternately shaken (approximately 68 cycles
min-1) and kept stationary for 2-min intervals until no
additional fish hatch in two consecutive 2-min periods; two or
three intervals of shaking are required. If a large percentage
( = 75%) of the embryos hatch readily, then hatch those in all
treatments, but discard those from the extra carrier-control
cup. If a good hatch is not obtained from the first cup
attempted, wait another day and try the second extra carrier-
control cup. After removing cups with larvae from the shaker,
dead embryos are removed, and after replacing the siphons in
the aquaria with 2 mm I.D. siphons that vary the water depth
from 5 to 7 cm, the incubation cups are placed in their
respective aquaria. The photoperiod is changed from OL:24D
to!2L:12D.
4. Fish - In this test are fed <24-hr-old Artemia nauplii twice
daily, except for the last 24 hrs of the experiment when no
food is provided. Equal volumes of a suspension of nauplii are
dispensed to each incubation cup. The suspension should be
sampled frequently and the number of Artemia in a volume
equal to that dispensed per cup at each feeding should
average 5,000 (Goodman et al., 1985a). Dead animals are
removed and recorded daily, and qualitative observations of
signs of poisoning are noted.
5. At termination of the experiment — Fish are killed by
immersion in ice water, then drained on paper towels,
individually weighed, and may be frozen for subsequent
analysis. Survival and growth data may be analyzed by one
way analysis of variance. Percentage survival data should be
arcsine transformed before analyses. Treatment means may
be compared by using Duncan's multiple-range test. A
significance level of a= 0.05 is used for all analyses.
34
-------
VI. Batch culture of the alga, Chlorella sp. and
mixohaline rotifer, Brachionus plicatilis.
A. Apparatus used.
1. Six to eight, 13 to 201 pyrex glass carboys and stoppers to fit.
2. Four to six, 41 pyrex glass flasks with cotton plugs.
3. High wattage thermostatically controlled immersion heater,
Thelco-Thermajust Model 15094 1000W-110V or equivalent.
4. Laboratory timer, up to 120 mins elapsed time and able to
accomodate immersion heater of 1000W-110V capacity.
5. Source of clean compressed air.
6. Analytical balance-top loader.
7. Microscope and hemacytometer.
8. Automatic pipettor - 0.01 to 1.0 ml capacity, preferably with
disposable tips.
9. Plankton netting ~60 um mesh to collect rotifers.
10. Tygon or equivalent tubing 3 and 10 mm I.D.
11. Glass tubing 3 and 10 mm O.D.
B. Media preparation and Chlorella sp. culture.
1. Refer to Figure 8 for a diagrammatic explanation of the
following procedure.
2. Clean carboys — Wash thoroughly with hot soapy water, rinse
with tapwater, then triple rinse with deionized or distilled
water.
3. Fill carboy -- Filtered (20 um) natural seawater, salinity 25 to
30 %o should be used. If an artificial seawater mix is used, the
reconstituted media should be handled according to
manufacturer's instructions and allowed to age for at least 7
days prior to pasteurization. The carboy is filled to the top lip.
4. Immersion heater -- Should be placed in carboy with
thermostat set at 70 to 80°C. Heater power supply is always
routed through an automatic timer as a safety precaution.
35
-------
o
AERATION
n IMMERSION
HEATER
HEAT WATER TO 70-80°C
FOR I HOUR
12-14 HOURS
STOPPER TO
INSTALL STOPPER FIT CARBOY
ADO
TRACE METALS
PHOSPHATE + NITRATE
VITAMIN MIX
FERRIC CHLORIDE
VENT
PROVIDE MODERATE
AERATION
AFTER 5 TO 7 DAYS
CHLORELLA CELL DENSITY
2 TO 3X107/1 THEN
ADD BRACHIONUS
AFTER 5 T07 DAYS
BRACHIONUS DENSITY
,000/1
PROVIDE CONSTANT
ILLUMINATION, II.OOOLUX
AND MODERATE AERATION
PROVIDE GENTLE
AERATION
FEED TO
LARVAL MENIDIA
Figure 8. Diagrammatic explanation of procedure for culturing
the mixohaline rotifer, Brachionus plicatilis.
36
-------
5. Seawater -- Should be heated to a thermostatically set
temperature of ~70 to 80°C. This temperature should be
maintained for 1 hr. A glass tube (3 mm O.D.) inserted into
the carboy (before heating begins) and attached to an air
line provides moderate aeration/circulation of the water
during the pasteurization process. This aeration/circulation
is essential to ensure even heating of media and to prevent a
thermocline from forming in the carboy.
6. After maintaining ~70 to 80°C for 1 hr -- Turn off the
immersion heater, let it cool down for 5 mins, then remove it.
The automatic timer system can be set to the proper elapsed
time to accomplish heat-up, pasteurization and cool-down on
an automatic basis after heating dynamics of the carboy-
immersion heater system are determined on the first few
carboys of seawater. Note: If a large autoclave is available it
may be used to sterilize culture media thus avoiding the
procedures outlined above in B. 4-6.
7. Gentle aeration ~ Should be maintained for 12 to 14 hrs until
the seawater has cooled to ambient temperature. The carboy
should then be moved to a convenient location where high
intensity lighting (> 11,000 lux) is continuously provided.
8. The aeration tube-siphon tube-vent system - This is wrapped
in clean aluminum foil and maintained in a drying oven at ~50
to 60°C, it is then removed from the oven and installed in the
carboy of pasteurized seawater.
9. Appropriate enrichment media — To grow Chlorella,
nutrients are then added to the carboy and allowed to mix for
5 mins. Enrichment media will differ according to the type of
seawater used, natural or artificial, and the locale where
collected. One media enrichment formulation is summarized
in Appendix H. This formulation may need to be modified for
optimal growth of Chlorella sp. under local conditions.
Additional formulations are available in Walsh and
Alexander (1980), Theilacker and McMaster (1971) and the
American Public Health Association, Standard Methods
(1985).
10. Moderate aeration — Should be provided to mix nutrients
and to keep Chlorella sp. cells in suspension.
11. Carboy - It should then be inoculated with 4 to 5 x 105
Chlorella sp. cells.
12. Chlorella sp. cells - They should be allowed to grow for 5 to 7
days depending upon the initial density of starter cells
added to the 13 to 201 carboy.
a. 5 ml samples of Chlorella sp. should be removed and cell
density determined using the hemacytometer until the
37
-------
culture becomes a bright green and contains 2 to 3 x 107
cells I-1. We generally take replicate 5 ml samples from the
carboy and make 4 to 8 counts per sample to determine cell
density.
C. Culture of B. plicatllis.
1. Starter cultures -- Brachionus plicatllis cultures are
maintained in static 4 1 glass flasks containing 2 to 3 1 of
Chlorella sp. with an initial cells density of 2-3 x 107 cells I-1.
2. Cultures -- Should be renewed at 7 to 10 day intervals by
innoculation of new pasteurized (sterilized) and nutrient
enriched culture media with Chlorella sp. When Chlorella sp.
cell densities grow to 2 to 3 x 107 cells 1-1, Brachionus is
added.
a. To determine Brachionus densities — The 4 1 glass flask is
shaken to ensure an even distribution of Brachionus. The
density of Brachionus may be determined by pouring 1 1 of
the media from the flask through ~60 mm plankton netting.
This sample is then washed into a 100 ml beaker (using a
squeeze bottle containing pasteurized seawater) and
pasteurized water is then added to the beaker to bring the
volume to 20, 50 or 100 ml, depending upon the
concentration of Brachionus present.
b. Five, 10 pi subsamples are then removed from the 100 ml
beaker. Each sample is placed in a deep well depression
slide or in the center of a clear glass crystallization dish.
One drop of a 10% neutral buffered formalin solution is
then added to each 10 ul sample of Brachionus.
c. Counts of Brachionus - Should be conducted at 15 to 30x
magnification using a dissecting microscope, and the mean
number of Brachionus determined for the five samples.
d. Back calculate - The number of Brachionus 100 ml-1 (or to
50 or 20 ml of concentrate) is calculated from the original 1
1 sample.
1. For example in five samples the x number of Brachionus
= 80 per 10 pi x 100 = 8000 ml-l
8000 ml-l x 100 ml = 800,000 H
8000 ml-l x 50 ml = 400,000 H
8000 ml-l x 20 ml = 160,000 1-1
e. Use 10,000 to 20,000 Brachionus I-1 to innoculate each 13
to 20 1 carboy of Chlorella sp. when the alga cell density has
reached 2 to 3 x 107 cells H.
3. Brachionus densities -- The densities should reach 150,000 to
300,000 H by the 5th to 7th day after a carboy is innoculated.
38
-------
The procedure explained in VI. C. 2. a. through e. is used to
quantify Brachionus in carboys.
a. When densities are > 200,000 I-1 we recommend that the
entire volume of the carboy be used over a 4 to 5 day period.
1. Daily use of 2 to 3 1 of Brachionus is recommended.
2. Daily counts of Brachionus I-1 are required to maintain a
record so that feeding rates can be adjusted (by dilution
or concentration of 1 1 samples). Recall that a feeding
rate of 10,000 Brachionus I-1 volume of holding aquaria
for Menidia sp. is recommended.
Caution:
4. Brachionus is not capable of synthesizing essential fatty acids
(EFA's) for marine fish in the quantities required by those
fish (Lubzens et al., 1985). The rotifers must therefore be fed
on algae that contain high levels of EFA's for marine fish
(e.g., Skeletonema, Tetraselmis, Chaetoceros, Isochrysis, or
marine Chlorella, but not Dunaliella or Phaeodactylum).
39
-------
VII. Hatching of brine shrimp, Anemia sp.
A. Chemical analyses-The strain of Artemia used should be
analyzed for chemical residues including heavy metals and
pesticides.
B. Apparatus required.
1. Four to six, 2 1 separatory funnels.
2. Several 500 ml beakers.
C. Hatching procedure.
1. Fill -- A 2 1 separatory funnel with approximately 1,800 ml of
20 urn filtered seawater or reconstituted artificial seawater
that has been aged for at least 7 days. The salinity should be
adjusted to 25 to 30 %o.
2. Add — Artemia cysts, 15 to 20 ml (dry measure) to the
separatory funnel.
3. After adding Artemia cysts — Clean air is vigorously bubbled
through a 1-ml pipette which is lowered through the neck of
the funnel until the tip rests on the bottom. Aeration keeps
the cysts and newly hatched Artemia nauplii in suspension.
Cysts should hatch in 24 to 36 hrs when the temperature is
27°C.
4. After 24 hrs - The pipette supplying air is removed. Allow
Artemia nauplii to settle to the bottom of the separatory
funnel. A light source placed near the bottom of the
separatory funnel will enhance the settling process. Empty
cysts will rise to the surface.
5. After approximately five mins, using the stopcock, collect the
nauplii into a 500 ml beaker with a 100 mm mesh screen
bottom. Discard the hatching water and rinse the nauplii into
a 500 ml beaker.
6. After another five mins, again collect the nauplii and rinse
into the beaker.
7. The nauplii are further concentrated by pouring the
suspension into a small cylinder which has one end closed
with 100 mm plankton netting.
40
-------
8. The concentrate is resuspended in 50 ml of appropriate
culture water, mixed well, and dispensed with a pipette.
9. Discard the remaining contents of the hatching vessel, wash
the vessel with hot soapy water, and rinse thoroughly.
10. Prepare fresh seawater for each new batch culture of
Artemia nauplii.
11. To have a fresh supply of Artemia nauplii daily, several
hatching vessels must be set up and harvested on alternate
days.
D. Enumeration of Artemia nauplii.
1. Suspensions of nauplii should be well mixed.
a. draw off five 1.0 ml aliquots of the suspension.
b. the Artemia nauplii in each 1.0 aliquot may then be slowly
pipetted onto white filter paper so that the 1.0 ml volume is
dispersed over a wide area of the paper.
c. Artemia nauplii are then enumerated by placing the filter
paper on a dissecting microscope. The mean number of
nauplii in five samples is then used in back calculations to
determine the volume of original suspension to be fed to
Menidia sp. or Leuresthes tenuis. Recall that a feeding rate
of 5,000 Artemia nauplii I-1 volume of holding aquaria is
recommended.
E. Nutritional quality of Artemia.
1. To the extent that the atherinid species discussed here are
marine, they require a marine diet. Marine and freshwater
fish have different nutritional requirements, especially with
regard to essential fatty acids. Watanabe et al., (1978) pointed
out that Artemia can be divided in two categories, those that
are adequate for marine organisms and those that are not,
based on their fatty acid composition. Different geographical
strains of Artemia vary in their nutritional quality for
Menidia (Beck et al., 1980; Beck and Bengtson, 1982) and a
single geographical strain can also vary over time (Leger et
al., in press).
41
-------
VIII. REFERENCES
American Public Health Association, American Water Works
Association, Water Pollution Control Federation. 1985. Standard
Methods for the Examination of Water and Waste water, 16th
Edition, 1268 p. American Public Health Association, Washington,
DC, 20005.
American Society for Testing and Materials. 1980. Standard
practices for conducting acute toxicity tests with fishes,
macroinvertebrates, and amphibians. In: Annual Books of ASTM
Standards pp. 1-25. ASTM, Philadelphia, PA.
Anderson, W.D., J.K. Dias, R.K. Dias, D.M. Cupka and N.A.
Chamberlain. 1977. The macrofauna of the surf zone of Folly Beach,
South Carolina. NOAA (National Oceanic and Atmospheric
Administration) Technical Report NMFS (National Marine
Fisheries Service) SSRF (Special Scientific Report Fisheries) 704.
Barkman, R.C., and A.D. Beck. 1976. Incubating eggs of the
Atlantic silverside on nylon screen. Prog. Fish Culturist. 38:148-
150.
Bayliff, W.H. 1950. The life history of the silverside, Menidia
menidia (Linnaeus). Contr. Chesapeake Biol. Lab., Publ. 90:1-27.
Beck, A.D., D.A. Bengtson and W.H. Howell. 1980. International
Study on Artemia. V. Nutritional value of five geographical strains
of Artemia: effects on survival and growth of larval Atlantic
silverside, Menidia menidia. In: The Brine Shrimp Artemia pp. 249-
259. Vol. 3 (G. Persoone, P. Sorgeluos, 0. Roels and E. Jaspers, eds.)
Universa Press, Wetteren, Belgium.
Beck, A.D. and D.A. Bengtson. 1982. International Study on
Artemia XXII. Nutrition in aquatic toxicology: diet quality of
geographical strains of the brine shrimp Artemia. In: Aquatic
Toxicology and Hazard Evaluation: Fifth Conference, ASTM STP
766 pp. 161-169. (J.G. Pearson, R.B. Foster and W.E. Bishop, eds.),
American Society for Testing and Materials, Philadelphia.
Bengtson, D.A. 1982. Resource partitioning by Menidia menidia (L.)
and Menidia beryllina (Cope) in two Rhode Island estuaries. Ph.D.
Thesis, University of Rhode Island, Kingston, RI, 214pp.
42
-------
Bengtson, D.A. 1984. Resource partitioning by Menidia menidia and
Menidia beryllina (Osteichthys: Atherinidae) Mar. Ecol. Prog. Ser.
18:21-30.
Bengtson, D.A. 1985. Laboratory experiments on mechanisms of
competition and resource partitioning between Menidia menidia
(L.) and Menidia beryllina (Cope) (Osteichthyes: Atherinidae). J.
Exp. Mar. Biol. Ecol. 91:1-18.
Bengtson, D.A., R.C. Barkman and W.J. Berry. In preparation.
Relationship between maternal size, egg diameter, time of spawning
season, temperature, and length at hatch of Atlantic silversides,
Menidia menidia.
Bigelow, H.B. and W.C. Schroeder. 1953. Fishes of the Gulf of
Maine. U.S. Fish Wildl. Serv. Fish. Bull. 53:1-577.
Blair, W.F., A.P. Blair, P. Brodkorb, F.R. Cagle and G.A. Moore.
1968. Vertebrates of the United States. 2nd ed. McGraw-Hill, Inc.
N.Y.,616pp.
Borthwick, P.W., J.M. Patrick, Jr. and D.P. Middaugh. 1985.
Comparative acute sensitivities of early life-stages of Atherinid
fishes to Chlorpyrifos and Thiobencarb. Arch. Environ. Contam.
Toxicol. 14:465-473.
Briggs, P.T. 1975. Shore-zone fishes in the vicinity of Fire Island
inlet, Great South Bay, New York. N.Y. Fish and Game, 22:1-12.
Cadigan, K.M. and P.E. Fell. 1985. Reproduction, growth and
feeding habits of Menidia menidia (Atherinidae) in a tidal marsh-
estuarine system in southern New England. Copeia 1985:21-26.
Cain, R.L. and J.M. Dean. 1976. Annual occurrence, abundance, and
diversity of fish in a South Carolina intertidal creek. Mar. Biol.
36:369-379.
Chernoff, B., J.V. Conner and C.F. Bryan. 1981. Systematics of the
Menidia beryllina complex (Pisces: Atherinidae) from the Gulf of
Mexico and its tributaries. Copeia 1981:319-335.
Clark, F.N. 1925. The life history of Leuresthes tenuis, an atherinid
fish with tide controlled spawning habits. Calif. Fish Game Comm.
Bull. 10:1-51.
Clark, J.R., J.M. Patrick, Jr., D.P. Middaugh and J.C. Moore. 1985.
Relative sensitivity of six estuarine fishes to carbophenothion,
chlorpyrifos and fenvalerate. Ecotoxicol. and Environ. Safety.
10:382-390.
Clay, W.M. 1975. The fishes of Kentucky. Kentucky Dept. of Fish
and Wildl. Resources. Frankfurt 416 pp.
43
-------
Cook, S.F. and R.L. Moore. 1970. Mississippi silverside, Menidia
audens, (Atherinidae), established in California. Trans. Amer. Fish.
Soc. 99:70-73.
Conover, D.O. and B.E. Kynard. 1981. Environmental sex
determination: Interaction of temperature and genotype in a fish.
Science 213:577-579.
Conover, D.O. and S. Murawski. 1982. Offshore winter migration of
the Atlantic silverside, Menidia menidia. Fish. Bull. 80:145-149.
Conover, D.O. and B.E. Kynard. 1984. Field and laboratory
observations on spawning periodicity and behavior of a northern
population of the Atlantic silverside, Menidia menidia (Pisces:
Atherinidae). Environ. Biol. Fish. 11(3):161-171.
Cox, P. 1921. List of fishes collected in 1917 off Cape Breton coast
and the Magdalen Islands. Contrib. Canadian Biol., 1918-1920,
11:109-114.
Dahlberg, M.L. 1972. An ecological study of Georgia coastal fishes.
U.S. Fish. Wildl. Serv. Fish. Bull. 70:323-353.
DeSylva, D.P., F.A. Kalber, Jr., and C.N. Schuster. 1962. Fishes and
ecological conditions in the shore zone of the Delaware River
Estuary, with notes on other species collected in deeper water. Univ.
of Delaware Mar. Labs. Inform. Ser. Publ. 5:1-164.
Echelle, A.A. and D.T. Mosier. 1982. Menidia clarkhubbsi n. sp.
(Pisces: Atherinidae), and all female species. Copeia 1982:533-540.
Ehrlich, K.F. and D.A. Farris. 1971. Some influences of temperature
on the development of the grunion, Leuresthes tenuis (Ayrcs). Calif.
Fish and Game. 57(l):58-68.
Elston, R. and B. Bachen. 1976. Diel feeding cycle and some effects
of light on feeding intensity of the Mississippi silverside, Menidia
audens, in Clear Lake, California. Trans. Amer. Fish. Soc. 105:84-
88.
Fisher, F. 1973. Observations on the spawning of the Mississippi
silverside, Menidia audens Hay. Calif. Fish and Game. 59(4):315-
316.
Fowler, H.W. 1945. A study of the fishes of the southern piedmont
and coastal plain. Acad. Nat. Sci. Philad., Mono. 7:1-418.
Gomez, R. and H.L. Lindsay, Jr. 1972. Occurrence of the Mississippi
silverside, Menidia audens (Hay) in Keystone Reservoir and the
Arkansas River. Proc. Okla. Acad. Sci. 52:16-18.
Goodman, L.R., D.J. Hansen, G.M. Gripe, D.P. Middaugh, and J.C.
Moore. 1985a. A new early life-stage toxicity test using the
California grunion, Leuresthes tenuis, and results with chlorpyrifos.
Ecotoxicol. and Environ. Safety 10:12-21.
44
-------
Goodman, L.R., D.J. Hansen, D.P. Middaugh, G.M. Cripe and J.C.
Moore. 1985b. Method for early life-stage toxicity tests using three
atherinid fishes and results with chlorpyrifos. In: Aquatic
Toxicology and Hazard Assessment; Seventh Symposium, ASTM
STP854 (R.D. Cardwell, R. Purdy and R.C. Bahner, eds.) pp. 145-
154, ASTM, Philadelphia, PA.
Gosline, W.A. 1948. Speciation in fishes of the genus Menidia.
Evolution 2:306-313.
Gunter, G. 1945. Studies on marine fishes of Texas. Publ. Inst. Mar.
Sci. KD:1-90.
Gunter, G. 1950. Distributions and abundance of fishes on the
Aransas National Wildlife Refuge, with life history notes. Publ.
Inst. Mar. Sci. 1(2):89-101.
Hildebrand, S.F. 1922. Notes on habits and development of eggs and
larvae of the silversides, Menidia menidia and Menidia beryllina.
Bull. U.S. Bur. Fish. 38:113-120.
Hildebrand, S.F. and W.C. Schroeder. 1928. Fishes of Chesapeake
Bay. Bull. U.S. Bur. Fish. 43(1):366 pp.
Hubbs, C. 1965. Developmental temperature tolerance and rates of
four Southern California fishes, Fundulus parvipinnus, Atherinops
affinis, Leuresthes tenuis, and Hypsoblennius sp. Calif. Fish and
Game. 51(2):113-122.
Hubbs, C., H. Bryan and J.F. Schneider. 1971. Developmental rates
of Menidia audens with notes on salinity tolerance. Trans. Amer.
Fish. Soc. 100:603-610.
Hubbs, C. 1976. The diel reproductive pattern and fecundity of
Menidia audens. Copeia 1976:386-388.
Hubbs, C. 1982. Life history dynamics of Menidia beryllina from
LakeTexoma. Amer. Midi. Nat. 107:1-12.
Johnson, M.S. 1975. Biochemical systematics of the Atherinid genus
Menidia. Copeia 1975:662-691.
Kendall, W.C. 1902. Notes on the silversides of the genus Menidia of
the East Coast of the United States, with descriptions of two new
subspecies. Rept. U.S. Comm. Fish and Fisheries for 1901:241-267.
Kuntz, A. and L. Radcliffe. 1918. Notes on the embryology and
larval development of twelve teleostean fishes. Bull. U.S. Bur. Fish.
35(849), for 1915-1916,1918:87-134.
Lagler, K.F., J.E. Bardach, R.R. Miller and D.R. Passino. 1977.
Ichthyology, 2nd ed. John Wiley and Sons. New York, N.Y. 506 pp.
Leim, A.H. and W.B. Scott. 1966. Fishes of the Atlantic Coast of
Canada. Fish. Res. Bd. Canada, Bull. 155:1-485.
45
-------
Leger, P., D.A. Bengtson, K.L. Simpson and P. Sorgeloos. In press.
The use and nutritional value of Artemia as a food source. In:
Oceanography and Marine Biology Annual Review, Vol. 24 (M.
Barnes, ed.) Aberdeen University Press, Aberdeen, Scotland.
Lubzens, E., A. Marko, and A. Tietz. 1985. De novo synthesis of fatty
acids in the rotifer, Brachionusplicatilis. Aquaculture 47:27-37.
Lucas, J.R. 1982. Feeding ecology of the Gulf silverside, Menidia
peninsulas, near Crystal River, Florida, with notes on its life
history. Estuaries 5:138-144.
Massman, W.H. 1954. Marine fishes in fresh waters of Virginia
rivers. Ecology 35(l):75-78.
McMullen, D.M. and D.P. Middaugh. 1985. The effect of
temperature and food density on survival and growth of Menidia
peninsulae larvae (Pisces: Atherinidae) Estuaries, 8(l):39-47.
Mense, J.B. 1967. Ecology of the Mississippi silverside, Menidia
audens, in Lake Texoma. Bull. Okla. Fish. Res. Lab. 6:1-32
Middaugh, D.P. and P.W. Lempesis. 1976. Laboratory spawning and
rearing of a marine fish, the silverside, Menidia menidia menidia.
Mar. Biol. 35:295-300.
Middaugh, D.P. 1981. Reproductive ecology and spawning
periodicity of the Atlantic silverside, Menidia menidia (Pisces:
Atherinidae). Copeia 1981:766-776.
Middaugh, D.P., G.I. Scott and J.M. Dean. 1981. Reproductive
behavior of the Atlantic silverside, Menidia menidia (Pisces:
Atherinidae). Environ. Biol. Fish. 6(3/4):269-276.
Middaugh, D.P., H.W. Kohl II and L.E. Burnett. 1983. Concurrent
measurement of intertidal environmental variables and embryo
survival for the California grunion, Leuresthes tennis, and Atlantic
silverside, Menidia menidia (Pisces: Atherinidae). Calif. Fish and
Game. 69(2):89-96.
Middaugh, D.P. and T. Takita. 1983. Tidal and diurnal spawning
cues in the Atlantic silverside, Menidia menidia. Environ. Biol.
Fish. 8(2):97-104.
Middaugh, D.P. and M.J. Hemmer. 1984. Spawning of the tidewater
silverside, Menidia peninsulae (Goode and Bean) in response to tidal
and lighting schedules in the laboratory. Estuaries 7:137-146.
Middaugh, D.P., P.G. Hester, M.V. Meisch and P.M. Stark. 1985.
Preliminary data on use of the inland silverside, Menidia beryllina,
to control mosquito larvae. Jour. Amer. Mosquito Control Assoc.
1(4):435-441.
Middaugh, D.P., M.J. Hemmer and Y.L. Rose. 1986. Laboratory
spawning cues in Menidia beryllina and Menidia peninsulae (Pisces:
46
-------
Atherinidae) with notes on survival and growth of larvae at
different salinities. Environ. Biol. Fish. 15(2):107-117.
Middaugh, D.P. and M.J. Hemmer. 1986a. Reproductive ecology of
the tidewater silverside, Menidia peninsulae (Pisces: Atherinidae)
from Santa Rosa Island, Florida. Copeia, in press.
Middaugh, D.P. and M.J. Hemmer. In preparation. Evidence of the
influence of environmental temperature on sex-ratio in the
tidewater silverside, Menidia peninsulae.
Miller, D.J. and R.N. Lea. 1972. Guide to the coastal marine fishes
of California. Fish Bull. 157. Calif. Fish and Game, Sacramento, CA.
Moffatt, N.M. and D.A. Thomson. 1975. Taxonomic status of the
Gulf grunion (Leuresthes sardina) and its relationship to the
California grunion (L. tenuis). Trans. San Diego Soc. of Nat. History
18(4):75-84.
Moffatt, N.M. and D.A. Thomson. 1978. Tidal influence on the
evolution of egg size in the grunions. (Leuresthes: Atherinidae).
Environ. Biol. Fish. 3(3):267-273.
Moyle, P.B., F.W. Fisher, and H.W. Li. 1974. Mississippi silversides
and log perch in the Sacramento-San Joaquin River system. Calif.
Fish Game 60(3): 144-147.
Mulkana, M.S. 1966. The growth and feeding habits of juvenile
fishes in two Rhode Island estuaries. Gulf Res. Rpts. 2(2):97-167.
Pearcy, W.G. and S.W. Richards. 1962. Distribution and ecology of
fishes of the Mystic River Estuary, Connecticut. Ecology 43(2): 249-
259.
Raney, E.G. 1950. Freshwater fishes. In: The James River basin,
past, present, future pp. 151-194. Va. Acad. Sci., Richmond.
Reynolds, W.W., D.A. Thomson and M.E. Casterlin. 1977. Responses
of young California grunion, Leuresthes tenuis, to gradients of
temperature and light. Copeia 1977:144-149.
Robbins, T.W. 1969. A systematic study of the silversides Membras
Bonaparte and Menidia (LinnaeusXAtherinidae, Teleostei).
Unpubl. Ph.D. dissertation, Cornell University, Ithica, NY. 282 pp.
Rubinoff, I. and E. Shaw. 1960. Hybridization in two sympatric
species of atherinid fishes, Menidia menidia (Linnaeus) and
Menidia berylllna(Cope). Amer. Mus. Nov. 1999:1-13.
Schimmel, S.C., D.J. Hansen and J. Forester. 1974. Effects of
Aroclor 1254 on laboratory-reared embryos and fry of sheepshead
minnows (Cyprinodon varlegatus). Trans. Am. Fish. Soc. 103:582-
586.
47
-------
Shenker, J. and J.M. Dean. 1979. The utilization of an intertidal
saltmarsh creek by larval and juvenile fishes: abundances, diversity
and temporal variation. Estuaries 2:154-163.
Shepard, P.P. and E.C. LaFond. 1940. Sand movements along the
Scripps Institution Pier. Am. Jour. Sci. 238:272-285.
Simmons, E.G. 1957. Ecological survey of the Upper Laguna Madre
of Texas. Publ. Inst. Mar. Sci. 4(2):156-200.
Sisk, M.E. and R.E. Stephens. 1964. Menidia audens (Pisces:
Atherinidae) in Boomer Lake, Oklahoma, and its possible spread in
the Arkansas River system. Proc. Okla. Acad. of Sci. 44:71-73.
Sisk, M.E. 1973. Six addditions to the known piscine fauna of
Kentucky. Trans. Ky. Acad. Sci. 30(3-4):54-59.
Smith, H.M. 1893. Report on a collection of fishes from the
Albermarle Region. Bull. U.S. Fish Comm. 11(1891):185-200.
Smith, P.W. 1979. The Fishes of Illinois. University of Illinois Press.
Urbana. 314pp.
Springer, V.G. and K.D. Woodburn. 1960. An ecological study of the
fishes of the Tampa Bay area. Fla. State Bd. of Conservation.
Professional Paper Series. Marine Laboratory, St. Petersburg. 104
PP-
Tagatz, M.E. and D.L. Dudley. 1961. Seasonal occurrence of marine
fishes in four shore habitats near Beaufort, NC, 1957-1960. U.S.
Fish, and Wildl. Serv. Spec. Sci. Rpt. Fish. 390:1-19.
Theilacker, G.H. and M.F. McMaster. 1971. Mass culture of the
rotifer Brachionus plicatilis and its evaluation as a food for larval
anchovies. Mar. Biol. 10:183-188.
Tilton, J.D. and R.L. White. 1964. Menidia beryllina from several
central Texas impoundments. Texas Jour. Sci. 16:120.
Walker, B.W. 1949. Periodicity of spawning by the grunion
Leuresthes tennis, an atherine fish. Ph.D. Thesis. Univ. of Calif. Los
Angeles. 166pp.
Walker, B.W. 1952. A guide to the grunion. Calif. Fish and Game.
38:409-421.
Walsh, G.E. and S.V. Alexander. 1980. A marine algal bioassay
method: results with pesticides and industrial wastes. Water, Air
and Soil Pollut. 13:45-55.
Watanabe, T., F. Oowa, C. Kitajima and S. Fujita. 1978. Nutritional
quality of brine shrimp, Artemia salina, as a living feed from the
viewpoint of essential fatty acids for fish. Bull. Jap. Soc. Sci. Fish.
44:1115-1121.
48
-------
Appendix A
Selected biogeographical data for occurrence of the
inland silverside, Menidia beryllina
State
Locale
Habitat
Rhode Island
Rhode Island
Massachusetts
Massachusetts
Connecticut
Connecticut
Delaware
Virginia
Virginia
Virginia
North Carolina
North Carolina
North Carolina
North Carolina
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Alabama
Mississippi
Mississippi
Louisiana
Louisiana
Louisiana
Louisiana
Louisiana
Louisiana
Arkansas
Tennessee
Kentucky
Illinois
Texas
Texas
Texas
Texas
Texas
Texas
Texas
Oklahoma
Oklahoma
Oklahoma
Oklahoma
California
California
California
Pattaquamscutt R.
Point Judith Pond
Massachusetts Bay
Quincy Bay
Mystic R. (lower)
Mill R.
Delaware R. (lower)
Potomoc R. (lower)
James R.
Rappahannock R.
(45 mi upstream)
Pasquotank R.
Edenton Bay
Perquiman's R.
Neuse R.
Palatka
L. Eustis
L. Weir
St. Johns R.
South Lake
L. Monroe
L. Jessup
Escambia Bay
Blackwater Bay
Perdido Bay
Gulf Shores
Escatawpa R.
Moon Lake
L. Pontchartrain
L. Angola
Wax Lake
Mississippi Delta
Chandeleur Islands
L. St. John
L. Chicot
Reelfoot Lake
Hamby Pond
Mississippi R.
L. Marble Falls
L. Buchanan
L. Inks
L. Brownwood
L. Texoma
Colorado R.
Laguna Madre
L. Texoma
Keystone Reservoir
Arkansas R.
Boomer Lake
Clear Lake
Sacramento River
Lexington Reservoir
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Estuarine
Riverine
Estuarine
Estuarine
Estuarine
Estuarine
Lake
Lake
Lake
Riverine
Lake
Lake
Lake
Estuarine
Estuarine
Estuarine
Estuarine
Riverine
Lake
Estuarine
Lake
Lake
Riverine/
Estuarine
Estuarine
Lake
Lake
Lake
Pond
Riverine
Lake
Lake
Lake
Lake
Lake
Riverine
Estuarine
Lake
Lake
Riverine
Lake
Lake
Riverine
Lake
Citation
Bengtson, 1984
Bengtson, 1984
Kendall, 1902
Robbing, 1969
Pearcy and Richards, 1962
Johnson, 1975
DeSylvaetal.,1962
Kendall, 1902
Raney,1950
Massman, 1954
Smith, 1893
Smith, 1893
Johnson, 1975
Tagatz and Dudley, 1961
Johnson, 1975
Chernoffetal., 1981
Chernoffetal.,1981
Chernoffetal., 1981
Kendall, 1902
Kendall, 1902
Kendall, 1902
Chernoffetal.,1981
Middaughetal., 1986
Chernoffetal.,1981
Johnson, 1975
Chernoffetal.,1981
Chernoffetal.,1981
Chernoffetal.,1981
Chernoffetal.,1981
Chernoffetal.,1981
Chernoffetal.,1981
Chernoffetal.,1981
Johnson, 1975
Chernoffetal.,1981
Johnson, 1975
Sisk.1973
Smith, 1979
Tilton and White, 1964
Tilton and White, 1964
Tilton and White, 1964
Tilton and White, 1964
Mense, 1967
Tilton and White, 1964
Simmons, 1957
Hubbs, 1982
Gomez and Lindsay, 1972
Gomez and Lindsay, 1972
Sisk and Stephens, 1964
Cook and Moore, 1970
Moyleetal., 1974
Moyleetal., 1974
49
-------
Appendix B
Recommended environmental variables and
feeding regimes for laboratory spawning of three
atherinid fishes
Environmental
Variable
Adult holding/spawning
Photoperiod 13L:11D
Intensity (lux)
175-300
Tidal signals/
interrupted
current velocity
Salinity (%«)
Water Temp. (°C)
none required
0-5
25
Food required
Tetramin Flakes
Artemia sp.
nauplii (optional)
ca. 150,000 I1
8 gat0800-0900 hr
4gat 1000-1100 hr
4 g at 1300-1400 hr
8 g at 1500-1700 hr
1 liter all 130 hr
Atherinid species
M. menidia
13L:11D
175-300
1200-1300 hrs
2400-0100 hrs
25-30
22 (N. lat.)
25(S.lat.)
8gat0800-0900 hr
4gat 1000-1100 hr
4 g at 1300-1400 hr
8 g at 1500-1700 hr
1 liter all 130 hr
M. peninsulae
13L:11D
175-300
1200-1300 hrs
2400-0100 hrs
25-30
25
8 gat 0800-0900 hr
4gatlOOO-1100hr
4g at 1300-1400 hr
8gatl500-1700hr
1 liter all 130 hr
50
-------
Appendix C
Recommended environmental variables and
feeding regimes for laboratory incubation and
larval rearing of four atherinid fishes
Environmental
Variable
Atherinid species
M. menidia M. pemnsulae L. tenuis
Embryo-larval rearing
Photoperiod 14L:10D
Intensity (lux) 11,000
Salinity (%.) 0-15
Water Temp. (°C) 25
Larval density I'1
(grow-out tanks)
Food required
Brachionus sp.
amount 1 '
vol. of
grow-out tanks
5-10
days 1-5
10,000 twice
a day
14L:10D
11,000
25-30
22 (N. lat.)
25(S.lat.)
5-10
days 1-5
10,000 twice
a day
14L:10D
11,000
25-30
25
5-10
days 1-5
10,000 twice
a day
14L:10D
11,000
2530
25
5-10
not required
not required
Artemia sp.
amount H
vol. of
grow-out tanks
day 6 through
end of holding
5,000 twice
a day
day 6 through day 6 through day 1 through
end of holding end of holding end of holding
5,000 twice
a day
5,000 twice
a day
5,000 twice
a day
51
-------
Appendix D
Selected biogepgraphical data for occurrence of the
Atlantic silverside, Menidia menidia
State
Maine
Maine
Maine
Maine
Rhode Island
Rhode Island
Rhode Island
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Massachusetts
Connecticut
Connecticut
New York
Delaware
Virginia
Virginia
Virginia
Virginia
Maryland
Maryland
Maryland
Maryland
North Carolina
North Carolina
North Carolina
South Carolina
South Carolina
South Carolina
South Carolina
South Carolina
South Carolina
Georgia
Georgia
Georgia
Georgia
Georgia
Florida
Florida
Florida
Florida
Florida
Locale
S. Portland
Scarborough Harbor
Todd Point
Casco Bay
Point Judith Pond
Pettaquamscutt R.
Bissell Cove
Massachusetts Bay
Vineyard Sound
Buzzard Bay
Nantucket Sound
Eel Pond
Great Harbor
Hadley Harbor
Katama Bay
Woods Hole
Salem Harbor
Noank
Pataguanset R.
Fire Island Inlet
Delaware R. (lower)
James R.
Rappahannock R.
Pamunkey R.
Oyster
Solomons Island
Drew Point
Green Holly Creek
Molly's Island
Moorehead City
River's Island
Neuse River (lower)
Cape Romain
Magnolia Beach
Cape Island
North Edisto R.
Edisto R. (lower)
Battery Island
Sea Island
Ogeechee R.
Sapelo Island
St. Simons Island
Jekyl Island
Ft. George Inlet
Malangos R.
Flagler Beach
New Smyrna Beach
Mosquito Lagoon
Habitat Citation
Estuarine Johnson, 1975
Estuarine Robbins, 1969
Estuarine Robbins, 1969
Estuarine Kendall, 1902
Estuarine Bengtson, 1984
Estuarine Bengtson, 1982
Estuarine Bengtson, 1982
Estuarine Bigelow and Schroeder,
1953
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Kendall, 1902
Estuarine Conover and Kynard, 1984
Estuarine Johnson, 1975
Estuarine Cadigan and Fell, 1985
Estuarine Briggs, 1975
Estuarine DeSylva et al., 1962
Estuarine Massman, 1954
Estuarine Massman, 1954
Estuarine Massman, 1954
Estuarine Johnson, 1975
Estuarine Bayliff, 1950
Estuarine Bayliff, 1950
Estuarine Bayliff, 1950
Estuarine Bayliff, 1950
Estuarine Kendall, 1902
Estuarine Hildebrand, 1922
Estuarine Tagatz and Dudley, 1961
Estuarine Fowler, 1945
Estuarine Fowler, 1945
Estuarine Fowler, 1945
Estuarine Middaugh, 1981
Estuarine Fowler, 1945
Estuarine Johnson, 1975
Estuarine Fowler, 1945
Estuarine Fowler, 1945
Estuarine Fowler, 1945
Estuarine Robbins, 1969
Estuarine Robbins, 1969
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Gosline, 1948
52
-------
Appendix E
Selected biogeographical data for occurrence of the
tidewater silverside, Menidia peninsulae
State
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Florida
Mississippi
Mississippi
Texas
Texas
Texas
Texas
Locale
Melbourne
Ft. Meyers
Shell Point
Southport
Pass-A-Grille
Mosquito Lagoon
Crystal R.
Cedar Key
Santa Rosa Island
Escambia Bay
Pascagoula
Horn Island
Galveston Bay
Aransas Pass
Leffy Ann
Capano Bay
Habitat Citation
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Johnson, 1975
Estuarine Chernoffet al., 1981
Estuarine Chernoffet al., 1981
Estuarine Chernoffet al., 1981
Estuarine ChernofTet al., 1981
Estuarine Middaugh and Hemmer,
1984
Estuarine Johnson, 1975
Estuarine Chernoff etal., 1981
Estuarine Chernoff etal., 1981
Estuarine Johnson, 1975
Estuarine Chernoff etal., 1981
Estuarine Chernoffet al., 1981
Estuarine Johnson, 1975
53
-------
Appendix F
Selected biogeographical data for occurrence of the
California grunion, Leuresthes tennis
State
California
California
California
California
California
California
California
California
California
California
California
California
California
California
California
California
California
California
California
Locale
Morrow Bay to
Cayucos
Ismo Beach
Santa Barbara
Malibu
Santa Monica
Venice
Hermosa Beach
Cabrillo Beach
Long Beach
Belmont
Huntington Beach
Newport Beach
Corona del Mar
Doheny Beach
Del Mar
Black's Beach
La Jolla
Mission Beach
Coronado Strand
Habitat
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Beach
Citation
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
Walker, 1952
54
-------
Appendix G
Recommended test parameters and feeding regimes
for conducting static or flow-through 96 hr acute
toxicity tests with 14 day-old atherinid fishes
Environmental
Variable M. beryllina
Photoperiod
Intensity (lux)
Salinity (%,)
Water Temp. (°C)
UL:10D
11,000
0-15
25
Atherinid species
M. menidia M. pemnsulae
14L:10D
11,000
25-30
22 (N. lat.)
25 (S. lat.)
14L:10D
11,000
25-30
25
L. tenuis
14L:10D
11,000
25-30
25
Feeding Requirements
Artemiasp
nauplii
20-30 fish -i
2 to 3 times
daily
20-30 fish -i
2 to 3 times
daily
20-30 fish i
2 to 3 times
daily
20-30 fish i
2 to 3 times
daily
55
-------
Appendix H
Enrichment media for seawater used to grow
Chlorella sp.
Nutrient Mixes Amount
TRACE METALS"
MnCl2-4H2O 361 mg
ZnCl2 42 mg
CuSO4-5H2O 8 mg
Na2MoO4-2H2O 5 mg
CoCl2-6H2O 8 mg
Glass distilled or deionized water 1.0 liter
IRON>>
FeCl-6H2O 480 mg
Glass distilled or deionized water 100 ml
NITRATES AND PHOSPHATES "
NaNO3 75 mg
NaH2PO4 6 mg
Glass distilled or deionized water 1.0 liter
Vitamin mix d
Thiamine hydrochloride 0.4 g
Biotin 1.0 mg
B12 1.0 mg
Glass distilled or deionized water 1.0 liter
a add 1.0 mil' of seawater media
b add 0.1 ml 1 -1 of seawater media
c add 1.0 ml 1 ' of seawater media
d add 1.0 ml I ' of seawater media
56
------- |