NOAA Technical Report NMFS 106

Marine Ranching
Proceedings of the Eighteenth
U. S. -japan Meeting on Aquaculture
Port Ludlow, Washington
18-19 September 1989
Ralph S. SVIjcek (editor)

U.S. Department of Commerce

February 1992

NOAA Technical Report NMFS
The major responsibilities of the National Marine Fisheries SeIVice (NMFS) are to monitor and assess the abundance and geographic distribution of fishery resources, tounderstand and predict fluctuations in the quantity anddistribution of these resources, and to establish levels fortheir optimum use. NMFS is also charged with the development and implementation of policies for managing national fishing grounds, with the development andenforcement of domestic fisheries regulations, with the surveillance of foreign fishing off U.S. coastal waters, andwith the development and enforcement of international
fishery agreements and policies. NMFS also assists thefishing industry through marketing seIVice and economic
analysis programs and through mortage insurance andvessel construction subsidies. It collects, analyzes, andpublishes statistics on various phases of the industry.
The NOAA Technical Report NMFS series was established in 1983 to replace two subcategories of the T echnical Report series: "Special Scientific Report -Fisheries"and "Circular." The series contains the following types ofreports: scientific investigations that document long-term

continuing programs of NMFS; intensive scientific reports
on studies of restricted scope; papers on applied fisheryproblems; technical reports of general interest intended toaid conservation and management; reports that review, in'considerable detail and at a high technical level, certain
broad areas of research; and technical papers originatingin economics studies and in management investigations.
Since this is a formal series, all submitted papers, except
those of the U.Sdapan series on aquaculture, receive peer
review and all papers, once accepted, receive professional
editing before publication.
Copies of NOAA Technical Reports NMFS are available free in limited numbers to government agencies, bothfederal and state. They are also available in exchange forother scientific and technical publications in the marine
sciences. Individual copies may be obtained for the U.S.
Department of Commerce, National Technical Information SeIVice, 5285 Port Royal Road, Springfield, VA
22161. Although the contents of these reports have notbeen copyrighted and may be reprinted entirely, reference
to source is appreciated.

Recently Published NOAA Techniclll Reports NMFS
90.

Elaunobraac:Ja... IiviDg resources: advance. in
the biology, ecology, II)'IIteJDatica, and the .tatua of
the &sheries, edited by Harold L. Pratt Jr., Samuel
H. Gruber, and Toru Taniuchi. July 1990, 518 p.

96.

Marble flora and fa1lJlA of the Eastern UnitedStates--Copepoda, Cyc1opoida: ArchiDotodelphyidae, Notodelphyidae, and Ascidicolidae, by Patricia
L. Dudley and Paul L. I11g. January 1991,40 p.

91.

Marble flora and fa1lJlA of the aortbeaatern United
Stat_Ec:hJnode......ta: Crinoidea, by Charles G.
Messing and John H. Dearborn. August 1990,30 p.

97.

Catalog of osteological collectio_ of aquatic DlaDlDla1a frODl Meldco, by Omar Vidal. January 1991,
36 p.

92.

GeDetica ill aquaculture: proceediDp of the aisteeDth U.S.-JapaD DleedDg OD aquacultureJ
CharlestoD, South CaroliDa, 20-21 October, 1987,
edited by Ralph S. Svrjcek. November 1990, 81 p.

98.

Marble mamma) .trandiDp ia the United StatesJ
proceediDg. of the .econd Dlarine DlaDlDlal.trandiag worbhopJ MiaDli, Florida, 3-5 DeceDlber, 1987, edited by John E. Reynolds III and Daniel
K. Odell. January 1991, 157 p.

93.

DistributiOD aad abuadaace ofjuveaile .aiDlonids
oB' CregoD and W. .hiDgtoD, 1981-1985, by William
G. Pearcy and Joseph P. Fisher. November 1990,83 p.

99.

Marille flora and fauDa of the Northeastern
United State.: Erect Bryozoa, by John S. Ryland
and Peter J. Hayward. February 1991, 48 p.

Au ecoDODlic:a guide to allocatioD of &sh .toc:b betweeD cODlDlercial aad recreatioaal &sherie., by
Steven F. Edwards. November 1990, 29 p.

100. Marine Sora aDd fauna of the Eastern United States:
DicyeDlida, by Robert B. Short. February 1991, 16 p.

94.

95.

Larval fi.h recnUweDt and re.earch iD the
Alneric..: proceediDp of the thirteeDth aDDuai
1arvaI &sh coafereaceJ Merida, Mesico, 21-26 May
1989, edited by Robert D. Hoyt. January 1991, 147 p.

101. Larvae of Dearshore fishes in oceanic waten Dear
Oahu, Hawaii, by Thomas A. Clarke. March 1991,
19 p.

NOAA Technical Report NMFS 106

Marine Ranching
Proceedings of the Eighteenth
u.s.Japan Meeting on Aquaculture
Port Ludlow, Washington
18-19 September 1989
Ralph S. Svrjcek
Publications Unit
Northwest and Alaska Fisheries Science Centers

Panel Chairmen:
Conrad Mahnken, United States
Hisashi Kan-no,japan

Under the Us.Japan Cooperative Prog;ramin Natural Resources (UJNR)

February 1992

U.S. DEPARTMENT OF COMMERCE
Robert Mosbacher, Secretary

National Oceanic and Atmospheric Administration
John A. Knauss, Under Secretary for Oceans and Atmosphere

National Marine Fisheries Service
William W. FoxJr., Assistant Administrator for Fisheries

Contents (continued)
Experimentation to improve recruitment of blood ark shell,
Scapharra brOUghtOitii, in the Seto Inland Sea

105

H.NAKAHARAThe Marine Ranching System: the integration of biology
and engineering technology

115

K.. TAYAA.HASEGAWA
Y. MASUI

Economic problems of the ~arine Ranching System

129

S. UMEZAWA

iv

Reproductive Endocrinology of the Shrimp
Sicyonia ingentis:
Steroid, Peptide, and Terpenoid Hormones
ERNEST S. CHANG, WILLIAM A. HERTZ, AND GLENN D. PRESTWICH*
Bodega Marine Laboratory,
University of California,
P.O. Box 247, Bodega Bay, CA 94923

ABSTRACT
Female reproduction in penaeid shrimp is carefully regulated by several different endocrine factors. Their precise modes of action have not yet been fully elucidated. Three
endocrine factors, each representing a different chemical class of hormones, have been
investigated in the penaeid shrimp Sicyonia ingentis in our laboratory: ecdysteroids, vitellogenesis-inhibiting hormone (VIH) , and methyl farnesoate (MF). Ecdysteroids (thesteroid molting hormones of arthropods; predominantly 20-hydroxyecdysone), are initially present in low levels (<10 ng/mg) in shrimp embryos. As development of theembryos nears time of hatch, the ecdysteroid levels increase to approximately 150 ng/
mg, indicating that they may be of embryonic origin and involved in embryonic development. An assay was developed for shrimp VIH, which presumably is a protein. Delay of
onset of the next reproductive cycle was observed following injection of sinus glandextracts into shrimp that had previously had their eyestalks removed. A photoaffinity
analog was synthesized for the putative shrimp reproductive hormone MF-a terpenoid.
This analog, farnesyl diazomethyl ketone (FDK) , was used to demonstrate the presenceof specific binding proteins for MF in shrimp hemolymph.

Introduction_

The control of female reproduction in penaeid(members of the superfamily Penaeoidea) shrimp ishighly complex. It appears that a number of environmental signals can influence different hormonal
factors which in turn regulate various aspects of the
reproductive process. The understanding of thisregulation is an area of intense research (see reviews
by Adiyodi 1985; Charniaux-Cotton and Payen 1988).
In addition to the basic biological studies in comparative endocrinology, there is great interest in theapplied aspects of this research.
Penaeid shrimp comprise one of the most economically important marine products both
domestically and worldwide (Rosenberry 1990).
While natural fisheries for shrimp have declined,
there has been a concomitant surge in aquatic culture of penaeids. One of the major problems
preventing optimization of the commercial culture of
shrimp is control of female reproduction.

We examined three areas of shrimp reproduction.
First is the role of the arthropod molting hormones,
ecdysteroids, in ovarian and embryonic development.
Although there have been a number of studies on therole of ecdysteroids in crustacean molting (Chang1989), relatively little is known about the action ofthese steroid molting hormones on ovarian and embryonic development.
Second, we investigated the activity of the vitellogenesis-inhibiting hormone (VIH). The initialobservations that provided a basis for the existence of
the VIH were made by Panouse (1943, 1944). He observed that, depending upon the molt stage, removalof the eyestalks from the shrimp Palaemon serratus
resul ted in accelerated ovarian developmen t andspawning. Similar observations have been made in a
number of natantians (see Chang 1992).

* Permanent address: Department of Chemistry, State University of
New York, Stony Brook, NY 11794-3400.

4

NOAA Technical Report NMFS 106

buffer, mixed, and boiled 4 minutes. Samples (45j..lL) were loaded onto a 0.75 X 150 X 150 mm denaturing polyacrylamide gel (SDS-PAGE, 12%acrylamide), stacked at 13 rnA per gel, and separated
at 10° C at 18 rnA per gel using a Tris-glycine pH 8.3
running buffer. Gels were stained (4 h) with
Coomassie Blue R250 and then destained (95%ethanol:H 2 0:acetic acid, 50:40: 10).
The destained gel was rinsed for 5 minutes withglacial acetic acid and then impregnated with
diphenyloxazole (15% PPO in glacial acetic acid) for
25 minutes. The acetic acid was poured off and thegel was treated with 50% PEG 2000 at 50-75° C for 30
to 40 minutes. The miniaturized gels (Mohamed etal. 1989) were dried and exposed to pre-flashed
Kodak XAR-5 x-ray film for 5-15 days at -75° C.

Results and Discussion_

Embryonic Ecdysteroids
The concentration profile of ecdysteroids in the hemolymph of Sicyonia ingentis is similar to tl'ose of
most other crustaceans that have been examined(Fig. 2; see Chang 1989). There are low levels during
postmolt and intermolt with a dramatic increase justprior to ecdysis (premolt). This peak falls back tobasal levels just prior to ecdysis. Approximately similar values were obtained in the shrimp Palaemon
serratus (Baldaia et al. 1984; Van Wormhoudt et al.
1986) .
Since ecdysteroids play such an important role inlarval, juvenile, and adult molting (Chang 1989), we
assayed extracts of developing embryos to determine
if these steroid hormones were also involved in embryonic development. Figure 3 shows the total RIA
activity of extracts of embryos at various times after
spawning (fertilization). Negligible levels of
ecdysteroids were present in the embryos at spawning, but the concentrations increased significantly asembryonic development proceeded. These data im-ply that there is relatively little maternal investmentof ecdysteroids in the eggs and that the increasinglevels of hormone observed in the extracts is likely
due to endogenous synthesis.
These data are in apparent contrast to observations
that our laboratory has made in the crab Cancer
anthonyi (Okazaki and Chang 1991). These crab embryos have relatively high levels of ecdysteroids at thetime of spawn (ca. 9 ng/mg wet weight) that decreaseduring embryonic development. This implies an embryonic utilization of the hormones during
development. These differences may be due to either
the dramatically different rates of development (s.
ingentis hatches after ca. 30 hours, C. anthonyi afterca. 35 days) or to differences in the chemical formsof the molting hormones. For example, conjugatedmetabolites of ecdysteroids have a much lower
affinity for the ecdysteroid antiserum and hence, if
present, would give the overall appearance of lesstotal ecdysteroid RIA activity. However, we favor the former explanation (differen tial rates ofdevelopment) .
Recently, data were presented that implicate a rolefor ecdysteroids in the mediation of embryogenesis
in the shrimp Palaemon serratus (Spindler et al. 1987).
In that species, although much lower concentrations
were measured (peaks of about 80 ng/g), a similar
hormone pattern was observed of low levels of
ecdysteroids at egg extrusion followed by increasinglevels near hatch.

Vitellogenesis-Inhibiting Hormone
S. ingentis is a useful species for the assay of VIH
because it undergoes several cycles of reproduction
without intervening molt cycles in the summer
months. Following a spawn, shrimp were injectedwith extracts of sinus glands obtained from winter(nonreproductive) female shrimp. A significant inhibition (P<O.OI) of ovarian development andspawning resulted. This effect was not observed inTable 1

Effect of shrimp sinus gland extracts on spawning duration.

Extract
nonsinus gland neural tissue
sinus glandsa P < 0.01 (Student's t-test).

Spawning Duration(days ± S.D.)

N

16.31 .:':: 1.31
19.93 ::':: 2.08 a

16
15

Chang et al: Reproductive Endocrinology of the Shrimp

control shrimp that were injected with nonsmus
gland neural tissue (Table 1) or summer shrimp thatwere injected with sinus gland extracts obtained fromsummer donor females (Chang and Hertz, un pub!.
data) .
We have observed differences in the peptide profiles from sinus glands of winter (quiescent) andsummer (active) shrimp following separation using
high-performance liquid chromatography (Changand Hertz, unpub!. data). We have purified theseshrimp sinus gland peptides and are currently assaying them for VIH activity.
There are reports that provide some chemical information on VIH. Most of these previous
experiments utilized a heterologous assay system.
Bomirski et a!. (1981) extracted a factor with a molecular weight of approximately 2000 from eyes talksof the crab Cancer magister. Their assay consisted ofmeasuring ovarian growth in the shrimp Crangon
crangon. A 5000 dalton factor was extracted from eyestalks of the spiny lobster Panulirus argus. It was
assayed by measuring ovarian growth in the fiddler
crab Uca pugilator (Quackenbush and Herrnkind
1983). A 7500 dalton peptide was isolated from sinus
glands of the American lobster Homarus americanus
and assayed in vivo in the shrimp Palaemonetes varians(Soyez et a!. 1987). The assay consisted of measuringoocyte diameters. Antisera were also raised against
this lobster VIH which cross-reacted with sinus glandextracts from a number of different decapods (Meusy
et a!. 1987). A 3300 dalton factor was characterizedfrom eyestalks of Penaeus setiferus as assayed in vitro
using fiddler crab ovaries. This assay utilized precipitation of radiolabeled leucine by antibodies that had
been generated against fiddler crab vitellogenin(Quackenbush and Keeley 1988).

Farnesyl Diazomethyl Ketone
Using the radiolabeled photoaffinity analog ofmethyl farnesoate, [3H]-FDK, we examined a numberof different tissues from several different species for
specific hormone binding. We examined both the cytosol and membrane fractions of these tissues.
Although we were unable to demonstrate cellularbinding proteins for [3H]-FDK from any tissue, we
consistently observed specific binding in the hemolymph. Figure 4 shows the effects of increasing
amounts of unlabeled MF in the presence of a constant amount of [3H]-FDK. The radiolabeled analog
forms a covalent bond with a binding protein of ca.
36,000 daltons and is effectively competed with a 250fold excess of the unlabeled hormone. In addition,

M

50x

lOx

15"

30"

II

2

1

41

25x

5

250x
500
500x
100x

M

o
.-200
'-92
..-66

..-45
..-36

..-29
'-24

.-20

'-14

Figure 4
Comparison of photolysis
times (15 s, 30 s, I min, 2min, and 4 min), and thequantity (fold excess) of
unlabeled methyl farnesoate(MF) used to displace the labeling of female shrimp
hemolymph
with
[3H]_
farnesyl diazomethyl ketone(photolysis time was 4 min) .
The band at ca. 36,000
daltons appears to be specifically labeled. The lanes
marked M500 and MO represent hemolymph from male
shrimp labeled with [3H]_
FDK with and without a
SOD-fold excess of unlabeledMF, respectively. Molecularweight markers (X 10-3 ) are
given on the right of the autoradiograph. No labeling isobserved at zero photolysis
time.

6

NOAA Technical Report NMFS 106

the effects of increasing the length of time of U.v.
irradiation is demonstrated. A high degree of attachment of [3H]-FDK to the hemolymph binding protein
is observed after only 15 sec (Fig. 4). The 36,000
dalton protein appears to be present in both female
and male shrimp (Fig. 4).
We have also recently utilized [3H]-FDK to characterize an analogous binding protein for MF in thehemolymph of adult female American lobsters(Prestwich et al. 1990). The lobster MF binding protein has an approximate molecular weight of 42,000.
Although a definitive hormonal role for MF has
not yet been established, these data suggest that MF
has a specific binding protein that may prevent its
rapid degradation and may facilitate its cellular action. The role of the crustacean MF binding protein
may be analogous to the insect juvenile hormone
binding protein.

Acknowledgments
We thank M. J. Bruce for technical assistance. Financial support was provided by NOAA, National Sea
Grant College Program, Department of Commerce,
under grant number NA85AA-D-SGI40, project number R/ A-68, through the California Sea Grant
College Program (to E.S.C.). We also acknowledge
Prof. J. Clegg and the University of California,
Bodega Marine Laboratory for Distinguished Research Fellow support to G.D. Prestwich. The NSF is
acknowledged for support of the ligand synthesis(Grants CHE-8809588, DCB-8509629, and DCB8812322 to G.D.P.).

CitationsAdiyodi, R.G.
1985. Reproduction and its control. In The biology of crustacea, vol. 9 (D.E. Bliss and L.H. Mantel, eds.), p. 147215. Acad. Press, Inc., Orlando.
Baldaia, L., P. Porcheron,J. Coimbra, and P. Cassier.
1984. Ecdysteroids in the shrimp Palaemon serratus: relations
with molt cycle. Gen. Compo Endocrinol. 55:437-443.
Bomirski, A., M. Arendarczyk, E. Kawinska, and L.H. Kleir holz.
1981. Partial characterization of crustacean gonad-inhibiting
hormone. Int. J. Invertebr. Reprod. 3:213-219.
Chang, E.S.
1989. Endocrine regulation of molting in Crustacea. Rev.
Aquatic Sci. 1:131-157.
1992. Endocrinology of shrimp and other crustaceans. In
Culture of marine shrimp: principles and practices (A.W.
Fast and L.J. Lester, eds.). Elsevier Science Publ., NY [npress.
Chang. E.S.. and J.D. O'Connor.
1979. Arthropod molting hormones. In Methods of hormone radioimmunoassay, 2nd ed. (B.M. Jaffe and H.R.
Behrman eds.), p. 797-814. Acad. Press. Inc., NY

Charniaux-Cotton, H., and G. Payen.
1988. Crustacean reproduction. In Endocrinology of Selected Invertebrate Types (H. Laufer and R.G.H. Downereds.), p. 279-303. Alan R. Liss, Inc., NY
Goodman, W.G., and E.S. Chang.
1985. Juvenile hormone cellular and hemolymph binding
proteins. In Comprehensive insect physiology, biochemisLry and pharmacology, vol. 7 (G.A. Kerkut and L.1. Gilbert,eds.), p. 491-510. Pergamon Press, Ltd., Oxford.
Laufer, H., D. Borst, F.C. Baker, C. Carrasco, M. Sinkus, C.C.
Reuter, L.W. Tsai, and D.A. Schooley.
1987. Identification ofajuvenile hormone-like compound ina crustacean. Science 235:202-205.
Meusy,J.-J., G. Martin, D. Soyez,J.E. van Deijnen, andJ.-M. Gallo.
1987. Immunochemical and immunocytochemical studiesof the crustacean vitellogenesis-inhibiting hormone(VIH). Gen. Compo Endocrinol. 67:333-341.
Mohamed, M.A., KA. Lerro, and G.D. Prestwich.
1989. Polyacrylamide gel miniaturization improves protein
visualization and autoradiographic detection. Anal.
Biochem. 177:287-290.
Okazaki, R.K, and E.S. Chang.
1991. Ecdysteroids in the embryos and sera of the crabs,
Cancl'T magister and C. anthonyi. Gen. Compo Endocrinol.
81: 174-186.
Panouse,J.
1943. Influence de l'ablation du pedoncule oculaire sur lacroissance de l'ovaire chez la Crevette Leander serratus.
Comptes Rendus Academie des Sciences, Paris 217:553-555.
1944. L'action de la glande du sinus sur l'ovaire chez laCrevette Leander. Comptes Rendus Academie des Sciences,
Paris 218:293-294.
Prestwich, G.D., MJ. Bruce, I. Ujvary, and E.S. Chang.
1990. Binding proteins for methyl farnesoate in lobster tissues: detection by photoaffinity labeling. Gen. Compo
Endocrinol. 80:232-237.
Quackenbush, L.S., and W.F. Herrnkind.
1983. Partial characterization of eyestalk hormones controlling molt and gonadal development in the spiny lobster
Panulirus argus. J. Crustacean BioI. 3:34-44.
Quackenbush, L.S., and L.L. Keeley.
1988. Regulation of vitellogenesis in the fiddler crab, Uca
pugilator. BioI. Bull. (Woods Hole) 175:321-331.
Rosenberry, B.
1990. World shrimp farming 1989. Aquaculture Digest, San
Diego, 28 p.
Soyez, D.,J.E. Van DeUnen, and M. Martin.
1987. Isolation and characterization of a vitellogenesis-inhibiting factor from sinus glands of the lobster, Homarus
ammcanus. J. Exp. Zoo I. 244:479-484.
Spindler, K-D., A. Van Wormhoudt, D. Sellos, and M. SpindlerBarth.
1987. Ecdysteroid levels during embryogenesis in the
shrimp, Palaemon serratus (Crustacea Decapoda): quantitative and qualitative changes. Gen. Compo Endocrinol.
66:116-122.
Ujvary, I., and G.D. Prestwich.
1990. An efficient synthesis of the crustacean hormone[12-'H]-methyl farnesoate and its photolabile analog[l3-'H]-farnesyl diazomethyl ketone. J. Labelled Compd.
& Radiopharm. 28:167-174.
Van Wormhoudt, A., P. Porcheron, and L. Le Roux.
1986. Ecdysteroids et synthese proteique dans l'hepatopancreas de Palaemon serratus (Crustacea, Decapoda) au cours
du cycle d'intermue. Bull. Soc. Zool. Fr. 110:191-204.

Hormonal Control of Reproductive Maturation in Penaeid Shrimp

ISAOYANO
National Research Institute of Aquaculture
Nakatsuhama Nansei-cho,Mie 516-01,Japan

ABSTRACTControl of reproductive maturation is a major problem to the development of commercial aquaculture programs for penaeid shrimp. Although hormonal control ofmaturation is well documented in oviparous vertebrates and insects, similar knowledge
for crustaceans is fragmentary. It has long been suspected that reproductive maturation
may be controlled by two antagonistic hormones, one that stimulates and the other that
inhibits. Ovarian maturation in penaeid shrimp has been induced and accelerated by
implantation of thoracic ganglion prepared from maturing female lobsters. This indicates that ovarian maturation may be induced by a gonad-stimulating hormone (GSH)
secreted by the thoracic ganglion of maturing females. Many workers have reported thatovarian maturation is regulated by a gonad-inhibiting hormone (GIH) from the X organsinus gland complex. This GIH probably has an antagonistic relationship with GSH
secreted by the thoracic ganglion. This paper further details our present understandingof the endocrine systems of penaeid shrimp and how they control reproductive maturation. It is suggested that the brain, mandibular organ, or ovary has a close relationshipwith the X organ-sinus gland complex or thoracic ganglion, which secrete GIH or GSH,
in regulating ovarian maturation in penaeid shrimp.

Introduction_

Control of reproductive maturation is a major
problem to the development of commercial aquaculture programs for penaeid shrimp. Controlling
reproduction in captivity could help to provide a reliable year-round supply of juveniles, serve indeveloping selective breeding programs, and be generally useful for obtaining disease-free spawners.
Eyestalk ablation has been used to mature female
shrimp in captivity in conjunction with the management of water temperature, photoperiod, lightintensity, areal density, sex ratio, and nutrition(Caillouet 1973; Lumare 1979; Yano 1984; Primavera
1985; Crocos and Kerr 1986). Knowledge of. hormonal control of reproductive maturation incrustaceans, however, is fragmentary. Although manyobservations of endocrine systems have been conducted on the inhibition of reproductive maturationby eyestalk hormone(s) since the pioneering work of
the 1940s (Panouse 1943, 1944), recent research has
focused mostly on organs (e.g., brain, thoracic ganglion, ovary, and mandibular organ) and theirfunctions, which are closely related with the releaseof gonad-stimulating factors or hormone (s) in crustaceans. This paper summarizes our present knowledgeof the endocrine systems of penaeid shrimp and how
they control reproductive maturation.

Eyestalk Hormone Effects
on Maturation_

Eyestalk ablation stimulates ovarian maturation inpenaeid shrimp, palinurid lobsters, and other decapod crustaceans (Caillouet 1973; Yano 1984; Yano,
unpub\. data). This treatment reduces the production of a gonad-inhibiting hormone (GIH), and thus
permits maturation of the ovaries in females. Many
workers suggest that reproductive maturation inpenaeid shrimp is regulated by a GIH from the X
organ-sinus gland complex in the eyestalks. In thiscomplex, the acidophilic sinus gland is connectedwith the axons from the neurosecretory cell of the X
organ, which is located in the medulla terminalis ofpenaeid shrimp (Yano, unpub\. data) and the other
7

10

NOAA Technical Report NMFS 106

Figure 5Thoracic ganglion, showing
neurosecretory cells in Penaeus
japonicus. Neurosecretory cells
contained basophilic granules
dispersed throughout the cytoplasm (Yano, unpub\. data).
Haematoxylin and eosin stain,
X65.

(ValVi)
5

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E

...

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QI

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4

C

,...

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QI
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III
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c

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QI

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~

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r-

""""

--""""

a!

0

RS

AG

TG

Figure 6

Rate of Vg increase in sera of maturing female Penaeus
japonicus, 48 hours after injection of abdominal (AG) andthoracic ganglion (TG) extracts. Vi: Initial Vg concentration; Va: Vg concentration 48 hours after injection; RS:
Ringer solution (Yano, un pub!. data) .

antagonistic relationship with GIH in regulating vitellogenesis (Fig. 7). Unilateral eyestalk ablation was
not effective in increasing serum Vg in maturing female P japonicus (Fig. 8, Yano, un pub!. data). Thissuggests that GIH may decrease quickly immediately
befure the initiation of vitellogenesis and then stay at
a low level until after the vitellogenesis is completed.
Therefore, eyestalk ablation may no longer be effective in regulating the production of GIH after
vitellogenesis has been initiated. On the other hand,
injection of thoracic ganglion extract, prepared fromvitellogenic females, was effective in increasing serum Vg, even in maturing females (Fig. 6). This
indicates that after initiation of vitellogenesis, higher
amounts of GSH, which are increased by injection ofthoracic ganglion extract, accelerate Vg synthesis andits release into the blood. This implies the possibility
that in penaeid shrimp, GSH levels may increase further with advancement of vitellogenesis, parallel to adecrease in the level of GIH (Fig. 7).
It is well known that biogenic amines release
peptide neurohormones from neuroendocrine struc-.----------------------------------------.
Vitellogenesis

Ql

Vitellogenesis~ HighQl

C

Figure 7

E

Model for the control of vitellogenesis inpenaeid shrimp. Gonad-stimulatinghormone (GSH) probably has an antagonistic relationship with gonadinhibiting-hormone (GIH) in regulating
vitellogenesis.

o

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Yano: Hormonal Control of Reproductive Maturation in Shrimp(Va/Vi)
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Eyestalks
intactFigure 8

Rate of Vg increase in sera of maturing female
Penaeus japonicus, 48 hours after unilateral eyestalkablation. Vi: Initial Vg concentration; Va; Vg concentration 48 hours after injection (Yano, unpub!. data).

(ValVi)E 25...::lQI
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Figure 9Rate of Vg increase in sera of control (0.1 f.Lg pure ethanol/g body weight) and treated female Penaeus japonicus,
48 hours after 17 a-hydroxy-progesterone (0.1 f.Lg/g bodyweight) injection. ET: Ethanol; 17P: 17 a-hydroxy-progesterone; Vi: Initial Vg concentration; Va: Vg concentration
48 hours after injection (Yano 1987a).

tures in several crustaceans (Fingerman 1985). Serotonin has been found to induce the release ofmolting-inhibiting hormone from isolated eyes talks(Mattson and Spaziani 1985). This implies the possibility that biogenic amines may stimulate the release11of GIH from the X organ-sinus gland complex incrustaceans.
Vitellogenin has been identified electrophoretically and immunologically in the haemolymph ofvitellogenic female crustaceans (Horn and Kerr
1969; Fielder et al. 1971; Wolin et al. 1973; Fyffe andO'Connor 1974; Yano 1987a; Yashiro 1989). Therefore, extra-ovarian tissue has been suspected for along time as the site of Vg synthesis in crustaceans. In
fact, evidence has been presented to show that Vg is
synthesized by the fat body or adipose tissue in amphi pods and isopods (Picaud 1980; Croisille andJunera 1980; Souty and Picaud 1981). Recently, several workers demonstrated that Vg is synthesized on alarge scale by the ovaries of crayfish (Lui et al. 1974),
fiddler crabs, Uca pugilator (Eastman-Reks andFingerman 1985), kuruma prawns (Yano and Chinzei
1987), and white prawns (Quackenbush 1989; Rankin
et al. 1989). Considering these observations, the siteof Vg synthesis in decapods is different from that in
isopods and amphipods. Vitellogenin is secreted into
the haemolymph after its synthesis and then accumulated in the developing oocytes as vitellin (Yano
1988). It is known that ecdysone, which is producedin the Y-organ, stimulates vitellogenesis in the isopod
Procellio dilatatus and amphipod Orchestia gammarella(Souty et al. 1982; Blanchet-Tournier 1982;
Charniaux-Cotton 1985). Recently, evidence has
been presented to show that 17Cl-hydroxy-progesterone stimulates Vg synthesis or release, or both, into
the blood in kuruma prawns (Fig. 9; Yano 1987a),fresh water prawn, Macrobrachium lanchesteri (George
and Khoo 1989), and black tiger shrimp, P monodon(Yashiro 1989). The hormone 17 Cl-hydroxy-progesterone is generally distributed in the ovary ofcrustaceans (Kanazawa and Teshima 1971). Junera etal. (1977) deduced the 'existence of a Vg-stimulating
ovarian hormone(VSOH)
which controls
vitellogenin synthesis in females of O. gammarella. It is
probable that 17 Cl-hydroxy-progesterone stimulates
Vg synthesis and release into haemolymph as a VSOH
in penaeid shrimp and other shrimp (Fig. 10). By
immunofluorescence, Vg was found to occur for the
first time in the follicle cells of the oil globule stage-I
oocytes in early developing ovaries, which actively
synthesize Vg (Yano and Chinzei 1987). The follicle
cells greatly expanded on the oil globule stage-I
oocytes (Yano and Chinzei 1987; Yano 1988). Therefore, the follicle cells are nominated as a possible cell
type responsible for ovarian Vg synthesis in kuruma
prawns. It is suggested that 17 Cl-hydroxy-progesterone, probably secreted from the ovary as a VSOH,
stimulates Vg synthesis in follicle cells and Vg release
into the haemolymph in penaeid shrimp (Fig. 10).
On the other hand, brain extracts, prepared from

12

NOAA Technical Report NMFS 106

tion in P japonicus (Laubier-Bonichon 1978; Yano
1984). High temperature (25° C) and long daylength(15 hours) induce Vg synthesis and secretion into theblood in kuruma prawn (Yano 1987a). Lumare(1981) demonstrated that even in ablated P japonicus
full sexual maturation did not occur below 17° C. Increasing ovarian development and rapid maturation
were observed as water temperature rose above 18° C(Yano 1987b). The change from 12 to 13 hours of
daylight was also observed to accelerate the ovarian
maturation (Yano 1987b). In un ablated P monodon, alonger photoperiod of 19 hours (light) did not stimulate ovarian maturation (Beard and Wickins 1980).
These findings indicate that the effect of GSH and
GIH on induction and inhibition of ovarian maturation or vitellogenesis is affected by water temperatureand photoperiod in female shrimp. It is probablethat the release of GSH and GIH from the thoracic
ganglion and the X organ-sinus gland complex may
be induced directly or indirectly by water temperature and light stimuli, via respectively the antennule(Barber 1961) and eye (Waterman 1961) in female
penaeid shrimp, as shown in Figure 10.

Temperature

I
Light

~

CitationsFigure 10
Scheme of the factors affecting vitellogenesis and associated endocrine patterns in penaeid shrimp. An:
Antennule; Br: Brain; Xo-Sg: X organ-sinus glandcomplex; Th: Thoracic ganglion; Fc: Follicle cell; 00:
Oocyte; Ov: Ovary; BH: Brain hormone; GIH: Gonad-inhibiting hormone; GSH: Gonad-stimulating
hormone; 17P: 17 a-hydroxy-progesterone.

maturing females induced Vg synthesis in P j(l,ponicus(Yano, unpub!. data). This suggests the presence of abrain hormone that stimulates the release of GSH inpenaeid shrimp (Fig. 10). Also, mandibular organ
implantation could stimulate vitellogenesis in the spider crab Libinia emarginata (Hinsch 1980). Laufer etal. (1987) found that methyl farnesoate, an
unepoxidated analog of the insect juvenile hormone
III, was synthesized by the mandibular organ of spider crabs, L. emarginata. However, the function of
methyl farnesoate is not known to control maturationin crustaceans.

Temperature and Light Effects
on Maturation_

It is well known that ovarian maturation is affected bywater temperature and photoperiod in female
shrimp. A photoperiod of 14-16 hours (light) andtemperature of 24-26° C stimulate ovarian matura-

Barber, S.B.
1961. Chemoreception and thermoreception. In The physiology of Crustacea Vol. 2 (T.H. Waterman, cd.), p. 109131. Acad. Press, NY.
Beard, TW., andJ.F. Wickins.
1980. Breeding of Penaeus monodon Fabricius in laboratory
recirculation systems. Aquaculture 20:79-89.
Bell, T A., and D. V. Lightner.
1988. Compound eye and associated organs. In A handbook of normal penaeid shrimp histology, p. 26-33. World
Aquaculture Society, Baton Rouge, LA.
Blanchet-Tournier, M.F.
1982. Quelques aspects des interactions hormonales entre lamue et la vitellogenesis chez Ie Crustace Amphipode
Orchestia gammarflla (Pallas). Reprod. Nutr. Dev. 22:325344.
Bomirski, A.M., A.E. Kawinska, and L. H. Kleinholz.
1981. Partial characterization of crustacean gonad-inhibiting
hormone. Int..J. Invertebr. Reprod. 3:213-220.
Caillouet, C.W.
1973. Ovarian maturation by eyestalk ablation in pink
shrimp Penaeus duorarum Burkenroad. Proc. World
Maricul. Soc. 3:205-225.
Carlisle, D.B., and F.K. Bart.
1959. The neurosecretory system of the head and thorax. In
Endocrine control in crustaceans (M. Abercrombie, P. B.
Medawar, and G. Sal, eds.), p. 13-39. The Univ. Press,
Cambridge.
Charniaux-Cotton, H.
1985. Vitellogenesis and its control in malacostracan
Crustacea. Am. Zool. 25: 197-206.
Crocos, P j., and J.D. Kerr.
1986. Factors affecting induction of maturation and spawning of the tiger prawn, Penaeus esculentus (Haswell), underlaboratory conditions. Aquaculture 58:203-214.

Yano: Hormonal Control of Reproductive Maturation in Shrimp
Croisille, Y., and H.Junera.
1980. Recherche du lieu de synthese de la vitellogenine chez
Ie Crustace amphipode Orehestia gammarella (Pallas). Demonstration, a l'aide d'anticorpos specifiques, de la presencede vitellogenine dans Ie tissu adipeux sous-epidermique defemaelles en vitellogenese secondaire. C. R. Acad. Sci.,
Paris 290:]487-1490.
Eastman-Reks, S., and M. Fingerman.
1984. Effects of neuroendocrine tissue and cyclic-AMP on
ovarian growth in vivo and in vitro in the fiddler crab, Uea
pugilator. Compo Biochem. Physiol. 79A:679-684.
1985. In Vitro synthesis of vitellin by the ovary of the fiddler
crab, Uea pugilator. J. Exp. Zool. 233: 111-116.
Fielder, D.F., K.K. Rao, and M. Fingerman.
1971. A female-limited lipoprotein and the diversity ofhemosyanin components in the dimorphic variants of thefiddler crab, Uea pugilator, as revealed by disc electrophoresis. Compo Biochem. Physiol. 39B:291-297.
Fihgerman, M.
1985. The physiology and pharmacology of crustacean
chromatophores. Am. Zool. 25:233-252.
Fyffe, W.E., and J.D. O'Connor.
1974. Characterization of quantification of a crustaceanlipovitellin. Compo Biochem. Physiol. 47B:851-867.
George, S., and H.W. Khoo.
1989. Effects of estrogens and progesterones on vitellogenin
levels in the haemolymph of the shrimp, Maerobraehium
lanehesteri. In Abstracts of the Fifth International Congressof Invertebrate Reproduction; 23-28 July 1989, Nagoya, Japan, p. 57. Publ. by Int. Soc. Invert. Reprod., Mainz, Germany.
Hagedorn, H.H., andJ.G. Kunkel.
1979. Vitellogenin and vitellin in insects. Annu. Rev.
Entomol. 24:475-505.
Hinsch, G.W.
1980. Effect of mandibular organ implants upon the spider
crab ovary. Trans. Am. Microsc. Soc. 99:317-322.
Hinsch, G.W., and D.C. Bennett.
1979. Vitellogenesis stimulated by thoracic ganglion implants into destalked immature spider crab, Libinia
emarginata. Tissue & Cell II :345-351.
Horn, E.C., and M.S. Kerr.
1969. The haemolymph protein of the blue crab, Callinectes
sapidus. I. Hemocyanins and certain other major protein
constituents. Compo Biochem. Physiol. 29:493-508.
Junera, H.,C. Zerbib, M. Martin, andJJ. Muesy.
1977. Evidence for control of vitellogenin synthesis by an
ovarian hormone in Orehestia gammarella (Pallas), Crustacea,
Amphipoda. Gen. Compo Endocrinol. 3] :457-462.
Kanazawa, A., and S. Teshima.
1971. In vivo conversion of cholesterol to steroid hormonesin the spiny lobster, Panulirus japonicus. Bull. Jpn. Soc.
Fish. 37:891-898.
Laubier-Bonichon, A.
1978. Ecophisioloie de la reproduction chez la crevett
Penaeus japonicus trois annees d'experience en milieu
controle. Oceanol. Acta. 1:135-150.
Laufer, H., D. Borst, F.C. Baker, C. Carrasco, M. Sinkus, C. Reuter,
L.W. Tsai, and D.A. Schooley.
] 987. The identification of a juvenile hormone-like compound in a crustacean. Science 235:202-205.
Lui, C.w., B.A. Sage, and J.D. O'Connor.
1974. Biosynthesis of lipovitellin by the crustacean ovary.
J. Exp. Zool. ]88:289-296.
Lumare, F.
1979. Reproduction of Penaeus kerathurus using eyes talkablation. Aquaculture 18:203-214.

13] 981. Artificial reproduction of Penaeus japonieus Bate as basis for commercial production of eggs and larvae. J. World
Maricul. Soc. ]2:335-344.
Mattson, M. P., and E. Spaziani.
1985. 5-hydroxytryptamine mediated release of molt-inhibiting hormone activity from isolated crab eyestalk
ganglia. BioI. Bull. (Woods Hole) 169:246-255.
Meusy,J.:J., G. Martin, D. Soyez,J.E. van Deijnen, andJ.-M. Gallo.
1987. Immunochemical and immunocytochemical studies of
the crustacean vitellogenesis-inhibiting hormone (VIH).
Gen. Compo Endocrinol. 67:333-341.
Otsu, T
1960. Precocious development of the ovaries in the crab,
Potamon dehaani, following implantation of the thoracic
ganglion. Annot. Zool.Jpn. 33:90-96.
Oyama, S.N.
1968. Neuroendocrine effects on ovarian development in
the crab Thalamita crenata Latreille studied in vitTO. Ph. D.
diss., Univ. Hawaii, Honolulu, Hawaii.
Panouse,J. B.
1943. Influence de I'ablation du predoncule oculaire sur lacroissance de I' ovaire chez la crevette Leander sel'ratus.
C. R. Acad. Sci., Paris 217:553-555.
1944. L'action de la glande du sinus sur l'ovaire chez lacrevette Leander. C. R. Acad. Sci., Paris 2]8:293-294.
Picaud, J.-L.
] 980. Vitellogenin synthesis by the fat body of Poreelio
dilatatus Brandt (Crustacea, Isopoda). Int. J. Invertebr.
Reprod.2:341-349.
Primavera,J.H.
1985. A review of maturation and reproduction in closed
thelycum penaeids. In Proceedings of the first international
conference on the culture of penaeid prawns/shrimps; 4-7
December] 984, Iloilo City, The Philippines (Y. Taki, J.H.
Primavera, and J.A. Llobrera, eds.), p 47-64. Japan Int.
Cooper. Agency, Tokyo, Japan.
Quackenbush, L.S.
1989. Yolk protein production in the marine shrimp Penaeus
vannamei. J. Crustacean BioI. 9:509-5]6Quackenbush, L. S., and W. F. Herrnkind.
]983. Partial characterization of eyestalk hormone controlling
molt and gonadal development in the spiny lobster,
Panulil'us argus. J. Crustacean BioI. 3:34-44
Rankin, S.M.,J.Y. Bradfield, and L.L. Keeley.
1989. Ovarian protein synthesis in the South American white
shrimp, Penaeus vannamei, during the reproductive
cycle. Int. J. Invertebr. Reprod. 15:27-33.
Souty, C., andJ.L. Picaud.
1981. Vitellogenin synthesis in the fat body of the marine
crustacea isopoda, Idotea balthica basteri, during
vitellogenesis. Reprod. Nutr. Dev. 21:95-102.
Souty, C., G. Besse, andJ.-L. Picaud.
] 982. Ecdysone stimulates the rate of vitellogenin release inhaemolymph of the terrestrial crustacean Isopoda Poreellio
dilatatus Brandt. C.R. Acad. Sci., Paris, 294:1057-1060.
Takayanagi, H., Y. Yamamoto, and . Takeda.
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Tata,J.R.
1978. Induction and regulation of vitellogenin synthesis by
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1961. Light sensitivity and vision. In The physiology ofCrustacea, Vol. 2 (TH. Waterman, ed.), p. 1-64. Acad.
Press, NY.

14

NOAA Technical Report NMFS 106

Wolin, E.H., H. Laufer, and D.F. Albertini.
1973. Uptake of the yolk protein lipovitellio by de'/eloping
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1984. Induction of rapid spawning in kuruma prawn, Penaeus
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1987a. Effect of 17 o:-hydroxy-progesterone on vitellogenin
secretion in kuruma prawn, Penaeus japonicus. Aquaculture
61:49-57.
1987b. Maturation of kuruma prawns Penaeus japonicus cultured in earthen ponds. In Reproduction, maturation andseed production of cultured species: p'roceedings of thetwelfth U.S.:Japan meeting on aquaculture; 25-29 October
1983, Baton Rouge, LA (C.]. Sinderman, ed.:" p. 37. NOAA Tech. Rep. NMFS 47.
1988. Oocyte development in the kuruma prawn Penaeus
japonicus. Mar. BioI. (Berlin) 99:547-553.

Yano, I., and Y. Chimei.
19ts7. Ovary is the site of vitellogenin synthesis in kuruma
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1990. Effects of human chorionic gonadotropin and unilateral eyestalk enucleation on vitellogenin induction andovary development in kuruma prawn, Penaeus japonicus. In
Shrimp culture in the world (C.C. Justo, ed.), p. 365370. Midori Shohou, Tokyo,Japan
Yano, 1., B. Tsukimura,].N. Sweeney, and].A. Wyban.
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Fabricius. Ph.D. diss., Univ. Philippines System. Quezon
City, Philippines.

Reproductive Physiology and Induced Spawning of Yellowtail(Seriola quinqueradiata)

HIROHIKO KAGAWA
National Research Institute ofAquaculture
NanseiMie 5I6-0I,Japan

ABSTRACT
For the large-scale production of the yellowtail Senoia quinqueradiata an understandingof its reproductive physiology and the ability to artificially control its reproduction arenecessary. This article briefly reviews recent studies on the reproductive physiology andhormonally induced spawning of the yellowtail. Plasma steroid hormone levels associatedwith ovarian development and the results of spawning induced by using human chorionicgonadotropin and gonadotropin releasing hormone are discussed.

Introduction_

In Japan, the yellowtail Senoia quinqueradiata is one of
the most highly regarded fish species for fresh consumption. In addition, its suitability for marine
aquaculture is good. The annual production of cultured yellowtail currently amounts to about 165,000
metric tons (approximately 70% of total productionof marine finfish aquaculture; DSI 1988). Curren tly,
wild juvenile fish are caught as seed for aquaculture.
For the large-scale production of the yellowtail andstable production of seed, an understanding of its
reproductive physiology and the ability to control its
reproduction are necessary. This article briefly reviews recent studies on the reproductive physiology
and hormonally induced spawning of yellowtail.

Reproductive Physiology_

After the fry stage, female yellowtail normally attainsexual maturity in floating net pens in about three
years. Oocyte diameter begins increasing in February.
The most advanced oocytes rapidly increase t~eir diameter after vitellogenesis starts in March and reachabout 700 /-Lm at the tertiary yolk globule stage in
April (Fig. 1). The yellowtail has an asynchronoustype ovary which contains oocytes at various
developmental stages ranging from the perinucleolus
to tertiary yolk globule stage in the spawning season(April and May). Since yolky oocytes still remain in
the ovary after ovulation, yellowtail appear to be capable of multiple spawnings during a spawningseason.
It is well known that estradiol-1713 induces
vitellogenin synthesis in the teleost liver. The increase in plasma estradiol-17-13 levels correlates wellwith increase of gonadal weights (Kagawa et a!.
1983). Changes in plasma estradiol-1713 levels in yellowtail (Fig. 1) also correlate with the developmentof the ovary; estradiol-1713 levels are constantly low(below 1 ng/mL) in the previtellogenic stage from
December to February and increase rapidly during
the vitellogenic period reaching a maximum level inthe spawning season at a concentration of about 9
ng/mL (Kagawa et aI., unpub!. data). These values
found in female yellowtail are of the same order as
those reported for other fish that have asynchronoustype ovaries (goldfish, Carassius auratus, Kagawa et al.
1983; common carp, Cypnnus carpio, Santos et al.
1986). The compound 170., 2013-dihydroxy-4pregnen-3-one (170., 2013-diOHprog) was first
identified as a maturation-inducing steroid in amago
salmon Oncorhynchus rhodurus (Nagahama 1987) andhas been known to induce final oocyte maturation in
many other fish. Plasma levels of this steroid dramatically increased at the time of oocyte maturation in
many teleost species. In yellowtail, an increase of 170.,
2013-diOHprog (Fig. 2) was also observed during thefinal oocyte maturation that was induced using hu15

16

NOAA Technical Report NMFS 106,-----------------------------------------,

----

10

o

0>

C

c::o..
I"'-

~

I

o

is

I

I~~'

E

5

ctl
....

.....
CJ)

I

I

700 E

I '\\ .

~/

W

•

I

•

I
~

0- _ _ -

.-

-

-

~

\

_-0"

Spawning
period...

_ _- - - - - - .

Dec

Jan

Feb

Apr

Mar

-o

May

\

::t

....
(1)
500 CDE
ctl

is300 .....
(1)
>(,)

o

0

100 0

Jun

Figure 1
Changes in plasma estradiol-1713 levels and oocyte
diameter during the oocyte
development in the yellowtail (Kagawa et aI.,unpubi. data).

,0"," A-

0"" Ovulation 1000 ?

1000
/

E

--

/
/

Ol
0.

---o
Ol

g

I
I

/

E°:::t,..0_0'
0-- 0--0------...

•Q)
Q)

is 500

500 E

<:0..
o
N
d
f'-.

i•

.---.- ,
/

•
o

«S

is
Q)

:>.

u

o

....

6 12
24 30 36
Hours after HCG injection

o

•
48

man chorionic gonadotropin (HCG) injection(Kagawa et aI., unpub!. data). Values for plasma 170.,
2013-diOHprog obtained from yellowtail are on thesame order as those reported for goldfish (Kagawa et
a!. 1983), but are very low compared with those reported in salmonids (Young et al. 1983). The
difference between plasma 170., 2013-diOHprog levelsof yellowtail and salmonids may be due to the lower
capacity of the yellowtail ovarian follicle to produce
this steroid as was suggested in goldfish (Kagawa etal. 1983). It is still uncertain whether 170., 2013-

Figure 2
Changes in plasma 170., 2013-dihydroxy-4pregnen-3-one (170., 2013-diOHprog) levels andoocyte diameter in the yellowtail after injectionof human chorionic gonadotropin (Kagawa etai., unpub!. data).

diOHprog is the maturation inducing steroid in yellowtail. Further studies on in vitro effect of thissteroid on oocyte maturation and its production in
the ovarian follicles are necessary.
There is little information on the reproductive
physiology of male yellowtail. Males mature very
early; sperm formation can be observed in the testisof fish only 1 year old. Little attention has been paid
to the collection of milt for fertilization because
sperm formation occurs naturally even under net-culture conditions. The only attempt to control male

Kagawa: Reproductive Physiology and Spawning of Yellowtail

17

Table 1

Number of ovulated eggs, fertilization rates, and hatching rates in the yellowtail treated with single intramuscular
injection of human chorionic gonadotropin.
Experiment
numberNo. offish used

No. offish
ovulated

No. of eggs
ovulated

% of
fert.

% of
hatching

1
2
3
4
5

3
15
17
15
11

3
13
14
11
10

1,710,000
9,900,000
4,460,000
7,290,000
8,210,000

49.1
69.6
72.6
62.5
32.5

14.6
49.8
51.1
39.2
21.2

Table 2

Number of spawned eggs, fertilization rates, and hatching rates in the yellowtail implanted with cholesterol pellet
containing the gonadotropin releasing hormone analog.
Spawning
date

No. of spawned
eggs% fertilized% hatched

May 1
2
3
4
5
6
8
9
11

217,000
630,000
882,000
623,000
1,477,000
2,212,000
949,000
2,562,000
98,000

71.4
41.6
25.9
52.1
16.1
12.2
19.7
13.5
0

71.4
41.6
25.9
27.3
8.8
4.9
8.7
7.5
0

reproductive physiology is the use of a single injection of HCG two days before fertilization, to ensurethat a sufficient volume of milt is collected.

Induced Spawning_

Although female yellowtail have ovaries that containoocytes at the tertiary yolk stage Uust prior to final
oocyte maturation) during the spawning season, they
do not ovulate and spawn in the floating net pensnaturally. It has been known that HCG induces spawning in many fish including marine species, such as sea
bream Sparus aurata (Gordin and Zohar 1978). In cultured female yellowtail (mean body weight
8.6 kg), ovulation is induced by a single intramuscular
injection of HCG at a dose of 600-700 IV/kg bodyweight (= 6,000 IV/fish). Mter HCG treatment, ovulation occurred within 24 to 48 hours. For practical
purposes eggs were collected between 50 and 60
hours after HCG injection by manually stripping theirabdomen. The number of eggs obtained by HCG
treatment (Table 1) averaged about 500,000/fish.

Eggs were fertilized using milt from male fish treatedwith 6,000 IV of HCG two days before fertilization.
Fertilized eggs were then incubated in a I-ton tankfor 2 days at 20° C. The mean fertilization rate wasabout 60% and the hatch rate was about 40%. Ovulation, fertilization, and hatch rates varied with the fishused because ovarian maturity at the time of injection
was often different between fish. Their ovaries often
contained many degenerated oocytes and only a few
oocytes at the tertiary yolk stage. Assessment of ovarian maturity by cannulating oocytes from the fish
before hormone treatment will improve these results.
As mentioned earlier, yellowtail have a potential tospawn several times during a spawning season; however, they do not spawn again if ovulation is induced
by HCG injection. One of the most important reasons for this phenomenon is the stress incurred byhandling while they are injected with hormone andtheir abdomens stripped to obtain ovulated eggs.
These stressful treatments probably induce the degeneration of yolky oocytes. Gonadotropin releasing
hormone (GnRH) and its analog are used successfully to induce spawning in a variety of fish (Marte et

18

NOAA Technical Report NMFS 106

al. 1988). Recently in our laboratory, attempts to use
an implanted GnRH analog pellet to reduce theamount of handling and induce multiple natural
spawning have been started.
Seven female yellowtail (mean body weight 8.0 kg)
were implanted with a cholesterol pellet of gonadotropin-releasing-hormone analog, des Glylo [D-Ala 6 ]LHRH ethylamide (Sigma) 1,000 J..lg/fish on 29
April. Cholesterol pellets (2 X 5 mm in size) whichwere composed of cholesterol powder, cocoa butter,
and GnRH-analog were intramuscularly implanted.
Females were reared with seven males in a 60 m 3 tankand the spawned eggs were collected by net every
morning. Collected eggs were incubated in a I-ton
tank for 2 days at 20° C.
Preliminary results (Table 2) show that implantation of a cholesterol pellet containing GnRH analog
induced natural spawning. Yellowtail spawned eggs
over 11 days; fertilization and hatch rates were relatively high during the first 4 days but decreased
rapidly from day 5-even though the number ofspawned eggs increased. In this experiment, we could
not know how many times one fish spawned during
the experiment. Total number of spawned eggs by
the GnRH-analog treated fish (1,378,000 eggs/fish)
was more than twice that of HCG-injected fish
(500,000 eggs/fish). Thus, it is possible that somefish probably spawned more than twice during theexperiment. Because fertilization and hatching rates
rapidly decreased during the experiment, h:rther
studies are necessary to improve the techinques for
obtaining eggs of high quality.

Acknowledgment
Thanks is extended to the staffs of the Komame(Kochi Prefecture) and Goto (Nagasaki Prefecture)

Branchs of the Japan Sea-Farming Association for
their kind collaboration and technical assistance inperforming this work. A part of the data in this paper
has already been published in the Annual working
reports of the Japan Sea-Farming Association(Shinko Bldg., 2-1, Kanda Ogawa-cho, Chiyoda-ku,
Tokyo 101 1987). Unpublished data from this laboratory is a result of collaboration with K. Hirose and S.
Arai who are gratefully acknowledged.

Citations_

Department of Statistical Information (DSI)
1988. Fisheries Statistics of Japan, 142 p. DSI, Ministry ofAgriculture, Forestry, and Fisheries, Government ofJapan.
Gordin, H., and Y Zohar.
1978. Induced spawning of Sparus aurata by means of hormonal treatment. Ann. BioI. Anim. Biochem. Biophys.
18:985-990.
Kagawa, H., G. Young, and Y. Nagahama.
1983. Changes in plasma steroid hormone levels during gonadal maturation in female goldfish Carassius auratus.
Nippon Suisan Gakkaishi 49: 1783-1787.
Marte, C.L., N. Sherwood, L. Crim, and]. Tan.
1988. Induced spawning of maturing milkfish (Chanoschanos) using human chorionic gonadotropin and mammalian and salmon gonadotropin releasing hormone
analog. Aquaculture 73:333-340.
~agahama, Y.
1987. 17C1., 2013-Dihydroxy-4-pregnen-3-one: A teleost maturation-inducing hormone. Dev. Growth & Differ. 29:1-12.
Santos, AJG., K. Furukawa, K. Bando, K. Aida, and l. Hanyu.
1986. Photoperiodic determination of preovulatory gonadotropin surge on set time in the carp Cyprinus carpio.
Nippon Suisan Gakkaishi 57:1167-1172.
Yuung, G., L. Crim, H. Kagawa, A. Kambegawa, and Y. agahama.
1983. Plasma 17C1.,2013-dihyroxy-4-pregnen-3-one levels during sexual maturation of amago salmon (Oncorhynchus
rhodurus): Correlation with plasma gonadotropin and in
vi/m production by ovarian follicles. Gen. Compo
Endocrinol. 51:96-105.

Gametogenesis of Triploid Bivalves with Respect to Aquaculture

KATSUHIKO T. WADA AND AKIRA KOMARUNational Research Institute ofAquaculture
NanseiMie 516-01,Japan

ABSTRACTTriploid bivalves have been induced in many species and are expected to be better inquality than diploid animals in aquaculture in relation to retarded gametogenesis(Stanley et al. 1981; Allen et al. 1982, 1986). Recently however, triploids of some bivalve
species have been reported to mature and to produce active sperm and ripe eggs. This
paper deals with the gametogenesis in triploid bivalves with respect to resolving problems
associated with their release when they are introduced into intensive culture in the sea.
We have been studying triploids of the Japanese pearl oyster Pinctada Jucata martensii(Uchimura et al. 1989; Wada et al. 1989; Komaru and Wada 1990) and the scallop
Chlamys nobilis (Komaru et al. 1988). Gametogenesis proceeds very differently in these
two species as seen during the seasonal histological observation of gonads. Retardationof maturation was observed in both sexes of the triploid scallop: neither sperm nor
mature eggs were detected (Komaru and Wada 1989). However, some triploid pearl
oysters produced sperm and ripe eggs in gonads that were less developed than those of
diploids (Komaru and Wada 1990). Sperm dissected from some of the triploid pearl
oysters were active during microscopic examination. The eggs dissected from triploids
showed germinal vesicle breakdown after treatment with ammonium sea water as seen in
diploid eggs. Sperm from triploid pearl oysters had about 1.3 times higher mean values
and a wider distribution range of relative D A contents than those from diploids. We
have not observed the spawning of triploid pearl oysters in the histological observationof animals cultured under natural conditions (Wada and Komaru unpubl. data). However, in the oyster Crassostrea gigas, abnormal DNA-content values of sperm dissectedfrom triploids have been reported and the possibility of genetic abnormality was suggested in the D-shaped larvae. This genetic abnormality may have resulted from theinsemination of normal eggs from a diploid female with sperm dissected from a triploidmale (Allen 1987; Allen and Downing 1990; Akashige 1990). In the natural area occupied by the intensive cultures, it is necessary to avoid genetic abnormality in the zygotes
from the triploids because such a zygote might affect the native stocks. If triploid animals
release functional gametes to the sea, they should be cultured in a restricted and closed
area to avoid damage to natural reproduction. Another proposal may be the introductionof triploids of exotic species into areas of the sea that they do not currently inhabit andwhere growth of the species could be expected after the introduction.

Citations_Akashige, S.
1990. Growth and reproduction of triploid Japanese oyster in
Hiroshima Bay. In Proceedings of the fifth international ~on­gress of invertebrate reproduction; 23-28 July 1989, Nagoya(M. Hoshi and O. Yamashita eds.), p. 461-468. Elsevier,
Amsterdam.
Allen, S.K.,Jr.
1987. Gametogenesis in three species of triploid shellfish:
Mya arenaria, Crassostrea gigas and Crassostrea virginica. In

Proc. world symp. on selection, hybridization and genetic
engineering in aquaculture, Vol. II; (K. Tiews ed.),p. 208-217. Bordeaux, 27-30 May 1986 Heeneman
Verlaggesellschaft, Berlin.
Allen, S.K.Jr., P.S. Gagnon, and H. Hidu
1982. Induced triploidy in the soft-shell clams: cytogenetic
and allozymic confirmation. J. Hered. 73:421-428.
Allen, S.K.Jr., H. Hidu, andJ. G. Stanley1986. Abnormal gametogenesis and sex ratio in triploid softshell clams (Mya arenaria). BioI. Bull. (Woods Hole)
170: 198-210.

19

20

NOAA Technical Report NMFS 106

Allen, S. K. Jr., and S. L. Downing
1990. Performance of triploid Pacific oysters, Cmssostrea
gigas: gametogenesis. Can.]. Fish. Aquat. Sci. 47:121~1222.
Komaru, A. and K. T Wada
1989. Gametogenesis and growth of induced triploid scallops Chlamys nobilis. Nippon Suisan Gakkaishi 55:447-452.
1990. Gametogenesis of triploid Japanese pearl oyster, Pinetada
fucata martensiL In Proceedings of the fifth international congress of invertebate reproduction; 23-28 July 1989, Nagoya(M. Hoshi and O. Yamashita eds.), p. 469-474. Elsevier,
Amsterdam.
Komaru, A., Y. Uchimura, H. leyama and K. T Wada
1988. Detection of induced triploid scallop, Chlamys nobilis,
by D A microfluorometry with DAPI staining.
Aquaculture 69:201-209.

Stanley,].G., S. K. Allen,Jr. and H. Hidu
1981. Polyploidy induced in the American oyster, Crassostrea
virginica, with cytochalasin B. Aquaculture 23:1-10.
Uchimura, Y., A. Komaru, K. T Wada, H. leyama, M. Yamaki. and
H. Furuta
1989. Detection of induced triploidy at the different ages oflarvae of the Japanese pearl oyster, Pinetada fucata martensii
by microfluorometry with DAP( staining. Aquaculture
76:1-9.
Wada, K. T, A. Komaru and Y. Uchimura
1989. Triploid production in the Japanese pearl oyster,
Pinetadafueata martensiL Aquaculture 76:11-19.

Increasing the Growth Rate of Abalone, Haliotis discus hannai,
using Selection Techniques

MOTUYUKI HARA AND SHOGO KIKUCHI
Tohoku &gional Fishery &search Laboratory
3-27-5 Shinhama, Shiogama
Miyagi 985, Japan

ABSTRACT
Recently, a large number of abalone, Haliotis discus hannai, seeds have been produced
by artificial fertilization and cultured in Japan. Most of the seeds have been released into
the sea to increase abalone resources. However, some people have tried to continue thecultivation of the abalone seeds to commercial size in tanks. These efforts have been
economically unsuccessful because of the typically low growth rate of this species. Therefore, it is necessary to improve their growth genetically. The genetic variation among lotsof the seeds produced from different parents were examined. Differences in growthamong the lots were observed for 190 days during rearing in the same tank. Moreover, totest the association between growth and isozyme genes, the Gallele frequency at the
Pgm-l locus was compared between the large and small shell size classes. A higher frequency in the large shell size class was observed in 8 of II lots. This result suggests thatgrowth in abalone is closely related to genetic factors. We also attempted to genetically
improve abalone. By selecting and reculturing the fastest growing individuals for two
generations in small scale culture in our laboratory. The results suggest a possibility of
improving the growth rate of cultured abalone using selection techniques.

Introduction_

Presently, many marine species are produced by artificial fertilization and are cultured in Japan. One of
the most impor tant species is abalone. Twenty tothirty million abalone seeds are produced every year
by over fifty farming centers (DSI 1990). Most of theseeds are cultured to the size of 10-30 mm in shelllength then released into the sea in order to increase
the abalone resource. Some people have tried to continue the cultivation of abalone seeds to commercial
size in tanks; however these efforts have not been
economically successful to date because the seeds exhibited low growth rates. Therefore, it is necessary to
improve growth genetically. Artificial spawning techniques, diets for juveniles, and rearing equipment
needed for abalone culture have been researched formore than 20 years (Kikuchi and Uki 1974;
Takahashi and Koganezawa 1988 ; Uki 1989), but thegenetic characters of artificial seed have been notstudied at all. It is known that variations in the shellsize exist within and between lots of abalone seed.
Such a phenomenon could be caused by both genetic
differences and environmental differences such as
food, temperature, and density of individuals.

Estimation of the
Parental Effects on Growth--'---_ _We reared several lots of abalone (Haliotis discus
hannai) seed in the same tank and compared thegrowth of individuals. Plastic film tags (Hara 1989)
were used to identify individuals under the culture
conditions. The influence of the tagging on growth
was tested. At the beginning of this study (day 0), themean shell length of tagged and non tagged individuals was 32.4 and 32.8 mm, respectively (Table 1).
After 50 days, the mean shell lengths were 35.8 and

21

22

NOAA Technical Report NMFS 106

Table 1Size of tagged and non-tagged seeds after 50 days rearing in the same tank.
Days in culture

TaggedNontagged-'? -'?--o 0-'? -'?

0nT"-'? -'?

o50

32.4 ::': 2.3
32.8::': 2.1

35.8::': 2.2
35.7::': 2.0-'? -'?--o 00-'? -'?

0

Cultivation for one year in each tankMeasured with day 0-

Collect about 40 individuals
froa each lot

and aix

Measured with day 50-

in saae tank after tagging

Measured with day 90-

Experiaental conditions
later teap.: 20±1"C

Measured wi th day 140 -Food : Diatoas . Laminaria
and Eisenia

Measured with day 190 -Tank size : 158xll0x60ca.l ton

35.7 mm. Thus, no difference in growth was observed
after 50 days rearing in the same tank.
To examine parental effects, we compared thegrowth among three lots of offspring, each produced
from different sets of parents. Figure 1 shows the experimental procedure. Each lot was produced by four
females and three males in separate I-ton tanks (158
X 110 X 60 em). After 1 year of cultivation, about 40
individuals were collected from each lot, tagged, andthen mixed in the same tank. Shell lengths were measured at about 50 days intervals. Seawatertemperature was about 20° C, and food was supplied
first by diatoms, then by Laminana and Eisenia spp.
Figure 2 shows the growth curve (by shell length)of each lot. At day 0 the mean shell length of lots A,
B, and C were nearly identical at 23.6, 23.3, and 23.8mm, respectively. As the abalone grew, differences inshell length became large among the three lots. After
190 days from the beginning of the experiment, themean shell length of lot B was significantly larger

Figure 1
Experimental procedure used to culture
abalone and timetable used in assessing
their growth.

than that of lot A (i-test, P<0.05). This difference indicates parental effects in cultured abalone andsuggests genetic deficiencies.
In order to clarify these results, differences ingrowth of offsprings were estimated among three parental pairs (A, B, and C) each divided into and threedifferent size classes (small, medium, and large). Figure 3 shows the frequency distribution of theiroffsprings' shell lengths. Average daily growth andthe coefficient of variance for each set of offspring isshown in Table 2 by size class. In the offspring of pairA, the average daily growth of the small class was 58.6/-Lml day, that of the medium class was 82.3 /-Lml day,and the growth of the large class was 111.4 /-Lml day.
In offspring B, the average daily growth for small,medium, and large classes were 143.0, 134.1, and127.7 /-Lml day, respectively. In the offspring of C, thesmall, medium, and large classes were 127.8, 120.5
and 134.6 /-Lml day, respectively. Thus, daily growthmeans of the offspring from pairs Band C were re-Hara and Kikuchi: Growth Rate of Abalone

23

these results, one could surmise that the growth was
related to genotypes of parents.

50
E
E

Association between Pgm-l Allele
in Isozyme and Growth

~40

.s::
....,

_

en

c::
QJ

....

r-

30

QJ

.s::
V1

20

o

50

190

140

90
Days
Figure 2

Growth of offspring from different parental stocks. Age of
abalone at day 0 was 1 year old.
markably higher than that of the siblings from pair A
in all size classes. These results support the hypothesis that there are parental effects on growth in all
size classes. Differences were also noted in the daily
growth among the size classes within the pair Bandthe pair C offspring. The coefficient of variance inthe offspring of pair B ranged from 19.9 to 27.1 %.
The coefficient of variance in those from pair C
ranged from 11.2 to 28.0%. Pair A offspring, whichshowed the greatest differences in daily growthmeans between the small and large size classes, also
showed larger coefficients of variance than pair B
and pair C offspring in all size classes. The small sizeclass of the pair A offspring had the lowest daily
growth and largest coefficient of variance. This observation might be caused by the existence of a large
number of individuals with low growth rates. From

A-88

40

'O'l

To test the association between genotypes andgrowth, the enzyme polymorphism of phosphoglucomutase (PGM) isozyme was used. Electrophoresis
was carried out using an 11 % starch gel in 15.5 mM
tris and 4.5mM citrate acid buffer. The electrodebuffer was 0.155M tris and 0.045M citrate acid (pH7.0). A voltage of 14.25 V jcm was applied for 6 hours
at 4° C. The detection of phosphoglucomutase on
the gel was done by the staining procedure in Show
and Prasad (1970). Figure 4 shows electrophoretic
pattern of PGM isozyme in abalone, and the phenotypic variation observed. There are two loci, namely
Pgm-l and Pgm-2. Appearance of Pgm-2 is not stable;
therefore, Pgm-l was used as the genetic marker inthis study. A, B, C, and D indicate names of alleles.
The Band C alleles at the Pgm-llocus were predominate in all the seeds from the 11 lots examined. The
C allele of the Pgm-llocus was selected as the marker
gene in the present experiment.
To test the association between genotype andgrowth, the C allele frequency among 11 lots was examined. Mter cultivation for about one year, theseeds were divided into three classes: large, medium
and small. The Gallele frequency at the Pgm-l locus
was compared between the large and small classes. A
higher frequency (8 lots out of 11) was observed in
the large class (Table 3). If growth is controlled by
the C allele at the Pgm-l locus, the Gallele frequency
would be universally high in the large class. If growthis independent of the allele at the Pgm-l locus, the C
allele's appearance is likely to be equal in both the

8-88

C-88

30

>,
u

c:

Q)

::::l
tT

20

Q)

'lJ...

10
I

M

0
18

24

28

32

18

I

I

S

L

22

26

Figure 3

I

M IL

MIL

S

30

Shell length (mm)

16

22

26

30

Frequency distribution of shelllength in offspring of parental
pairs A-88, B-88, and C-88. S =small size class; M = medium
size class; L = large size class.

24

NOAA Technical Report NMFS 106

Table 2
Daily growth (I-Lm/day) and coefficient of variance in three size class of offspring fromthree parental pairs after 190 days of laboratory culturing.
Parentalpair

Offspring size class
Medium
Large

Small

A-88

58.6
(79.0%)
143.0(19.9%)
127.8(28.0%)

B-88
C-88

82.3
(57.1%)
134.1(23.8%)
120.5(30.7%)+

111.4(41.7%)
127.7(27.1%)
134.6(11.2%)A

B
C

Pgm -1

0

-

A

B

Pgm-2

C

BIB

C/C

AlB

A/C

BID

B/C

C/O
Figure 4
Phenotypic variations of PGM isozyme observed in abalone.

Genotype of Pgm-1

Table 3
C-allele frequencies at the Pgm locus of large and small classes.
Lot

Large class

A-85
B-85
C-85
A-87
B-87
C-87
D-87
E-87
A-88
B-88
C-88

0.330
0.315
0.465
0.919
0.466
0.497
0.662
0.787
0.216
0.497
0.500

(427)a
( 62)
(187)
(301)
(326)
(301)
(339)
(338)
(117)
(149)
( 93)

Small class
>
>
>
>
<
<
>
>
>
>
<

0.216
0.235
0.365
0.872
0.525
0.505
0.587
0.720
0.201
0.485
0.564

(162)
(249)
(230)
( 82)
(100)
(107)
(104)
(134)
( 67)
(132)
(110)

a Number of individuals examined.

classes. It could be tentatively concluded that the association between the Gallele frequency and growth
is related to the linkage of the phosphoglucoC'lutase
isozyme gene with the genes related to growth.

Growth of Selected Abalone_

The results above suggest that growth in abalone isclosely related to genetic factors. If this is true, anHara and Kikuchi: Growth Rate of Abalone

1st Generation

80

T

/

195 rm/day

1

E
E

I

60

.c

Selection oflarge sized seeds

.....

0'1

40

OJ

.c

3 d G

r

·

•

t

OJ

....-

I

•

240

c:

2ndGenerah

25

Vl

20

Selection oflarge sized seeds

1.

1Y

enera~

o

,

I

8

10

14

18

Months

Large sized seeds
used in this experiment

Figure 6
Growth of abalone offspring selected for superior growth rate.

Process of selecting experimental seeds.
Figure 5
Process used to select abalone seeds for experimentation.

Table 4
Average daily growth (i-Lm/day) of seeds from parents selected for high growth rate and commercial seeds.
Shell size(mm)Seed aSelectedControl

188

< 155
< 141

20-30
30-50
30-70
50-70
>70

240

< 147
195

a Maximum values of commercial seed growth reported in the past (Uki et al. 1981).

effect should be expected when selecting for the
maximum growth rate in abalone. The effect of selection for shell length was thus tested. Figure 5 shows
the process of selecting abalone for this experiment.
The original lot (first generation) were large-sized
seed taken from a fishermen's cooperative farming
center in Japan. The first generation was grown in
culture and produced offspring. The large-size classof these offspring (second generation) was grown in

culture and the large-size class offspring were similarly obtained. These offspring were also cultured for8 months. The large-size class from this lot was usedin this experiment.
The results are shown by the growth curve in Figure 6. The average shell length increased from 21 to
84 mm over a duration of 10 months (from age 8 toage 10 mon ths). While abalone were from 20 to 30
mm in shell length, the average daily growth was \88

26

NOAA Technical Report NMFS 106

f.Lm/day. From 30 to 70 mm in size, the daily growth
averaged 240 f.Lm/ day and those over 70 mm more
over grew about 195 f.Lm/ day.
These daily growth rates were compared to thoseof commercial seed (Table 4). The average daily
growth from 20 to 30 mm in shell length was 188
f.Lm/day in this experiment, while the maximum daily
growth in commercial seed was less than 155 f.Lm/ dayas reported by Uki et al. (1981). The daily growthrate of the selected seed from 30 to 70 mm in size was
240 f.Lm/ day, while commercial seed ranging from 30
to 50 mm and 50 to 70 mm in shell length grew at a
rate of only 141 and 147 f.Lm/ day, respectively.
Clearly, the daily growth in the present experiment
was larger than that reported in the past.
This result indicates that selective breeding is useful for improving the growth rate of cultured
abalone. It is difficult to obtain stable conditions inrearing experiments. Therefore, it is necessary to repeat this or a similar rearing experiment in order to
get a general conclusion.

Acknowledgments
The authors would like to thank Y Fujio and A.
Kijima for critically reading this manuscript. This
study was supported by grants to the Agriculture, Forestry, and Fisheries Research Council Secretarin.

Citations_

Department of Statistical Information (OSI)
1990. Fisheries Statistics of Japan. OSI, Ministry of Agriculture, Forestry, and Fisheries, Governmen t of Japan. (In English.)Hara, M.
1989. Effect of parents on growth in the abalone seeds,
Suisanikushu 14:39-42. (In Japanese.)
Kikuchi, S. and N. Uki
1974. Technical study on the artificial spawning of abalone,genus Haliotis. Part II: effect of irradiated sea water withultraviolet rays on inducing to spawn. Bull. Tohoku Reg.
Fish. Res. 33:79-86.
Takahashi, K. and A. Koganezawa.
1988. Mass culture of Ulvella lens as a feed for abalone,
Haliotis discus hannai. In New and innovative advances inbiology/engineering with potential for use in aquaculture:proceedings of the fourteenth U.S.-japan meeting on aquaculture (A.K. Sparks, ed.) p. 29-36. NOAA Tech. Rep.
NMFS 70.
Uki, N.
1989. Abalone seedling production and its theory (3). Int.
J. Aquat. Fish. Technol. 1:224-231.
Uki, N.,J.F. Grant and S. Kikuchi
1981. Juvenile growth of the abalone, Haliotis discus hannai,
fed certain benthic micro algal related to temperature.
Bull. Tohoku Reg. Fish. Res. Lab., 43:59-64.
Show, C.R. and R. Prasad1970 Starch gel electrophoresis of enzymes-a compilationof recipes. Biochem. Genet. 4:297-320.

Ovarian Development in the South American White Shrimp,
Penaeus vannamei

SUSAN M. RANKIN,jAMES Y BRADFIELD and LARRYL. KEELEY
Laboratories for Invertebrate Neuroendocrine Research
Department ofEntomology
Texas A&M University
College Station, TX 77843-2475

Abstract
Ovarian development is characterized in part by an accumulation of several polypeptides (protein subunits). These polypeptides become the most abundant components ofthe mature ovaries. Thus, the factors that regulate synthesis of these polypeptides aremajor factors that control egg development. In the South American white shrimp,
Penaeus vannamei, these polypeptides are synthesized by the ovaries, presumably under
the direction of one or more hormones. We are using contemporary techniques ofmolecular biology and peptide chemistry to identify the factors which inhibit or promote
synthesis of these important reproductive polypeptides. Our characterization of thesefactors will lead to the developmen t of analogs to promote· reproduction under mariculture conditions. In this paper we describe ovarian maturation in shrimp and review ourprogress in (1) promoting an understanding of the intricate biological activities that
culminate in spawning and (2) exploiting this knowledge to regulate reproduction in
intact broodstock females under mariculture conditions.

Introduction: the Problem_

Achievement of the full economic potential of theshrimp mariculture industry depends on the successful domestication of the shellfish, along with genetic
selection for desired traits such as rapid growth, orhigh tolerance for changes in temperature, salinity,or water quality. The key to domestication lies in enhanced, controlled reproduction by the broodstock
animals.
To date, efforts to domesticate shrimp have beenhampered, at least in many of the U.S. firms, by thepractice of harvesting wild stock and promoting reproduction by eyestalk ablation ( i.e., surgicalremoval of an eyestalk). This practice has several
drawbacks: 1) continual introduction of wild stock as
the basis for the next generation does not allow genetic selection and risks introduction of shrimp
predators and pathogens; 2) many eyestalk-ablated
animals fail to reproduce, which reduces total yieldand predictability of numbers of progeny; and 3) eyestalk ablation also reduces the longevity and total
fecundity of the broodstock.

Thus, a fundamental problem in the shrimp mariculture industry is the lack of predictable, abundant
supplies of offspring of known heritage. The resolution of this problem lies in a multidisciplinary,multidimensional approach that involves the cooperation of physiologists, chemists, molecularbiologists, and producers. We have an obligation
to combine our efforts to relieve this problem by developing biotechnology and by establishing rearing conditions (appropriate temperatures, light intensities, and diets) that induce reproductive
development.
Here we describe ovarian maturation in shrimp
and our progress in establishing the tools necessary
for identification of the hormones that affect reproduction. Furthermore, we describe our efforts inestablishing the basis for bioengineering these organisms to suit the needs of mariculture. We represent
several disciplines working together to solve a common problem: understanding shrimp reproductionand exploiting that understanding to improve shrimp
production.

27

28

NOAA Technical Report NMFS 106

..
..
.

"cu
Q,.
>.

CO

y = 41.998 + 341.23x + 1942.8xA 2 RA2 = 0.756

500 r------------------.
400

>

o

Q)

300

Q.

- 200
Cl

-..
E
c

100

Q)

o

£l.

or-......,.--,.-.....,...--.-...,.....-~-....--~

0.0

0.1

0.2

0.3

Oocyte diameter (mm)0.4

Figure 1
The total amount of protein in the ovaries as a function of oocyte diameter in P. vannamei. Each point
represents the measurement for a pair of ovaries
from a single broodstock female. (Rankin et al.
1989.) Diameters below the resolution of the dissecting microscope are graphed at "0.0".

Morphology: Identification of the Players _Biochemical Analysis of Ovarie"'-s_

Animals. Broodstock Penaeus vannamei, the species
most favored for commercial mariculture in the
United States, were obtained from Laguna Madre
Shrimp Farm (Los Fresnos, TX) and Sea Critters(Tavernier, FL). Most of the broodstock females were40-60 g and unilaterally eyes talk ablated to promote
reproductive development (Rankin et al. 1989).
Eyestalk ablation is presumed to promote reproductive development by removing a gonad inhibiting
hormone (see Charniaux-Cotton and Payen 1988).
The eyestalks synthesize, store, and release a numberof hormones, with one or more of these presumably
suppressing reproduction. The bluish-white sinus
gland is the neurohemal organ that stores andreleases neurohormones in the eyestalk (CharniauxCotton and Payen 1988).
The mature ovary runs the length of the abdomen,
surrounds the hepatopancreas, and extends into thehead region (Bell and Lightner 1988). During maturation, the ovaries progress from a clear, empty
appearance, to white, then usually to a creamy yellow
as they near maturity. There is some variation, however, in the color of vitellogenic ovaries; they may
appear greenish or orange instead of yellow.
The hepatopancreas is analogous to the liver of avertebrate, serving as a general center of intermediary metabolism and as a site for storage of reserves.
Compared to the mature ovary, it is a relatively discrete and small organ, comprised interiorly of a vast
number of tubules that are surrounded and held inplace by the highly pigmented sheath (Bell andLightner 1988).

Oocyte diameter increased from below the resolutionof the dissecting microscope ("0", ie., <0.01 mm) up
to about 0.3 mm. Total protein in pairs of ovaries
increased from < 5 mg to about 400 mg (Fig. 1;
Rankin et al. 1989). The protein measurements
shown in Figure 1 are the sum of a number of different kinds of proteins. Fortunately, ovarian growth ismarked by the accumulation of a few specific proteins that we designate as the major ovarian proteins.
Factors that regulate the synthesis of these particular
proteins are therefore major elements that control
ovarian development.
To identify these major ovarian proteins, we usedas our initial criteria that they should be absent fromprevitellogenic (immature) ovaries, and become increasingly prominent, to strikingly abundant innearly mature ovaries. We used sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS PAGE)
to separate polypeptides, subunits of proteins. Thisapproach is unorthodox in that procedures for identifying major ovarian protein usually include salt
precipitation and column chromatography prior to
or instead of SDS PAGE. We intentionally omitted
precipitation and chromatography to avoid potential
loss of major protein candidates. The potential for
loss of major proteins was great in Penaeus vannamei,
because solubility in some buffers was low (Rankin etal. 1989).
The results of SDS PAGE of ovaries in varying
stages of reproductive maturity are shown in Figure 2(Rankin et al. 1989). Ovaries in each stage had manydifferent polypeptides. Most of the polypeptides wereRankin et al.: Shrimp Ovarian Development

M r )(10-3

205-[

12 3
I

116-

66-

45-

weight polypeptides that we designated the major
ovarian polypeptides. Yano and Chinzei (1987) havedemonstrated that the ovary of Penaeus japonicus is asite of synthesis of ovarian proteins, whereas Tom etal. (l987b) suggest an extraovarian source inParapenaeus longirostris.
Not shown are hepatopancreas tissue samples;
those appeared to lack the high molecular weight
polypeptides. We also tested whether the hepatopancreas made and secreted these polypeptides fortransport to the ovary by similarly incubating thehepatopancreas and then monitoring the culture medium for the presence of secreted polypeptides.
Using this strategy, we could not detect synthesis of
the high molecular weight polypeptides in the medium after incubation (Fig. 2C). Thus, the hepato-

1 23

i

t

•

----'"

--

1

29-

A

B

29

c

2

3

kb
9.5.2

Figure 2

SDS-PAGE of P. vannamei ovarian and hepatopancreatic
polypeptides during the gonadotrophic cycle. (A)Coomassie blue-stained ovarian samples: Lane 1,
previtellogenic ovary; lane 2, ovary at the onset of vitellogenesis; lane 3, ovary in mid to late vitellogenesis. (B)Autoradiograph of the same samples incubated with [35 S]_
methionine. (C) Autoradiograph of [ 35 S]-labelled
polypeptides in the culture medium after incubation ofhepatopancreas from shrimp in mid-late vitellogenesis.
Bracket indicates the major polypeptides (-175-200 kD)
synthesized and'-accumulaled by ovaries during yolk formation. (Rankin et al. 1989.)
found in ovaries of each of the three stages examined. However, several polypeptides were prominent
in developed ovaries and absent from immature
ones. These polypeptides were 175-200 kDa, and met
our criteria for major ovarian polypeptides.
The relative insolubility of the major ovarianpolypeptides suggested that in P vannamei, either theovary was making the polypeptides itself, or it was
modifying them after they were taken up. We investigated the origin of these polypeptides using
autoradiography (Rankin et al. 1989). Tissues wereincubated in a culture medium with 35S-me thionine,then the protein was extracted and separated by SDSPAGE.
The autoradiographic visualization of the polypeptides synthesized by the ovary is shown in Fig. 2B. It
appears that most of the polypeptides that were
present in the ovary (Fig. 2A) could also be synthesized there. It clearly synthesized the high molecular

4.4-1

2.4-

I
1.4~

0.3Figure 3

Total RNA samples (5 f.Lg/lane)from mid-reproductive cycle P.
vannamei tissues were denatured, separated by electrophoresis in agarose, transferred
to nylon membrane, and hybridized with the cloned andlabelled 3 kb ovarian eDNA.
Lane 1, ovary; lane 2, hepatopancreas; lane 3, muscle. RNA
size markers are indicated atleft. (Bradfield et al. 1989.)

30

NOAA Technical Report NMFS 106

Construction of Expression Plasmid

Insertion sitet

B-gal promoter

PUR291~

pUR 291

B-gal code,' ."
,

"
"
,

,

,

"

B-gal promoter

,

,

I

I

I

Figure 4
I

I

II
II,
I

I

I
III

"
pUR 291 ~~~~l/\..""'---PUR 291
B-gal code Shrimp polypeptide code

1

"

,

,

2

116-

Expression vector pUR
291 was linearized nearthe 3'-terminus of the
J3-galactosidase coding
region, and ligated inphase with the 3 kb
shrimp ovarian cDNAfor production of a fusion protein.

pancreas does not appear to synthesize these polypeptides. Similarly, in P japonicus, yolk protein is not
immunoprecipitated from the hepatopancreas (Yanoand Chinzei 1987). These results do not preclude
other major contributions by the hepatopancreas toovarian development, such as lipid synthesis necessary for yolk, as has been suggested by Castille andLawrence (1989).

97Beginning Genetic Analysis of Ovaries _ _

66-

45-

Figure 5
E. coliJMlOl was transformed with plasmidpUR 291, and plasmid-encoded J3-galactosidase was induced by addition of
isopropyl-J3-0-thiogalactopyranoside. Cellextracts were separated by SOS-PAGE(7.5%) and stained with Coomassie Blue.
Lane I, native pUR 291; lane 2, recombinant pUR 291 containing the 3 kb ovarian
eDNA. The heavy band at 116 kDa in lane
I is unfused J3-galactosidase. The band at
205 kDa in lane 2 is a fusion consisting of
J3-galactosidase linked to a polypeptide encoded by the ovarian eDNA. (Bradfield etal. 1989.)

The future contribution of genetic engineering toaquaculture lies in the production of genetically altered individuals with phenotypes that are well suited
to aquacultural conditions. Realization of the benefits of genetic engineering depends on thegeneration of complementary DNA (eDNA) libraries,
identification of genes and gene products, and on
judicious use of molecular technology. We have begun this long process in P vannamei and have used
modern recombinant genetic techniques to explore
the processes of ovarian development, again, with thegoal of promoting the understanding and eventual
manipulation of those events.
Our genetic analysis began with the constructionof an ovarian eDNA library (Bradfield et al. 1989). To
accomplish this, mRNA was purified from total RNA
that was isolated from the ovaries. The mRNA wasthen used to make the eDNA library. The eDNA for
one of the major ovarian polypeptides was isolated,cloned, and used as a probe to identify tissues active
in synthesis of that polypeptide. This is a very sensitive assay for determining tissue sources of particularpolypeptides.
The tissue source and the size of this highly expressed transcript were made visible using northernhybridization. In this procedure, RNA was extracted

Rankin et al.: Shrimp Ovarian Development

31

Figure 6
Localization of the 200 kDa ovarian polypeptide using immunocytochemistry. 5 flom ovarian sections were incubated with
(Panel A) preimmune rabbit Immunoglobin G (IgG) and (Panel B) IgG from the rabbit inoculated with the gel-purified
fusion polypeptide (see Fig. 2). Immunoreaction was visualized with an alkaline phosphatase-linked second antibody. Arrows
indicate cortical granules. Scale bars = 100 flom. (Bradfield et al. 1989.)

from various tissues (muscle, hepatopancreas, andovaries) from vitellogenic females, separated on an
agarose gel, transferred to nitrocellulose (which immobilizes the RNA), then hybridized with theradioactive DNA probe representing a major ovarian
mRNA in P vannamei. The result was made visible byautoradiography.
Only the ovary had detectable levels of this major
ovarian polypeptide RNA (Fig. 3). The size of thetranscript was - 6.5 kb. This is very large, but ofcourse the major ovarian polypeptide is also verylarge. The mRNA for this polypeptide was not detected in muscle or hepatopancreas of either
previtellogenic or vitellogenic females. Thus, we had
confirmed by genetic analysis some of our previous
results from in vitro experiments.
We then made a genetically engineered pOlypeptide consisting in part of shrimp ovarian polypeptide
and in part, bacterial f3-galactosidase (f3-gal). To dothis, the 3 kb cloned eDNA representing a portion of
the 6.5 kb ovarian transcript was inserted into theplasmid pUR 291, in order to get high level expression of a fusion polypeptide consisting ofplasmid-encoded f3cgal, linked to the eDNA-encoded

product (Bradfield et al. 1989).
Figure 4 shows diagrammatically how this wasdone, illustrating the promoter region, the f3-gal
code and the insertion site. The recombinant plasmid was then inserted into E. coli. The geneticallyengineered fusion polypeptide was - 205 kDa: -115
kDa f3-gal and -90 kDa P vannamei ovarian pOlypeptide. Figure 5 shows that fusion polypeptide as it
appeared on an SDS gel. It is only by coincidence
that the fusion polypeptide was about the same sizeas the ovarian polypeptide from which it was in part
derived.
In summary, we established a eDNA expression library from shrimp ovary, and from that, isolated an
ovarian polypeptide eDNA. That gene was inserted
into E. coli, for production of the genetically engineered fusion product. The genetically engineered
protein was then injected into rabbits for productionof polyclonal antibodies.
Immunocytochemistry on 5 !Lm paraffin sections
was used to determine the cellular localization of thismajor ovarian polypeptide (Bradfield et al. 1989).
The darkly-staining regions, which indicated immunoreactivity, were located in the cortical special-

32

NOAA Technical Report NMFS 106

Figure 7
Localization of a 175 kDa ovarian polypeptide using immunocytochemistry. 5 f.Lm ovarian sections were incubatedwith IgG from the rabbit inoculated with gel-purified ovarian polypeptide. No immunoreactivity was detected incontrol samples (see Fig. 6, panel A). Immunoreaction was
visualized with an alkaline phosphatase-linked second antibody. Scale bars = 100 f.Lm.

izations (Fig. 6). Thus, the cortical specializations at
the periphery of the oocyte contained the 200 kDa
major ovarian polypeptide.
Cortical specializations, also called cortical rods or
cortical bodies, are membrane-bound organelles thatare assembled during oocyte development and become associated with the cell membrane in mature
eggs. In response to one or more stimuli (such as
contact with water at spawning, or fertilization), thecortical specializations are rapidly extruded to form alayer of material that encompasses the egg. In
shrimp, the layer is rapidly dissipated into the surrounding seawater (Clark and Lynn 1977; Clark et al.
1980). Cortical-specialization composition, function
and regulation are largely unknown. It is speculated
that they prevent polyspermy or serve as an environmental protectant.
The club-shaped cortical specializations of P.
vannamei (Figs. 6 and 7) and other shrimp(Duronslet et al. 1975; Clark and Lynn 1977; Clark et
al. 1980; Anderson et al. 1984; Tom et al. 1987b; Belland Lightner 1988;, Tan-Fermin and Pudadera 1989)

are extraordinarily prominent. Because these corticalspecializations are highly abundant and large, shrimp
provide a beautiful model system for studying regulation of synthesis of major cellular organelles.
In contrast to the 200 kDa ovarian polypeptide, a
175 kDa polypeptide, appears to be a major component of the yolk (Fig. 7). In this case, gel-purified
polypeptide, rather than genetically engineered
polypeptide was used to generate polyclonal antibodies in rabbits. Yolk polypeptides serve as food for thedeveloping embryos. This polypeptide is larger thanyolk polypeptides described for other decapods (Luiand O'Connor 1977, 1976; Zagalsky 1985; EastmanReks and Fingerman 1985; Tom et al. 1987a). In
p. longirostris, yolk polypeptides are 45 and 66 kDa,
repectively (Tom et al. 1987a).
In summary, we have demonstrated that two major
high molecular weight polypeptides in vitellogenic
shrimp ovaries are immunologically distinct, andserve quite different functions for the developing
embryos. We now have probes for these two major,
distinct ovarian polypeptides which may or may not
be regulated by the same factors. Both of thesepolypeptides are highly abundant and both are essential to egg development.

Conclusions
Now that we have identified several importantpolypeptides that accrue in the ovary during thecycle of egg development and have demonstrated
that the ovary synthesizes them, we are in a position
to pursue factors that regulate the production of
those important polypeptides. One goal is to identify
and characterize the hormones that affect reproduction and then to determine the mechanisms by which
the hormones act. We anticipate that this work, inturn, will lead to the development of stable hormone
analogs which will promote controlled, predictablereproduction by intact females of known bloodlines.
We have an obligation to bring people together tosolve our common problem-understanding reproduction-and to exploit that understanding to help
industry improve both production and profits. Thisrequires not only physiologists, but also chemists andmolecular biologists as well-all working with producers to achieve our common goal.

Acknowledgments_

This work was supported by institutional grant NA
85AA-D-SDI28 to Texas A&M University by the National Oceanic and Atmospheric Administration'sRankin et al.: Shrimp Ovarian Development

Sea Grant Program, and by the Texas Advanced Technology Research Program. The research was
conducted at the Texas Agricultural Experiment Station.

Citations
Anderson, S.L., E.S. Chang and WH. ClarkJr.
1984. Timing of postvitellogenic ovarian changes in theridgeback prawn Sicyonia ingentis (Penaeidae) determinedby ovarian biopsy. Aquaculture 42: 257-271.
Bell, T.A., and D.V. Lightner.
1988. In A handbook of normal penaeid shrimp
histology. Allen Press, Lawrence, KS.
Bradfield,J.Y., R.L. Berlin, S.M. Rankin, and L.L. Keeley.
1989. Cloned cDNA and antibody for an ovarian cortical
granule polypeptide of the shrimp, Penaeus vannamei. BioI.
Bull. (Woods Hole) 177:344-349.
Castille, F.L., and A.L. Lawrence.
1989. Relationship between maturation and biochemicalcomposition of the gonads and digestive glands of theshrimps Penaeus aztecus Ives and Penaeus setiferus (L.). J.
Crustacean BioI. 9:202-211.
Charniaux-Cotton, H., and G. Payen.
1988. Crustacean reproduction. In Endocrinology of selected invertebrate types, p. 279-303. Alan R. Liss, Inc, NY.
Clark, WH.Jr., andJ.W Lynn.
1977. A Mg++ dependent cortical reaction in the eggs ofpenaeid shrimp. J. Exp. BioI. 200:177-183.
Clark, WH.Jr.,J.W Lynn, A.I. Yudin, and H.O. Persyn.
1980. Morphology of the cortical reaction in the eggs ofPenaeus aztecus. BioI. Bull. (Woods Hole) 158:175-186.
Duronslet, M.j., A.1. Yudin, R.S. Wheeler, and WH. ClarkJr.
1975. Light and fine structural studies of natural and artificially induced egg growth of penaeid shrimp. Proc. Meet.
World Mari. Soc. 6:105-122.

33

Eastman-Reks, S.B., and M. Fingerman.
1985. In vitro synthesis of vitellin by the ovary of the fiddler
crab, Ucapugilator: J. Exp. ZooI. 233:111-116.
Liu, C.H., and O'Connor,J.D.
1976. Biosynthesis of lipovitellin by the crustacean ovary.
Part 11: Characterization of and in vitro incorporation ofamino acids into the purified subunits. J. Exp. Zool.
195:41-52.
1977. Biosynthesis of crustacean lipovitellin. Part Ill: the incorporation of labeled amino acids into the purified
lipovitellin of the crab Pachygrapsus crassipes. J. Exp. Zoo I.
199:105-108.
Rankin, S.M.,J.Y. Bradfield, and L.L. Keeley.
1989. Ovarian protein synthesis in the South American white
shrimp, Penaeus vannamei, during the reproductive
cycle. Invertebr. Reprod. Dev. 15:27-33.
Tan-Fermin,J.D., and R.A. Pudadera.
1989. Ovarian maturation stages of the wild giant tiger
prawn, Penaeus monodon Fabricius. Aquaculture 77:229242.
Tom, M., M. Goren, and M. Ovadia.
1987a. Purification and partial characterization of vitellin fromthe ovaries of Parapenaeus longirostris (Crustacea, Decapoda,
Penaeidae). Compo Biochem. Physiol. 87B:17-23.
1987b. Localization of the vitellin and its possible precursors
in various organs of Parapenaeus longirostris (Crustacea,
Decapoda, Penaeidae). Int. J. Invertebr. Reprod. and Dev.
12:1-12.
Yano, 1., and Y. Chinzei.
1987. Ovary is the site of vitellogenin synthesis in kuruma
prawn, Penaeus japonicus. Compo Biochem. Physiol.
86B:213-218.
Zagalsky, P.F.
1985. A study of the astaxanthin-lipovitellin, ovoverdin,
islolated from the ovaries of the lobster, Homarus gammarus(L.). Compo Biochem. Physiol. 80B:589-597.

Changes in Hatchery Rearing and Release Strategies Resulting
from Accelerated Maturation of Spring Chinook Salmon
by Photoperiod Control

WALDO S. ZAUGG*
National Oceanic and Atmospheric Administration
National Marine Fisheries Service
Northwest Fisheries Center
Coastal Zone and Estuarine Studies Division2725 Montlake Boulevard East, Seattle, WA 98112

JACK E. BODLE AND JACK E. MANNING

u. S. Fish and Wildlife Service
Little "White Salmon National Fish Hatchery
Cook WA 98605, USA

Abstract
Early maturation and spawning of adult spring chinook salmon (Oncorhynchus
tshawytscha) was induced by controlling photoperiod at the Little White Salmon National
Fish Hatchery. Tagged groups of progeny from these adults were released as subyearlingsin May and June, and again at their normal release time as yearlings in April of thefollowing year. Fish released as subyearlings were recovered as adults 1) in a ratio of 1:3.5
to fish released during the same years (1984 and 1985) as yearlings; 2) at a higher ratioof males to females than yearlings; 3) with reduced numbers of precocious males Uacks)
compared with yearlings; 4) predominantly as 4-year-old fish, similar to fish released as
yearlings; and 5) at a larger size at the same age than adults released as yearlings. In spiteof the lower return of adults from subyearling releases, this is a cost- effective method of
supplementing the yearling release program and provides an excellent means of augmenting the run.

Introduction_

Historically, the abundant runs of chinook salmon(Oncorhynchus tshawytscha) were used as an importantfood source and for barter among Native Americans
inhabiting the Columbia River Basin. Later, as settlers
immigrated into the basin, these fish became important to commercial harvesting and recreational
fishing and were soon overharvested. Presently,
stocks of chinook salmon are depleted in most of thebasin because of blocked and degraded habitat; artificial production in hatcheries has become essential* Present address: National Marine Fisheries Service, Cook Field
Station, Cook WA 98605, USA•

for the maintenance of harvestable numbers ofadults. Methods used in the hatcheries to rear andrelease juvenile chinook salmon vary considerably,
depending to some degree upon physical facilities
and available water supplies of the individual hatcheries, but more importantly upon the subspecies ofchinook salmon being reared.
Adult chinook salmon can be found in the Columbia River system, migrating to native waters or
rearing facilities, in nearly every month of the year.
There are, however, three principal groups: spring,summer, and fall chinook salmon, so designated tocorrespond approximately to their upstream migration and spawning times. These groups are describedon the next page:

35

36

NOAA Technical Report NMFS 106
Salmongroup

Migrationtime

SpringJan-Maychinook
Summer
Jun-Augchinook
Fall chinook Aug-Dec

Spawning
time

Length oftime in river

Jul-Sep

4-6 morths

Sep-Nov

3 months

Sep-Jan

1 monthSpring chinook salmon remain longer in fresh water before spawning than the other major fish groups
and for much of this time exist in excellent condition. They are highly prized by recreational andcommercial freshwater fishermen. However, serious
problems arise when these fish move into hatcheryholding ponds early in the spawning run and must be
held for an extended time before spawning. Such
confinement often results in high prespawning losses
due to stress, physical injury, and disease. At the
Little White Salmon National Fish Hatchery (Little
White Salmon NFH) in Washington State, measures
have been taken to reduce some of the mortality associated with long-term holding by accelerating
maturation with controlled photoperiods. Early
spawning not only reduces prespawning mortality but
allows earlier hatching of eggs. The resulting progeny experience greater growth because of longer
rearing times and manifest physiological and behavioral characteristics of developing smoIts from 10 to
11 months prior to the demonstration of such characteristics in progeny from adults spawned undernormal conditions.
This report provides information on releases andsubsequent adult recoveries of tagged subyearlingand yearling spring chinook salmon from the Little
White Salmon NFH. Some preliminary information
has been reported previously (Zaugg et al. 1986).

Materials and Methods_

The Little White Salmon NFH is located on the Little
White Salmon River (Washington) about 2 km fromits confluence with the Columbia River. Adult springchinook salmon returning to holding ponds at thehatchery by mid-May were moved into portions of
raceways covered by a metal building (9.9 X 13 m)
equipped with six sodium vapor lamps (Lucalox G.£.,
LU 150/55) located 3.1 m above the water's s·.nface.
They were then subjected to a reduced photoperiod
schedule that began with 12 hours of light per day
and decreased at a rate of 30 minutes per week until
spawning occurred about mid-July (Zaugg et al.
1986). After hatching, fry were reared inside in tanks
containing water ranging from about 7 to 9° C. In

November they were transferred to outside ponds
where water temperatures ranged from 4 to 7° C. In
late March and early April 1983-85, groups of approximately 50,000 juveniles each were tagged withcoded wire tags and held for release as subyearlingsin either Mayor June of the same spring or as yearlings in April of the following year (1984-86) as
indicated below (brood year in parenthesis):Year

TaggedReleased as
subyearlings

1983
1984
1985
1986

+ (82)+ (83)+ (84)+ (82)+ (83)+ (84)

Released as
yearlings+ (82)+ (83)+ (84)

Comparisons of adult returns between groups released in the same spring (different brood years)
could only be made for 1984 and 1985, as in 1983
only tagged subyearlings were released and in 1986
only tagged yearlings were released.
Fish were fed Oregon Moist Pellets (aMP)throughout the rearing period. Groups of tagged
subyearlings that, for six weeks prior to release, hadreceived aMP to which 7% NaCI (on a dry weightbasis) had been added, were liberated with May releases in 1984 and 1985. Thus, three groups ofsubyearlings were released in each 1984 and 1985;
two in May (one salt-fed) and one inJune.
Information on the post-release migratory performance of subyearlings released in 1983 was obtained
from captures in beach seines and mid-river purse
seines at the upriver boundary of the estuary Gones
Beach, Oregon) after fish had migrated 186 km(Dawley et al. 1985; Zaugg et al. 1986). Seining operations were not conducted after the 1983 season.
Information on adult recovery was obtained from adata base of the Pacific States Marine Fisheries Commission, Portland, Oregon. Weights of recoveredtagged adults were estimated from lengths by comparison with a length/weight relationship determined in 1989 on a group of 72 adults held at thehatchery for antibiotic injections.

Results_

Figure 1 compares the abbreviated adult holding andjuvenile rearing times that result from photoperiod
treatment of adults with times for controls that undergo normal spawning and juvenile rearing. Under
the accelerated spawning program, juveniles were released as subyearlings in both May and June. Progeny
from the early spawn began feeding earlier and, consequently, were larger on any given date thereafter(Table 1; Zaugg et al. 1986).

Zaugg et al.: Hatchery Rearing and Release Strategies for Chinook Salmon

Feb ··...···May

lun··-- Aug

Adult migmtion

At hatchery

37

NORMALACCELERATED

Figure I
Comparison of spring chinook salmon adult handling and juvenile rearing between normal and advanced photoperiod
schedules at the Little White Salmon National Fish Hatchery. Adults returning to this hatchery probably enter the ColumbiaRiver no earlier than February and are normally spawned in August. For release as yearlings, juveniles from normallyspawned adults were reared in the hatchery until their second April. For subyearlings, juveniles from treated adults that were
spawned inJuly were released as subyearlings the following May andJune.
Table 1
Historical spawning, hatching, and fry information for spring chinook salmon from the Little White Salmon National
Fish Hatchery (Zaugg et al. 1986).

Group
Con trol (1976-79)
Photoperiod (1980-82)

Spawning
dates

Mean wt (g)at first feeding

Date offirst feeding

Mean wt (g)on I October

11-17 August
16-22 July

0.32
0.27

10-28 Dec
29 Oct-12 Nov

13.8
24.4

Table 2Juvenile migration of subyearling spring chinook salmon from the Little White Salmon National Fish Hatchery (1983),
and adult recoveries.
Releasedate

Mean
wt (g)

Beach
seine

4 May 1983
24June 1983

6.7
10.3

37
0

Number caught a
Purse
seine

Migration
rate (km/day)

7
89

12
16

Adult recoveryb
Number
Percent
14
23

0.03
0.05

a Number caught adjusted for fishing effort (Dawley et al. 1985).
b Numbers adjusted to a release of 50,000; includes commercial and sport fishery catches and hatchery returns (source: Pacific States

Marine Fisheries Commission data base).

38

NOAA Technical Report NMFS 106

Table 3
Adult recoveries from juvenile salmon released from the Little White Salmon National Fish Hatchery in 1984 and 1985.
Release yearand group
1984
Yearling
Subyearling(salt fed b)
Subyearling(controls)
Subyearling
1985
Yearlings
Subyearling(salt fed~
Subyearling(controls)
Subyearling

Adult recoveries a

ReleaseDate

Wt (g)

Number

Percent

19 Apr
7 May

36.6
7.0

109
5

0.22
0.01

7 May

7.0

11

0.02

22 Jun

11.7

30

0.06

17 Apr
6 May

43.7
7.6

266
104

0.53
0.21

6 May

7.1

101

0.20

20 Jun

10.5

68

0.14

a Based on 50,000 released; data includes commercial and sport fishery catches and hatchery returns (source: Pacific States Marine

Fisheries Commission data base).
b Fed a diet containing an additional 7% (dry wt) NaCI for 6 weeks prior to release.

Table 4
Numbers of tagged female and male adult salmon returning to the Little White Salmon National Fish Hatchery fromreleases in 1984-85.
Females

Males

Female/maleYearling releases
less jacks

199
199

89
75

2.24
2.65

Subyearling releases
less jacks

158
158

119
118

1.33
1.34

During the first year of the study, juveniles releasedin June were larger, migrated faster, and produced
more adults than those released in May (Ta'::>le 2).
Fish released in June were caught exclusively in the
mid-river purse seine as they reached the lower river,whereas those released in May were captured primarily in the beach seine.
More adults were recovered from yearling thansubyearling releases made in 1984 and 1985 (Table3). Subyearlings released in June 1984 were recovered as adults in higher numbers than those releasedin May, whereas the opposite was true forsubyearlings released in 1985. The overall rate of return for all groups was much higher for fish released
in 1985 than for fish released in 1983 or 1984 (Tables
2and3).
Sex determinations on adults returning to thehatchery from tagged fish released in 1983-85 indicated a greater survival of females (Table 4). The

female to male ratio for adults returning fromsubyearling releases was nearer to 1: 1 than the sameratio for adults returning from yearling releases.
Only one precocious male (jack) was observed out of
119 males (1 %) returning from subyearling releases,
whereas 14 jacks out of 89 males (16%) returned
from yearling releases.
Examination of the age distribution of tagged
adults from both the fishery and hatchery returns
showed that fish released as either subyearlings or
yearlings were recovered predominantly as 4-yearolds (Table 5). Age-4 adults from subyearling releases were equivalent in size to age-5 adults returning to
the hatchery from yearling releases (Table 6), bothage groups having spent about 3 years in the ocean.
Table 7 presents the ratios of numbers and weights
of adults recovered from yearling releases in 1984
and 1985 to those of subyearlings released in thesame years. The generally lower total weight ratios

Zaugg et al.: Hatchery Rearing and Release Strategies for Chinook Salmon

39

Table 5
Numbers of adult salmon (% of total in parentheses) taken in the fishery and returning to the Little White Salmon
National Fish Hatchery, according to age in years.
Age
Release year and group

4

3

2

5

1983
Subyearling (May)Subyearling (June)

14 (100)
24 (89)

(4)

2

(7)

1984
35 (36)I (9)10 (35)62 (63)5 (100)
IO (91)19 (65)
173
80
79
54

(68)
(82)
(81)
(81)

69 (27)(I)

13 (5)
18 (18)19 (19)12 (18)I (0.3)14 (4)
61 (17)

235
285

(67)
(82)

104 (29)
2 (0.6)(I)

Yearling (April)
Subyearling (May, salt)
Subyearling (May, control)
Subyearling (June)

1985
Yearling (April)
Subyearling (May, salt)
Subyearling (May, control)
Subyearling (June)
TotalsYearling
SubyearlingI
Table 6
Mean fork lengths in centimeters (# of fish measured) of adult salmon returning to the Little White Salmon National
Fish Hatchery, according to age in years.
Age
Release year and group

3

2

4

5

1983
Subyearling (May)Subyearling (June)

89
87

(II)
(20)
(63)
(4)
(7)
(19)

90 (30)72 (I)
71 (10)

76
92
90
88

56 (13)73 (18)71 (17)73 (12)77 (156)
89 (65)90 (61)90 (52)89 (58)

66

(I)

61

(I)

101

(2)

1984
Yearling (April)
Subyearling (May, salt)
Subyearling (May, control)
Subyearling (June)

1985
Yearling (April)
Subyearling (May, salt)
Subyearling (May, control)
Subyearling (June)
TotalsYearling
Subyearling

46 (I)

46 (I)

56 (14)77 (209)89 (88)72 (59)89 (239)

101(compared to the total number ratios) reflects thelarger average size of recovered adults from subyearling releases (86.3 cm fork length, 7.74 kg, n = 348)
to yearling released fish (79.7 cm fork length, 5.95kg, n = 348).

Discussion~

Controlling photoperiod regimes to which adults are
exposed is an effective way of accelerating matura-(2)

tion and spawning in salmonids (MacQuarrie et al.
1978, 1979; Whitehead et al. 1978a, 1978b; Eriksson
and Lundqvist 1980; Bromage et al. 1982). Johnston
et al. (1990) have shown that it is possible to mature
Atlantic salmon (Salmo salar) at nearly any time of
the year by making appropriate photoperiod adjustments. At the Little White Salmon NFH, the use ofadvanced photoperiods on returning adult chinook
salmon accelerated maturation and spawning by 4 to
5 weeks, which has resulted in the accomplishment of

40

NOAA Technical Report NMFS 106

Table 7
Ratios of numbers and estimated weights of adults recovered from yearling releases to those of adults from subyearling releases.

Release yearand group
1984
Yearling
Subyearling(May, salt)
Subyearling(May, control)
Subyearling(June)
Subyearling average
1985Yearling
Subyearling(May. salt)
Subyearling(May. control)
Subyearling(June)
Subyearling average

Number ofrecoveredadults a

Relative numberof recovered adults(yearJing/subyearling)

Estimatedtotal wt (kg)of adults b

Relative weightof recovered adults(yearling/subyearling)

Averageadultwt (kg)

109
5

21.8

677
48

14.1

6.2
9.6

11

9.9

91

7.4

8.3

30

3.6

198

3.4

6.6

15

112

7.5

266
104

2.6

1,552
796

2.0

5.8
7.7

101

2.6

782

2.0

7.7

68

3.9

532

2.9

7.8

91

703

7.7

Totals c

Yearling (1984+ 1985)
Subyearling

375
106

3.5

2,229
816

2.7

5.9
7.7

a Based on release of 50,000; numbers include commercial and sport fishery catches and hatchery returns (Pacific States Marine

Fisheries Commission data base).
b Weights estimated from length/weight relationships determined in 1989 from 72 returning adults.
C

Averages of the three 50,OOO-fish release groups for each year are totaled and compared with the sum of values obtained from thetwo yearling release groups.

two original goals: 1) to reduce prespawning mortality by decreasing the adult holding time and 2) toproduce larger yearling smolts because of a longer
rearing time.
However, an unanticipated benefit resulted fromthe incorporation of photoperiod-accelerated spawning into the hatchery production scheme-that ofsmolt development in progeny during their firstspring of rearing. Development of smolt-like characteristics, such as increased silver coloration, finclarification, a dark band in caudal fins, crowding at
the outlet screens, increased gill Na+-K+ ATPase activities, and active seaward migration, suggested thatthese fish could possibly be released 10 to 11 months
earlier than normal and survive to adulthood (Zaugg
et al. 1986). This study was designed to test whether
such early releases of subyearlings could be used toeffectively augment current production and releaseof yearling juveniles. Adult recoveries from tagged
subyearlings released in the first year of the study(1983) were low (0.03-0.05%). However, no tagged
yearlings were released that year, so a comparison ofrelative recoveries from the two year classes was impossible. Adult recoveries from subyearling releases
made in the second year (1984) were also low (0.010.06%), but these could be compared to adultrecoveries from yearlings released at their normal
time in April of the same year (0.22%). Adult recoveries from both subyearlings and yearlings that werereleased in 1985 were much higher (0.14-0.21 and
0.53%, respectively).
Although numbers of recovered adults were small,
it is nevertheless apparent that the release ofsubyearlings resulted in a ratio of adult females tomales more closely approximating 1 (1.33) than did
the release of yearlings (2.24). In addition, very few
jacks were observed in hatchery returns from subyearling releases (1 of 119 returning males), whereas
there were 14 jacks of 89 males that returned fromyearling releases. This greater proportion of adultZaugg et al.: Hatchery Rearing and Release Strategies for Chinook Salmon

males, and the lack of jacks, results in a greater opportunity to spawn individual pairs of fully matured
adults, a practice presently being used to control disease.
Eighty-two percent of adults recovered fromsubyearling releases were age 4, whereas 67% of recoveries from yearling releases fell into this age
group. This is a majority, however, for each releasegroup. Age-4 adults from subyearling releases averaged 12 cm longer than age-4 adults from yearling
releases, and were estimated to be about 1.75 kg
heavier. This size differential, which was a characteristic of the adults in each age category, translated
into an estimated ratio of 2.7 for the combined
weight of adults recovered from yearling releases tothe combined weight of adults recovered fromsubyearling releases made in 1984-85. The ratio of
total numbers of recovered adults from these sametwo groups was 3.5.
Although the ratios of adults returning from yearling/subyearling releases varied considerably for thevarious groups released in the 2 years that comparisons could be made (1984 and 1985), the results
suggest that it would be necessary to release three tofour times as many subyearlings as yearlings to recover an equivalent number of adults. We estimatethat three to four times as many subyearlings as yearlings could be reared in the same pond space, but fora much shorter rearing time and with lower production costs. In addition, the larger size of adults
returning from subyearling releases and the reduced
number of jacks makes this strategy even more economically beneficial. The release of subyearlingprogeny of adult spring chinook salmon matured
early through the use of photoperiod control is aneffective hatchery practice at the Little White Salmon
NFH. A similar program might be used at otherhatcheries to provide an effective method of maintaining and augmenting spring chinook salmon runsin the Columbia River Basin or elsewhere.

Citations

41_

Bromage, N.R., C. Whitehead,]. Elliot, B. Broton, and A. Matty.
1982. Investigations into the importance of daylength on thephotoperiod control of reproduction in the female rainbow
trout. In Proceedings of the international symposium on
reproductive physiology of fish; 2-6 August 1982,
Wageningen, The Netherlands, (C.Jj. Richter and H.J. Th.
Goos, eds.), p. 233-237. Pudoc Press, Wageningen.
Dawley, E.M., R.D. Ledgerwood, and A.L.Jensen.
1985. Beach and purse seine sampling of juvenile salmonids
in the Columbia River estuary and ocean plume, 1977-83.
Vol. II: Data on marked fish recoveries, 397 p. NOAA
Tech. Memo. NMFS F/NWC-75. Northwest Fisheries Center, 2725 Montlake Blvd. E., Seattle, WA 98II2. NTIS PB85187722.
Eriksson, L.a., and H. Lundqvist.
1980. Photoperiod entrains ripening by its differential effectin salmon. Naturwissenschaften 67 (4) :202.
Johnston, C.E., S.R. Farmer, RW. Gray, and M. Hambrook.
1990. Reconditioning and reproductive responses of Atlantic
salmon kelts (Sa/rna safar) to photoperiod and temperature
manipulation. Can.]. Fish. Aquat. Sci. 47:701-710.
MacQuarrie, D.W.,].R. Markert, and W.E. Vanstone.
1978. Photoperiod induced off-season spawning of coho
salmon (Oncorhynchus kisutch). Ann. BioI. Anim. Biochim.
Biophys.18(4):1051-1058.
MacQuarrie, D.W., W.E. Vanstone, and].R. Markert.
1979. Photoperiod induced off-season spawning of pinksalmon (Oncorhynchus gorbuscha). Aquaculture 18(4):289302.
Whitehead, c., N.R. Bromage, B. Breton, and R. Billard.
1978a. Effects of altered photoperiod on serum gonadotropin levels and spawning in female rainbow trout. ].
Endocrinol. 79(2):29-35.
Whitehead, C., N.R. Bromage,].R.M. Forster, and Aj. Matty.
1978b. The effects of alterations in photoperiod on ovariandevelopment and spawning time in the rainbow trout (Sa/rna
gai7-dneri). Ann. BioI. Anim. Biochim. Biophys. 18(4)10351043.
Zaugg, W.S.,].E. Bodle,].E. Manning, and E. Wold.
1986. Smol t transformation and seaward migration in O-age
progeny of adult spring chinook salmon (Oncorhynchus
tshawytscha) matured early with photoperiod control. Can.
]. Fish. Aquat. Sci. 43:885-888.

Induced Spawning of Japanese Eel with LHRH-A Copolymer Pellet
KEIJI HIROSE
National Research Institute of Aquaculture
NanseiMie 516-01, Japan

Abstract
A new hormone therapy was tested for inducing maturation and ovulation in theJapanese eel Anguilla japonica. A luteinizing hormone-releasing hormone analogue(LHRH-A) copolymer pellet, which shows a slow and constant release of hormone over
75 days, was used in this study. The fish were treated with eight weekly injections ofsalmon pituitary and human chorionic gonadotropin (HCG), after which time the
LHRH-A pellets were implanted in the fish. Final maturation and ovulatiofl was inducedin the mature fish at 15-29 weeks by injection with 17a-hydroxyprogesterone (17aOHprog) and HCG; the treated fish spawned naturally for 2 days. This hormone
treatment schedule could induce a high success of spawning because it eliminates thestresses incurred by repeated injections of hormones. The best result was obtained in fish
kept at a low water temperature of 15° C.

Introduction_

Recently, eel consumption has increased, reaching 70
thousand metric tons (t) per year in Japan. However,the production of eel elvers in natural waters has
been decreasing for the past 20 years (Statistics andInformation Department, Ministry of Agriculture,
Forestry, and Fisheries 1990). Presently, there is a
shortage of seed which is barely covered by imports.
The development of artificially induced breeding
techniques of eel have long been needed in Japan.
Thus, the controlled reproduction of the Japaneseeel Anguilla japonica has been attempted for a longtime. Ten to fifteen years ago, several research groups
succeeded in inducing spawning of the eel by treatment with fish pituitary (Yamamoto et al. 1976;
Motonobu et al. 1976; Satoh 1979). Since then, there
have been almost no excellent results on the induced
spawning of eels. In those previous experiments, thespawning rate is very low, 5-10%. Final maturationand ovulation are not well controlled and the larvae
do not survive over 10-15 days. In general, catadromous type eels (silver eel A. japonica) are most widelyused for induced spawning. The initial state of theirovaries is immature just before or shortly after initiation of yolk formation. If we wish to induce ovarian
maturation and spawning in immature eels, a relatively long-term hormone therapy is necessary.

However, repeated administration of exogenous hormones results in gonadal atresia, which may becaused by stress and the production of an undesirable antibody. The repeated treatments of exogenous
hormones may be related to the failure to induce
ovulation, low spawning rates, and also problems in
egg quality. To avoid stress and undesirable resultsfrom the repeated treatments, we developed a copolymer pellet containing luteinizing hormone-releasing hormone analogue (LHRH-A), which
provides a slow and constant release of hormone overa long period of time and which succeeded in inducing final maturation and ovulation when implanted
in ayu (Plecoglossus altivelis) (Yoshida et al. 1986;
Hirose et al. 1990). In this study, the author tested
the potency of this pellet for inducing maturationand spawning in Japanese eels.

Materials and Methods_

The cultured eels used for this study were over 10
years old and were raised at the Inland station of the
National Research Institute of Aquaculture. Before
beginning the experiments, fish were transferred tothe main station of the institute and held in a 1-t
flow-through seawater aquarium with temperature
constant at 15 or 20° C.

43

44

NOAA Technical Report NMFS 106

Treatment schedule A-Weekly treatment with chumsalmon pituitary (30 mg/fish) and HCG (300 IU/fish) for the first 8 weeks. Only chum salmon pituitary was administered thereafter until ovulation was
induced artificially.

-78°C

W CO r -rayGO

5X10 5 rad)

Molecular sieve 3A

25mg

LHRH-A50)J9Treatment schedule B-Weekly treatment with chumsalmon pituitary (30 mg/fish) and HCG (300 IU/fish) for 8 weeks followed by implantation of the
LHRH-A polymer pellet. The fish were not treatedwith pituitary and HCG after implantation of the pellet until ovulation was induced.

polymer(2G: 14G)
Figure 1
Schematic diagram for preparation of LHRH-A copolymer
pellet using the methods of Yoshida et al. 1986. Luteinizing
hormone-releasing hormone analog (LHRH-A, 50 fLg) dissolved in phosphate buffer was absorbed to 25 mg ofmolecular sieve 3A. Following evaporation, molecular sieve
3A containing hormone was charged into a glass ampoule.
A hydrophobic diethylene glycol dimethacrylate (2G) andhydrophilic polyethylene glycol #600 dimethacylate (l4G)
monomer mixture was also charged into the ampoLie. The
ampoule was sealed off under nitrogen gas and irradiatedwith gamma-rays from a Co 60 source at 5 X 105 rad/hr, at_78 0 C. The co-monomer in the supercooled state was completely pOlymerized by the irradiation. The product wasthen removed from the ampoule mold.

Hormones
Luteinizing hormone-releasing hormone analogue,
des Gly'O-[D-Ala 6 J LHRH ethylamide (Sigma), humanchorionic gonadotropin, (HCG, Teikoku Hormone
MGF. Co. LTD., Tokyo) and chum salmon
Oncorhynchus keta pituitary (caught in Iwate Prefecture) were used. The schema for the preparation of adry-copolymer composite in rod form (1.6 mm in diameter and 10 mm long) is shown in Figure 1.
Detailed preparation methods of the copolymer pellet by radiation-induced polymerization are describedby Yoshida et al. (1986). In vitro experiments showedthat the copolymer pellet released LHRH at a relatively cons tan t rate over 75 days (Hirose et al. 1990).
In this study, a copolymer pellet containing 50 ILg
LHRH-A was used. The pellet was implanted intramuscularly. Chum salmon pituitary and HCG werealso given weekly to induce maturation and spawningof the eels.

Treatment schedule C-Weekly treatment with chumsalmon pituitary (30 mg/fish) and HCG (300 IU/fish) for 8 weeks followed by chronic treatment with
the LHRH-A polymer pellet, which was implanted in
the 8th week. Chum salmon pituitary was administered at a level of 30 mg/fish for the first 8 weeks: 20
mg/fish thereafter. No HCG was given after 8 weeks
until ovulation was induced.
Both experiments were conducted in the I-t tankusing groups of three female fish per treatment. Experiment I was held at 20° C and involved treatments
A and B only. Experiment II was held at 15° C andinvolved treatments A, B, and C.
Body weight was measured weekly and changed
little until final maturation was induced. A somewhat rapid increase in body weight occurs during
final maturation and ovulation due to hydration ofoocytes. Therefore, 17cx-hydroxyprogesterone (17cxOHprog 1 mg/fish) and HCG (1000 IU/fish) wereadministered to the mature eel when it showed aslight increase in body weight just prior to the rapid
increase in body weight. Ovulation and spawning
were induced within 2-3 days following the treatment
with I7cx-OHprog and HCG. For males, six or sevenweekly injections with HCG (300 IU/fish) wereenough to induce spermiation.

Results and Discussion_

At the initiation of the experiments the cultured eels
had not yet attained the primary yolk globule stage
and their gonadal somatic indexes were below 2.0.
After eight weekly injections of chum salmon pituitary and HCG, the fish ovaries had oocytes in the
primary yolk globule stage. At this time, LHRH-A
polymer pellet was implanted to the eel.

Treatment Schedules

Experiment I

Two experiments involving the following three treatment schedules were undertaken (see Fig. 2).

In the treatment schedule A group (Table 1), one(D-3) of three fish spawned after 15 of the weekly

Hirose: Induced Spawning ofJapanese Eel

45

Hormone treatment schedule
symbol

A

I'-

...8I

16

~I-

salmon pituitary 30mg

spawning..

SP~salmon pituitary 30mg
HCG

+ HCG 300 IU
LHRH-A polymer

B

i

t-

. .P- __ ---polymer1.....
salmon pituitary 30mg

+ HCG 300 IU
LHRH-A polymer

C

!

16

I-- - -----J''''''--

1-1

salmon pituitary 30mg

salmon pituitary 20mg

+ HCG 300 IU~-

17. -hydroxyprogesterone( 1mg/fish)and HCG( 1OOOIU/fish)were used for inducing
ovulation and spawning.

Figure 2
Hormone treatment schedules for inducing maturation and spawning in Japanese eel in Experiments I and II. Experiment I
was held at 20° C and involved treatments A and B only. Experiment II was held at 15° C and involved treatments A, B, and
C. HCG = Human chorionic gonadotropin; SP = Chum salmon pituitary. Time on line marked in weeks. 17ahydroxyprogesterone (1 mg/fish) and HCG (1000 IV/fish) were used to induce ovulation and spawning.

Table 1
Experimental results from experiment I which was performed at 20° C. See text for descriptions of treatment schedules.

Fish
no.
Treatment schedule A
D-I% increaseof body
weightGonadalsomatic
index

Time ofinjection(weeks)Remarks

606

8.7

33.4

16

over-mature

2

627

30.5

35.2

16

over-mature

3

656

a

19.7+

15

spawned, unfertilized

545

0

19.7

20

dead, final
maturation

2

561

24.5

16

spawned, larvae
survived 7 days

3

633

26.3

20

spawned, larvae
survived 7 days

Treatment schedule B
E-l

a -

Body
weight(g)= data not collected.

46

NOAA Technical Report NMFS 106

Table 2
Experimental results from experiment n, which was performed at 15° C. See text for description of treatment schedules.
Body
weight(g)% increaseof body
weightGonadalsomatic
index

Time ofinjection(weeks)

691

13.2

12.3

16

dead, final
maturation

2

1019

30.0

a

26

over-mature

3

1713

-7.5

20.5

31

yolf deposition

1492

13.3

37.8

24

ovulated, artificial
fertilization

2

988

33.3

29

spawned, larvae
survived 8 days

3

1250

12.9

27

spawned, larvae
survived 12 days

22

dead, mature

Fish
no.
Treatment schedule A
F-lTreatment schedule B
G-I

Treatment schedule C
H-I

a -

1600

12.8

25.5

2

1400

11.4

34.5

3

1579

16.0

Remarks

18

dead, larvae

27

spawned, larvae
survived 8 days

= data not collected.

injections, although the eggs were not fertilized. The
others quickly developed to an overly-mature stateand could not spawn. In the treatment schedule B
group, two (E-2 and 3) of three fish spawned naturally and the resulting larvae survived for 1 week.
Female fish developed to a mature or spawning state
after 16-20 weeks of injections.

Experiment II
In the treatment C group (Table 2), one fish (H-3)
spawned at 27 weeks, while the other fish died when
they became completely mature. In the schedule B
group, two of the three fish spawned naturally; theother one ovulated earlier at 24 weeks at which time
artificial fertilization was performed. Some of theschedule B larvae survived for 8-12 days but they
could not ingest any live organisms (rotifers or fertilized oyster eggs). Treatment schedule A fish could
not spawn. The lower temperature used for experiment II (15° C) lengthened the duration of theweekly injections to 24-29 weeks.
Based on the results from two experiments, treatment schedule B performed at 15° C is the best
method for inducing maturation in tht> Japanese eel.

A combination of 17o.-0Hprog and HCG was given to
the mature fish to induce final maturation and ovulation and, thereafter, they were expected to spawn
naturally within 48 hours. The processes of final
maturation and ovulation proceeded very quickly at awater temperature of 20° C. At that temperature thefish showed a very rapid increase in body weight andhigh gonadal somatic index, changing into an overly
mature state.
Figure 3 shows the frequency of egg diameter frommature eels kept at 20° C (treatment schedule C in apreliminary experiment). The most developed 00cytes, which were in the migratory nucleus stage andcontained many small oil droplets, were 0.7-0.8 mm
in diameter. Following ovulation, the oocytes quickly
changed into an overly mature state, as seen by thepresence of eggs that now contained only a few large
oil droplets and were not fertilized (Hirose, unpubli.
data). Moreover, ovulation was not always precededby final oocyte maturation when the mature fish were
kept in water temperatures over 20° C. Thus, hightemperature conditions may cause the production ofpoor quality eggs in a very short time. Mature fish
should therefore be kept in somewhat cool water(15° C). If gravid eels are cultured under the lower
temperature condition, body weight does not in-Hirose: Induced Spawning ofJapanese Eel

47

Dec. 22 , 1988

symbol17Q(-OHprog

15

+ HCG
W.T.20·C

105Dec. 24 , 1988

>- 10
Co)
C

G»
:::I

.r:r
G»

5

I&.

1.2 (mm)Egg diameter
Figure 3

Frequency distribution of oocytes in the mature eel treated with 17a-hydroxyprogesterone (l7a-OHprog) and HCG at
20° C.(Hirose, un pub!. data). Fish were observed to mature using treatment schedule C in a preliminary experiment. Oocyte
samples were obtained using a polyethylene cannula inserted into the urogenital pores of the eels. Thereafter, the oocytes
were observed under a microscope. The mature oocytes (0.7-0.8 mm in diameter) had germinal vesicles (GV) and manysmall oil droplets. After hormone treatment, the oocytes rapidly changed into an overly mature state (shaded portion) in 2
days; germinal vesicle breakdown (dotted circle) and a few large oil droplets were also present-a typical feature in overly
mature eggs.
crease as rapidly during final maturation, as shown inprevious reports (Sugimoto et aI. 1976; Yamauchi
and Yamamoto 1982). Consequently, a final injectionof HCG and 17a-OHprog for inducing ovulation istimely when given to mature fish, and a high frequency of natural spawning can be anticipated.
In general, it is said that the stress of handlingduring hormone injection decreases the efficiency ofhormone treatment and may cause rapid atresia of
the gonads (Hirose 1980; Billard et aI. 1981). This
LHRH-A polymer pellet has a long hormone delivery
time and is especially useful for inducing maturationin fish requiring a series of hormone treatments overa long time period. The ability to induce a high percentage of treated eels to spawn is of great impor-tance. In this study, a high frequency of maturationand spawning was induced using hormone treatment
schedule B in experiment II (15° C) followed by an
injection of 17a-OHprog and HCG. If we can increase the spawning frequency in the hormonetreated fish, the artificial propagation of eels will beachieved in the future.

Acknowledgment
I am deeply grateful to Dr. M. Yoshida of Japan
Atomic Energy Research Institute for kindly supplying the LHRH-A copolymer pellet.

48

NOAA Technical Report NMFS 106

Citations_Billard, R., C. Bry, and C. Billet.
1981. Stress, environment, and reproduction in teleost fish.
(A.D. Pickering, ed.), p.185-208. Acad. Press, NY.
Hirose, K.
1980. Effects of repeated injections of human chorionic gonadotropin (HCG) on ovulation and egg qualities in theayu, Plecoglossus altivelis. Nippon Suisan Gakkaishi 46:813818.
Hirose, K., H. Kagawa, M. Yoshida, M. Kumakura, and H.
Yamanaka.
1990. Application of LHRH copolymer pellet for inductionof final oocyte maturation and ovulation in ayu Plecoglossus
altivelis.
ippon Suisan Gakkaishi 56: 1731-1734.
Motonobu, T., K. Yamashita, and H. Oka.
1976. Memoir on the hatched larva of the Japanese eel,Anguilla japonica, from the adults matured artificially. Bull.
Shizuoka Pref. Fish. Exp. Station 10:87-90. (In Japanese.)
Satoh,H.
1979. Towards the complete cultivation of Japanese eel. Iden
(Genetics) 33:23-30 (in Japanese).
Statistics and Information Department, Ministry of Agriculture,
Forestry, and Fisheries.
1990. Gyogyo-yoshokugyo-seisan-LOukei-nenpou. Tentative
English title: Fisheries statistics ofJapan, p.181 and 268. (InJapanese.)

Sugimoto, Y., Y. Takeuchi, K. Yamauchi, and H. Takahashi.
1976. Induced maturation of female Japanese eels (Anguillajaponica) by administration of salmon pituitaries, with noteson changes of oil droplets in eggs of matured eels. Bull.
Fac. Fish. Hokkaido Univ. 27:107-120.
Yamamoto, K., M. Nakamura, H. Takahashi, and K. Takano.
1976. Cultivation of larvae ofJapanese eel. ature 263:412.
Yamauchi, K. and K. Yamamoto.
1982. Experiments on artificial maturation and fertilizationof the Japanese eel (Anguilla japonica). In Proceedings ofthe international symposium on reproductive physiology
of fish (C.JJ. Richter and H.J. T. Goos, eds.), p. 185189. Center for Agricultural Publishing and Documentation, Wageningen, the Netherlands.
Yoshida, M., M. Asano, I. Kaetu, K. Imai, H. Yuasa, H. Nakayama,
K. Shida, K. Suzuki, K. Wakabayashi, and I. Yamazaki.
1986. Pharmacological response in male rats with controlled
release formulations of luteinizing hormone-releasing hormone agonist. Polymer Jour. 18:287-296.

Reproductive Physiology of Sablefish (Anoplopoma fimbria) with
Particular Reference to Induced Spawning
IGOR I. SOlAR, EDWARD M. DONALDSON, IAN J. BAKER,HELEN M. DYE, ALEXANDRA VON DER MEDEN, and JACK SMITH
Department ofFisheries and Oceans
Biological Sciences BranchWest Vancouver Laboratory
4160 Marine Drive
West Vancouver, B.G. V7V 1N6 Canada

Abstract

This paper summarizes information on the reproductive physiology of sable fish
(Anoplopoma fimbria) obtained during five years of studies on the induced ovulation of

captive female sablefish. The ability to induce ovulation and spawning in captive
broodstock is considered critical for the development, expansion, and diversification of
aquacultural operations on the Pacific coast of Canada into indigenous marine species.
Results are discussed on the use of partially purified salmon gonadotropin and several
mammalian luteinizing-hormone releasing hormone analogues (LHRHa); the ovarian
hydration response; the effect of administering LHRHa by different routes on plasma
estradiol levels; and the use of a dopamine antagonist. Other preliminary and unpublished data are also presented.

Introduction_

Sablefish (Anoplopoma fimbria), also known as
blackcod (Fig. 1), range along the North American
coast from the northeastern Bering Sea to Baja California. Its depth distribution ranges from thecontinental shelf to about 1500 m (McFarlane andBeamish 1983). Spawning, which takes place at
depths of 300-700 m, lasts from January to Marchand peaks in February (Mason et al. 1983, Fujiwara
and Hankin 1988).
The sablefish is considered a prime candidate forcommercial aquaculture because of its adaptability toconfinement and its indiscriminate feeding behavior.
The development of methods for spawning sablefish
in captivity and for rearing the larvae are expected tofacilitate the future establishment of mariculture operations for this species on the west coast of Canada.
Early studies on the culture of sablefish were dependent on the impoundment of wild juveniles, andstudies of larval rearing have relied upon the captureof wild maturing broodstock and the stripping of
their gametes at sea (McFarlane and Nagata 1988).
Neither of these practices would be feasible or desirable as a basis for commercial culture.

The Pacific Biological Station (PBS), Nanaimo,
B.C., initiated studies on sablefish mariculture in theearly 1970's (Kennedy 1972). These studies have continued using a multidisciplinary approach both at the
PBS and at the West Vancouver Laboratory since1985. The present review summarizes our findings todate in the area of induced spawning.

Maturation of Captive Fish_

Sablefish do not usually undergo final maturation in
captivity. Since 1985, only 1 of over 150 females ripened spontaneously (fish in stock group, without
treatment) at the Pacific Biological Station. Althoughsablefish are known to be yearly spawners in the wild,
only one individual maintained in captivity from one
spawning season to the next responded to hormone
therapy a second time. The inability of females toundergo ovarian maturation in captivity has been attributed to enviromental and nutritional factors.
Males, on the other hand, frequently mature andspermiate without endocrine treatment.
Hormonal treatments applied to maturing captive
fish in our laboratories from 1985 to 1989 resulted in

49

50

NOAA Technical Report NMFS 106

Figure 1

Adult female sablefish. Anoplopomafimbria.

an ovulatory response in 59.4% of the experimental
animals while no fish in the control groups ovulated.

Hormonal treatments_

Several compounds have been assayed for their ability to induce spawning in sablefish. Our study
conducted in 1985 (Solar et al. 1987) demonstrated
for the first time the feasibility of inducing ovarian
hydration and ovulation in sablefish (65% ovulation
response). In this first study, we utilized a singlt intraperitoneal (ip) injection of 1.0 mg/kg body weight(bw) of partially purified salmon gonadotropin (SGGlOO) or 0.2 mg/kg bw of GlylO [D-Ala 6 ) LH-RH
ethylamide luteinizing hormone releasing hormone
analogue (LHRHa), a potent synthetic analogue ofmammalian LHRH. Limited success (19.0% ovulation. response) was achieved the following year using
two doses of SG-GI00 (1.0 and 0.5 mg/kg bw), SGGI00 in combination with LHRHa (0.5 ± 0.2 mg/kgbw), or LHRHa as a primer followed by a second
dose (0.1,0.2 mg/kg bw) three days later. Autopsy ofseveral unresponsive females at the end of the latter
experiment revealed that most had immature ovaries.
In 1987, the effect of desGlylO [D-Ala 6 ) LH-RH
ethylamide alone or in combination with domperidone, a dopamine antagonist, was investigated(Solar et al. 1988). Results suggested that dopaminergic control of gonadotropin release in this species isless apparent than in certain other teleosts (Cyprinids, Peter et al. 1986; catfish, van Oordt and Goos
1987). A second experiment in the same year investigated the effect of injections of D-Ala6 LH-RH or
microencapsulated D-Trp 6 LHRHa (Solar et al.
1988). Prior to the experimental work conducted in
1987, fish sex was tentatively identified by measuring
plasma estradiol-1713· levels. These preliminary determinations were later confirmed by ovariancatheterization.
A preliminary study conducted in 1988 investigated the influence of certain environmental
parameters (water temperature and light) on the induction of ovulation. Maintenance of the fish inflowing chilled seawater (3-4° C) and darkness forabout three months prior to hormonal treatment did

Solar et al: Reproductive Physiology of SableHsh

E
S

25
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A
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20

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51

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DAYS
-G-. LHRH INJECTION

%

25

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E

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I
G
H

15

T
I
N
C
R
E

A
S
E

-*- TREATED (N-13)-e- LHRH PELLET-B- CONTROL (N-7)

Figure 2

Plasma estradiol levels in female sablefish in response to intraperitoneal injection or cholesterol pellet implantation of des-GlylO[D-A1a6 ]LHRH ethylamide. (Solar et aI., unpub!. data 1989.)'H'V' CONTROL

I

10

5

O ! B = - - L . - - - - ' ' ' ' - - - - " - - - - L - -_ _L--_---l'-----_---l

28

24

20

16

12

8

o

4

DAYS PRIOR TO OVULATION

%

(13)

~

100

80
(9)

0
V

U
L

A
T
I

0
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60
Figure 3
40

20

O

0

t

5

10

15

20

DAYS POST-INJECTION

2530(A) Mean percent body weight changes in female sablefish treated with desGlylO [D-A1a 6 ]LH-RH elhylamide to induce ovulation, relative
to body weight at the time of injection. (B) Cumulative percent ovulation response insable fish shown in A. Numbers in brackets indicate actual number of fish ovulated andspawned to the date. Arrow indicates the timing of treatmen t. (Solar et aI., unpub!. data
1989.)

52

NOAA Technical Report NMFS 106

DAYS
30

*

* *
---- "'-- ...
z

20

"-

.. ------ ...............

*. _- ........ -....... -~a...

...............

------------ *

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---

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"

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1.00

1.05

1.10

1.15

1.20

1.25

INITIAL EGG SIZE

not improve the ovulation response relative to fish
held on a natural photoperiod at 9-10° C (I. Solar eta!. un pub!. data 1988).
In 1989, changes in plasma estradiol levels during
the latency period (time span between drug administration and ovulation) were investigated following
treatment with D-Ala 6 LHRHa by ip injection andslow-release cholesterol pellet implantation. Both injection and implantation of the 5% D-Ala6LHRHa
pellet at the same dose (0.1 mg/Kg bw) induced ovulation in 100% of the experimental animals (n = 13).
while none of the control fish (n = 7) ovulated.
Changes in plasma estradiol levels measured by radioimmunoassay were followed at 4-day intervals
during the 24 days postinjection. In both treatments
there was an increase of the estrogen level whichpeaked at day 4 and decreased to levels similar tocontrols on day 8. The increase in plasma estradiol
was more pronounced in the injected group than inthose that had received the slow-release pellet (Fig.
2) (Solar et a!. un pub!. data 1989). In a second experiment, during the same year, the oral
administration of 1.5 mg/kg bw D-Ala6 LH-RHa in
two doses (1.0 and 0.5 mg/kg bw, respectively, 11days apart) resulted in 75% of the fish ovulating
within 18 days from the first treatment. The timecourse of uptake and net plasma presence of LHRHa
during an 8-hour period following delivery wa also
determined thereby providing the first evidence for

1.30

1.35

1.40

Figure 4
Relationship between egg diameter (mm) at the time of
injection and the duration of
the latency period (days to ovulation) in hormonally induced
female sablefish (r = 0.8). Dotted lines represent 95% confidence limits for mean predictedvalues. (Solar et aI., unpubl.
data 1989.)

biologically active nonapeptide absorption by the fishgut (Solar et a!. 1990).

Ovarian Hydration and Latency Period _ _The female fish responding to hormonal treatment
have typically shown protrusion of the uro-genital papilla, gradual distention of the abdominal region,
and an increase in total body weight of about 25% of
the weight at the time of injection. The increase inbody weight is normally slow during the days immediately following D-Ala 6 LH-RHa injection and very
rapid during the three days prior to ovulation (Fig.
3). The increase in weight is due to oocyte hydration.
The eggs, which at the time of injection are small andopaque (mean diameter 1.25 ± 0.1 mm), became
larger and transparent (2.2 ± 0.1 mm) at the time ofovulation (Solar et a!. unpub!. data 1988).
Latency period has been variable ranging from six
days to close to a month (mean 15.8 ± 8 days). Statistical analysis showed that egg diameter at the time of
injection and the number of days to spawn (latency
period) are negatively correlated (r = 0.8) (Fig. 4)(Solar et a!. unpub!. data 1989).

Artificial Fertilization and Incubation_

The fecundity of female sablefish ranges from about
60,000 to close to one million eggs, depending on

Solar et al: ReprOductive Physiology of Sablef"lSh

fish size (Mason et a!. 1983). The average female produces close to 200,000 eggs (G.A. McFarlane and
M.W. Saunders, PBS unpub!. data). Success in fertilizing the eggs obtained from induced sablefish has
been variable and generally lower than the fertilization rate of gametes obtained from mature sablefish
collected from the wild during the peak spawningseason. During the 1989 season, the best fertilization
rates (those which were also followed by normal development to the 8-cell stage) were 62.4 ± 7.1 % for
eggs from captive induced females and 52.6 ± 3.7%
for those from wild non induced female. Average fertilization rates, however, were 25.0 ± 17.6% (wild)and 14.5 ± 17.4% (induced) O. Jensen, PBS, pers.
commun.June 1989).
Sablefish eggs are very fragile and easily injured
during culture and therefore require special incubation techniques (Alderdice et a!. 1988a, 1988b). The
mature fertilized eggs are clear and normally remainsuspended in the water column. The incubation period from fertilization to hatch lasts about 12 days at
6° C (McFarlane and Nagata 1988).

Acknowledgments
Thanks are due to Dr. R.J. Beamish and Mr. G.A.
McFarlane for making the fish used in these studies
available to us and for continued support. We also
thank J.OT. Jensen and E. Groot for allowing us toquote unpublished fertilization and incubation data.

Citations
Alderdice D.F.,J.0.T.Jensen, and F.PJ. Velsen.
1988a. Preliminary trials on incubation of sablefish eggs
(Anoplopoma fimbria). Aquaculture 69:271-290.
1988b. Incubation of sablefish (Anoplopoma fimlma) in a system designed for culture of fragile teleost eggs.
Aquaculture 71:271-283.

53

Fujiwara, S., and D.G. Hankin.
1988. Sex ratio, spawning period, size and age at maturity ofsablefish (Anoplopoma fimbria) off northern California.
Nippon Suisan Gakkaishi 54:1333-1338.
Kennedy, W.A.
1972. Preliminary study of sablefish culture, a potential new
industry. J. Fish. Res. Board Can. 29:207-210.
McFarlane, G.A. and RJ. Beamish.
1983. Biology of adult sabJefish (Anoplopoma fimbria) in waters off western Canada. In Proceedings of the International Sablefish Symposium. Alaska Sea Grant Rep. 838:59-80.
McFarlane, G.A., and W.O. Nagata.
1988. Overview of sablefish mariculture and its potential forindustry. In Proceedings of the 4th Alaska Aquae. Conf.;
18-21 Nov., Sitka, Alaska (S. Keller, ed.), p. 105-120.
Alaska Sea Grant Report 88-4.
Mason,J.C., R.J. Beamish, and G.A. McFarlane.
1983. Sexual maturity, fecundity, spawning, and early life history of sablefish (Anoplopoma fimbria) off the Pacific coast of
Canada. Can.J. Fish. Aquat. Sci. 40:2126-2134.
Peter, R.E., J.P. Chang, C.S. Nahorniak, R.J. Omeljaniuk,
M. Sokolowska, S.H. Shish, and R. Billard.
1986. Interactions of catecholamines and GnRH in regulation of gonadotropin secretion in teleost fish. Recent Prog.
Horm. Res. 42:513-548.
Solar, 1.1., 1.J. Baker, and E.M. Donaldson.
1987. Effect of salmon gonatropin and a gonadotropin releasing hormone analogue on ovarian hydration and ovulation in captive sablefish (Anoplopomafimbria). Aquaculture
62:319-325.
Solar, LT., Ij. Baker, and E.M. Donaldson.
1988. Induced ovulation in sablefish (Anoplopomafimbria) using gonadotropin releasing hormone analogues. In Proceedings of the 3rd. lnt. Symp. on Reprod. Physiol. of Fish;
2-7 Aug. 1987, D.R.Idler, L.w. Crim and J.M. Walsh (editors) St. John'S, Newfoundland. Mem. Univ., Newfoundland.
Solar T.L, E. McLean, 1.J. Baker, N.N. Sherwood and E. M.
Donaldson.
1990. Induced ovulation of sablefish (Anoplopomafimbria) following oral administration of desGly'O(D-Ala 6 ) LH-RH
ethylamide. Fish Physiol. Biochem. 8:497-499.
van Oordt, P.G.Wj., and Hj.Th. Goos.
1987. The African catfish, Clarias gariepinus, a model forthe study of reproductive endocrinology in te!eosts.
Aquaculture 63:15-26.

Geoduck Culture in Washington State:
Reproductive Development and Spawning
J. HAROLD BEATTIE and C. LYNN GOODWIN
Washington State Department ojFisheries
Point Whitney Laboratory
Brinnon, WA 98320

Abstract
The Washington State Department of Fisheries operates and maintains a hatchery forthe geoduck clam (Panopea abrupta). The purposes of this hatchery are to provide clamseed to plant subtidal areas which commercial harvesters have depleted, and to eventually double the present annual harvest from 2,000 to 4,000 metric tons. Hatcheryoperations occur from March through July, centering on the peak of natural spawning inlate spring. We induce spawning by elevating seawater temperatures to 15° C and increasing algal concentrations. In our hatchery, females release an average of 2 to 10 million
eggs per spawning.

Introduction_

The geoduck, Panopea abrupta, is the largest burrowing clam in the world. These clams live about a meterdeep in the sedimen t and have long siphons which
they can extend up their burrows for feeding, respiration, and excretion (Fig. I). Extensive populationsof geoducks occur intertidally and subtidally in Washington State throughout Puget Sound and Hood
Canal. The average weight for adults taken from
Washington waters is about 0.8 kg; however, records
show specimens in excess of 5 kg (Anderson 1971).
The Washington State Department of Fisheries(WDF) has established a maximum sustainable yield
estimate for geoducks in Washington State at 2,000
metric tons (t) per year. Geoducks are very long
lived; Shaul and Goodwin (1982) recorded ages ofadults in excess of 100 years. Buried beyond thereach of natural predators, having no known diseases, and exhibiting low recruitment, geoducks tend
to form stable, uniformly sized populations.
The commercial harvest of subtidal geoducks in
Washington State began in 1970. Prior to that year,
state regulations permitted only personal use of aharvest from hand digging. During the late 1960's,
surveys by the WDF documented extensive subtidal
beds of geoducks in Puget Sound. In 1970, the Washington State legislature changed the regulations toallow commercial subtidal harvest of these clams.
This act not only created the commercial fishery forthis resource but designated the WDF and the Washington State Department of Natural Resources(DNR) as cooperative managers. The state sells theright to harvest subtidal tracts of geoducks to companies whose divers excavate the clams with water jets.
A portion of the proceeds from these lease fees areused to fund compliance, management, patrol, andenhancement efforts by the DNR and the WDF.
In 1972, the WDF began conducting subtidal surveys of harvested tracts. By 1975, the results of thesesurveys showed that, although in many areas recruitment was sufficient to maintain healthy populationlevels, in some areas there was little or no recruitment. Thus, in 1982, the DNR and WDF decided to
implement an enhancement program which would
replenish poorly recruiting areas.
Central to this enhancement eff6ft was the establishment of a hatchery and nursery program. Using
data from Point Whitney Laboratory research(Goodwin 1973, Goodwin et al. 1979), this program
adapted shellfish hatchery and nursery techniquesdeveloped for oysters and hard shell clams. Moving
through experimental, pilot, and commercial scale
phases, the enhancement project targeted a goal of
doubling the annual harvest of geoducks from 2,000
to 4,000 t.

55

56

NOAA Technical Report NMFS 106

Reproductive Development

I:;

/j

..

~'~"

.. . .

//

;1

~>.

l
./

Figure 1
A cut away drawing of a geoduck in its
natural habitat. Shell height is 16 em.
(From King 1986.)_

Conducting a successful hatchery program depends
on understanding the physiology and timing of reproduction. Like most other mollusks, geoducks haveno external indicators of gender. They are dioeciousand sexually intransmutable; hermaphroditic individuals are rare. The gonadal ducts are intertwinedwith the intestinal mass, and together they form twolarge spheroids which lie on each side of a medial
line (Fig. 2).
When they are approximately 3 years old, malegeoducks become sexually mature; female geoducks
begin egg production when they are about 4 years
old (Anderson 1971). When mature, the gonadal tissues begin a cycle of development and maturation
which leads to spawning and to tissue resorption or
both. They continue this cycle for most of their lives.
Sloan and Robinson (1984) found reproductively active specimens that were 105 years of age. Anderson(1971) and Goodwin (1976) found ripe geoducks inat least eight months of the year from November
through June.

Anterior
Right Valve~~~~

Gonad ----....,.'+1-+Foot-Hfo---",..

MantlectenidiaFigure 2
The gross morphology and location of the gonads in gcoducks. Shell length is 12 em. Drawing by Don Velasquez, Univ. Washington.

Beattie and Goodwin: Geoduck Culture in Washington State

57

Figure 3
Histological section showing theearly active phase of male gonadwith only a few pockets of spermshowing. (Scale bar = 100 f.Lm;
stain = Harris' hematoxylin andeosin.)

Figure 4
Histological section of a ripe
male, follicles packed with
sperm. (Scale bar = 100 f.Lm;
stain = Harris' hematoxylin andeosin.)

Histology: Spermatogenesis

Histology: Oogenesis

In. the early stage (September through December),the reproductive follicles are small and possess only
pockets of sperm (Fig. 3). When the animals reachmaturation (November through June), the follicles
are packed with sperm (Fig. 4). Once spawning commences (December through July), the follicles
partially empty (Fig. 5); and when spent (Marchthrough August), the follicles partially collapse andconnective tissue proliferates in the gonads.

In the early stages (September through November), ovacytes are just beginning to form in theovarian follicles (Fig. 6). During the late active stage(September through January), ovacytes begin to fill
the follicles, clinging to the ovarian walls by a nutritive stalk (Fig. 7). When females are ripe (November
through June), the eggs detach and float free in thelumen (Fig. 8). By the partially spent stage Qanuary
through July), fewer eggs exist and there are very few,

58

NOAA Technical Report NMFS 106

Figure 5
Histological section of a partiallyspawned male. (Scale bar = 100
fLm; stain = Harris' hematoxylin
and eosin.)

Figure 6
Histological section of a female
geoduck gonad. A few ovacytes
are just showing. (Scale bar =100 fLm; stain = Harris' hematoxylin and eosin.)

if any, developing ova. Figure 9 illustrates a nearlyspent female in which the ovarian follicles are empty
and leucocytes may be presen t (May through August).

Reproductive Cycle_

Male geoducks begin gonadal development during
September. By November some are ripe. Thoughsome spawn earlier, they start mass spawning inMarch as indicated by the first occurrence of spent

animals. By August, they are entirely spawned andthe spawning cycle begins anew (Fig. 10). Femaleswith ripe gonads occur from fall until early summer;
however, mass spawning only begins in April (again
as evidenced by the occurrence of spent animals),
and is completed by August (Fig. 11). The temporalwindow for spawning in the animals examined byAnderson (1971) began from the time of first spawning in January and continued until August and thatthe m~ority of spawning occurred in May and June.
In obtaining and spawning geoducks for our hatchery work, we have found that animals from southBeattie and Goodwin: Geoduck Culture in Washington State

59

Figure 7
Histological section of the lateactive stage of female geoduck.
(Scale bar = 100 fJ-m; stain = Harris' hematoxylin and eosin.)

Figure 8
Histological section from a ripe
female geoduck gonad. Eggs arefloating free in the lumen.
(Scale bar = 100 fJ-m; stain =Harris' hematoxylin and eosin.)

Puget Sound ripen two to three weeks earlier thanthose from north Puget Sound. We attribute this phenomenon to the warmer waters and high productivity
characteristic of south Puget Sound.

Spawning in the Hatchery
To obtain mass spawnings earlier than they occur innature (March and April), we accelerated gonadal
development in February. We ripened thebroodstock in five tanks (1.2 X 2.2 X 0.6 m). Each ofthese tanks can hold 160 geoducks. The conditions in
the tank were the following: flow-through ambient(l0° C), unfiltered sea water and a constant densityof about 5,000 cells/mL of cultured diatoms(Chaetoceros mulleri). After two to four weeks in these
conditions, geoducks will usually spawn. The males
are slightly precocious: even though a 1: 1 proportion
exists between the sexes among the geoducks, we often found that a disproportionate number of males
spawned in the early season. In March 1990, for example, of the 684 animals under culture 302 males
spawned, but only 178 females released eggs. By

60

NOAA Technical Report NMFS 106

Figure 9

Histological section from anearly spent female geoduck. A
few residential ova are evident.
(Scale bar = 100 jJ.m; stain =Harris' hematoxylin and eosin.)SPAWNING CYCLE· MALE GEODUCK

100
90

80
70

60
%

50
40

30
20

Figure 10

10

o
sep

WJ spent

oct

nov

dec

II1II partiallyspawnedjanfeb

II1II ripe

mar

apr

may

jun

july

aug

o late active • early active

April, naturally occurring gonadogenesis precludes
the need for a ripening period. During this time, we
could bring animals directly to the hatchery for
spawning, and males and females would spawn in approximately equal numbers. We generally attemptedto spawn a group of at least 100 animals at a time. On
average, only 20 to 50 percent of the females
spawned well. Each female will spawn from 2 to 10
million eggs. Thus, an average spawn will result in 60
million eggs.

Area graph depicting the annual
reproductive cycle of male geoducks as represented by thepercentage of animals in eachgonadogenic stage at the time ofsampling. The preponderance of
spawning in nature occurs between the first and last occurrences of spent animals. (Datafrom Anderson 1971.)

To induce geoducks to spawn, we stopped the water flow and increased algal density in the spawning
tank to a range of 100,000 to 200,000 cells/mL. Sincealgae in culture live at 18 to 20° C, this influx of algae
causes a rise in water temperature in the spawning
tank. Mter the temperature reaches 15° C, we begin aflow of 14-15° C seawater.
When spawning commenced, we determined thesex of each animal by observing its spawn: In aflashlight beam, sperm give the appearance of milkBeattie and Goodwin: Geoduck Culture in Washington State61

SPAWNING CYCLE - FEMALE GEODUCK

100
90
80

70
60
0/0

50

40
30
20
10

o
sep

oct

nov

dec

jan

feb

mar

apr

may

jun

jUly

aug

r-----------------'---------'-----=----'-----'~:....--=_=:

I i [Ej spentl

III partially

III ripe

0 late active•

early active

spawned_mixed with water, whereas the eggs look like dust
suspended in the air. We removed the spawning animals from the tank and placed them into several
spawning trays (30 X 46 X 15 cm) filled with 1415° C seawater keeping males separate from females.
Once they had completely spawned, we removed
the females and fertilized the ova with a few milliliters of sperm suspension from the male spawningtray(s). For cleaning the fertilized eggs we used two
screens. Each of these screens is constructed bystretching and attaching precision woven nylon fabric(Nytex®) across one end of a 38-cm diameter PVC
tube. Using 14° C seawater, we gently washed the zygotes through a 130-micron screen onto a 35-micron
screen, thus removing larger and smaller particles
and excess sperm. One of the problems we encountered with using high densities of phytoplankton to
induce spawning was that, while spawning, the well
fed geoducks produced excessive amounts of feces
and pseudofeces. This material mixed with the eggs,
and made washing through the 35-micron screens a
tedious process.
After they were washed, we placed the fertilized
eggs into a 20-liter container of 15° C seawater. To
monitor development, we microscopically examinedsamples for polar bodies and first cleavage. When wewere assured that normal development was occurring, we placed the zygotes in a 20,000 L tank of
14° C seawater.
At 14° C, embryonic development was completed
in 72 hours and ended at the first veliger form. The
veliger larval period lasts 25 days at 17° C. After they
had metamorphosed, we moved the geoduck juve-

Figure 11
Area graph depicting the annual
reproductive cycle of female geoducks as represented by thepercentage of animals in eachgon:ldogenic stage at the time ofsampling. The preponderance of
spawning in nature occurs between the first and last occurrences of spent animals. (Datafrom Anderson 1971.)

niles to a sand substrate nursery. Over the next 90
days, they grew to seed size: an average of 9 mm shellheight. During the past four years, the WDF geoduck
hatchery has produced and planted 18 million seed.
In a good growing area, geoducks will grow to a 0.8kg mean harvest weight in five to six years. Thus, withan expected field survival of between 0.5 and 1.0%,
we anticipate that we will have enhanced the resource by between 70 and 140 t by the time theseanimals reach harvest size.

Acknowled~ents--------------------The authors thank Alex Bradbury, of the Washington
Department of Fisheries for his assistance with themanuscript.

Citations_

Anderson, A.M. ,Jr.
1971. Spawning, growth and spatial distribution of the geoduck clam, Panope generosa (Gould) in Hood Canal,
Washington. Ph.D. diss., Univ. Washington, Seattle,
133 p.
Goodwin, C.L.
1973. Effects of salinity and temperature on embryos of thegeoduck clam (Panope generosa). Proc. Nat. Shellfisheries
Assoc. 63:93-95.
Goodwin, C. L.
1976. Observations on spawning and growth of subtidal geoducks (Panope geul'1'Osa, Gould). Proc. Nat. Shellfisheries
Assoc. 65:49-58.

62

NOAA Technical Report NMFS 106

Goodwin, C.L., W. Shaul, and C. Budd
1979. Larval development of the geoduck clam (Panope
generosa, Gould). Proc. Nat. Shellfisheries Assoc. 69:73-76.
King,jJ.
1986. Juvenile feeding ontogeny of the geoduck Panope
abrupta (Bivalvia: Saxicavacea), and comparative ontogeny
and evolution of feeding. Ph.D. diss., Universjty ofVictoria, Victoria, BC, 281 p.

Shaul, W., and C.L. Goodwin
1982. Geoduck (Panope generosa: Bivalvia) age as determinedby internal growth lines in the shell. Can. J. Fish Aquat.
Sci. 39:632-636.
Sloan, N.A. and S.M.C. Robinson
1984. Age and gonad development in the geoduck clam
Panope abrupta (Conrad) from southern British Columbia,
Canada. J. Shellfish Res. 4(2):131-137.

Ecology of Sargassum spp. and Sargassum Forest Formation

JUN-ICHI TSUKIDATE
Nansei National Fisheries Research Institute
Maruishi 2-17-5, Ohno Saeki
Hiroshima 739-04,Japan

Abstract
Sargassum spp. are distributed in the coastal waters of Japan from the northernmost
island of Hokkaido down to the southern island of Kyushu. The forests they create in the
sublittoral zone of the sea are important spawning and rearing grounds for fish andshellfish. A study of Sargassum spp. was conducted to support a project of creating Sargassum forests. Methods for producing these forests are described. The majority of species
started growth in autumn as the water temperature began to fall. The release of oospores
was also temperature dependent and varied between species. Many species matured
between May and June when the rising water temperature approaches 18° C. Contrary
to previous findings, oospores were released irrespective of the tide. Both mono- andmulti-species forests were created and maintained for two years in Hiroshima before
being invaded by other seaweed species.

Introduction_

Sargassum spp. (Fig. 1) are members of the brownalgae family and are among the most highly developed seaweeds in both their morphology and life
cycle. Morphologically, they have a holdfast, stipe,
and lamina and thus resemble a terrestrial tree. Having no asexual stage in their life history (Fig. 2), eachplant is a gametophyte producing both female (00spores) and male gametes (spermatozoids). They are
distributed throughout Japan from the northernmost
island of Hokkaido down to the southern island of
Kyushu (Arasaki 1984). Most species grow in a wide
range of water temperatures and form forests in the
sublittoral zone of the sea that are known to be
spawning, nursery, and feeding areas for many kindsof fish, shellfish, and other marine organisms.
This paper describes the growth and maturation ofSargassum spp. and the propagation of Sargassum forests with artificial reefs and seeded string.

Oospore Liberation and
Growth of Sargassuml--_

The optimum time for oospore liberation variedamong species with respect to season and water temperature (Table 1). At a research area in Fukuoka

Prefecture, Cystophyllum sisymbrioides was the first to
release oospores at 14° C in March. Sargassum horneri,
S. micracanthum, S. tortile, and S. muticum released
oospores at around 15° C in April (Okuda 1981).
Many species matured from May to June when thewater temperature rises to 18° C and higher.
It is generally thought that oospores are discharged at high tide but we found oospore werereleased irrespective of the tide (Okuda 1981, 1982)(Fig. 3). S. micracanthum liberated oospores every seventh or eighth day, S. patens at 3-day intervals, andmost other species every fourth day.

Sargassum Forest Formation_

Most Sargassum spp. start to grow in autumn when
the water temperature begins to fall. Figure 4 shows
the growth of several species on a wet weight basis at
one of the research areas near Hiroshima, where aforest spreads for about 1.1-1.4 ha (Takaba andMizokami 1982). S. horneri and S. tortile grows profusely here from November to the following January.
The largest standing crop for the four species was
15 kg wet weight/m 2 in 1981, and production is estimated to be 15-21 kg wet weight/(m 2 • yr). The
growth of S. ringgoldianum in another area located off
Tokushima Prefecture is shown in Figure 5 (Nakahisa
63

64

NOAA Technical Report NMFS 106

Stem-

Holdfast--

Figure 1
Morphology of Sargassum.

and Kojima 1982). This species matures in autumn ataround 22° C with the largest growth observed in October just before maturation. The standing crop at
this time was 5-6 kg wet weight/m 2 .
The environmental characteristics of the water inthe research field off Yashiro Island are summarizedin Table 2. (Takemoto et al. 1984). The temperature
at the bottom is lower than that at the surface inAugust, although no difference in temperature wasfound in November. Salinity was observed to be alittle low in August. Dissolved oxygen concentrations
were around 5 mg/L and saturated in both Augustand November. Turbidity changed from 0.2 to l.Q
ppm depending on the tide and current. Chemical

oxygen demand ranged from 0.15 to 0.85 ppm. Nutrient concentrations fluctuated during the year.
Dissolved inorganic nitrogen was 1-2 J.Lg-atoms/L in
August and 4-5 J.Lg-atoms/L in November.

Mono-Species Culture
Figure 6 shows an outline of the artificial reefs placedin the sea for the various Sargassum forest formations(Komoto et al. 1987). Fifty concrete blocks wereplaced in both the eastern (E) and western (W)zones of the study area. The E-zone was close to anatural Sargassum forest, while the W-zone was about

Tsukidate: Ecology of Sargassum Forest Formation

65

Fertilized egg

0". 9,~'oph",
FusionJI\,

Spermatozoid

Y'"09

\

Conceptacle

Figure 2
Life Cycle of Sargassum spp.

Table I
Maturing season of Sargassum spp. and coastal watertemperature (Okuda 1982)

SpeciesMonth and
Temperature

Cystophyllum sisymtmoides

Late March
14° C

Sargassum horneri

Middle andlate April
15-16° C

S. micracanthum

S. tortile
S. muticurn

S. patens
S. hemiphyllum

Early andmiddle May
18° C

S. thunbergii

Middle andlate May
19-20° C

Hijikia fusiforme

late May andearly June
2]0 C

S. ringgoldianum

Early and middle
October
22° C

50 m from it. The natural forest was dominated byseveral species of Sargassum and Ecklonia cava. The
seasonal variation in the number of individuals of
S. patens is shown in Figure 7 (Yoshikawa andTsukidate 1984, 1985). Mortality was very high during the first three months; afterwards the number ofplants stayed at the level seen in the natural forests.
The seasonal variation in the total length of S. patens
is shown in Figure 8 (Yoshikawa and Tsukidate 1986).
The total length increased toward the following
spring, reaching an average of about 70 cm. Length
declined reaching its smallest size of 20 cm in October, followed by another increase to between 50 and
60 cm in February. Thus the maximum growth stage
was found between winter and spring. Figure 9 shows
the seasonal variation in wet weight of S patens(Yoshikawa and Tsukidate 1986). The growth pattern
was identical to that of total length. These forests
diminished in the summer months but enlarged
again during this winter to spring period. However,in the third year, they did not enlarge their biomass
because of the invasion of other seaweeds such as
Ecklonia cava and Padina arborescens. Thus, Sargassum
forests were created and maintained in a roughly
1,000 m 2 area with artificially prepared reefs by using
seeded string for at least two years.

66

NOAA Technical Report NMFS 106:~ rWL
1:: I lIl-1J1lllJL57

f

1

C1I

I

III

70 J
25

1
11

100 I

-c
C1I

III

co

~

o
C.

o
25

~

25

25
5
2

-11111_

2

3

4

5

Sargassum confusum

Sargassum muticum

DID

U

IUD 011

Sargassum hornerimnSargassum micracanthum----Ill I.llL-.wIlIDll-l...IDI IllI

Cystophyl/um sisymbrioides

am

~L
~

Hizikia fusiforme

Sargassum hemiphyl/um-WlIDnL-_UDJ,-?fi.

Sargassum patens

om

I

t__

+"

'0

_

_

mL

25t

tQ

Sargassum thunbergii

lIIIl_ _- l~

III

o
o

_

.Ill!!

1

C1I_.J!!!:

6

nrn

um

7

8

l

om ftJB
9 10 11 12011

Sargassum tortile

Figure 3
Interval of oospore liberation in
Sargassum spp. (Okuda 1982).

Interval period (days)

10

J\

1

o Sargassum horneri
X Cystophyllum sisymbrioides

o Sargassum patens
Do Sargassum tortile

+oJ

..r.

0'1 .........

• _N

~~
0'1

2

+oJ

Q)~

~--

1
G-

-0

Jan.

Mar.

0

May

July
1981

Sept.

Nov.

1982

Figure 4
Seasonal variations in wet
weight of Sargassum spp.
(Takaba and Mizogami
1982).

Tsukidate: Ecology of Sargassum Forest Formation

•

5

o

67

sall/assum ringgoldianum
sall/assum homeri

0.1

0
May

Sept.

July

Nov.

Jan.

Mar.

1982

1981

Figure 5
Seasonal variations in wet weight of' Sargassum
ringgoldianum(Nakahisa and Kojima 1982;
Nakahisa et al. 1983).

Table 2
Environmental characteristics of the research field in Yashiro Island. (Takemoto et al. 1984.) S = surface; B = bottom.
DO = dissolved oxygen; COD = chemical oxygen demand; DIN = dissolved inorganic nitrogen. tr = trace.
Water
depth
(m)

Layer
(m)

3.5

B

7.0

B

7.0

B

4.0

B

11:52

2.5

B

11:55

4.0

B

12:00

12.0

B

12:05

3.5

B

12:08

9.0

B

SurveyseasonSurveystation

Surveytime

1983
Aug. 30

L-4-4

10:39
10:37
10:56
10:54
11:02
11:00
11:11
11:09

L-4-8
L-12-4
L-12-8
1983
Nov. 30

S
S
S
S

L-4-2

S

L-4-4

S

L-4-8

S

L-12-4S

L-12-8

S

Water
temp.

SalinityDO(0C)(0/00)(mg/L)

pH

Turbidity
(ppm)

25.4
24.8
25.6
24.4
25.4
23.9
25.4
24.7

32.12
32.18
32.12
32.28
32.18
32.48
32.12
32.30

5.57
5.60
5.48
5.22
5.80
5.58
6.07
5.55

8.12
8.15
8.10
8.10
8.12
8.15
8.12
8.15

0.18
1.12
0.10
0.85
0.15
7.20
0.23
0.65

0.64
0.37
0.85
0.24
0.14
0.24
0.32
0.16

0.22
tr
0.12
0.10
tr
0.17
0.11
0.11

1.75
1.85
1.69
1.73
1.11
2.04
0.95
0.95

1.22
1.27
0.55
0.70
0.75
0.82
0.42
0.43

0.37
tr
tr

0.53
0.58
1.14
0.90
0.36
0.85
0.53
0.52

16.6
16.8
16.4
16.8
17.0
16.8
16.4
16.7
17.0
16.8

32.88

5.62
5.72
5.58
5.59
5.56
5.58
5.56
5.50
5.87
5.37

8.15
8.15
8.20
8.19
8.20
8.15
8.20
8.20
8.20
8.20

0.20
1.00
0.45
0.95
0.55
1.65
0.85
0.60
0.40
0.45

0.43
0.48
0.53
0.77
0.51
0.46
0.58
0.45
0.56
0.42

0.42
0.42
0.48
0.38
0.42
0.42
0.39
0.42
0.39
0.40

3.67
4.41
6.47
3.54
4.83
4.70
3.94
3.76
3.56
3.73

1.80
2.25
2.21
1.53
2.40
2.43
2.08
1.72
1.38
1.41

0.83
0.88
0.84
0.87
1.00
0.97
0.83
0.84
0.95
1.00

1.04
1.28
3.42
1.14
1.43
1.30
1.03
1.20
1.23
1.32

32.88
32.90
32.94
32.91
32.91
32.86
32.94
32.92

Multi-Species Culture
Two species of Sargassum out of three, namely S. patens, S. piluliferum, and S. ringgoldianum werecombined and planted on the c<;mcrete blocks at thesame time. Seeded strings with embryos werewrapped around the blocks alternately. Thus, as
shown in Figure 10, eight strings with S. patens andeight with S. piluliferum resulted (Komoto et al. 1987).

P03-P
COD
(ppm) (J.l.g-at/L)

DI
NH 4-N
N0 2-N
N0 3-N(J.l.g-at/L) (J.l.g-at/L) (J.l.g-at/L) (J.l.g-at/L)

tr
tr
tr
0.13
tr

The ratios in wet weight of S. piluliferum to S. patens are seen in Figure 11 (Komoto et al. 1988).
S. piluliferum dominated the artificial reefs in the firstyear and also demonstrated a superior regenerative
ability in the second and third years. In the first year,
S. piluliferum accounted for 74.7% of the combined
wet weight of the two species, 86.5% in the second
year and 87.7% in the third year. Figure 12 shows theratios in wet weight per single concrete block of

68

NOAA Technical Report NMFS 106

r--------------------------------o m
20
50
oJ

88

o

,''fij.. . . . .. .,'Iii';

,

lj:£!

I
I

'§j
/lJr:,;

,:

,~

I

o

.........../

00
CD

Mono-speciesw..I _------.,.
0,rd'---'

'c

00
00

'0

r'oifffil I~cl/__,II.a.8 --.:1

Multi-species

i
I

I

'" "

1.-

.n{

8'

~!
!l

E

I

111 Mono-species

°nl

88

r) · Y"h;,."L_

Figure 6
An outline of the artificial reef forseaweeds off Yashiro Island(Komoto et al. 1987). W = western zone of the study area; E =eastern zone of the study areajn
the Seto Inland Sea, off Yashiro
Island .

•
300

o

200

\

•o w

•

\

\

\.\

\

100

'b.......

• __e'..'0----------- 0 ""'-.

'",~

""" •

.J:)""",

o_~~~.~~

Aug.

1983

Oct.

Dec.

Feb.

1984

Apr.

June

Aug.

Oct.

Dec.

Figure 7
Seasonal variations in number of individuals per 75 cm string of Sargassum
patens (Yoshikawa and Tsukidate 1984,1985) . • = western zone of the study
area; 0 = eastern zone of the study
area.

Tsukidate: Ecology of Sargassum Forest Formation

69

80
70
60
..c
+-'

C'

c~

~

!

/

50

I
I
I

~E
_U

"'~

0

I-

•o w

!

40

!

I

!

30

20

()

/

.....-~..,.~.-

10.-.~~-

o
Aug.

1983

Oct.

Dec.

Feb.

Apr.

June

Aug.

Oct.

Feb.

Dec.

1984

Apr.

June

Aug.

•

~o

0----0.

Oct.

Dec.

1985

Figure 8
Seasonal variations in total length of Sargassum patens (Yoshikawa and Tsukidate 1986) . • = western zone of the study area;= eastern zone of the study area.

oS. piluliferum and S. patens to the invading seaweeds(Komoto et al. 1987, 1988). S. piluliferum accounted
for 77.6%-86.2% of all seaweeds in the first and second years and then dropped to 47.9% in the thirdyear with the invasion of other seaweeds. Therefore,
the forests were not solely composed of Sargassum but
also of other seaweeds by the third year. Among theother seaweeds, Ecklonia cava first appeared in thesecond year, joined by Padina arborescens and Hypnea
charoides, all of which grew profusely during the thirdyear.
The natural vegetation of Sargassum spp. on theartificial reefs was surveyed during culture of the two
species on the non seeded control blocks (Fig. 10). As
seen in Figure 13, S. horneri was the prevailing specieson the control reefs the first year, probably because it
matures earlier than any other species and grows fast
(Komoto et al. 1987, 1988). The following year,
S. tortile grew vigorously and became dominant.
S. micracanthum also grew markedly in the second andthird years. It is assumed that the annual Sargassum
spp. settle first followed by the perennial ones.

Conclusion
Sargassum forests were created and maintained in anexperimen tal area using artificially prepared reefs
with seeded string for at least two years using bothmono- and multi-species cultures. However, in thethird year the forests diminished and were overcomeby the invasion of other seaweeds. Therefore, more
seedlings must be added in the third year and otherseaweeds eliminated in order to maintain Sargassum
forests for a longer time. Only a small amount ofgrazing by herbivores was observed in the Seta Inland Sea. This is thought to be one of the reasons
why we succeeded in forming Sargassum forests.

Citations_Arasaki, S.
1984. Seaweed forests in Japan. In Moba and Marine organisms (T. Hisamune, ed.), p. l-ll. Report ofJapan Fisheries Resources Conservation Association, Zenkoku Choson
Kaikan, ll-35, Nagatacho 1, Chiyoda-Ku, Tokyo 100.

70

NOAA Technical Report NMFS 106

200

.~

•

•

o

f"'" "\.

ff.'"".. . .

fiW"'b'" •.••

.Q\

100

•
........

\'t57

6

"-"'D

•

Aug.

Dec.

Oct.

1983

Feb.

Apr.

June

Aug.

Oct.

Dec.

1984

Feb.

Apr.

June

Aug.

Oct.

Dec.

1985

Figure 9
Seasonal variations in wet weight of Sargassum patens (Yoshikawa and Tsukidate 1986) . • = western zone of the study area;= eastern zone of the study area.

oA

LJ UJ [.1] D
EJ ~ ~ D'"c:~...

~

'"
c:

0
0>

B

232

~

0

Q.. 'C:

Vi Vi

75 em

V) V)

Vi Vi

DDDD
D [] [J D-75 em-S. ringgoldianum

75 em

_____
- - - 75 em - - -

-

75 em

Control

S. patens
S. piluliferum

Figure 10
A schematic diagram of artificial reefs for growing Sargassum spp. off Yashiro Island, Seta Inland Sea (Komoto et al. 1987).
Blocks were planted with seeded strings as follows: Y = S. Patens; M = S. piluliferum; 0 = S. ringgoldianum; C = no seeded string.
Blocks with mixed species were arranged within the reef as shown in (A) and wrapped with alternating strings of each species
as shown in (B).

Tsukidate: Ecology of Sargassum Forest Formation

First year

7I

Second year-,,'OJ,,~:patens

S. patens(13.5%)

S. piluliferum(86.5%)
S. piluliferum(74.7%)

Third year
S. patens(12.3%)

S. piluliferum(87.7%)

Figure 11
Comparison of Sargassum patens to S.
piluliferum (in percent wet weight) on
an artificial reef (Komoto et al. 1988).

First year

Second year
Other seaweeds

S. micracanthum(3.2%)(6.8%)

S. patens(6.4%)

S. tortile(0.1%)
Unknown(0.1%)

S. piluliferum(77.6%)

S. piluliferum(86.2%)

Third year

S. piluliferum(47.9%)

S. micracanthum(1.0%)

Figure 12
Comparison of Sargassum patens to
S. piluliferum to other Sargassum spp.
(in percent wet weight) on an artificial reef. (Komoto et al. 1987, 1988).

72

NOAA Technical Report NMFS 106

First year

Other seaweeds

S. piluliferum(3.1%)(1.2%)

Other seaweeds

S. micracanthum(2.2%)

Unknown(0.3%)u'..
Second year

(1.0%)

piluliferum(21.9%)

Unknown(4.5%)

S. tortile(49.5%)

S. horneri(93.2%)

S. mkracanthum(23.1%)

Third year
S. piluliferum(3.5%)
S. micracanthum(25.4%)H~~~

S.tOrtile~(61.7%)~

S. horneri(0.1%)

Figure 13
Comparison among Sargassum spp.
(in percent wet weight) on an artificial reef. (Komoto et al. 1987,
1988).

Komoto, Y, T. Iwamoto, and H. Matsuura.
1987. Studies on the control of Sargassum forests. Report ofMarine Ranchinp; Program, Nansei Natl. Fish. Res. Inst.,
Hiroshima, p. 17-33. (In Japanese.)
1988. Studies on the con trol of Sargassum forests. Report ofMarine Ranching Program, Nansei Natl. Fish. Inst.,
Hiroshima, p. 23-42. (In Japanese.)
Nakahisa, Y, and H. Kojima.
1982. Ecology of Sargassum ringgoldianum and S.
giganteifolium. Report of Marine Ranching Program, Nansei
Nat!. Fish. Res. Inst., Hiroshima, p. 71-85. (In Japanese.)
Nakahisa, Y, H. Kojima, and N. Tanimoto
1983. Ecology of Sargassum "inggoldianum and S.
giganteifolium. Report of Marine Ranching Program,
Nansei Nat!. Fish. Res. Inst., Hiroshima, p. 99-115. (In Japa-

saceae. Report of Marine Ranching Program, Nansei Natl.
Fish. Res. Inst., Hiroshima, p. 101-110. (In Japanese.)
Takaba, M., and A. Mizogami
1982. Seasonal fluctuation of Sargassum communities' andvertical distribution of Sargassaceae at Kuroshima Islandin the western Akinada. ]. Hiroshima-Ken Fish. Exp. Station. 12:33-44.
Takemoto, K., S. Takayama, S. Yoshioka, Y Komoto, and T. Miyago.
1984. Sargassum forest formation by artificially preparedseedlings. Report of Marine Ranching Program, Nansei
Natl. Fish. Res. Inst., Hiroshima, p. 51-74. (In Japanese.)
Yoshikawa, K., and]. Tsukidate.
1984. Sargassum forest formation by setting artificial
reefs. Report of Marine Ranching Program, Nansei Natl.
Fish. Res. Inst., Hiroshima, p. 75-89. (In Japanese.)
1985. Sargassum forest formation by setting artificial
reefs. Report of Marine Ranching Program. Nansei Natl.
Fish. Res. Inst., Hiroshima, p. 69-86. (In Japanese.)
1986. Sargassum forest formation by settling artificial
reefs. Report of Marine Ranching Program, Nansei Natl.
Fish. Res. Inst., Hiroshima, p. 55-70. (In japanese.)

nese.)
Okuda, T.

1981. Settlement mechanisms on the embryo of Sargassaceae. Report of Marine Ranching Program, ansei Natl.
Fish. Res. Inst., Hiroshima, p. 105-117. (In Japanese.)
1982. Settlement mechanisms on the embryo of Sargas-

Salmon Gonadotropins

PENNY SWANSON
School ofFisheries
University of Washington
Seattle, WA 98195'

ABSTRACTControl of gonadal function by pituitary gonadotropins (GTHs) is a general feature ofvertebrate reproduction. In most tetrapods, gonadal function has long been known to be
regulated by two GTHs: follicle-stimulating hormone (FSH) and luteinizing hormone(LH) (Licht et al. 1977). Luteinizing hormone, follicle-stimulating hormone, and a third
pituitary hormone, thyroid-stimulating hormone (TSH), are chemically related. All threeconsist of an ()( and 13 subunit, which interact noncovalently and are both glycosylated. In
mammals it has been found that the ()( subunits ofTSH, LH, and FSH are identical withina species, whereas the 13 subunits are hormone specific and structurally conserved between species (Pierce and Parsons 1981).
The question of whether fish reproduction is regulated by one or two pituitary GTHs
has been controversial for nearly two decades. A single GTH, sometimes referred to as
maturational GTH, has been isolated from several teleost species: chinook salmon,
Oncorhynchus tschawytscha (Breton et al. 1978); common carp, Cyprinus carpio (BurzawaGerard 1971); silver carp, Hypophthalmichthys molitrix (Chang et al. 1988a, 1990); pike eel,
Muraenesox cinereus (Huang et al. 1981; Liu et al. 1989); tilapia, Orechromis mossambica(Farmer and Papkoff 1977); Mrican catfish, Clarias gariepinus (Goos et al. 1986); andAtlantic croaker, Micropogonias undulatus (Copeland and Thomas 1989). Maturational
GTH has been thought to regulate all aspects of gametogenesis (see Burzawa-Gerard
1982; Fontaine and Dufour 1987). The single type of GTH which has been consistently
isolated from the teleost species so far examined, appears to be chemically related totetrapod FSH and LH because of its glycoproteic and subunit nature. In contrast, Idler
and colleagues (see Idler and Ng 1983) prepared two GTH fractions from pituitaries offour teleost species; one adsorbed to Concanavalin-A Sepharose and stimulated gonadal
steroidogenesis (Con A II) while the other did not adsorb to Concanavalin-A Sepharoseand stimulated in vivo vitellogenin uptake by ovarian follicles (Con A I). Con A II showssome chemical similarity to classic tetrapod pituitary GTHs, whereas Con A I does not;Con A I is low in carbohydrate content, and its subunit nature has not been demonstrated.
More recently two pituitary GTHs, GTH I and GTH II, which are distinctly different
from each other in chemical characteristics and structurally homologous to tetrapod FSH
and LH, have been isolated from chum salmon (Oncorhynchus ketal (Kawauchi et al. 1989;
Suzuki et al. 1988 a,b; Itoh et al. 1988) and coho salmon (0. kisutch) (Swanson et al.
1991). Chum salmon GTH I 13 and GTH II 13 subunits have only about 31% amino acidsequence identity to each other. Amino acid sequence comparisons of the chum salmon
GTH 13 subunits to those of bovine FSH (bFSH) and bovine LH (bLH) revealed that the
GTH I 13 subunit has slightly greater sequence identity to the bFSH 13 subunit (41 %) than
to the bLH 13 subunit (35%), whereas the GTH II 13 subunit has greater identity to thebLH 13 subunit (42%) than to the bFSH 13 subunit (38%). Comparisons of the eDNA
sequences of the chum salmon GTH 13 subunits to the cDNAs of the bLH 13 and bFSH 13
subunits demonstrate the same structural relatedness (Sekine et al. 1989). The amino

I Present address: School of Fisheries, University of Washington, Seattle, WA 98195 and Northwest Fisheries Science Center, National Marine
Fisheries Service, 2725 Montlake Blvd. E., Seattle, WA 98112.

73

74

NOAA Technical Report NMFS 106
acid sequences of chinook salmon (Trinh et al. 1986), common carp (Chang et al.
1988b), silver carp (Chang et al. 1990), and pike eel GTH 13 subunits (Liu et al. 1989)
show about 40% sequence homology to mammalian LH 13 subunits, and about 75%
sequence identity to the chum salmon GTH II 13 subunit. Therefore, it is apparent thatboth GTH I and GTH II have structural homology to tetrapod FSH and LH, and thesingle GTH molecule previously isolated from several teleost species is structurally similar
to chum salmon GTH II. GTH I had not been identified in previous studies.
The physiological distinction between GTH I and GTH II is not as clear as the chemical distinction. Studies done to date so far indicate that GTH I and GTH II can stimulate
gonadal steroidogenesis. Suzuki et al. (1988c) found that chum salmon GTH I and GTH
II were equally potent in stimulating in vitro estradiol 17-13 production by vitellogenic
amago salmon (0. rhoduTUs) ovarian follicles. However, chum salmon GTH II was foundto be two times more potent than chum salmon GTH I in stimulating 170.hydroxyprogesterone production by ovarian thecal layers and 170.,2013-dihydroxy4-pregnen-3-one production by granulosa layers in the presence of 170.hydroxyprogesterone (Suzuki et al. 1988c). Coho salmon GTH I and GTH II stimulated
in vitro estradiol 17-13 and total androgen production by juvenile coho salmon ovarianand testicular tissue, respectively, in a similar dose-dependent manner (Swanson et al.
1989). In recent studies using in vitro incubations of coho salmon testicular fragments, it
was found that coho salmon GTH I and GTH II were equipotent in stimulating 11ketotestosterone secretion U. Planas, School of Fisheries, Univ. of Washington, Seattle,
WA, pers. commun. September 1989). The only clear difference between the biologicalactivities of GTH I and GTH II was the GTH I stimulation of vitellogenin uptake byrainbmy trout (0. mykiss) oocytes both in vivo and in vitro (Tyler et al. 1991). In the invitro studies by Tyler and colleasues, GTH I was roughly 100-times more potent than
GTH II whereas in vivo GTH II was not active compared to GTH 1, which doubled therate of vitellogenin uptake. Distinctly different actions of GTH I and GTH II in malesalmon have not been found and should be the subject of future investigations.
Although salmon GTH I and GTH II appear to have similar steroidogenic activities
when tested in vitro, blood and pituitary levels of these two GTHs vary significantly
during reproductive development. GTE I was the predominate GTH in the plasma andpituitary of vitellogenic/spermatogenic rainbow trout, whereas GTH II was the predominant GTH at the time of final reproductive maturation (Suzuki et al. 1988d). Swanson etal. (1989) found that in prespermatogenic and previtellogenic (prepubertal) coho
salmon, GTH I was the only GTH detectable in the plasma. Recent analysis of plasma
levels of GTH I and GTH II in coho salmon during the final year of reproductive maturation have shown an extended increase in GTH 1 from May through October during
vitellogenesis/spermatogenesis, and a decline at the time of spawning (late November or
early December). On the other hand. plasma GTH II levels were consistently at low or
nondetectable levels throughout vitellogenesis/spermatogenesis, and increased dramatically at the time of spawning (Swanson et al. unpublished). Therefore, the physiologicalrelevance of the steroidogenic activity of GT11 II in prepubertal or even vitellogenic/
spermatogenic coho salmon is questionable since it does not appear to be present insignificant levels in the plasma during this time.
Immunocytochemical studies of salmonid pituitary gonadotrophs have revealed a lacka co-localization of the two GTHs; this is contrary to what has been found for LH and
FSH in tetrapods. Nozaki et al. (1990a) demonstrated that antisera directed against coho
salmon GTH I 13 and GTH II 13 subunits stain two distinctly different gonadotroph celltypes in the pars distal is of salmonids and do not stain thyrotrophs, sommatotrophs,
lactotrophs, corticotrophs, or melanotrophs. Moreover, in an ontogenetic study of rainbow trout pituitary gonadotrophs, Nozaki et al. (1990b) showed that GTH I-producing
cells are present prior to puberty and increase in number during vitellogenesis; GTH IIproducing cells do not appear until after the onset of spermatogenesis/ vitellogenesis
and are greater in number than GTH I-producing cells at the time of final reproductive
maturation. These data, in addition to data on blood levels of GTH I and GTH II, suggest
relationships of GTH 1 with gonadal growth and GTH II with final maturation of the
gonads similar to those of FSH and LB in mammals. More detailed investigations of thespecific roles of GTH 1 and GTH II in salmonid reproductive development are necessary
to determine the physiological importance of the dual gonadotropin system in fish.

Swanson: Salmon Gonadotropins

Citations_

Breton B., P. Purnet, P. Reinaud
1978. Sexual differences in salmon gonadotropin. Ann.
BioI. Anim. Biochim. Biophys. 18:739-765.
Burzawa-Gerard E.
1971. Purification d 'une hormone gonadotrope hypophysaire de poisson ti~leosteen, la Carpe (Cyprinus carpioL.). Biochimie (Paris) 53:545-552.
1982. Chemical data on the pituitary gonadotropins andtheir implication to evolution. Can. J. Fish. Aquat. Sci.
39:80-91.
Chang y'S., FL. Huang, C.T Chen, TB. Lo
1988a. Isolation and properties of the pituitary gonadotropin from silver carp (Hypophthalmichthys molitrix). In t. J.
Pept. Protein Res. 31:150-156.
Chang y'S., C.J. Huang, FL. Huang, TB. Lo
1988b. The primary structures of carp gonadotropin subunits deduced from cDNA nucleotide sequences. Int..J.
Pept. Protein Res. 32:556-564.
Chang y'S., C.J. Huang, FL. Huang, C.S. Liu, TB. Lo
1990. Purification, characterization, and molecular cloningof gonadotropin subunits of silver carp (Hypophthalmichlhys
molitrix). Gen Compo Endocrinol. 78:23-33.
Copeland P.A., P. Thomas
1989. Purification of maturational gonadotropin from Atlantic croaker (Micropogonias undulatus) and development of ahomologous radioimmunoassay. Gen. Compo Endocrinol.
73:425-441.
Farmer S.W., H. Papkoff
1977. A teleost (Tilapia mossambica) gonadotropin that resembles luteinizing hormone. Life Sci. 20:1227-1232.
Fontaine Y.A., S. Dufour
1987. Current status of LH-FSH-like gonadotropin infish. In Proceedings of the third international symposium
on reproductive physiology of fish
(D.R. Idler, L.W. Crim,
andJ.W. Walsh, eds.) p. 48-56. Memorial Univ. Newfoundland, St. John's Newfoundland, Canada.
Goos H.J.T, R. De Leeuw, E. Burzawa-Gerard, M. Terlou, CJJ.
Richter
1986. Purification of gonadotropic hormone from the pituitary of the Mrican catfish, Clarias gariepinus (Burchell), anddevelopment of a homologous radioimmunoassay. Gen.
Compo Endocrinol. 63:162-170.
Huang, F.L., C.J. Huang, S.H. Lin, TB. Lo, H. Papkoff
1981. Isolation and characterization of gonadotropin
isohormones from the pituitary gland of pike eel(Muraenesox cinereus). Int.J. Pept. Protein Res. 18:69-78.
Idler D.R., TB. Ng
1983. Teleost gonadotropins: Isolation, biochemistry andfunction. In Fish physiology, Vol. 9A (WS. Hoar, OJ.
Randall and E.M. Donaldson, eds.), p. 187-221. Acad.
Press. NY.
ltoh H., K Suzuki, H. Kawauchi
1988. The complete amino acid sequences of 13 subunits of
two distinct chum salmon GTHs. Gen. Compo Endocrinol.
71:438-451.

75

Kawauchi H., K Suzuki, H. ltoh, P. Swanson, M. Nozaki, N. Naito,
Y. Nagahama
1989. Duality of salmon pituitary gonadotropins. Fish
Physiol. Biochem. 7:29-38.
Licht P., H. Papkoff, S.W. Farmer, C.H. Muller, H.K Tsui, D. Crews
1977. Evolution of gonadotropins structure and function. Recent Prog. Horm. Res. 33:169-248.
Liu C.S., F.L. Huang, Y.S. Chang, TB. Lo
1989. Pike eel (Muraenesox cinereus) gonadotropin. Aminoacid sequences of both a. and 13 subunits. Eur. J. Biochem.
186:105-114.
ozaki M., . Naito, P. Swanson, K Miyata, Y. Nakai, Y. Oota, K.
Suzuki, H. Kawauchi
1990a. Salmonid pituitary gonadotrophs I. Distinct cellular
distributions of two gonadotropins, GTH I and GTH
II. Gen. Compo Endocrinol. 77:348-357.
Nozaki M., N. Naito, P. Swanson, WW Dickhoff, Y. Nakai, K.
Suzuki, H. Kawauchi
1990b. Salmonid pituitary gonadotrophs II. Ontogeny of
GTH I and GTH II cells in the rainbow trout (Salmo
gairdneri irideus). Gen. Compo Endocrinol. 77:358-367.
Pierce J.G., TF. Parsons
1981. Glycoprotein hormones: structure and function. Ann. Rev. Biochem. 50:465-495.
Sekine S., A. Saito, H. Itoh, H. Kawauchi, S. ltoh
1989. Molecular cloning and sequence analysis of chumsalmon gonadotropin cDNAs. Proc. Nat. Acad. Sci.
86:8645-8649.
Suzuki K, H. Kawauchi, Y. Nagahama
1988a. Isolation and characterization of two distinct gonadotropins from chum salmon pituitary glands. Gen. Compo
Endocrinol. 71 :292-301.
1988b. Isolation and characterization of subunits of two
distinct gonadotropins from chum salmon pituitary
glands. Gen. Compo Endocrinol. 71:302-306.
1988c. Steroidogenic activities of two distinct salmon gonadotropins. Gen. Compo Endocrinol. 71:452-458.
Suzuki K, A. Kanamori, H. Kawauchi, Y. Nagahama
1988d. Development of salmon GTH I and GTH II
radioimmunoassays. Gen. Compo Endocrinol. 71:459-467.
Swanson P., M.G. Bernard, M. Nozaki, K Suzuki, H. Kawauchi,
WW Dickhoff
1989. Gonadotropins I and II in juvenile coho salmon. Fish
Physiol. Biochem. 7: 169-176.
Swanson P., K Suzuki, H. Kawauchi, W.W. Dickhoff
1991. Isolation and characterization of two coho salmon gonadotropins, GTH I and GTH II. BioI. Reprod. 44:29-38.
Trinh KY., N.C. Wang, C.L. Hew, L.W Crim
1986. Molecular cloning and sequencing of salmon gonadotropin beta subunit. Eur.J. Biochem. 159:619-624.
Tyler C.,J.P. Sumpter, H. Kawauchi, P. Swanson
1991. Involvement of gonadotropin in the uptake ofvitellogenin into the vitellogenic oocytes of the rainbow trout,
Oncorhynchus mykiss. Gen. Compo Endocrinol. 84:291-299.

Breeding Season of Japanese Scallop off theEastern Coast of Hokkaido

HIROSHI ITO
Hokkaido National Fisheries Research Institute(formerly: Hokkaido Regional Fisheries Research Laboratory)
Fisheries Agency
The Ministry of Agriculture, Forestry and Fisheries
116 Katsura-koi, Kushiro City, Hokkaido, 085JAPAN

Abstract
The breeding ofJapanese scallops, Patinopecten (Mizuhopecten) yessoensis, in coastal beds
off east Hokkaido was estimated using a gonadal-somatic index. Although the breeding
season differs by locality, it begins when bottom temperatures reach about 5° C in thebeds. Therefore the bottom temperature indicates the beginning of scallop breeding.

Introduction_

Mariculture of the Japanese scallop, Patinopecten(Mizuhopecten) yessoensis, developed rapidly in the
mid-1960s owing to development of methods formass seed production using natural spats (Tsubata
1982). After the 1970s, scallop mariculture production rapidly increased, mainly off the coast of
Hokkaido in northern Japan (Figs. 1 and 2; Ito
1988). In 1988, Japanese scallop production
amounted to 341,618 metric tons. Although the scallop fishery has become a large industry, the reliabilityof scientific information concerning scallops remains
uncertain (Ito 1989).
Japanese scallop mariculture begins with the collection of wild spat. Spat collection relies upon
realtime information on the breeding occurrence of
scallop breeding: the subsequent larvae reflect theearliest stage of scallop life (Fig. 3: modified from
Yamamoto 1964; Maru 1972, 1976, 1978; Osanai
1975; Kawamata et al. 1981). The breeding season of
the scallop in various coastal beds has in the past
been roughly monitored by coastal fisheries cooperative associations. Results in 1982 are shown in Figure
4 (Ito 1990). Among the scallop culturing areas off
Hokkaido, breeding occurred first in the Japan Sea,
second in the Okhotsk Sea, and third in the Nemuro
Straits. It is thought that a relationship exists between
the gonado-somatic index and the surface tempera-ture, although scallop live on the bottom. Similarly
observed results have already been reported(Kinoshita 1934; Yamamoto 1951). At about 8° C surface temperature (Fig. 5), some- scallops might begin
breeding; however, bottom temperature is not always
parallel to that of the surface. Currently, informationon breeding is vague, whereas industrial spat collection requires accurate data on the key phenomena.
In fact, industrial monitoring methods for wild spatcollection are not established even now. Therefore,
the author created a special team for this researchand surveyed the breeding season, focusing on wild
spat collection.

Materials and Methods_

Japanese scallops were collected from three localities
in the Nemuro Straits, east Hokkaido (Fig. 6). Depths
of survey si tes were determined acoustically: 13 m off
Rausu, located in the north in the Nemuro Straits; 21
m off Shibetsu in the middle portion of the Nemuro
region; and 12 m off Bekkai, which is located in thesouthern portion of Nemuro Bay. Specimens were
sampled every week from a wild population of 4- to 7year-olds at the site off Rausu by diving, from similar
wild population off Shibetsu using a scallop dredge,
and from a cultured population of 3-year-olds off
Bekkai using a scallop dredge. Water temperature at

77

78

NOAA Technical Report NMFS 106~ 14SoE~OKHOTSK

SEA

Figure 1
Main mariculture areas in Japan and natural distribution In east Asia of the Japanese scallop, Patinopecten(Mizuhopectl'n) yl'ssoensis.

every survey time was observed from surface to bottom at intervals of 2 m with the use of athermometric electrode. This field study began in
mid- to late April 1982 at every site and finished in
May/July 1982 as breeding was completed at eachsite. Gonads and soft body tissues were measured by
wet weight. The gonado-somatic index was estimatedwith the following equation.
gonado-somatic index = gonad weight X
100/soft body weight

Results and Discussion_

Changes in the gonado-somatic indexes of males andfemales were nearly synchronized, although details
showed they were slightly different (Fig. 7). The index values at each site were lower in April andincreased in May. When the index decreased suddenly after reaching a maximum, we estimated thatbreeding occurred. Using this index, we also estimated that the time of breeding varied by locality. It
began in early June off Rausu in the north (NemuroIto: Breeding Season of Japanese Scallop

79• TOTAL:JAPANo HOKKAIDOPRODUCTION: METRIC TONS
350,000
1910-23:Temporary high LeveL catch period
192~-32:StabLe high LeveL catch period
1933-~~:Extreme high LeveL catch period
19~5-68:StabLe Low LeveL catch period
1969-76:Hanging cuLture deveLopment period
1977-88:Sowing and hanging cuLture mass
production period

Figure 2
Annual productionof the Japanese scal2000
lop, Patinopecten (Mizuhopecten) yessoensis, in-" Japan, 1910-1988.

o
1900
y

1950
E
A

R

L-

Straits), in mid:June off Shibetsu in the middle area,
and in mid-May off Bekkai in the southern portion of
the Nemuro region. Although the breeding season
differs by locality, it begins when the bottom temperature reaches about 50 C in every bed. Therefore,
the bottom temperature seems to indicate the beginning of scallop breeding. This is new information on
the Japanese scallop. In the past, scallop researchers
focused on the midpoint of the breeding season. In
this paper the author is proposing a new idea, thatearlier generation spat can be of great advantage tothe scallop mariculture industry. Earlier generation
spat grow to a larger size than those collected later.
The larger scallop also market at a better price. So,
knowledge of the beginning of the breeding season isbeneficial information. In addition, the author suggests that research focus on new concepts (Ito 1989)
for scallop mariculture which includes an examination of the biology of breeding.

Acknowledgments
The author wishes to express sincere thanks to thefollowing co-workers of the research team: Toshikazu
Fujimoto, Takao Sasaki, Haruo Moriya, Tohru Ikeda,
Tadashi Abe, Yukiyasu Nakata, and Yoshiji Kobayashi
for their expertise and friendship.

Citations_

Ito, H.
1988. Sowing culture of scallop in Japan. In New and innovative advances in biology/engineering with potential
for use in aquaculture: proceedings of the fourteenth
U.S.:lapan meeting on aquaculture (A.K. Sparks, ed.),p. 63-69. NOAA Tech. Rep. NMFS 70.
1989. Concept in mariculture of Japanese scallop. Scallop(2), Hotatekai (Kushiro), p. 89-97. (In Japanese.)
1990. Some aspects of offshore spat collection of Japanesescallop. In Marine farming and enhancement: proceedings of the fifteenth U.S.:lapan meeting on aquaculture(A.K. Sparks, ed.), p. 35-48. NOAA Tech. Rep. NMFS 85.
Kawamata, K., Y. Tamaoki and A. Fuji.
1981. Gonad development of the cultured scallops in Funka
Bay. J. Hokkaido Fish. Exp. Stn. (Hokusuisi Geppo)
38:132-146. (In Japanese.)
Kinoshita, T.
1934. Relation between breeding of the Japanese scallop andwater temperature. Report of the Hokkaido Fish. Res.
Stn. (Hokusuisi Junpo) 233:3-8. (In Japanese.)
Maru, K.
1972. Morphological observations on the veliger larvae of ascallop, Patinopecten yessoensis Uay). Sci. Rep. Hokkaido
Fish. Exp. Stn. 14:55-62. (In Japanese, English abstr.)
1976. Studies on the reproduction of a scallop, Pati~opecten
yessoensis Uay)-l. Reproductive cycle of the cultured
scallop. Sci. Rep. Hokkaido Fish. Exp. Stn. 18:9-26. (InJapanese, English abstr.)
1978. Bludies on the reproduction of a scallop, Patinopecten
yessoensis Uay)-2. Gonad development in l-year-old
scallops. Sci. Rep. Hokkaido Fish. Exp. Stn. 20:13-26. (InJapanese, English abstr.)

80

NOAA Technical Report NMFS 106

J

SC:-he~d

E!SSP

o~8WIIlng

I

Breeding

3-SP
tall SO-60p}GOnSd Index~(demersal egg)

Unfertilized egg
80 P (70-100 p)

1,~preeursor to ,moni toring thplanktonic
larva

Ferti lizatlon

Fertilized egg

1

Developing embryo

Polar lobe stage

Larvalmonitoring

Firat c lesvage
Second cleavage

Third cleavage
Fourth cleavage
Blastula

Age> Prodi ssoconch stage

GastrulaOyr
4 days
Trochophore larvs
sfter fertllizatlonl
S-7 days

30-35 days
40 days
70-80 days
4-5 months

Early vellger larva

1
Late vellger larva(Pedlvellger)<lSOp

D-shaped larva
Esrly umbo-stage lsrva

150-190

Umbo-s tage larva
EarlY full_grown larva

190-240

I
I

Spatfallprediction}24O<

Full-grown larva!

Planktonanalysis}

2S0-280p<

At taehed spat

1
Bottom Inhabl tlng spattJuvenllspat} collection

1 cm<

C'J

Early grown stage~

Intermediate
cuI ture

Late growing stage

De~eneratlng stage
Un~ifferentlated atsgs

Age

Diiterentlated atage(Srx differentiated stage)~

I yr

Reetlng stage

Adult

1 Ii!

Ea~r1Y growing stage

Early growing stage

Late growing

Late growing stage

~

stage

l

Malturlng stage

MaLrlng stage~Jeedlng stag. _ - - - - . .

G;\!edlng stagel

Hangl ng eu !turaAge

Resting stage

2yr

EarlY growing stage•l
Resting stagel
Early growing stageandLa t. growl ng s tago

Late growing stage

Spent stage

older

~

l

~

uturlng stag.

Life cycle of scallop

Dissoconch
stage>Sowingor cui tura

Spent stage.

during stage

Culture method

Figure 3
Life cycle of the Japanese scallop, Patinopecten (Mizuhopecten) JPssoensis, with the note of culture methods (modified from
Yamamoto 1964; Maru 1972, 1976, 1978; Osanai 1975; Kawamata et al. 1981).

Ito: Breeding Season ofJapanese ScallopJAPAN• • ••

I

•

81

•

SEA

••

COAST

SARUFUTSU-. •••• . ••OOMU

Lt:3 II a
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':

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.::

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JUL.

Figure 4
Changes in gonado-somatic index (gonad weight X 100/softbody weight, %) of the Japanese scallop, Palinopecll'n(Mizuhopeclen) yessoensis, off
the coasts of Hokkaido, 1982.
Specimens were mainly 3- and4-year-olds (Ito 1990).

82

NOAA Technical Report NMFS 106

JAPAN:SEA IMASHI"E
COASTI:~t31tmE =t====~~==~r==1FJ:=:\==\:=='r=~~=\==\==lI==t=:J=I=I=I=f=J~~

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COAST I
I

J
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COAST:

JAN.

Figure 5
Changes in surface temperature of the coastal water related with the locality of Hokkaido in 1982 (Ito 1990).

0

145 E

OKHOTSK~fTi'£"fi'dIOn.I miN
10

20km

OCEAN

Figure 6
Survey sites (A, B, C) in Nemuro Straits, east
Hokkaido. A = site off Rausu. B = site off
Shibetsu. C = site off Bekkai.

Ito: Breeding Season ofJapanese Scallop

BEKKAISHIBETSU

il'~~~!? ~ I p~a~b ~, ~
I

APR.

MAY

I

JUN. JUl.

T1~~ OF YEAR

~WA~~~]~\~

~ ~
CI

-

' " "" ,

I

A

83

I

[~ ~, ij ~

M

A

0
10

20

M

~ '~

lOf\~" :oFigure 7
Changes in the gonado-somatic index of Japanese scallops, Patinopecten (Mizuhojiecten) yessoensis, and the water temperaturein Nemuro Straits, 1982 (Ito 1990).

Osanai, K.
1975. Seasonal gonad development and sex alteration in thescallop, Patinopecten yessoensis. Bull. Mar. BioI. Stn.
Asamushi. Tohoku Univ. 15(2):81-88.
Tsubata, F.
1982. A history of the fisheries researches of the scallop,
Patinopecten yessoensis, in Mutsu Bay. Published by Aomori
Prefecture (Aomori Prefecture government publication),
120 p. (In Japanese.)

Yamamoto, C.
1951. Ecological study on the spawn ing of the scallop, Pecten(Patinopecten) yessoensis in Mutsu Bay. Bull. Japan. Soc. Sci.
Fish. 17(2):53-56. (In Japanese, English synop.)
1964. Scallop culture in Mutsu Bay. Suisan Zoyoshoku
Sosho (Aquaculture series in fisheries) 6 (Tokyo), 77
p. (In Japanese.)

A Better Method for Oyster Farming in JapanTETSUO SEKI
Oyster Research Institute211 Higashi Mohne Karakuwa
Miyagi 988-05, Japan

Abstract

The Japanese oyster farming industry experienced a remarkable change with the development of the raft culturing method. Recently, northern Japanese oyster farmers havebeen facing difficulty in maintaining the quality of oysters. It was concluded that thisproblem originated with certain methods utilized by the present oyster farmers of theraft culture system. In order to resolve the problems, new farming practices were developed in which the introduced flat oyster Ostrea edulis was used at the Oyster ResearchInstitute. A new cultch-based seed collecting method for single oyster seed production
and a multilayered, compact hanging method were devised to better utilize the coastal
water column for oyster farming. The recent progress of this strategy is described anddiscussed.

Introduction_

The Oyster Research Institute (OR!), a private nonprofit foundation, was established in 1961 at Mohne
Bay in Miyagi Prefecture. Funding for the Institute
was initially derived from private company sources in
Sendai. These companies hoped to redevelop the traditionally important oyster industry in Miyagi
Prefecture and to encourage the related studies ofDr. Takeo Imai, a professor at Tohoku University, by
establishing an industry-based research laboratory.'
The Oyster Research Institute pioheered seed production and farming of the Japanese abalone species
Haliotis discus hannai (Seki 1980; Seki andCuthbertson 1988) and the French oyster Ostrea
edulis (Imai 1967; Imai 1971). The initial aim of introducing French oysters was to establish an exotic
oyster industry in Miyagi Prefecture. Although large
amounts of French oyster seed were produced, theoysters were not popular wi th the J apan~se consumer.
However, recent changes in Japanese eating habits
show an increased demand for luxury seafood.
The aim of French oyster research at OR! was targeted at providing innovative alternatives for oyster
farmers in northern Japan such as improved technological sophistication and ways to better utilize
associated coastal waters.

New methods of collecting and culturing oyster
seed were introduced in 1989; this paper describes
these developments and discusses the relevant background issues.

Recent Problems with Oyster Farmingin Northern Japan_

Since 1950, substantial developments in oyster farming in Hiroshima and northeastern Japan have been
experienced. These developments are due primarily
to the popularity of raft culture and the introductionof long-line systems (Fujiya 1970; Imai 1971; Kafuku
and Ikenoue 1983). Despite such developments, theindustry now faces a number of problems. The increasing popularity of scallop farming due to its
greater returns has stagnated oyster production in
many areas. A gradual increase in the average age ofoyster farmers together with increasing water pollution in growing areas has further contributed to thedecline of the industry.
The introduction of styrofoam flotation to oysterlines gave much greater buoyancy to the system. As aresult, farmers tended not to remove fouling organisms such as mussels from the support lines. Rather
than removing long lines from the water and drying

85

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NOAA Technical Report NMFS 106

out the fouling material, it was found to be mucheasier to simply lower the system further into the water by adding more line. This treatment leaves theoyster in a cool, dark environment where their foodsource, phytoplankton, is less abundant. Such a practice occurs despite the fact that the oyster's natural
habitat is the brackish intertidal zone where temperatures fluctuate more rapidly and food is moreabundant. Failure to remove competitors such asmussels, cionas (sea squirts), and hydroids(tubeworms) further compounds the problem by reducing the amount of food plankton available to theoysters and by restricting light penetration to thedeeper zones and thereby reducing phytopl<,nkton
reproduction.
As a result of the above situation, culture time isextended beyond one year because of slower growth,which in turn affects the spawning cycle because of alack in cumulative temperature required for maturation. A deterioration in taste during autumn also
occurs as a result of unspawned eggs brought about
by a delay in glycogen accumulation. Japanese oysters
are gradually losing flavor compared with the same
species harvested in other countries where the traditional sowing methods are still applied. The result isa lower price for the local product, which creates fUl'ther disincentive for Japanese farmers to maintainproper husbandry practices.
Farmed oysters in the coastal water column on the
Pacific side of northern Japan now utilize a totallydifferent phytoplankton population and have different, corresponding energy flow dynamics to those
harvested prior to the development of oyster culture
techniques. The time has arrived for a detailed analysis of current culture practices and a complete
reassessment of the direction of oyster culture in the
future.
In shallow areas less than 3 m deep, favorable conditions for the culture of oysters can still be found;higher water temperatures and more abundant phytoplankton during spring and autumn are just two of
the favorable parameters associated with these typesof culture grounds. A decrease in salinity, which occurs during the rainy season, is also an importantfeature. The culture of oysters in this zone combinedwith proper practices to control fouling wiP result ina quality product that is acceptable in the market
place.
It may be argued that such a practice will result in
lower production; while this may be correct, there
will be a corresponding shortening of growing timefrom 3 to 1.5 years and there is always the prospect ofincreased prices for a better quality product. Once
this is achieved the devel0pment of more efficientfarming techniques can be employed to fully utilize
the shallow waters.

French Oyster Farming: Seed Collection
and Industry Planning_
Rather than attempt to adapt existing techniques, a
totally new system was developed for the French oyster industry.
Although French oysters require higher salinity
than Crassostrea gigas, the new techniques establishedfor French oyster farming can be applied to the euryhaline Japanese oyster.
Improved seed collection technique was the first
factor considered because the current system provides extra space for competitors, which by harvest
time outweigh the oysters. The low survival rate of
attached spat (less than 10%) (Akio Oshino, ORI,
pers. comm un. Dec. 1988) further con tri bu tes tohigher levels of fouling organisms. One obvious solution would be to culture oyster seed alone using
single oyster collection techniques without substrate.
This, however, would require the use of very fine netting to hold an early-stage juvenile and could resultin a much worse fouling problem.
The use of crushed mussel shell as a substrate alleviated the above problems. A total of 500 g of shellparticles with sizes ranging from 10 to 20 mm wereused as substrate. These were spread in a monolayeron 20 plastic trays measuring 352 X 204 X 35 mm.
The trays are manufactured to hold small fish fordisplay purposes in supermarkets. The walls and bottom of each tray have 8-mm mesh pores. One set ofcollectors, consisting of 20 trays, was used to holdapproximately 60,000 shell pieces. The 20 trays weretightly stacked and tied with strings, which were set
through four corners to form a vertical hanging. A
total of 40 collector sets were introduced into a set
tank measuring 2.0 X 1.5 X 1.0 m. Along with thecollectors, approximately 1.5 million competent larvae were introduced with an attachment rate of two
seed per piece of shell. During two days of settling
time, the water temperature was maintained at 20° Cwith an electric heater and aeration. Approximately
6UO,000 French oyster seed of 20-mm size were collected in this manner in 1989. The survival rate withthis method was much higher and fouling was considerably less compared with the earlier use of scallop
shell (Oshino and Seki, unpubl. data). Thinning ofseeds in the trays was not required until the fourth
month when sorting was also undertaken. Foulingcontrol was achieved by soaking the seeds in 9% sodium chloride enriched seawater once at a month forSeki: Oyster Farming in Japan

10 minutes (Oshino and Seki, unpubl. data). The
seed production rate using this method was 14.8%
compared with 8.7% with the previous scallop cultch
method. Further work needs to be done to determine the optimal timing for removal of foulingorganisms, the best growing densities, and the associated labor requirements.
Consideration needs to be given to better hanging
techniques that would lead to better yields and lower
labor costs. Current raft licensing and associated culture densities are controlled by the relevant fisheries
cooperatives in each farming area. The Karakuwa
fisheries cooperative, which controls rafts adjacent to
ORI, allow 180 hanging lines to each raft; raft size islimited to 5.4 X 9 m. The length of each culture line
is variable dependent on water depth. From theabove information it is estimated that a combinationof 200 oysters and associated fouling organisms occur
in each cubic meter of the water column. Clearly, if
the existing planktonic food was available only to theoyster cultures, the total biomass of the culture could
be greatly increased. Because culturing space is at apremium, it appears logical that greater production
can only be achieved through better utilization of thewater column.
For example, each oyster farmer can expect a return of 600,000 yen or $4,138 from each raft in thethird year of production assuming 24,000 oysters/
raft. This equates to an annual gross income of
200,000 yen, or $1,379/raft. With an average raftnumber per farmer in northern Japan of 20, it can beseen that the annual income from oyster farming is
not sufficient to sustain a farmer and his family. This
current situation can be overcome by starting a program to educate farmers in the improved practices
previously mentioned.
Among the commercially manufactured containers, plastic trays with a 9-mm mesh size are the best
available to hold oysters economically. Each 705 X
462 X 57 mm container can hold 50 adult oysters
and a string of 10 containers can house 500. Thus,
the same raft area in the same depth of water column
can house about 70,000 oysters. The smaller size ofthe hanging medium allows for more efficient handling in terms of fouling control. Currently, work isunderway to determine such parameters as the optimum fouling control immersion period and the useof aeration to maintain adequate water movement.

87

In summary, Japanese consumers are appreciating
more exotic, rare, and higher quality seafoods suchas the French oyster-even the price is higher (thehigher price received for French oysters together
with the advanced growing system is a means to attract existing farmers into the redevelopment of theJapanese oyster industry). If the existing oyster industry is to survive and prosper, it must react to changes
in market forces. The development of the French oyster, together with associated advanced culture
techniques, is one such example of the need forchange in the existing industry.
Biological developments, such as triploid oysters,
are less practical in Japan because of the seasonal
tastes of the Japanese people. It is more likely that
biological advances will result in environmental
enhancement. It is absolutely vital that increased effort be placed in the area of water quality improvement in order to prevent the contamination ofoyster farming grounds and to maintain a safe, clean
environment for further development of the Japanese oyster industry.

Citations_

Fujiya, M.
1970. Oyster farming in Japan. Helgol. Wiss, Meeresunters,
20:464-479.
Imai, T.
1967. Mass production of molluscs by means of rearing thelarvae in tanks. VenusJpn.J. Malacol., 25:159-167.
1971. Through culture and its advance. Koseisha-Koseikaku,
Tokyo, 454 p. (In Japanese.)
Kafuku, T., and H. Ikenoue.
1983. Modern methods of aquaculture in Japan. Developments
in aquaculture and fisheries science, Vol. II. Kodansha
Ltd., Tokyo and Elsevier Scientific Publ. Co., Amsterdam,
216 p.
Seki, T.
1980. An advanced biological engineering system for abalone seed production. In Int. symp. coastal Pacific marine
life; 15-16 October 1979, Bellingham, WA, p. 45-54. Avail.
from Library, W. Wash. State Univ., Bellingham, WA 98225.
Seki, T. and A.S. Cuthbertson.
1988. Recent advances in abalone culture. In Proceedings
for the Aquaculture International Congress and Exposition,
Vancouver Trade & Convention Center, Vancouver, B.C.;
6-9 September, 1988, Vancouver, Canada, p. 377384. (Avail.) Aquacult. Int. Congo British Columbia Pavilion Corporation Box 31, 770 Pacific Blvd. South Vancouver,B.C., Canada.

Hormonal Regulation of Reproduction in Female Crustacea

HANS LAUFER and ELLEN HOMOLA
University oj Connecticut
Department oj Molecular and Cell Biology
Storrs, CT 06269-3125
andThe Marine Biological Laboratory
Woods Hole, MA 02543 U.S.A.

MATTHEW LANDAU
Deparlmenl of Marine Science
Slocklon Slale College
Pomona,lVJ.08240

Abstract
Basic knowledge of female reproduction is of primary importance to crustacean aquaculture. Our approach has been to investigate crustacean reproduction utilizing theapproach of comparative endocrinology. This paper reviews the hormonal regulation of
reproduction, focusing on eyestalk factors and the mandibular organ, and the target
tissues affected by their secretions. This approach has lead to the finding of methyl
farnesoate (MF), an unepoxidated juvenile hormone, in the circulatory system. MF is
produced by the mandibuiar organ (MO), a homologue of the insect corpus allatum,
and appears to be a crustacean juvenile hormone. Eyestalk ablation enhances crustacean
reproduction by removal of a gonad inhibitory hormone (GIH). Eyestalk removal in the
spider crab, Libinia emarginala, stimulates egg production and vitellogenesis innonreproductive females and increases MF production by the MO. Addition of sinus
gland extracts to Mas inhibits MF synthesis in vitro. Thus, there is a mandibular organ
inhibitory hormone in the eyestalk which may be similar or identical to GIH. The actionof GIH then may be on target tissues such as the hepatopancreas and ovary or may also
be on the MO. Whether this is the major action of GIH or one of its actions remains the
subject of current investigations. Additional interactions influencing the female reproductive system such as stimulatory factors from the brain and thoracic ganglion as well as
the role of biogenic amines are considered briefly. Other possible interactions mentioned in the literature are also indicated in this brief review of crustacean reproduction.

Introduction_

Decapod crustaceans represent a large, diverse biological group with significant potential as anaquacultural resource. Large-scale penaeid shrimp
culture industries currently exist in Asia and Central
America. Crayfish are grown in the United States, Europe, and Australia, and prawns of the genus
Macrobrachium are cultivated in many tropical environs. The culture of lobsters and other species may

become viable endeavors in the future. In somecases, such as the culture of penaeid shrimp, the production of sufficient healthy seed organisms has
hindered farming. A lack of understanding of the
reproductive process has forestalled selective breeding. We feel that a better understanding of thehormonal mechanisms that regulate the reproduction of these valuable resources is fundamental tosuccessful aquaculture.
During the past two decades our understanding ofcrustacean reproductive endocrinology, especially

89

90

NOAA Technical Report NMFS 106_sinus glandcommissural-medulla terminalis~

y organX organpostcommissuralkcl-----mandibular organ

organ

18

.; , '. ~ '-subesophageal ganglion

pericardialorgans~;::> "....,I;:..II~~. . .~:. . ~ ·~~..=-;--thoracic second roots

thoracic ganglion'iLsl"I~::~;:~n~~~;I~anglia
IIr1 ~('J!
cannecllve.
&

Figure 1

Major endocrine and neuro-endocrine structures of generalized
female Crustacea. Included are
the organs important for female
reproduction, the eyestalk sinus
gland x-organ, the mandibularorgan, V-organ, and thoracic
ganglion.
that of the female, has grown steadily. This is in parta result of the use of the comparative approach,whereby our understanding of other crustacean
groups and their close relatives, the insects, is applied
to the decapods. Major sites of endocrine activity are
shown in Figure 1. In this report we concentrate on
eyestalk factors and the mandibular organ, and thetarget tissues affected by their secretions. Additional
endocrine interactions will be discussed as they relate
to the reproductive process.

A Crustacean Juvenile Hormone_

The role of the terpenoid hormones, collectively
known as the 'Juvenile hormones" (IHs) or juvenoids(Fig. 2), has been well established in insect reproduc-tion. The JHs are produced in a pair of small endocrine glands, the corpora allata (CA), usuallyposterior and ventral to the brain. The JHs appear toplaya major part not only in the development of theinsect larval stages (hence the name 'Juvenile") but
also in the regulation of reproduction (Downer andLaufer 1983), first by stimulating the ovary to matureand then by stimulating the fat body to make yolk
proteins. Further, JH seems to alter the oocyte membrane so that the large yolk proteins synthesized by
the fat body can be taken up by the developing egg.
Since both arthropod subphyla, the Insecta andCrustacea, are already known to regulate moltingwith identical hormones, 20-hydroxyecdysone(Karlson 1956; Hampshire and Horn 1966), we (efLaufer et al. 1987c) speculated that the crustacea
might also have a functioning JH. This speculation

Laufer et al.: Hormonal Regulation of Reproduction in Female Crustacea

91

METHYL (2E,6E)-FARNESOATE

Figure 2(10B)JUVENILE HORMONE III (R= CH )S
(10B)JUVENILE HORMONE III ACID (R=H)

was supported by a considerable literature. Gomez etal. (1973) found that the cyprid to spat metamorphosis of the barnacle, Balanus galeatus, could be
inhibited by two structural analogs of the JHs,
methoprene and hydroprene. Similar studies werecarried out by Cheung and Nigrelli (1973) andTighe-Ford (1977). Landau and Finney (1977)
showed thatJH analogs are active, but that mevalonicacid, a precursor of JH, had no effect. Furthermore,precocene II, a compound that destroys the CA in
insects and thereby halts the production of JH, wasshown to strongly inhibit the hatching of barnacle
embryos treated in the early stages of development(Landau and Rao 1980). Such effects were revealedin other crustaceans as well. JH analogs were demonstrated to cause morphological abnormalities inmegalopa larvae of the crab Rhithropanopeus harrisii(Costlow 1977), and to alter the morphology andlength of development of the American lobster
Homarus americanus (Hertz and Chang 1986). Injection of a juvenile hormone into larval H. americanus
results in delayed metamorphosis and morphometricvariability in newly metamorphosed animals(Charmantier et al. 1988).
Beside affecting development, JHs and their analogs were also shown to affect reproduction.
Inhibition of reproduction in the cladoceran Daphnia
magna was shown by Templeton and Laufer (1983),
and inhibition of oogenesis and spermatogenesis
were demonstrated in the mud crab R. harrisi (Payen
and Costlow 1977), and the spider crab Libinia
emarginata (Hinsch 1981). Paulus and Laufer (1982)

The structural formula of insect juvenile hormone UHlII) and its metabolic breakdown
product, JH acid, are compared with a similarmolecule, methyl farnesoate (MF), present incrustacean hemolymph and synthesised by themandibular organs.

and Paulus (1984) showed that when methoprene
was injected into intact and ablated female crabs,
Carcinus maenas, the ovaries became enlarged.
While these experiments demonstrated that Crustacea were sensitive to JH, it was not established thatcrustaceans possessed endogenous juvenoids. Using
an insect bioassay, Schneiderman and Gilbert (1958)
detected some JH activity in the eyestalks of Crustacea, but the chemical nature of the extract was notinvestigated. When thoracic ganglia or mandibularorgans (MOs) were implanted into immature L.
emarginata, there was a stimulation of vitellogenesis(Hinsch and Bennett 1979; Hinsch 1980). Based onmorphological studies, the MO was suggested as apossible homolog to the insect CA (Chaudonneret
1956; Le Roux 1968; Byard et al. 1975). If the MO
was a structural homolog of the CA, it was reasoned(Laufer et al. 1987c) that it might produce one of thejuvenile hormones or a similar compound.
Methyl farnesoate (MF), the unepoxidated form of
JHIII (Fig. 2), was detected in the hemolymph of L.
emarginata using gas chromatography/mass spectrometry and selected ion monitoring (Laufer et al.
1987c) and in the hemolymph of crayfish Orconectes
virilis and H. americanus as well (Tsukimura et al.
1989). When the MOs of crustaceans were incubatedin physiological saline supplemented with a labelled
precursor, [methyPHJ methionine, it was identified as
the source of the MF (Laufer et al. 1986, 1987c; Borst
et al. 1987). Mandibular organs from 12 species of
Crustacea are known to produce MF, most are
Decapoda, but also a barnacle, Balanus nubilus, pro-

92

NOAA Technical Report NMFS 106- -......~Positive effect----~Inhibitory effect-- -I---Effectofeyestalk

Y -organ(ecdysone)

Induces secondary sex charactersij
duces MF. Based on a similar set of in vitro studies,
Tobe et al. (1989) has suggested that farnesoic acidrather than MF is the major product of the MO in
the mud crab Scylla serrata although only MF was detected in the hemolymph.
In the insects, JH produced by the CA is reversibly
bound to a "carrier protein" (Downer and Laufer
1983), which functions to protect the hormone fromenzymatic degradation and to increase its solubility
in the hemolymph. Likewise, it appears that a similarprotein, of about 40 kd, exists in the hemolymph ofthe lobster (Prestwich et al. 1990) and perhaps inother crustaceans. In L. emarginata, MF does not appear to be converted to JHIII in the hemolymph orby a variety of assayed tissues; it is, however, catabolized to farnesoic acid by a number of tissues,
especially the hepatopancreas, but not by the hemolymph (Laufer and Albrecht 1990).
Like JH in insects, the production of MF seems to
be related to reproduction. In female L. emarginata.
the rate of MF secretion by MOs in vitro is correlated

Figure 3
Major endocrine glands and their target tissues
involved in crustacean female reproduction. Indicated with solid lines and arrows arestimulatory effects. Inhibitory interactions areindicated with dashed lines and arrows. Notethat the inhibitory interactions from the eyestalksinus gland x-organ complex can be removed byeyestalk ablation. These include both the inhibitory effects of the molt inhibitory hormone(MIH) on the V-organ which produces ecdysones
and the gonad inhibiting hormone (GIH),
which according to the literature may be similaror the same as the vitellogenin inhibitory hormone (VIH) and which may also inhibit themandibular organ (MO-IH) as well as other target tissues such as the ovary and hepatopancreas(HP). The brain and thoracic ganglion (ThG),
according to some reports, may also stimulate or
inhibit reproduction, or do both. Biogenic
amines such as serotonin and octopamine havebeen shown to inhibit the MO.

with the reproductive cycle; during the reproductive
season, they produce eggs in a regular cycle every
19-20 days. The MO is least active immediately after
the eggs are layed and most active during vitellogenesis; it continues its activity until just before theembryos hatch and oviposition of a new brood commences (Laufer et al. 1987c). Hinsch (1980)
stimulated ovarian development in juvenile female
spider crabs by implanting adult MOs, so the MOseems to be a stimulus for the vitellogenic cycle in L.
emarginata. Vogel and Borst (1989) injected MF into
eyestalkless female L. emarginata, which according to
the findings of Panouse 1943 would enlarge theirovaries, and measured the change in hemolymph
vitellogenins by enzyme-linked immunosorbent assay(ELISA); a single injection (0.5 to 2.0 ng of MF)
cansed a modest rise in the hemolymph protein titer.
Similarly. Paulson and Skinner (1988) suggested that
MF increased the in vitro synthesis of specific integumentary tissue proteins of the land crab Gecarcinus
lateralis. The proteins stimulated were different from

Laufer et al.: Hormonal Regulation of Reproduction in Female Crustacea

those stimulated by 20-hydroxyecdysone, while JH
stimulated fewer proteins than MF.

Relationship of the Mandibular Organ andEyestalk to Reproduction_
The observation by Panouse (1943) that eyestalk ablation enhances ovarian growth, vitellogenesis, andoviposition has already been mentioned here. Thus,
reproduction in crustaceans appears to be under the
inhibitory control of an eyestalk factor known as the"gonad-inhibiting hormone" (GIH), which may be
the same as the "vitellogenin-inhibiting hormone"(VIH) (Brown and Jones 1949; Quackenbush andHerrnkind 1981; Soyez et al. 1987). The eyestalk is asource of many other hormones, including the "moltinhibiting hormone" (MIH) which directly inhibits
ecdysterone production by the V-organs in culture(Mattson and Spaziani 1985, a and b; Schoettker andGist 1990) (Fig. 3). Recently a molt-inhibiting hormone has been purified and its amino acids
sequenced from the lobster H. americanus. The peptide has 61 % sequence identity with crustacean
hyperglycemic hormone, (CHH) from Carcinus
maenas, and shows significant CHH activity when injected into H. americanus (Chang et al. 1990). It
appears that crustaceans alternate between production of MIH and GIH (Anilkumar and Adiyodi 1980;
Chang 1984). Fyhn et al. (1977) suggested that inbarnacles 20-hydroxyecdysone may be the functional
GIH, and Kallen and Meusy (1989) have advanced
the theory that GIH is similar in structure, but distinct from CHH. Quackenbush and Herrnkind(1983) and Charniaux-Cotton (1985) demonstrated
that MIH and GIH could be separated chromatographically.
Presumably because the eyestalk is the site of GIH
production, the ablation of the eyes talk may cause
increased growth of the ovarian tissue (Panouse
1943; Brown and Jones 1949; de Leersnyder andDhainaut 1978). From an aquacultural perspective,
eyestalk ablation may not be a desirable method of
increasing the stock's reproductive potential. The resulting embryos often are of inferior quality(Anilkumar and Adiyodi 1985; Choy 1987) because
either hormonal imbalances are created by eyestalkablation or GIH release is uncontrolled, or both. It
has been suggested that the targets of GIH are theovaries and hepatopancreas which are the sites ofyolk protein synthesis (Fig. 3) (Paulus 1984; Paulus
and Laufer 1987; Quackenbush 1989). Quackenbush
and Keeley (1987) showed that partially purified eyestalk extracts from the shrimp Penaeus vannamei

93

could inhibit crab Uca pugilator ovarian synthesis ofvitellogenin in vitro, while Eastman-Reks andFingerman (1984) also found that V. pugilator eyestalk extracts inhibit protein synthesis in cultured
ovaries of the crab.
Alternatively, GIH may have non-ovarian targets, orin fact there may be more than one eyestalk factor
which inhibits ovarian growth. In the insects, the production of JH is regulated by neuropeptides:
allatotropins which stimulate the synthesis or release,or both, of hormones (Kataoka et al. 1989), andallatostatins and allatohibins which inhibit hormone
synthesis/release (Girardie 1983; Woodhead et al.
1989). If there is synthesis of one of the inhibitors,
thus preventing JH production and release, there will
be no ovarian development; that is, the allatohibins
and allatostatins are functional insect analogs of crustacean GIH. Laufer et al. (1986 and 1987a,b) foundthat a water-soluble, heat-stable eyestalk factor(s) inhibits the in vitro synthesis of MF (Fig. 3). Based on
its extraction properties, it seems likely that this factor is a small peptide. We have termed this peptidethe "mandibular organ-inhibiting hormone" (MOIH). Laufer et al. (1986, 1987a, b, and c), showedthat the levels of MF in the hemolymph of the spider
crab increased dramatically after eyes talk-ablation,
which was also observed in H. americanus and O.
virilis (Tsukimura et al. 1989).
In addition to the eyestalk GIH/MO-IH, there isanother, the "gonad-stimulating hormone" (GSH)that is reported to be produced in the thoracic ganglion, and which may contribute to the control ofreproduction (Fig. 3). Otsu (1960, 1963) first suggested its existence because eyestalk ablation caused
precocious ovarian growth in adult crabs, Potamon
dehaani, but not in juveniles; he reasoned that not
only was the absence of GIH required for ovarian
growth, but the presence of a stimulatory hormone
was also necessary. When adult thoracic ganglia were
implanted in eyestalk-ablated juveniles the ovaries began to mature. These experiments were confirmed byHinsch and Bennet (1979) using L. emarginata.
Gomez (1965) found that the thoracic ganglion, as
well as the brain, stimulated growth of reproductive
tissue in Paratelphusa hydrodromous; similar results
were reported by Takayanagi et al. (1986) in the
shrimp Paratya compressa. Extracts of the thoracic ganglia of V. pugilator sampled during their reproductive
season stimulated ovarian growth in intact and eyestalk-ablated crabs, but extracts of the thoracic
ganglia of crabs outside the reproductive season resulted in inhibition of ovarian growth in ablatedorganisms (Eastman-Reks and Fingerman 1984).
More recently Yano et al. (1989) reported in a small

94

NOAA Technical Report NMFS 106

number of cases that implants of H. americanus
thoracic ganglia into non-reproductive Penaeus
vannamei stimulated ovarian maturation.

Mechanisms of Methyl Farnesoate
Regulation and Action
In an effort to understand the inhibitory action of
the eyestalk on the MO, we surveyed the literature on
the control (secretion or synthesis) of JH by insect
neuropeptides. A peptide extracted from the insect
corpora cardiaca (CC) was reported by Applebaumand Moshitzky (1986) to inhibit yolk production in
Locusta migratorioides; this peptide reacted with an antibody to adipokinetic hormone (AKH). Since AKH
is very similar in structure to crustacean "red-pigment-concentrating hormone" (RPCH) (Fernlund
andJosefsson 1968; Gade 1990), and because inhibition of yolk protein synthesis might be the result of a
substance that acts directly on the insect CA, it was
decided that the effect upon MF synthesis should bedetermined. Unexpectedly at 10-6 M RPCH significantly stimulated, rather than inhibited, the synthesisof MF by the MOs of crayfish Procambarus clarkii(Landau et al. 1989). We could mimic the effect of
RPCH by replacing it with the Ca 2 + ionophore
A23187, and synthesis could be inhibited by culturing
the tissue in Ca 2+-free media or including lanthanum,
which replaces Ca 2 + on cell surfaces (Weiss 1974), in
the culture medium. Lambert and Fingerman (1979)
had suggested that the RPCH might act as a Ca 2+
ionophore, and Ca 2 + seems to be involved in theregulation of JH synthesis by the CA (Aucoin et al.
1987; Kikukawa et al. 1987; Dale and Tobe 1988).
Furthermore, we found that "pigment dispersing hormone" (PDH) (Rao et al. 1985) at 10-7 M significantly
inhibited MO synthesis of MF in P. clarkii (Landau etal. 1989). It is interesting to note that Mangerich etal. (1986) found cells in the thoracic ganglia of the
crab Carcinus maenus that contained RPCH-like molecules using immunocytochemical techniques.
Kravitz and collaborators (Beltz 1988) reported
that the biogenic amines, serotonin and octapamine,playa significant role in determining mating behavior in the lobster H. amencanus. The synthesis of MF
by crab MOs also appears to be regulated in part bycertain biogenic amines. Disaggregated MO cells
from Libinia emarginata appear to be unaffected bydopamine and are only slightly inhibited byoctopamine; however, serotonin inhibited MF synthesis by 20-35% at 10-8 M, suggesting that it may
function as a neuroregulator (Homola et al. 1989).
Octopamine stimulates JH synthesis from Locusta

migratoria CA in vitro (Lafon-Cazal and Baehr 1988).
Biogenic amines are known to affect the levels ofcAMP, and to a lesser extent cGMP, in the insect corpora cardiaca, the site of AKH synthesis (Pannabecker
and Orchard 1986; Gole et al. 1987). Tsukimura et al.
(1986) treated lobster MOs with eyestalk, brain, andthoracic ganglion extracts; they found that the eyestalk extracts significantly increased the level ofcGMP, but not cAMP, in the MO, while the otherextracts had no effects.

Other Hormones Regulating
Reproduction_

The molting hormones, ecdysteroids, are known toplaya role in insect reproduction, and therefore may
act in a similar fashion in crustaceans. We have already alluded to the apparent molting-reproduction
antagonism. Blanchet et al. (1979) found 20hydroxyecdysone in the ovaries of the amp hi pod
Orchestia gammarellus and speculated that, althoughmolting hormones seem to influence the growth ofthe oocytes, vitellogenesis itself was unaffected. It waslater demonstrated that if the Y-organ of O.
gammarellus is destroyed at the beginning of the molt
cycle the ovary will not develop, and if it is destroyed
during ovarian growth the synthesis of vitellogenins
stops (Meusy and Charniaux-Cotton 1984). Suzuki(1986) also found that the Y-organ was required for
oocyte growth in the isopod Armadillium vulgare. In
the decapods, Lachaise et al. (1981) showed an increase in the levels of another molting hormone,ponasterone A, in ovaries of the crab Carcinus maenas
during ovarian maturation. We have shown an accumulation of ecdysones in Libinia emarginata oocyteswith release of ecdysteroids during embryogenesis(Laufer and Deak 1990).
Testosterone, progesterone, and pregnenolone,
the vertebrate sex steroids, have been identified in
the gonads and hemolymph of Astacus leptodacylus
and H. americanus (Burns et al. 1984 and Ollevier etal. 1986, respectively). Couch et al. (1987) found significant levels of estradiol and progesterone in theMOs of H. americanus, and Yano (1985, 1987) reported that vitellogenesis in two species of shrimp,
Metapenaeus ensis and Penaeus japonicus, could be
stimulated by injections of progesterone and 17-ahydroxyprogesterone. Vitellogenin synthesis in theisopod, Jdotea balthica basten can be stimulated by human chorionic gonadotropin (hCG) (Souty and
Picaud 1984); in the prawn Crangon crangon, vitellogenesis was also stimulated by hCG (Bomirski and
Klek-Kawinska 1976), and in another prawn, Caridina

Laufer et al.: Hormonal Regulation of Reproduction in Female Crustacea

catadhari, it not only stimulated growth of oogonial
and follicular cells but also increased the rate of yolk
deposition in the growing oocytes (Sarojini and
Persis 1988). A molecule very similar to hCG has
been identified in the prawn Palaemon serratus withthe aid of a radioimmuno-assay with an hCG antibody (Toullec and Wormhoudt 1987). Follicle-stimulating hormone (FSH) and luteinizing hormone(LH) were also shown to increase the rate of ovariandevelopment in the shrimp Crangon crangon(Zukowska-Arendarczyk 1981).

Acknowledgments
The research reported here was supported in part by
grants from the Sea Grant College Program, NOAA,
NA-AA-D-SG101, a National Research Service Award
from NIH, and the Lady Davis Trust. We wish also to
acknowledge the artistic talents of Mary Jane Spring
and the advice and technical assistance of Dr. FrankMauri of the University of Connecticut Biotechnology Center.

Citations_

Anilkumar, c., and K.C. Adiyodi.
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Reproduction in Cultured White Sturgeon, Acipenser transmontanus

GARY P. MOBERG and SERGE I. DOROSHOV
Department ofAnimal Science and Aquaculture and Fisheries Program
University of California, Davis
Davis, CA 95616

Abstract
The value of sturgeon flesh and the demand for caviar makes the culture of sturgeon
an exciting opportunity for aquaculture worldwide. However, a m~or problem in successfully adapting sturgeon to aquaculture has been the failure to produce mature domestic
broodstock for egg and milt production. Even the industrial-scale sturgeon hatcheries in
the U.S.S.R., which have been in operation since the late 1950s, producing fingerlings
for mitigation, depend upon harvesting wild broodstock for eggs and milt. Recently in
California, several private aquaculture ventures have begun to raise white sturgeon(Acipenser transmontanus) for the commercial market. While successful in producing marketable fish, they have not been able to raise a reliable source of domestic female
broodstock and must continue to harvest wild broodstock for eggs. Females raised inculture remain reproductively immature until at least 10 years of age (the age of theoldest cultured females in California). Reproduction in these females is arrested justprior to the initiation of vitellogenesis. In this report, we discuss our observations of the
reproductive maturation of captive sturgeon broodstock and review our previous efforts
to manipulate their endocrine system to promote earlier sexual maturation. It appears
that the hypothalamic-pituitary-gonadal axis of the cultured females is not secreting
sufficient gonadotropins to induce the full maturation of the reproductive system. We
have attempted to induce further reproductive maturity by hormone treatment. While wehave been successful in inducing vitellogenesis in these immature cultured females by
estradiol administration, the ovarian follicles will not incorporate the vitellogenin.

Introduction_

Because of the commercial value of its flesh and theuse of its roe for caviar, the demand for sturgeon has
been intense and has resulted in a severe exploitation of wild sturgeon stocks worldwide. The impact of
this commercial exploitation on the sturgeon population has been further exacerbated by the fish's low
reproductive rate. The females of most of the commercially valuable sturgeon species do not reach
reproductive maturity until 15 to 20 years of age.
Once a sturgeon has spawned, she may not reproduce again for several years. Further reducing
fecundity is the sensitivity of these animals' reproduction to a variety of environmen tal disturbances,
especially pollution and the damming of the rivers
where they migrate to spawn. The combined result offishing pressure and the low reproductive rate has
been a dramatic decline of sturgeon stocks through-out the world, with some species approaching extinction. One approach to assuring the survival of these
species is to raise sturgeon in aquaculture operations,providing sturgeon for both mitigation and as asource of flesh and caviar.

History of Sturgeon Culture
By the turn of the century, commercial exploitationof sturgeon in Russia had already severely diminishedwild stocks. Russia has had the largest resources ofsturgeon species in the world, where they reach unusually high abundance in the continental brackish
water of the historic Sea of Tetis, the Caspian, Aral,
and Azov Seas. The latter two are now lost as a sturgeon habitat, serving as one of the worst examples ofenvironmental mismanagement. The largest sourceof sturgeon, the Caspian Sea, still holds substantial

99

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NOAA Technical Report NMFS 106

sturgeon resources, but their fate and future commercial exploitation are endangered by the loss ofnatural reproduction and the increasing pollution of
the sea.
The first attempt to artificially reproduce anadromous sturgeon was made in Russia by Borodin in
1898 (Borodin 1898). The first experimental hatchery operations were established in the Caspian Sea
basin (Volga and Kura Rivers) in 1937-38. Hcwever,industrial-scale sturgeon hatcheries did not evolve
until the late 1950's when all major spawning rivers
were dammed and most of the spawning grounds forsturgeon were lost (Kozin 1964). Research lagged onsturgeon nutrition and development of intensive juvenile culture, however, resulting in over dependenceof Soviet sturgeon culture on unreliable methods
that even today require the harvesting of wild stockfor eggs and milt. Currently the U.S.S.R. 's sturgeon
culture system annually produces 70-100 million
fingerlings, which are used to stock the Caspian andAzov Seas, with an estimated survival to adulthood of
1-3% (Marti 1979). It is clear that this approach toculture has succeeded in temporarily maintaining thecommercial exploitation of wild sturgeon in the
U.S.S.R. (approximately 20,000 metric tons annual
catch). However, ecological changes in the CaspianSea, environmental pollution, and the continuationof the commercial "caviar" fishery may endanger existing stocks.
Extensive commercial fisheries for sturgeon in
North America were established during 1880--90 in
the Delaware River, Great Lakes, and along the Pacific Coast. The commercial catches peaked during
1890-1900, almost approaching the levels caught inRussia; however, the catch rates rapidly declined toinsignificance by 1910 (Ryder 1890; Bajkov 1949;
Harkness and Dymond 1961; Semakula and Larkin
1968; Galbreath 1985; Smith 1985). Since the earlypart of this century, regulations for most North
American sturgeon stocks here prohibited or effectively limited commercial catch. Nevertheless, there
is still no single example of complete stock recovery
to previous historical levels. It is also interesting thatwhile the commercial fishery for sturgeon has practically ceased in North America, the sport fishery,especially for the white sturgeon in the Pacific Northwest, has become a major user of sturgeon resources.
The dramatic and rapid decline in sturgeon
fisheries prompted early attempts to artificially feproduce sturgeon for stock replenishment. Hatchery
work with atlantic (Acipenser oxyrhynchus) , lake (A..
fulvescens) and shortnose (A. brevirostrurn) sturgeons
was pioneered during 1890-1910 (Ryder 1890; Post
1890; Stone 1901; Carter 1904; Meehan 1909; review

by Leach 1920). These attempts were only partly successful because of the difficulties wi th capturing
ovulatory females and incubating the highly adhesive
sturgeon eggs. Further development of sturgeon culture in orth America apparently waned until the
1960's, when hormonally induced spawning was successfully used to breed paddlefish Polyodon spathula(Purkett 1963). Interest in sturgeon culture revived
in the early 1980's when lake, atlantic, and white sturgeon (A. transmontanus) were spawned and raised to
the fingerling stage in various university laboratories
and state hatcheries (Smith et al. 1980; Doroshov etal. 1983; Czeskleba et al. 1985). At this time, only theartificial propagation of the white sturgeon has been
established as a complete technology that includes
induced spawning, egg incubation, and intensive
rearing of juvenile and adult fish (Doroshov 1985;
Conte et al. 1988).
The role of sturgeon culture in the restoration andreplenishment of North American stocks is still insignificant, in spite of the potentially high efficiency ofhatcheries to produce sturgeon juveniles. Most efforts in wild stock management are directed atenvironmental protection and the stringent controlof the predominant recreational fisheries. The major
stimulus for current attempts at developing sturgeon
culture originates from the development of commercial aquaculture in the United States and Western
Europe.

Culture of White Sturgeon
in California_

The Pacific Northwest is the home for two sturgeon species-the white sturgeon and the green
sturgeon, A. medirostris. The biology of green sturgeon is practically unknown. Although important insome geographic areas, this species is believed to beof inferior value as a food fish. In contrast, the meatof the white sturgeon has a very high market value
and this species is still harvested by a small commercial fishery in the lower Columbia River. The whitesturgeon is taken by sport-fishermen throughout the
Pacific Northwest. As a result, its harvest by recreational fisheries in the Columbia and Sacramento
Rivers has reached a rate equal to that of the earlier
exploitation by commercial fisheries (Galbreath1985), resulting in a significan t threat to the wild
population.
White sturgeon stocks live in the estuaries of threemajor rivers, Fraser, Columbia, and Sacramento. Thisis one of the largest sturgeon species, reaching amaximum recorded size of 1800 pounds (Moyle and_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Moberg and Doroshov: Reproduction in Cultured White Sturgeon

Cech 1988). Males reach first sexual maturity at 1015 years of age, while females mature at 15-20 yearsof age. Ripe fish migrate into the rivers and spawn
from March (Sacramento River) to June (Columbia
and Fraser Rivers), at water temperatures of 1416° C. Spawning grounds are now limited to rivers
below the dams, although there are small reproducing populations in the reservoirs on the ColumbiaRiver (Galbreath 1985).
Preliminary information was recently obtained on
the reproductive cycle of white sturgeon females in
the San Francisco Bay (Chapman et al. 1987;
Chapman 1989; Doroshov et al. 1990). The most important findings were the high individual variability
in the age of first ovulation (12-22 years) and theapparently biennial vitellogenic cycle which appears
to be strictly controlled by seasonal factors (photoperiod and, probably, temperature). It was also notedthat the females undergoing vitellogenesis exhibited
elevated plasma concentrations of estrogen.
The culture of white sturgeon in California was initiated in the early 1980's by several commercial
ventures. Wild fish are captured during their spawning migration in the Sacramento river. Ova andsemen are obtained by injecting fish with either carp
pituitary extracts or synthetic gonadotropin-releasing
hormone analog (Lutes et al. 1987; Conte et al.
1988). The fertilized eggs are incubated in jars; larvae and fingerlings are raised in tanks on salmonid
diets (Hung 1989).
Initially, the market for these small scale culture
operations was the sale of fry to other growers and to
aquarium retailers. During the past three years, several commercial ventures have established thegrowout of high market-value food fish. Sturgeon areraised in tanks, raceways, and earthen ponds andmarketed (primarily to restaurants) at body weight
6-8 kg as 2-3 year-old fish. The growout system is also
well established in several Western European ventures that produce highly prized smoked whitesturgeon (such as Agroittica Lombarda, Calvisano,
Italy). The white sturgeon is superior to many otherspecies in the high quality of their meat, their acceptance of artificial diets, their tolerance to highd~nsity and, most importantly, their fast growth rates
at water temperatures in the range of 20-23° C.
The major problem in developing a white sturgeon
culture has been the lack of domestic broodstocks
and dependence of fish farmers on wild broodstock
capture, which is not only unreliable but severely restricted by government regulation. As a result, several
aquaculture ventures in California have initiated therearing of captive broodstocks obtained from hatchery-produced fish (Fl generation) and they now

101

retain a substantial number of fish, the oldest ofwhich have reached 10 years of age and a bodyweight of 50 kg (Anonymous 1990). These fish areraised at low density in outdoor tanks and raceways
and are fed different types of salmonid diets(Silvercup, Rangen, Moore-Clark). In 1987, eight
farms started collaborative research with the University of California at Davis, in an attempt to establish
artificial reproduction in captive stocks and to improve sturgeon broodstock management. The major
objective of this research is to decrease the age at
which the females reach reproductive maturity.

Reproductive Maturation of
Captive Broodstock_

During the past two years we have performed biopsies on the gonads and collected plasma from morethan 1,000 cultured sturgeon. Paraffin sections of gonadal tissue, stained by hemotoxylin and eosin(males) or by periodic acid-Schiff stain (females),have been examined microscopically. These samples
have provided us with the first insight into the gonadal maturation of cultured sturgeon and has
enabled us to identify the major problems that arepreventing these animals from achieving normal reproduction in culture.
About 90% of the captive males mature by 3-4
years of age at a body size of 5-6 kg. The stages oftesticular development are shown in Figure 1, (Athrough C). Spermatogenesis of these cultured males
is synchronous, occurring during the summer andearly fall. By October-November, testicular cysts contain ripe spermatozoa, and it is possible to induce
spermiation by hormone injections. The quality of
captive male semen does not differ from the qualityof the semen collected from wild males (Chapman1989), and most farmers today use only captive males
for their hatchery production. It appears that the testicular cycle is annual and that the cultured males
spermiate each year. The only reproductive problem
that has been observed in cultured males is an apparent negative effect of warm (above 16° C) temperature on the final phase of the testicular cycle.
These temperature conditions are often observed insome California farms using underground water supplies that have a constant temperature of 18-20° C.
The colonies of males maintained under such conditions appear to undergo rapid testicular regression
and will not spermiate during the following spring
spawning season U. Michael, Sierra Aquafarms,
Elverta, CA 95626; and Ken Beer, The Fishery, Galt,
CA 95632, pers. commun. 1989).

102

NOAA Technical Report NMFS 106

Figure 1
Selected stages of gametogenesis in captive white sturgeon: (A), (B) and (C) show development of testicular germ cells(hematoxylin and eosin stains, microscope magnification 315x); (D) and (E) show development of the ovarian follicle
from previtellogenic (0) to early vitellogenic (E) stage, the latter observed in very few captive females (magnification50x); (F) peripheral area of early vitellogenic follicle possessing differentiated granulosa layer and egg chorion (periodic
acid-Schiff stain, magnification 315x). All fish were sampled in November 1988, at age 3-5 years (males) and 5-7 years(females).

In contrast to the males, most of the captive females remain reproductively immature for up to 10
years of age, which is the age of the oldest cultured
females in California. These captive females havereached a body size (40-50 kg) which is well above
the size of wild females when they first reach reproductive maturity (Chapman 1989). The great

majority of cultured females stop reproductive maturation just prior to the initiation of vitellogenesis.
Their ovaries complete gonial proliferation by four
years of age, but the ovarian follicles remain in arefractory condition characterized by the failure todifferentiate granulosa cells (Fig. 1, d through f).
This reproductive block could result from a number,

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Moberg and Doroshov: Reproduction in Cultured White Sturgeonof environmental and husbandry factors. We are currently focusing on the role of seasonal fluctations inwater temperature and nutrition of the broodstock as
the two most important factors that may be affecting
reproduction in these animals.

Endocrine Manipulationof Vitellogenesis_

Since the sexual maturation of the cultured female
sturgeon is arrested at the previtellogenic stage, wehave attempted to induce vitellogenesis and trigger
further ovarian maturation by hormone manipulation (Moberg et al. 1990). We implanted cellulose
pellets containing various doses of estradiol into fourto seven year-old white sturgeon females that werepart of the first generation of wild broodst~ck
spawned at the University of California at Davis.
These fish had been raised and maintained in outdoor freshwater tanks and fed commercial trout
broodstock diet (Murray Elevators). The estradiol implants not only elevated plasma concentrations ofestrogen in these females, but also stimulated the synthesis of vitellogenin, raising the plasma concentration of vitellogenin to levels that were 6 to 10
times higher than the mean values observed invi tellogenic wild females (Chapman et al. 1987). In
spite of these high concentrations of plasma
vitellogenin, there was no evidence of any uptake ofvitellogenin by the ovarian follicles, suggesting to us
that the ovarian follicles were not sufficiently developed to incorporate the vitellogenin found in theplasma.
These findings suggest to us that the failure of thecultured female sturgeon to mature beyond theprevitellogenic phase reflects a failure of the hypothalamic-pituitary-gonadal (HPG) axis to initiate thenext stage of reproductive maturity. Since folliculardevelopment as well as estrogen synthesis is dependent upon the HPG axis to secrete sufficient
gonadotropins to stimulate the next stage of ovariandevelopment, the failure of both events to occur inthe cultured females suggests that insufficient gonadotropin is being secreted in these animals. To
address this problem, we sought to stimulate gonadotropin secretion by treating the animals withsynthetic gonadotropin releasing hormone(GnRHa), either alone or in combination with estradiol administration. GnRHa treatment by itself did
not induce estrogen synthesis and as a result had noeffect on vitellogenin synthesis. The combined treatment of estrogen and GnRHa did result insignificantly higher plasma concentrations of estrogen than did estradiol treatment alone, suggestingthat this combined treatment may have stimulated an103

increased secretion of endogenous gonadotropins
which in turn stimulated endogenous estrogen synthesis. Regardless, the combined estrogen and
GnRHa treatment still did not initiate sufficient follicular differentiation (i.e., granulosa cells) to permit
the incorporation of vitellogenin. Since this combined treatment lasted for only 73 days, it is possiblethat a longer treatment period might have initiatedfurther follicular development (Moberg et al. 1990).
From our current understanding of the endocrine
control of sturgeon reproduction, it seems that manipulation of the endocrine system still is the mostviable approach to inducing reproductive maturationin the cultured female sturgeon. However, to accomplish this task, it will be necessary to refine thehormone treatment practices.

Conclusions
The value of sturgeon flesh coupled with the demandfor caviar makes the culture of sturgeon an exciting
opportunity for aquaculture not only in California,
but throughout the world. The adaptation of this species to aquaculture is not only necessary for the full
realization of its market potential, but may be themost viable strategy to insure the survival of thisunique group of fish. While remarkable progress has
been made in developing procedures to culture sturgeon, one major problem remains, full reproductive
maturation of the cultured females within a timeframe that makes sturgeon culture a viable economic
enterprise.
The failure of the cultured females to reach full
reproductive maturity appears to result from failureof the pituitary to secrete sufficient gonadotropins to
induce final ovarian follicular development. The
most viable approach to solving this problem seems
to be the utilization of hormone treatments to either
induce the secretion of gonadotropins or to duplicate the effects of these pituitary hormones. The
success of such hormone therapy would not only establish full reproduction in culture but would
provide a means to induce reproduction in females
at a much younger age than that which occurs in thewild, making the culture of sturgeon a more economically feasible enterprise.

Acknowledgments_

This work is a result of research sponsored in partby NOAA, National Sea Grant College Program, Department of Commerce, Under grant number
NA85AA-D-SGI40, project number R/A-73, throughthe California Sea Grant College Program, and 10
part by the California State Resources Agency.

104

NOAA Technical Report NMFS 106

Citations_

Anonymous.
1990. Third annual report of domestic white sturgeon
broodstock research and development program. Aquaculture and Fisheries Program, Univ. California, Davis, September, 1990,36 p.
Bajkov, A.D.
1949. A Preliminary Report on the Columbia River
Sturgeon. Oregon Fish Comm., Portland, Res. Brief
2:3-10.
Borodin, N.A.
1898. Experiments on artificial insemination of sturgeon
eggs and other biological observations conducted on the
Ural River in Spring 1987 [in Russian]. Vestnik Rybopromyschlennosti, St. Petersburg, Vol. 6-7.
Carter, E.N.
1904. Notes on sturgeon culture in Vermont. Trans. Am.
Fish. Soc. 33: 60-75.
Chapman, F.A.
1989. Sexual maturation and reproductive parameters ofwild and domestic stocks of white sturgeon, Acipenser
lransmonlanus. Ph.D. diss.. Univ. California, Davis.
Chapman, F.A., R.L. Swallow, and S.l. Doroshov.
1987. Ovarian cycle of white sturgeon (AcipenSfr
lransmonlanus). In Proc. of 3rd int. symp. on reproductive
physiology of fish. St. Johns, Newfoundland, Canada; 2-7
August 1987 (D.R. Idler, L.w. Crim, and J.M. Walsh, eds.),
p. 196. Memorial Univ. Newfoundland.
Conte, F.S., S.1. Doroshov, P.B. Lutes, and E.M. Strange.
1988. Hatchery manual for the white sturgeon, Acipenser
lransmonlanus. Coop. Ext. Univ. California, Div. Agriculture and Natural Resources, Publication 3322, 104 p.
Czeskleba, D.G., S. AveLallemant, and T.F. Thuemler.
1985. Artificial spawning and rearing of lake sturgeon,
Acipenser fulvescens, in Wild Rose State Fish Hatchery, Wisconsin, 1982-1983. In North American sturgeons: biology
and aquaculture potential (F.P. Binkowski and S.l.
Doroshov, eds.), 79-86 p. Dr. W. Junk. Publishers,
Dordrecht, Netherlands
Doroshov, S.l.
1985. Biology and culture of sturgeon, Acipenseriformes. In
Recent advances in aquaculture, Vol. 2 (J.F. Muir and RJ.
Roberts, eds.), p. 251-274. Croom Helm, London andSydney.
Doroshov, J.N., J.P. Van Eenennaam, F.A. Chapman, and S.1.
Doroshov.
1990. Histological study of the ovarian development in wild
white sturgeon, Acipenser lransmonlanus. In Proc. of 1st int.
symp. on sturgeon, Bordeaux, France; 2-5 Oct., 1989. Centre d'Etudes du Machinisme Agricole du Genie Rural desEauxet Forets, France. (In press.)
Doroshov, S.l., W.H. Clark Jr., P.B. Lutes, R.L. Swallow, KE. Beer,
A.B. McGuire, and M.D. Cochran.
1983. Artificial propagation of the white sturgeon, Acipenser
lransmonlanus. Aquaculture 32: 93-104.
Galbreath,J.L.
1985. Status, life history and management of Columbia River
white sturgeon, Acipenser lransmonlanus. In North American
sturgeons: biology and aquaculture potential (F.P.
Binkowski and S.l. Doroshov, eds.), p. 119-126. Dr. W.
Junk. Publishers, Dordrecht, Netherlands.
Harkness, Wj.K, andJ.R. Dymond.
1961. The lake sturgeon: the history of its fishery and problems of conservations. Publication of Fish and Wildlife
Branch, Ontario Dep. of Lands and Forests, Canada, 121 p.

Hung, S.S.O.
1989. Practical feeding of white sturgeon. Aquaculture
Magazine 15:60-62.
Kozin, N.l.
1964. Sturgeon of the USSR and their artificial propagation.
Uses. Nauchno-issled. Inst. Morsk. Rybn. Khoz. Okeanogr.,
Trudy, 52, p. 21-57.
Leach, G.C.
1920. Artificial propagation of sturgeon, review of sturgeon
culture in the United States. Reports U.S. Fish Commission, 1919:3-5.
Lutes, P.B., S.l. Doroshov, F. Chapman, J. Harrah, R. Fitzgerald,and M. Fitzpatrick.
1987. Morpho-physiological predictors of ovulatory successin white sturgeon, Acipenser lransmonlanus. Aquaculture
66:43-52.
Marti, Y.Y.
1979. Development of sturgeon management in the southern seas of the USSR. In Biological resources of inland waters of the USSR, p. 73-85. (Pub!. House Nauka, Moscow.
Meehan, W.E.
1909. Experiments in sturgeon culture. Trans. Am. Fish.
Soc. 39:85-91.
Moberg, G.P., S.1. Doroshov, F.A. Chapman, KJ. Kroll, J. Van
Eenennaam, andJ.G. Watson.
1991. Effects of various hormone implants on vitellogenin
synthesis and ovarian development in cultured white sturgeon, Acipenser lransmonlanus. In Acipenser (P. Williot, ed.)p. 389-399. CEMAGREF-DICOVA cedex, France. ISBN#2853622088.
Moyle, P.B., andJ.J. CechJr.
1988. Fishes, introduction to ichthyology. Prentice Hall,
NJ, 559 p.
Post, H.
1890. Sturgeon experiments in hatching. Trans. Am. Fish.
Soc. 19:36-40.
Purkett, C.A.
1963. Artificial propagation of paddlefish. Trans. Am. Fish.
Soc. 90:125-129.
Ryder,J.A.
1890. The sturgeon and sturgeon industries of the eastern
coast of the United States, with an account of experiments
bearing upon sturgeon culture. U.S. Fish Commission
Bull. 8:231-328.
Semakula, S.N., and P.A. Larkin.
1968. Age, growth, food and yield of the white sturgeon
Acipenser lransmonlanus of the Fraser Rier. J. Fish. Res.
Board Can. 25:2589-2602.
Smith, T.1.J.
1985. The fishery, biology, and management of Atlantic sturgeon, Acipenser oxyrhynchus, in North America. In North
American sturgeons: biology and aquaculture potential.
(F.P. Binkowski and S.l. Doroshov, cds.), p. 61-71. Dr. W.
Junk. Publishers, Dordrecht, Netherlands.
Smith, T.lj., E.K Dingley, and D.E. Marchette.
1980. Induced spawning and culture of Atlantic sturgeon.
Prog. Fish-Cult. 42:147-151.
Stone, L.
1901. Sturgeon hatching in the Lake Champlain basin. Trans. Am. Fish. Soc. 30:137-143.

Experimentation to Improve Recruitment of Blood Ark Shell,
Scapharca broughtonii, in the Seto Inland Sea
SATOSHI UMEZAWA *Japan Sea Regional Fisheries Research InstituteSuido-cho, Niigata 951,Japan

Abstract
The blood ark shell Scapharaca broughtonii is one of the most important bivalves in the
coastal fishery ofJapan. Recently, fishery hauls of blood ark shell have been decreasing.
The blood ark shell research group of the Marine Ranching Project has studied technologies to artificially propagate spawning stocks of these species in order to increase
recruitment in the natural fishery grounds. This paper reviews some of our efforts tounderstand the biology and ecology of ark shells and to develop techniques to improve
spat collection and prevent predation from starfish.

Introduction_

The blood ark shell Scapharca broughtonii is one of themost important shellfish of the coastal fishery of Japan. Its distribution extends from southern
Hokkaido to Kyushu in shallow bays and gulfs withmuddy bottoms. Recently the fishery has decreased
owing to excessive fishing pressure and the fluctuation of the natural level of the resource. Reduction in
the broodstock levels of ark shell are occurring because of the large number of premature shells
harvested in the wild.
Over the past decade mass production techniques
yielding about 10 million spats annually per hatcheryhave been established in Yamaguchi and other prefectures. First, broodstock is collected by selecting
spawners from the commercial catch of small trawlers
during late March to early April. The selectedbroodstocks are then reared in small tanks without
feeding for several months, becoming mature when
the water temperature rises to 20° C. The spawners
discharge eggs and sperm during late June to earlyJuly with temperature stimulation. Naturally spawned
eggs are collected and maintained in a larval rearing
tank, and newly hatched larvae are initially fed with
Pavlova lutheri or Cheatoceros spp. for two months. The
seeds are harvested from the tank when they reach
1 mm in shell length in September. Subsequently, thespats are reared in lantern nets until they reachabout 30-mm in shell length in the following year,
from March to July. These 30-mm seeds are used forreleasing into the fishery grounds and for cage cultures. In both released and cage-cultured ark shells,
one' can expect to harvest the shellfish two years after
birth. Because it takes over three years for blood ark
shells to mature sexually, these stocks will have nochance to contribute to reproduction. Therefore, thespawning stock is usually very small and may not produce a significant number of offspring.
The blood ark shell research group of the Marine
Ranching Project (MRP), which consists of the
Nansei Regional Fisheries Reseach Laboratory, theNational Research Institute of Aquaculture, and theInland Sea Fisheries Experimental Station of
Yamaguchi Prefecture, has studied technologies tocultivate spawning stocks of blood ark shell. By artificially propagating seeds to an age greater than two
years old, this group had attempted to increase recruitment in the natural fishery grounds (Fig. 1).
This paper outlines the research activities andsome results of the blood ark shell research group in
the MRP.

* Present address: Seikai National Fisheries Research Institute,
Kokubu, Nagasaki: 850,japan.

105

106

NOAA Technical Report NMFS 106
Hatcherylarval and seedling rearing!;"j\\\tFishery ground~o

Broodstock culture

Figure 1
Schematic drawing of aquaculture of the blood ark shell. Methodology of artificial broodstock culture is studied
in the Marine Ranching Project.

Environmental Conditions for the
Cultivation of Artificial Broodstock_

Figure 2 shows the location of the pilot farms for thecultivation of spawning stocks in the Kasado Bay at
Yamaguchi Prefecture. This bay was selected as a
farming ground because 1) this area has been usedpreviously to culture the species, 2) eggs and larvae
discharged from the broodstock are expected to remain in the bay because of its topographicalcharacteristics, and 3) the local fishermen's association cooperated in our survey and investigation.
The artificially propagated seeds must be reared
more than two years for use as broodstock; therefore,
the farming ground must provide environmental
conditions for good growth and high survival rates
during cultivation.
Cage culture of 1- to 3-year-old shell was tried at six
stations in Kasado Bay to investigate the growth andsurvival rates of blood ark shell (Umezawa et al.

1985). Water quality and bottom conditions were also
surveyed at each station. Figure 3 shows the results ofcage culturing l-year-old shell during the 5-month
period lasting from May to October. High survival
rates and good shell growth are seen at Station 13 incomparison with the other stations. Two- and 3-yearold shells have similar survival rates.
The characteristics of the environmental conditions at Station 13 are as follows:0• Water temperature is about 1 C or more higher
during the spring to early summer period than at theother stations. The water depth is shallower, about
10 m at the station.
• Water transparency is high.
• Grain size of bottom sediment is larger, about 1020jLm mainly.
• Organic content levels of sediment are low.

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Umezawa: Recruitment of Blood Ark Shellr"/0,-,0i"...--/\rea en I argedjr,,'''. ~

C.

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107

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• The percentage of chlorophyll-a in the phyto-pigment (chlorophyll-a plus pheopigment [pheophytin and pheophorbide]) is large, in the bottom
water.
It is suggested that the proper environmental conditions for blood ark shell propagation includes
higher water temperatures during spring to earlysummer, not exceeding 25° C continually. The requirement for bottom conditions is sandy mud
without high organic matter concentrations.

Prevention of Predation by Starfish_

High mortality occurs frequently for a few weeks after
the release of the blood ark shell seeds. One of thecauses is predation by starfish. Therefore, preventionof predation by starfish is essential for the cultivationof broodstock. Two methods were examined. One

Figure 2
Map of Kasado Bay and station
numbers of the environmental
survey (1-18). Pilot farms for thecultivation of blood ark shell arelocated at stations 9 and 10.

was the elimination of starfish with the use of traps.
The other was protection of the blood ark shell with
capsules until the broodstock reached a large enoughsize to avoid predation (Takami and Koumoto 1986).
Figure 4 shows the arrangement of the experimental lots for predation prevention at Station 9 in
Kasado Bay. Lot A shells were protected by two typesof starfish traps (shown in Fig. 5), which surrounded
the ark shell release area. Lot B, blood ark shells
were maintained in the capsules shown in Figure 6
for protection from starfish predation. Lot C shells
were a control group which received no protection
from predation.
Starfish are usually present in the experimental
area before the blood ark shells are released. A large
number of starfish swarmed gradually after the release-the greatest concentration of them near theexperimental area occurring in July. From September
to October few starfish are found. In lot A, 96.5% of
swarmed starfish were removed from the ark shellrelease area; type II was more useful than type I.

108

NOAA Technical Report NMFS 106

0=37

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10 20 30 <10 50 60

Total weight (g)'--------------_.~

Figure 7 shows the survival rate of blood ark shellin these experinments from May to October. During
May to the middle ofJuly, I-year-old shell mortality ispresumed to result from starfish predation because
the size of starfish caught in this period was large
enough to prey upon the blood ark shells. After the
middle ofJuly, the decrement in survival rate was notdependent on starfish predation because all of theexperimental lots experienced almost the same decline in survival. Two- and 3-year-old shells have noneed for prevention from starfish predation starfish
because the control experiment indicated a high survival potential for older shells.

Figure 3
Composition of total weight of blood ark shellafter five months of culture at six stations in
Kasado Bay. Within the three groups of two stations each, data for 23 May 1984 are shown inbottom graph, data for 24 October 1984 areshown in upper graph (Umezawa et al. 1985).
n=number of specimens.

When blood ark shells are released, those over 2
years old need no protection from predation. However, I-year-old shells are still vulnerable to predation
at release time, such that about 30% survival can be
expected even when methods to prevent starfish predation are used.

Plan against High Mortality
in the Summer_

From summer to early autumn, a high mortality offarmed I-year-old shell has occurred frequently. The

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Umezawa: Recruitment of Blood Ark Shell

109

I

1*." t** ** ** * *t * l* t t*
*t t****
f t ttl·UiUUU****.H".HUH"**k*****".U.. ******tt".,,******
**"*1
*UUUU

>****

Starfish trap

~

m
~

Lot A
(Removed starfish
by traps)~

ttl~t * t
*tt

j*tt

t*

It

*.......... ,,,U

Lot C
(Without traps and capsules)

*t
t *
**
**

**

Lot B
(Protected by capsules)

Figure 4

Configuration of experimental system to prevent starfish predation on blood ark shells during 14 May to 27
November in 1985. Each * indicates an individual starfish removed by a trap. Eachindicates anuncaptured starfish in the study area. D indicates a type-I starfish trap and 0 indicates a type-II trap asshown in Figure 5 (Takami and Koumoto 1986).

*

cause of this mortality is estimated to be the effect ofhigh water temperature, low dissolved oxygen or increased sulphide levels (Ishida et al. 1977;
Hamamoto 1981; Umezawa et al. 1984). It is important to avoid this high mortality during thecultivation of blood ark shell stock.
High-density hanging cultures with lantern nets
were tested to counter high mortality in the summer(Umezawa et al. 1987). One hundred artificially cultured blood ark shell seeds were maintained in eachof the four lantern nets, which were hung at a depthof 10m in the sea. Four cage cultures, each of which
had 50 seeds, were placed on the sea bottom. From
May to October, the high density hangings achieved
high survival rates of 94-100% compared with thoseof cage cultures of only 8-32% (Table 1). Thus, highdensity hanging is useful for avoiding high mortality
in summer. This method has been repeated successfully for several years (Umezawa et al. 1987).

Sexual Maturation of
Planted Broodstock_

Table 2 shows histological observations of gonad development in blood ark shells cultured from planted
seed in Kasado Bay for one to three years(Numaguchi et al. 1985). The phases of gonadal development are divided into five stages: stage I,
immature or sex unknown; stage II, early develop-

110

NOAA Technical Report NMFS 106_~35cm~r

Entrance \_&~ ~~Entrance

35cm

1

18cm

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~60cm~

Type I+--45cm~Typen

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Figure 5

Structure of traps designed
to remove starfish in lot A(Takami and Koumoto 1986).

Nylon net

45cm

1

t

18cm

11-ooI1----t- Vinyl pipeJ.

Figure 6

Structure of capsule to protect ark shell from predation of starfish in lot B (Takami and Koumoto1986).

Capsulement; stage III, late development; stage IV, mature or
spawning; and stage V, after spawning. One-year-old
shells do not mature in the first year. All 2-year-old
shells were immature in May-some male gonads developing after June. As for females, only one
individual matured in October. Three-year-old shells
did not mature in May. Mter June, the gonads developed in both males and females, and from August to
September many individuals reached the spawningphase. The spawning season is estimated to range
from July to September and the peak season is assumed to be from late August to early September.
Table 3 shows the estimated egg number for 10 3year-old parents. Three-year-old shells, which range
from 65 to 75 mm in shell length and from 77 to 91 g
in total weight, hold from 0.5 to 12 million eggs, averaging 5.5 million per individual.

Larvae and Spats Collection
from the Natural Environment_

In order to know the period, distribution, and number of larval blood ark shells, Norpac-net tows wereconducted at six stations in Kasado Bay and six stations in the coastal waters off Hikari close to the eastside of Kasado Island (Takami and Koumoto 1987).

Eight species of bivalve larvae were collected in
Kasaso Bay, and 11 species were collected off the
Hikari coast. Figure 8 illustrates the number of ark
shell type larvae at Kasado Bay and Hikari. There
were two species of ark shell (Scapharca broughtonii
and S. subcrenata) , but neither species could be specifically identified by larval examinations. In Kasado
Bay, 3.1 individuals/m 3 of ark-shell type larvae appeared in late July, and fewer were collected in
September and October. Off the coast of Hikari, 2.0
individuals/m 3 were present in July and August, rising to 15.3 individuals/m 3 in iate September.
Two types of seed collectors, the hanging and bottom types, were set at two stations in Kasado Bay andone station off the coast of Hikari (Takami andKoumoto 1987). Hanging seed collectors consisted ofa lantern net containing used fish net or nylon net.
Bottom seed collectors which were located on thebottom hung the same lantern net by means of aniron frame (Fig. 9). Figure 9 shows the number ofblood ark shell spat deposited on the two types ofseed collector. In the hanging type collector, more
blood ark shell spats are deposited as water depth
increases. The number of spats deposited in the bottom type is greater than that of the hanging type.
Spats collected in mid-December measured 1-6 mmin shell length, indicating that the larvae appearedUmezawa: Recruitment of Blood Ark Shell

III

100

100
90

25 1-year-old shells/m 2

90

80

80

70

70

60

60

50

50

40

40

30

30

20

20

10

10

50 1-year-old shells/m 2

ltl

>

'2:

0

May JunJul

Aug Sep

Oct Nov0
May JunJul

Aug Sep

Oct Nov

::l
VI

+-'

C
OJ
U
OJ

...
a..

100
90
80
70
60

~
20 2-year-old shells/m

50

100
90
80
70
60
50

40

40

30

30

20

20

10

10

o L---'_--L_-'-_.l---1_--L_-'

0
May Jun1985

20 3-year-old shells/m 2

2

Jul

Aug Sep

May JunOct NovJul

Aug Sep

1985

and attached onto the collector from late September
to October.
These larval and spat collection results were obtained in the investigation for 1986. The origin of
these spats was probably derived from the natural
population because the planted broodstock in
Kasado Bay was still immature in 1986. The results ofan identical investigation conducted in 1987
(Ouhashi and Koumoto 1988), showed marked improvement in the number of the ark shell type larvae
compared to the previous year. These findings suggest that some fraction of collected larvae were
spawned from the planted blood ark shell parents.
However, the number of deposited spats in the seed

Oct Nov

Figure 7
Survival rate of blood ark
shell by age and densityfrom experiments to prevent
starfish predation. Opencircles denote lot A where
two types of starfish traps
were used; closed stars denote lot B where shells wereshells protected by a capsule;closed circles denote lot C
where seed were released
without the protection oftraps or capsules (control)(Takami and Koumoto1986). The figures in eachgraph shows intial shelldensity.

collectors were smaller in 1986, indicating that spatcollection is a major problem to be solved at the pilot
scale.

Citation_

Hamamoto, S.
1981. Basic studies on the mortality factor and resisting
power of an ark shell, Scapharca broughtonii(SCHRENCK). Scientific Report of Kagawa Prefectual
Fisheries Experimental Station 18:1-19. (In Japanese.)
Ishida, M., I. Hayashi, and H. Ushima.
1977. Experiment on cage culture of blood ark shell. Annual Report of Fukuoka Prefectual Buzen Experimental Station 49-52. (In Japanese.)

112

NOAA Technical Report NMFS 106

Table 1Comparison of growth and survival between high-density hanging cultures in lantern nets and cage cultures on thesea bottom in 1986 (Umezawa et al. 1987).
High-density hanging culture

Cage culture at bottom

Date

No. of shells/net

Mean total wt. (g)No. of shells/net

Mean total wt. (g)May 13

100
100
100
100
100

6.8
6.9
6.8
6.8
7.0

50
50
50
50

8.1
7.3
7.1
6.1

Oct. 13

98
94
100
97

23.6
25.6
23.8
20.7

14
16
9
4

16.4
15.6
15.4
12.1

Table 2Frequency (%) of blood ark shells in each phase of gonad development in 1- to 3-year-old shells in 1984 (Numaguchi
et al. 1985). Stages I-V are phases of gonad development: Stage I, immature or sex unknown; Stage II, early development; Stage III, late development; Stage IV, mature or spawning; and Stage V. after spawning.
l-yr-old
11

IV

1Il

11

V

<3

<;>

<3

<;>

<3

<;>

<3

100
100
100
100
100
100
100

0
0
0
0
0
0
0

0
0
0
0
0
0
0

0
0
0
0
0
0
0

0
0
0
0
0
0
0

0
0
0
0
0
0
0

0
0
0
0
0
0
0

0
0
0
0
0
0
0

100

0

0

0

Sampling date
May 23
Jun 5
Jun 25
Jul4
Jul24
Aug 20
Sep 17
Oct 312-yr-old
<;>

<3

0
0
0
0
0
0
0
0 0 0 0 0

100 0
85 15
55 45
55 15
35 55
90 10
55 45
80

0

1Il
<;>

<3

3-yr-old
IV

<;>

<3

0 0 0 00 0 0 00 0 0 0o 15 o 15
o 10 0 0
0 0 0 00 0 0 00 0 0 0

11

V
<;>

<3

<;>

0 0
0 0
0 0
0 0
0 0
0 0
0 0
o 10 10
0
0
0
0
0
0
0

<3

III
<;>

<3

<;>

IV

<3

V

<3

<;>

<;>

100 0 0 0 0 0 0 0 055 25 20 0 0 0 0 0 055 30 5 10 0 0 0 0 020 50 10 10 10 0 0 0 0
5 25 5 45 20 0 0 0 0
0 0 05030 15 5 0 0
10 0 0 0 o 10 10 60 10
90 0 0 0 0 0 0 5 5

Table 3
Shell length. weight and egg number of 3-year-old blood ark shell (Numaguchi and Tanaka 1987).
Weight (g)No. of eggs

Sam lin No.

Shell length
(mm)

Meat

Gonad

(Xl0 5)

1
2
3
4
5
6
7
8
9
10

71
77
71
67
70
70
66
65
75
72

78
87
81
70
80
77
86
88
91
86

0.36
0.86
1.59
1.93
2.89
3.51
3.72
4.09
5.25
5.97

5.4
7.1
16.1
54.0
5.5
70.6
76.5
115.5
78.9
119.3

Average

70

82.5

3.02

54.9

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Umezawa: Recruitment of Blood Ark Shell

113

0 15.3
II

II
II

~
I'
I I

I'

I I

I
I

t

I

10

I

I

W
.c:
VI

I

I

~

L..

I
I

ltl

,

"U

o

o

:::c

I
I
I

'0
CII

~

~

Hikari

I

I
I
I

5

'0
CII

9

I

E

I
I

\

L..

.0
::J

Z

Kasado bay

I

~'\

I

I
I
I
I

:

I
I
I

I
Jul

Aug

Sep

Oct

1986

Om

Sm

Jun

1986

~Ka~d:Q

~b

o

\
\

., ~ • O'"i.~

0
Jun

\

Jul

\

Aug

Figure 8
Seasonal distribution of ark
shell type larvae at Kasado
Bay (water depths are about
15 m) and the coastal waters off Hikari (water
depths are about 20 m) as
collected with a Norpac net.
= mean tows from all
depths combined at each ofsix stations. • = mean forthe middle of water depth
to upper layer (Takami andKoumoto 1987).

Sep

Oct

Hikari

I

1

I

I
I

I
I

I
I

I

I

I

I

I

I

b~~
I

I

I
1

I
1

I

1

I
I
I

I

ffi ~~
I
I

10m

I
I

I
I
I

15m

20m

~~~~1bFigure 9
Number of blood ark shell spats deposited in seed collectors at different depths in Kasado Bay and off the coast of Hikari. Numbers in
the triangles predict the total spat number of 21 hanging cages.
Those with frames predict the total spat number of 12 cages locatedon the bottom (Takami and Koumoto 1987).

114

NOAA Technical Report NMFS 106

Numaguchi, K., and Y Tanaka.
1987. Acceleration technique of maturation and spawning of
the ark-shell, Scapharca broughtonii, in farming
ground. Marine Ranching Project Progress Report, Bay
Scallop and Blood Ark Shell 7:55-59. (In Japanese.)
Numaguchi, K., S. Funakoshi, and K. Wada.
1985. Maturation and spawning of cultured ark shell,
Scapharca broughtonii. Marine Ranching Project Progress
Report, Bay Scallop and Blood Ark Shell 5:71-75 (In Japanese.).
Ouhashi, H., and Y Koumoto.
1988. Technique for collecting ark-shell seed from artificial
broodstock. Marine Ranching Project Progress Report, Bay
Scallop and Blood Ark Shell 9:73-83. (In Japanese.)
Takami, T., and Y. Koumoto.
1986. Studies on the artificial propagation and control of the
predators in the ark shell. Marine Ranching Project
Progress Report, Bay Scallop and Blood Ark Shell 6:97-106.
(In Japanese.)

1987. Technique for collecting ark-shell seeds from artificial
broodstock. Marine Ranching Project Progress Report,
Bay Scallop and Blood Ark Shell 7:61-72.
Umezawa, S., K. Nogami, and O. Fukuhara.
1984. Relation between high mortality and some environmental conditions for ark shell, Scapharca broughtonii in cage
culture. Bull of the Nansei Regional Fisheries ResearchLaboratory 16:231-244. (In Japanese.)
1985. Environmenta! conditions and releasing density of ark
shell in the artificial farming ground. Marine Ranching
Project Progress Report 5:57-69. (In Japanese.)
Umezawa, S., S. Arima, K. Oochi, and O. Fukuhara.
1987. Farming the broodstock and environmental conditions for natural seeding of ark-shell. Marine Ranching
Project Progress Report, Bay Scallop and Blood Ark Shell
7:73-81. (In Japanese.)

The Marine Ranching System: The Integration of Biology
and Engineering Technology

HIROYUKI NAKAHARA
Research Institute for Ocean Economics
Marufuji Bldg., 3-1-10 Shinbashi
Minato-Ku, Tokyo 105,Japan

Abstract
This paper discusses a variety of concepts concerning aquaculture, including fish-farming and marine ranching with respect to their purpose, the ownership of the fish,participants, the ability to measure resource production, the technologies introduced todate, and future trends. Approaches to engineering systems, which should be designed tomeet with the biological aspects, are also examined along with some examples. Aquaculture, which used to be deployed in rather small, calm and protected bay areas, needs
improved hardware and techniques in order to be performed in more offshore areas ona larger scale. The expansion of fish farming should be accompanied with new environmental control technologies:technologies designed to enhance primary production andother related engineering capabilities. To integrate the efforts of biologists and engineers is the best way to create a successfull marine ranching system. Some of theexperiences reported in this paper are based upon research and development activities.
For example, seabed improvement efforts have been made which make use of civil engineering and water quality improvement to fit the needs of a target species during acertain stage of its growth to enhance feeding. This was done by making use of water
movement control technologies and by introducing new materials and structures for theattachment of food organisms.

Introduction_

Japan is the largest fishing nation in the world with a
total catch in 1985 of 12.2 million tons QFA 1987;
MAFF 1987). Additional fishery products valued atabout five billion dollars were imported while exports
totalled one billion dollars. Because the local offshore fishery in the waters around Japan consists
mainly of sardines and because growing international
constraints are making contributions from long-distance fishing harder to maintain, efforts are focusing
on coastal fishery production to keep up with thegrowing demand for high-quality fishery products.
The coastal fishery production increased from 2.24
million tons in 1965 to 2.71 million tons in 1975, andto 3.36 million tons in 1985. However, within these
totals, the percentage from fishing boats, the traditional style of harvest, remained constant at around 2
million tons during these twenty years. The increases
were realized by the marine net-pen fishery andaquaculture. (Figs. 1 and 2; DSI 1984) The contribu-

tion from aquaculture has risen five times during thisperiod. Sea-farming, which refers primarily to the
propagation of cultured seed that are released into
the natural environment, has been performed formore than twenty years. Typical of this effort is therelease of two billion chum salmon (Oncorhynchusketa). The mass production of other seed is now possible with kuruma prawn (Penaeus japonicus) , blue
crab (Portunus trituberculatus) , scallop (Patinopecten
yessoensis) , abalone (Haliotis discus hannai) , and others.
Based upon these cultivation activities, the newconcept of The Marine Ranching System has beendeveloped in recent years, which makes use of industrial engineering technologies (RlOE 1981-88). Someof them are in a very conceptual stage while others
are being experimentally executed on-site. This paper described the status quo of these efforts inJapan,
which are combining biological knowledge with engineering technologies.

115

116

NOAA Technical Report NMFS 106

X lOG TON

12
11

10

..t5;

9

3,i;~t\:.~;{;l.!.i-t .

-:~,~.~~

8..: ....

,~,~~I~S~:: :.':. :.'.

........

-~,:.~~~:".' ...

6...... '.

4'~hi;,j~~<f~~\~<~-i,~i}:¥/i~;~X;.!.'..

AQUACULTURE

7

o
60

65

70

75

90

34

Figure 1
Trends of fishery production in Japan (Fisheries Statistics of Japan, Ministry of Agriculture,
Forestry, and Fisheries 1984).

INLAND & FRESHWATER-AQUACULTURE

13·(Mi II ion metri c tons)

11.

Inland water 0.6%)
fi sheri es· cui tures
Marine cultures(8.7%)

10 •Coastal fisheries
07.7%)

12 •

9 •
8·
7·
6 •
5 •

Offshore fisheries(54.3%)
4 •
3 •
2 •

Distant water
fisheries(17.8%)

1•

O·
1974

'80

'84

Figure 2
Total catch by type of fishery, 1974-84(Fisheries Statistics of Japan, Ministry ofAgriculture, Forestry, and Fisheries 1984).

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

Definition of Aquaculture, Sea-farming,and Marine Ranching

117

Aquaculture_

Throughout the long history of mankind, fishinghas meant the taking of live fish from the sea in amanner analogous to hunting on land. In the past,
we caught more fish from the sea by sailing all over
the world using high-tech electronic instrumentation. Conversely, we are now developing aquaculture
techniques that are analogous to cattle breeding. In
Japan, both freshwater and marine aquaculture havebeen developed for a long time. Carp (Cyprinus
carpio), eel (Anguilla japonica) , chum salmon (0.
keta), rainbow trout (0. mykiss), and sweetfish or Ayu(Plecoglossus altivelis) are the major species cultured
in freshwater. Marine aquaculture mainly covers laver(Porphyra), oyster (Crassostrea gigas), yellowtail (Seriola
quinqueradiata), sea mustard (Undaria), scallop(Patinopecten yessoensis), and red seabream (Pagrus major).

The term "aquaculture" generally means the cultivation of fish in the protected sea area and is frequently
symbolized by the net~cage (Fig. 2). This process usually starts with the collection of eggs from either
natural or artificial environments, then rearing thejuveniles, usually with feeding until harvest time. The
most important characteristics of aquaculture are 1)
that the ownership of fish is quite clear, 2) that human control is possible throughout the whole life of
the fish although conditions within the net cages aresupported by nature, and 3) that the final fruits ofthe production are countable (Table 1).

Sea-Farming_

Sea-farming involves the propagation of cultured
seeds and fry into adults and their release into theTable 1
Major characteristics of aquaculture and fish-farming in Japan.
Aquaculture

Fish-farming

SectorPrivatePublicMajor species

Salmon (Oncorhynchus ketal
Red seabream (Pagraus major)
Kuruma prawn (Penaeus japonicus)
Yew abalone (Nordotis discus hannai)
Flounder (Paralichthys olivaceus)

Area

Laver (Porphyra)
Oyster (Crassostrea gigas)
Yellow-tail (Smolaquinqueradiata)
Scallop (Patinopecten
yessoensis)
Red Seabream (Pagrus major)
Protected sea

Open sea

PurposeDirect production

Resource increase

Human contrOl

Total period(until harvesting)

Until releasing

Ownership

Fish producer
Net cage owner

None

Bait

FeedableNon-feedable(food organism in nature)

Production

Countable

Technology

Net cage structure
Formula feed

Future trend

LargerDeeperMore offshore
Biotechnology

To be included in total catch(by traditional fishing operation)
Seeding production
Tagging
Artificial reef
Survival rate improvement
Environmental control
Primary production enlargement
Biotechnology

118

NOAA Technical Report NMFS 106

I

Fish

Shellfish

&

J.

I

I Spawning Ground I

I Catch of Adult Fish I

I

I

Egg

Fry

I

Collection

I

I Artificial Hatching
I

J,

I

Alevin Rearing&

I

Releasing

J.,

I
t..

("

Artificial
Fishing ~~Ground~I

I Limitation ofI: Season, Area

:

I

I
I

I
I

I

Production~=)

Total CatchI~t Control byI~,

I & Fish Size

I

I

I

I

I.

I
I INDIRECT

,'

,:
II

I
I

I~

Prevention of Fish Diseases

f9

Nutritionr~

Feeding~1 Enhancement of Primary Production

r4

.£

-

Fish Farming

Aquaculture-

I
I

RESOURCE I
ENHANCE- I
I
MENT&MANAGEMENT I

,

I

1

II'

Conventional
Fishing

OffsboIie~. Large-scale
Net Cage~1 Protection from Harmful Elements r~

I·
II

I RESOURCEI MANAGEMENTr~

Environmental Control& Management~,

I

Cultivation

Monitoring11
r- -- - - - - ---,~~ Fishing Gear, ~I

in Nature

Harvest fromFish-Preserve

Harvest at sea)
I

I

I

I

Figure 3

J,

I

TOTAL

FISH

PRODUCTION

sea. Obviously they cannot be counted except when
they are tagged.
Released fish belong to no one individual, instead
they contribute to the increase of the fishery resources and can harvested by anyone. For thisreason, sea-farming is done by the public sector inJapan. This type of cultivation is highly dependent on
the carrying capacity of the natural sea itself. Compared with aquaculture, human measures to improvetheir growth in the natural environment have had arather limited effectiveness and the artificial reef isalmost the only one effective method among them.
The final product of sea-farming is included in the
total number of fish caught by the traditional fishing
operations. In the case of chum salmon, it is easy todetect the contribution of this human effort whereas

I

Concept of the Marine
Ranching System (ResearchInstitute for Ocean Economics).

in other species it is quite difficult to assess the effectiveness quantitatively.
Both aquaculture and sea-farming are consideredvital parts of the marine ranching system, but when
we discuss their use in this progam, we are speakingof projects that are technically upgraded in various
perspectives. In the case of aquaculture for example,
the near future may bring the use of a netless caging
system, which utilizes acoustic, optical or some otheradvanced technology. Moreover, large floating structures to hold net cages of required size and numbers,
may be used to create offshore preserves which canbe moved if necessary.
Sea-farming also needs to step into a new age withvarious environmental control technologies created
for the system. We need multiple technologies from

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

Increase in the entire quantity of marine
resources by reducing natural enemies anddeaths.
I. Increase in thesurvival r:ttio or flshe
and shellfishes

hpanese horsemackerel~

119

Studies on an efficient integrated production
system for each marine product----t-Studies on new fisheriessystems
Complelt resource
culture technolo!yIV.

Yello~' tails

M:asu salmon
Arame Mh7-cseaweed ~Complu production
technology thou combines the properties or
ecology and those orwltenlKalime Jeawee(A 19ae)

~fis

..

~2
Ark shells

II.
Environment conlrol lechnolo!ySe3l11ater environment

Sea bollom
envuonment~e:lSunnt

Studies on the technology for
optimizing seawater and sea
bottom. the places of growthand life of marine creatureslechnolor,yMonitoring for maintenanceof environment

Prevention of dmeases offished and shellfishes

Figure 4
Flowchart illustrating the research of the Marine Ranching Program. (Research Council of Agriculture, Forestry, andFisheries).

civil, mechanical, electric and other engineering disciplines to be developed by industrial communities
outside of fisheries. Furthermore, to increase theseedling survival rates in the sea, biotechnolbgy will
be of great help in producing healthier seed thanever before. Of course the total system of marineranching should be combined within the appropriate
legal and social frameworks (Fig. 3).

Marine Ranching Program_

In the government sector, the Research Council ofAgriculture, Forestry and Fisheries, collaboratingwith the Fisheries Agency and its national fishery research institutes, started a 9-year research program,called the "Marine Ranching Program" (MRP), in
1980. During the first 3 years, the ecological study of

120

NOAA Technical Report NMFS 106(Stage of
Nutrition)

Addition of
Resource(e.g. net pens)

Addition of
Resource(e.g. Fry &Alevin Releasing)(already
being used)

Surplus of Primary Production(Utilization of Local(Bait Fish
Relocation)
Excess Resource)

Primary Production(already being used)

Enhancement ofPrimary Production(e.g. Artificial Upwelling
Promotion of Photosynthesis)

Primary". Production------------------------.------------------------------ ----------1---------4 ----------------------------- ---------------

1 0 - - - - - - - - - - - Existing Food Chain - - - - - - - - - - + - - - Enlarged Food Chain
Increase in Harvest
Due to Added Resources[Conventional Type]

Naturally Generated
Increase in Harvest
Due to Increase inPrimary Production

mrnill shows irx:rease of fishery resources. (!loth sides are exaggerated for better understrux:lil-g)

Figure 5
Structure of the marine food chain.

various target species and preliminary study on environmental control and disease prevention were
executed. The second 3-year period was devoted tothe study of production systems, and the last 3 years
to the examination of multiple-species ranching systems. The selected target species were masu salmon(0. masou) , horse mackerel (Trachurus japonicus),
bluefin tuna (Thunnus thynnus), flounder, bay scallops, and seaweed (along with abalone and others).
Each represents a different pattern of life in the sea,thus covering the categories of highly migrating,open-sea migrating, non migrating, and sedentaryfish and algae (Fig. 4). This program has been done
mainly by considering the biological aspects, butsome study to integrate it with engineering technologies has also been done by a group studying themarine ranching system organized within the Research Institute for Ocean Economics (RIOE), a
research institution which examines various ocean
uses and development and to which the author belongs. The approach used for help identify
engineering tasks within the marine ranching system
examined by RIOE is described below.

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

,

I

121

,

-t 0I 'SUNLIGHT--, ----~PLANKTON

Figure 6
Scheme of engineering on the
key parameters of the food chain
to enhance primary production(Research Council of Agriculture,
Forestry, and Fisheries).

Approach to the Marine Ranching
System Engineering_

Food Chain Scheme Approach
When we consider utilizing the sea's potential, or consider the addition of recoverable resources into it, aview from the perspective of the food chain offers us asuggestion. Such an effort involves expanding the
pyramid shaped hierarchy of the food chain as shownin Figure 5. To better understand the diagram, it
should be kept in mind that both sides of the pyramid
structure have been exaggerated somewhat.
The right hand side shows that artificial measures
should be taken in order to increase primary production by making use of the unused but potentiallyvaluable resources of the sea. One idea is to pump up
nutrient-rich deep-sea water to certain sea areas, suchas semiclosed bays and thus form an artificial upwelling system. Another idea is to transmit sunlightinto the dark lower layer of the sea to activate photosynthesis. These ideas are creating great challenges
which can only be solved with new engineering technologies. They are based upon the basic concept thatthe nutrients and the sunlight are among the criticalfactors which create optimum environmental conditions (Fig. 6) as described in below.
The left hand side of Figure 5 shows a direct approach to increasing resources by adding them
directly into the sea. Aquaculture represents this approach of adding the new resource into amanageable sea area so that all the added resource
can be harvested. In the case of yellowtail, for example, the feeding bait is sardine which represents
the fruits of primary production somewhere else in
the sea. In this approach, engineers may focus on thedevelopment of large scale structures to support fish
cages which must be strong enough to cope with theharsh environment of the deeper, more offshore sea
areas. The direct release of resources to the sea isanother type of approach. In this case, we expect thata portion of the added resource will be harvested in
the future by conventional fishing operations. This
represents sea-farming, namely, the propagation andrelease of juveniles. The natural environment supports this enhancement scheme if we assume that theexisting carrying capacity is sufficient. In this case,less effort has been necessary from the engineeringcommunity.

122

NOAA Technical Report NMFS 106(F 0 0 Dl(SECURITY)SUNLIGHT~ NUTRIENTS/BAIT SPECIES ~/PREDATIONDISEASEFISHING/OPERATIONSEngineering can control the level of nutrients andsunlight for baitfish. In addition, they can put improve security from predators by controlling
environmental conditions or by changing the predators' attention to other attractive things. Environmental conditions such as water movement, andseabottom conditions can be managed by civil andmechanical engineering technologies, to create favorable environmental conditions for juvenile fish.

IMPROVEMENT OFSURVIVAL RATE\TEMPERATURE

DISSOLVEDOXYGENSEAWATE~MOVEMENT
SALINITY(HEALTH)

Variations of Artificial
Upwelling System~DEPTH_SEABOTTOM CONDITIONS(HOUSEl

Figure 7
Critical factors for the improvement of survival of marine
resources.

Critical Factors Approach_

When we turn our eyes to the vital elements, or critical factors for fish growth and suvival, we find thatseveral important factors can be controlled by humanbeings to some degree. In the case of chum salmon, a
migratory species, the production increases havebeen made by accumulating fingerling releases overmany years without the use of additional specific art.ificial measures to control environmental factors. ThIs
success therefore, must also have been supported by
a combination of favorable conditions like the abundance of food sources in the northern Pacific Ocean,the rather small number of harmful elements, andthe mobility resulting from the swimming capacity ofthis species.
However, when we consider nonmigratory and sedentary species, the effect of pr~dators. andenvironmental factors are much more InfluentIal on
the survival rate, especially during the juvenile stage.
The critical factors for the survival of a fishery resource depends upon the species and its stage of
growth. In addition, when we discuss engineer.ing approaches, some basic considerations can be glve.n bymaking analogies with the lives of the human beIngs.
At the very least, we need food to eat, good health togrow, a house in which to sleep, and securi~y to live.
The corresponding factors for fish are baIt to ~at,security from predators, and a comfortable chemIcal
and physical environment (Fig. 7).

Various artificial upwelling technologies are illustrated in Figure 8. Among these is a land-based
system using pumped up nutrient-rich, deep seawater, which is already operating in Hawaii. This system
is also well known as a part of OTEC (Ocean Thermal Energy Conversion), which generates electricity
by using the temperature difference between surface
and deep seawater. We can therefore put the deep
water not only in the natural sea area but also in
artificially made ponds, and land-based tanks as has
been done in Hawaii. In Japan, a similar system exclusively for the cultivation of fish will soon start near
Cape Muroto in Kochi prefecture.
Another unique system utilizes deep water pumped
up to a surface barge. Partly intended for the direct
use of the nearby net cages which are attached to thebarge, its primary purpose is to supply a sprinkler
system which disperses the pumped up water onto
the surface to bolster primary productivity andbaitfish abundance in the area. Designed to be completed within a couple of years in the Toyama Bay,the system moderates the cold temperture of thepumped deep water taken from the depth of 300 m
by mixing it with warm surface water.
Through this field test, the optimum ratio of cold
deep water to warm surface water will be surveyed
and its effectiveness to increase the fishery production will be estimated. The estimation index is one of
the most important items to be examined. The bay
has a steep seabed, is halfway closed by Noto Peninsula, and holds cold deep water rich in nutrients.
This unique natural condition can only be seen in
Toyama Bay along the Sea of Japan coast and in
Suruga Bay along the Pacific coast. The Toyama ~ayproject should be of great interest from both the bIOlogical and the engineering point of view, althoughmany difficulties exist, including the method of theassessment of its effectiveness. However, this challenging project is said to be the first attempt at an
offshore floating-type upwelling system in the world.
Seabed civil engineering systems are approaches un-

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

ON-LAND

123(HAWAII. KOCHI,JAPAN)

~rt/
. ...

ARTIFICIAL '
POND

...

{
OFFSHORE
STRUCTURE
PIPELINE(PUMP UP)~~~1
J,

l~~:I-CLOSEDNATURALSEA

OFFSHORE OPEN *(TOYAMA,JAPAN)
SEA

t .•

1111I1111111111. /jl
MEMBRANE----TEXTILE SCREEN (HOKKAIDO.JAPAN-~Z~~ZONTAL)~

r:!.>.k

SEA BEDSTRUCTURE

11I!l1I11I11II
A4=SF

ARTIFICIAL DIKE

(FLOW CONTROL)GRAVITYi

CONCRETE

Figure 8

Various artificial upwellingsystems.

* PLANNING STAGE

der development to control water movement. A horizontal method of flow control is being deployedexperimentarily with a textile screen in the shallow
water off Hokkaido. an artificial seabed dike, whichmakes use of the fly ash, is also under examination.
Concrete seabottom structures which channel
deepwater flow up to the surface have also been experimentally employed in Aomori prefecture on asmall scale. In Ehime Prefecture a greater magnitudeof scale was accomplished requiring a budget of hundreds of millions of yen. A rubber dike to create an
artiRcial tideland is also being executed. Furthermore, subsurface structures for the creation of
artificial seabeds that employ available sunlight are
another idea promoting the multilevel utilization of
the water column (Fig. 9). Many new materials previ-

ously unfamiliar to fishery-related people are being
introduced among these efforts.

New Materials_

Materials are being introduced in the fishery-related
efforts mentioned above, including fly ash, a by-product continuously produced by coal-fired power
plants, and rubber, which most people equate withcar tires. Quite unique, new materials are also being
used for other marine ranching engineering. Two examples are described here. First is "Polyethtel," which
is used to construct a man-made seaweed-like tree.
Used in combination with a concrete artificial reef, it
upgrades the reef's attractiveness to marine life by

124

NOAA Technical Report NMFS 106

Rubber Dike.

Tideland/High-tide~S:;::;;:::::;'1'"."~'f" ::,g-------r-~-Loy-tide~ ?: ....

Rubber dike deflated~~~~~~~----t-. . . J~

-Y-.o

~

Tideland

Shore line

Artificial Seabed Dike

Rubber Dike

,'/

-O~
/ I '

¥.; t-lII

it;,1

,,1,/

Sunlight col1ect--::O~
ing bu.oy/11\'......... 0///

Sunlight Transmission System

STSS (Single-point Tension-leg
Subsurface Structure)
Figure 9
Examples of marine ranching technologies.

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

Frame of Artificial Reef

Completion of Artificial Reef

Attached Artificial Seaweed made by Polyethtel(Brand Name is "Bush-Absorber")Installing Operation

Installed on the Seabed

Figure 10
Artificial seaweed forest formation using new materials (Wakachiku Construction Co., Ltd., Tokyo,Japan).

125

126

NOAA Technical Report NMFS 106

_

'7.

..
~.=J. ~:::::::=====~d=.~-, ... ...~......~ .•.. ~~-Net Type.: ..

-_ ...
Mound-Covering Typesz...,......~~)..L- .'..":...
'~.,~.<, ... ~ ••.

~---.---

Suspended-Board TypeMulti-Board Type

Figure 11
Artificial seaweed forest formation using new materials (Chateau Marine Survey Co., Ltd., Miyakojima,
Osaka, Japan) .

providing a substitute for the attachment of algae(Fig. 10). This effort was made five to eight years ago.
The other example is a composite material of polyethylene and calcium carbonate that is made intotwine and boards. The twine is used to weave a nettype structure for seaweed attachment, while theboards are used to create an artificial seabed at any
depth. Any number of decks can be installed in midwater by anchoring or pile fixing (Fig. 11). This new
material was patented recently.

Engineering Measures Basedupon Biological Requirements
Two examples of the examination process used tomodel the development of engineering technologies
which will contribute to the increase of resources areshown in the cases of flounder and ark shells (Tables
2 & 3). The charts seek to identify critical factors
which determine survival, growth, and reproductive
rates. During the transformation period of flounder

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Nakahara: The Marine Ranching System

127

Table 2
Basic idea of engineering measures upon biological requirements
Flounder (Paralichthys olivaceus).

HabitatLife stage
Larval(floating life)

SeawaterSeabottom

Flounder(Paralichthys
olivaceus)

Prevention of dispersion(by controlling seawater
movement)
Improvement of
seabottom condition

Supply of bait

Adult(Same as above)(Same as above)

Spawning(Same as above)(Same as above)

Juvenile(seabottom life)

Other
species(Same as above)Protection
from predators

Table 3
Basic idea of engineering measures upon biological requirements
ark shell (Scapharca broughtonii).

HabitatLife stage

Seawater

Floating

Prevention of dispersion(by controlling seawater
movement to accumulate
larvae)Otherspecies

Supply of bait
Formation of attaching
structures(Same as above)

Seabottom

Improvement of
seabottom condition(Same as above)

Spawning(Same as above)(Same as above)

Attaching andtransformation toseden tary type

(Same as above)

Seabottom

Flounder(Paralichthys
olivaceus)

from the floating life stage to the seabottom life
stage, bait species change and water currents frequently disperse eggs and larvae to unfavorable
seabottom conditions. Both are critical situations that
affect the mortality of juveniles. These requirements,
determined by biological study, suggest that engineering measures should be taken to control theseawater movement in order to prevent egg dispersion and to improve the seabottom condition. In theProtection frompredators

Protection frompredators

case of ark shells, similar approaches could be considered.

Conclusion
The planning and development of engineering technologies should basically be supported by biologicalstudies, otherwise successful results can not be ex-

128

NOAA Technical Report NMFS 106

pected nor obtained. All aspects of the engineeringmust have a biological reason and endorsement.
Even then, the engineering side must be always
humble to accept the demands of nature. Biologists
must also keep themselves open- minded to the newtechnologies in other industrial communities. Making use of industrial engineering capabilities for thedevelopment of enhanced fishery resources is ofgreat significance for marine ranching systems.

Acknowledgment_

This paper is based upon an intensive study being
done by the research committee on the marineranching system, which was organized in 1981, andwhich is still active now. This committee has beenworking together under a research contract with the
Research Council on Agriculture, Forestry, and Fisheries, which consists of some thirty companies amongthe 120 member firms of the RIOE.

Citations
DSI (Department of Statistics and Information)
1984. Fisheries statistics of Japan 1984, DSI. Ministry of Agriculture, Forestry, and Fisheries, Government of Japan. (InEnglish.)JFA Oapan Fisheries Association)
1987. Fisheries ofJapan 1987. (In English.)
MAFF (Ministry of Agriculture, Forestry, and Fisheries)
1987. Annual Report on Japan's Fisheries, Fiscal 1986,
Summary. (In English.)Research Committee on marine ranching system (Research Institute for Ocean Economics)
1981. Model study on Environmental Monitoring system formarine ranching. (In Japanese.)
1982. Marine Ranching Program: enlargement method of favorable environmental conditions and monitoring system.
(In Japanese.)
1983. Marine Ranching System: a model study. (In Japanese.)
1984. Marine Ranching System: a Preliminary assessment.
(In Japanese)
1985. Study on enrichment method of the marine ranching
ground. (In Japanese.)
1986. Study on ranching ground use development by cultivation technology (I), with I) development of data based on
marine ranching, 2) examination on the integration of biology and engineering technologies. (In Japanese.)
1987. Study on ranching ground use development by cultivation technology (II) with examination on the integration of
biology and engineering technologies. (In Japanese.)
1988. Study on ranching ground use development by cultivation technology (III) with examination on the integrationof biology and engineering technologies. (In Japanese.)Economic Problems of the Marine Ranching System
KATSUOTAYANational Research Institute ofFisheries Science
5-5-1 Kachidoki, Chuo-ku
Tokyo 104, JapanAKIRA HASEGAWA
Tokyo University ofFisheries
4-5-7 Konan, Minato-ku
Tokyo 108, JapanYOSHIO MASUITokyo University ofAgriculture
1-1-1 Sakuragaoka, Setagaka-ku
Tokyo 156,Japan

ABSTRACTThe major challenge facing the Marine Ranching Project is to increase fishery production in an economically efficient manner. To accomplish this, new technology such as
improved seed production methods is required, along with an integrated approach that
includes well planned development of local economies as well as total resource management and harvest regulation. This report analyzes the work of the Marine Ranching
Project on the production and release of masu salmon (Oncorhynchus masou) by examining the relationship between production and release costs and the rate of smoltification.
In addition, survey data show that most of the masu salmon taken in Hokkaido from
October to March are juveniles under 600 g, and that the economic value of the fisherycould be raised considerably if the target fish were allowed to reach an adult size of 1,500
g before harvesting. The report also notes that the masu salmon fishery plays an important role in the regional economies of northern Japan, and that seed release and harvest
regulations must be designed to meet the economic and social needs of the various
regions and communities that depend heavily on the resource.

Introduction_

The most important tasks now facing the masu
salmon (Oncorhynchus masou) Marine Ranching
Project are the economically efficient production andrelease of smolt seed. At present, many researchers
are developing new seed production methods andstudying the ecology and physiology of masu salmon.
This report analyzes three methods of masu
salmon seed production and release: autumn releaseof 1+ year seed obtained from wild fish (hereafterreferred to as autumn release); spring release of 1+
year seed obtained from wild fish (spring release);and spring release of 0+ year seed obtained fromhatchery-raised fish (hatchery spring release). The
economic merits, demerits, and other problems associated with each method are analyzed and discussed.

- This report also analyzes the market price mechanism for masu salmon and evaluates the economic
benefits of managing the fishery through restrictions
on the harvest of undersized immature fish. Finally,
the report reviews the role of the masu salmon
fishery in the northern prefectures of Hokkaido,
Aomori, Iwate, Akita, and Niigata and discusses waysof coordInating the Marine Ranching Project with local variables such as types of fishing gear, level offishing effort, and fishery production costs. Data on
monthly harvest rates and average fish body weights
were collected from the Prefectural Fisheries Experimental Stations of these governments.
The research was divided into the following segments: I) Comparative study of masu salmon seedproduction costs at the Prefectural Hatchery Stations
at Iwate; 2) Economic comparison of autumn release,

129

130

NOAA Technical Report NMFS 106

Table I
Seed production cost of masu salmon per fish (in yen). (Iwate Prefecture Office 1974.)
AutumnreleaseSpringreleaseHatchery spring
release

2.0

2.0

2.0

(2.5)a

4.69
7.81

9.38
15.63

7.5
12.5

Power/fuel

0.75

2.0

1.2

Consumable ilems

0.75

1.5

1.2

Labor

5.28

6.36

4.2

Transportation

1.25

2.5

2.0

Maintenance

2.0

8.0

3.2

Survival ralio

0.5

0.4

0.6

Total cosl per fish
(1.5)
(2.5)

18.72
21.84

34.74
40.99

22.63
27.63

Eggs
Feed (1.5)aaparentheses contain feed conversion ralios of 1.5 (minimum) and 2.5 (maximum).
Conversion ratio; (A-B)/C X 100,
where A; Final weighl (g/fish)
B; Start weight
C; Feed weight.

Spring release

Autumn release

100

Hatchery spring release

100

100

Feed conversion rate
a=1.5
b=2.5
c=b-a

:I

--- --------i--- -- ----T-----

50

50

50

I

b

I

a

I
I

.

b

a

o

50

80

c
100

b

I

a

c
80

50

0

100 <X)

c
0

50

80

100 (%)

Smal tificatioo ratia(%)

Figure 1
Ratio of seed production costs to the smoltification ratio in masu salmon. Solid lines (a and b) represent various feed
conversion rates: a = 1.5; b = 2.5. Line c indicates the values for b-a.

spring release, and hatchery spring release methods
for the production and release of masu salmon, focusing on smoltification and recapture ratios; 3)
Collection of distribution and price data in various

regions with attention paid to fish size and the unit
weight market price; and 4) Investigation of fishinggear, fishing effort and fishery production costs formasu salmon in various regions.

Taya et 31.: Economic Problems of the Marine Ranching System

Analysis of Economic Efficiency.......
~

7(A) Autumn release

VI

.1:

(B) Spring releasp

:l

-0
tll

6

(C) Hatchery spring release

0'1

c

·E

...

:l

5

OJ

L-

'+-

0

OJ

...

0'1

4

------- -----,

tll

C
OJ

u
L-

3

OJ

a..

I

:

B

-----------r-- I

2------------f------:
I
I
I
I
I
I

I

•

I
I
I

I
I

I
I

,

o

•

A

I

_ _ --1-

50

80

too

Smoltification ratio (%)

Figure 2
Percentage of returning adults needed in relation to thesmoltification ratio. A=autumn release; B=spring release;
and C=hatchery spring release.

_

The approximate seed production costs expected using each of the three methods are shown in Table 1(Iwate Prefecture Office 1974). Costs are lowest for
the autumn release method and highest for thespring release method. The most important consideration, however, is the combined production andrelease costs per fish in relation to the smoltification
ratio which is graphed for each method in Figure 1,
where each curve indicates a different feed conversion rate. If the smoltification ratio (x) is assumed tobe constant between methods and varies between 50
and 80%, the costs per fish are as follows:•cI

between 18.72/xand 21.48/xyen for the autumn
release method,
between 22.63/ x and 27.63/ x for the hatcheryspring release method, andbetween 34.74/x and 40.99/x yen for the spring
release method.

In actual practice, however, the smoltification ratio
tends to be around 50% for the autumn and spring
release methods, and around 80% for the hatcheryspring release method. When this is taken into consideration, the spring hatchery release method isshown to be the most economically efficient in terms
of production and release costs.
Production and release costs are only one part ofthe overall economic aspects of the Marine Ranching

Table 2
Fry production cost and value of returning fish for chum salmon.

Year group

Investment
salmonranching(million yen)

Production
cost (yen)of salmon
fry/fish1962-65
Average374-540465

1966-70
Average131

Cost of returning salmon (yen)Production cost of
returning salmon
per kg/market
price X 100(%)

per fishper kgMarket price(yen/kg) ofreturning salmon

0.99-1.60
1.35

86.7-241.9
16.51

24.8-69.1
47.2

360-545464

5.8-13.1
9.95

476-767619

1.34-2.90
1.91

70.5-126.0
85.7

20.1-36.0
24.5

439-725586

3.0-6.2
4.29

1971-75
Average992-2489
1455

1.72-3.10
2.58

76.8-139.6
111.6

21.9-36.3
31.9

609-1029
891

2.5-5.76
3.72

1976-79
Average2423-5806
3698

4.02-6.65
5.04

3.5 kg/fish

132

NOAA Technical Report NMFS 106

Table 3
Seed production cost and value of returning fish for masu salmon. (Iwate Prefecture Office 1974.)

Type

Production cost (yen)per fish

Returnratio

Cost (yen) ofreturning masu
salmon per fish

Cost (yen) ofreturning masu
salmon per kg

Market priceof returning
fish (yen/kg)

Cost of returnmasu salmon/Unit
price X 100 (%)

18.72
34.72

x
x

18.72/x34.74/x12.48/ x23.16/x

1500
1500

0.83/x1.54/ x22.63

x22.63/x

15.09/ x

1500

1.01/xAutumn release
Spring release
Spring hatcheryrelease30(A) Autumn release(B) Spring release(C) Hatchery spring release0'';:::;

20

....
~

QJ

:J
~

>
:>
VI
0

u

10

10

20
Recovery ratio

System. Recapture ratios also playa vital role. If themarket price of recaptured fish is assumed to be
2,250 yen/fish, fishery production costs are assumed
to be negligible, the marketing commission rate is
assumed to be 5%, and the conversion ratio assumed
to be 2.0, then the upper and lower ranges of therecovery ratio (y) for each method (as it varies withthe smoltification ratio x) can be calculated as follows:•••

autumn releasemethodspring releasemethodhatchery spring
release method

y = 257.5/ x and y = 94.9/ x,
y = 480.8/xand y = 177.1/x,
andy = 319.2/ x and y = 117.6/ x.

30%

Figure 3
Relationship of the recovery ratio to the cost!
value ratio for masu salmon (Iwate Prefecture
Office 1974.)

Figure 2 plots smoltification ratio versus recovery
ratio for each method. From the data in Figure 1 it
appears that the hatchery spring release method,with an 80% smoltification ratio as compared to only
50% for the other two methods, is most efficient. In
reality, however, the recovery ratio for this method
tends to be lower than for the other two, and thus
research is required to improve the recovery ratio forspring hatchery release fish.
One way of evaluating the economic efficiency of
the Marine Ranching Project is to compare production and release costs with the value of the recovered
fish. Production and release costs as a percentage ofthe unit weight value of recovered fish for chumsalmon (0. keta) are shown in Table 2. Because production costs are a small fraction of the value of a

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Taya et al.: Economic Problems of the Marine Ranching System

133

50
Toyama

%
80

%

A

40

60
30

40 ~------+--+-~---------------j

20 +-+-I--I'----+--t-------\-t-----t-----

20 ~---_H_----;;L-----\''t-+_-----------__j 10 +-hH---f-------'\--t--'r-----

F

M

A

M

J

15

17

13

7

J

A

s

o

Month

5

N

D

F

M

A

M

J

J

Month

Figure 4
Monthly harvest patterns for masu salmon in 1986. (A) shows average body weight of salmon harvested in Hokkaido andHonshu prefectures (X 100 g). 0 = Aomori; + = Hokkaido; X = Iwate. (B) shows average body weight of sal.mon harvested
in Honshu prefectures; 0 = Niigata; • = Akita; X = Toyama; V = Ishikawa.

returning salmon, a recovery ratio of even 2% produces an increase in the economic value of thefishery. The same calculations are shown in Table 3
for masu salmon for each release method; these results are graphed in Figure 3. If the target figure of a
15% recovery ratio were achieved for all methods,
then the cost/value ratio (cost per returning salmon,
value is market price per fish) would vary between 5
and 10%, which is a desirable result. Even if the ratio
is calculated at the present recovery rate of 5%, thecost/value ratios would vary from 15-30%, indicatingthat artificial seed production projects have the potential for contributing to the masu salmon fishery.
Seasonal harvest patterns of masu salmon for selected prefectures in 1986 are shown in Figure 4. In
Hokkaido, most of the fish harvested between October and March are juveniles weighing under 600 g.
The current market value of such undersized masu
salmon is only 500 yen/kg (Fig. 5). The market valueof adult fish over 1,500 g, however, rises to 1,500 yen/
kg. Thus, if these undersized juveniles are allowed tomature before harvesting, the value per fish will risefrom 250 yen (500 g at 500 yen/kg) to 2,250 yen(l,500 g at 1,500 yen/kg). These figures represent anine-fold increase in value. Obviously, an increase ineconomic value of the fishery can be obtained if thejuveniles harvested in Hokkaido are allowed to mature before being harvested elsewhere. Calculations
show that if the fishery were closed from November
to February, an increase of 23% (from 7.6 billion yen
to 9.4 billion yen) could be obtained (Fig. 6). If theclosure were extended to include the entire period
from October to March, the increase would rise to
31 % (from 7.6 billion yen to 10.0 billion yen). A
problem arises in that fishermen in Hokkaido andelsewhere depend heavily on income from sport
fishing during this period and would certainly oppose any move to close or restrict the fishery. Thus,
an integrated plan is required that maximizes theoverall economic value of the fishery while protecting the interests of local harvest groups.

Role of Masu Salmon in Local
and Regional Economies
Our investigations showed that although there is a
noticeable lack of stability in catch volume, masu
salmon play an important role in both the local andregional economies of Hokkaido, Aomori, Iwate, and134

NOAA Technical Report NMFS 106

_

2000

~arch

February

1500

Cl

January~........
C

QJ

2::
QJ
U

.~

1000

Log(p)-2. 484+0. 625Log(BII)

500 L

o

__----l~.I....-------..l'-----~:::_---__::_!;:::_---500

1000

1500

2000

Body weight~

Sport fishing income~

Commercial fishery income

Figure 5
Relationship between priceand weight of masu salmon(Kamuenai Fisheries Cooperative 1987).

800 - l - - - - - - - - - {

c
~
c

~

600

I

<IJ

E

8
c

400

200

o..L~~~:-,.L~~~~~~~-'"-------'

Present

ClosedClosed

Nov-Feb

Oct-MarFigure 6
Evaluation of benefitsof fishery restrictionsin (Hokkaido Salmon
Hatchery 1988).

_ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ Taya et at.: Economic Problems of the Marine Ranching System

135

Table 4
Estimated landings ofmasu salmon in Japan (1984) (in metric tons).
Pacific Ocean
Hokkaido
HonshuJapan Sea
Hokkaido
HonshuOffshorefishing
Coastalfishery
Total
landings
Eacharea
Percentage

187.6

744.1

418.3

562.4

1162.4

224.0

143.4

2.1

750.0

1906.5

642.5

143.4

Total
Hokkaido
Honshu

605.9

744.1

43.5

832.0

1305.82.1

43.5

1437.92049.9

785.7

2.1

43.5

3487.9

76.17

22.53

0.06

1.25

100.0

1200

18.0

1100

16.5

1000

15.0

900

13.5

~

800
700
Ql

OkhotskSea

2656.5(%)

E

NemuroStraits

600

12.0
10.5
9.0

E
::l

15

>

CQl

>.

c

~

500

7.5

e

400

6.0

.2

300

4.5

Ql

ro

>

200

hlate
100

~
1976

77

'.

?

,

_ _ _ _- - . . . - : : - - - - - - - : - . : : . : ••: •:•;0'••,:..--------...~

78

79

8]

80

·····Ishikawa
83
82

84

85

1.5

86

Year

Figure 7
Masu salmon landings of the coastal fisheries by weight (metric tons) and value (value assumed at 1,500 yen/kg).

Niigata prefectures. The total annual catch fluctuates
between 3,500 and 4,000 metric tons (t), the average
price varies from 1,000 to 1,500 yen/kg, and the total
value of the fishery ranges from 35 to 60 billion yen/
year. The geographical distribution of the catch for1984 is shown in Table 4.

Hokkaido accounted for 41 % (1,438 t) and
Honshu for 59% (2,050 t) of the total harvest. Seventy-six percent (2,656 t) of the catch was harvested
in the Japan Sea, 22% (785 t) in the Pacific Ocean,
1.2% (43 t) in the Sea of Okhotsk, and 0.06% (2 t) in
the Nemuro Straits region. Clearly, the Japan Sea

136

NOAA Technical Report NMFS 106

Table 5

Family fishery income 1985 (1,000 yen). t = metric tons. Ministry of Agriculture, Forestry, and Fisheries, government ofJapan (1985).
Familyfishery

fishinggross income

fishing
expensesfishing
net income

net income
rate (%)

Japan Sea area
I t (under)
1-3 t
3-5 t
5-10 tBag netAverage

1,557
3,017
7,208
14,859
6,668
4,212

667
1,641
4,636
11,786
4,410
2,680

890
1,376
2,572
3,073
2,258
1,532

57.1
45.6
35.6
20.6
33.8
36.3

Pacific Ocean area
I t (under)
1-3 t
3-5 t
5-10 tBag netAverage

1,859
3,581
7,470
10,926
5,544
4,220

773
1,301
4,415
7,219
2,611
2,241

1,076
2,280
3,055
3,707
2,933
1,979

57.8
63.6
40.8
33.9
52.9
46.8

fishery is by far the most important, accounting for
40 billion yen in total value as compared to 8-12 billion yen for the Pacific Ocean fishery.
At the national level, coastal fisheries harvested
39% (1,349 t) and offshore fisheries 61 % (2,137 t) of
the total catch. The figures for the Pacific Ocean
fishery, were 53% (418 t) for coastal and 47% (367 t)
for offshore areas; while in the Japan Sea the major
percentages were reversed, with coastal yields 35%(931 t) and offshore harvests at 65% (1,724 t). Catchstatistics by region and fishery are shown in Table 4,
and Figure 7 graphs the volume and value of landings by prefecture for the 10-year period from 1976
to 1986.
Fishing methods for masu salmon were found tovary widely, and include gillnet, bottom trawl,
longline, pole and line, and setnet techniques. In theJapan Sea region of Hokkaido (e.g., Shako tan Peninsula area) sportfishing was found to be an importantsource of income for about 45 vessels. This
sportfishing industry harvests a large percentage ofundersized juveniles, and as such adversely affects theefficiency of the masu salmon Marine Ranching
Project.
Although landings in prefectures such as
Yamagata, Ishikawa, and Toyama are relatively small,masu salmon is none the less a valuable local resource and an important ingredient in many
traditional seafood dishes. Family income by vessel
size is shown in in Table 5 for fishing families in thenorthern Japan Sea and northern Pacific Ocean regions (MAFF 1985). Net family income from fisheries
varies from 0.890 to 3.073 million yen in the Japan
Sea, and from 1.076 to 3.707 million yen in the Pacific Ocean. Because the market price of masu
salmon is relatively high, this fish represents an important source of income for these fishing families.
Thus, investment in masu salmon marine ranchingfacilities will be a crucial element in developing the
local and regional economies in these areas.

Citations
Hokkaido Salmon Hatchery.
1988. Annual report of the Hokkaido salmon hatchery. (InJapanese.)
Iwate Prefecture Office.
1974. Exploitation program of masu salmon, p. 24-25. (InJapanese.)
Ministry of Agriculture, Forestry, and Fisheries.
1985. Annual report on fisheries economics (family
fishery). (In Japanese.)