<figure><img src="image_1.png" /></figure>

HAL
open science

Control of reproduction and fish farming

Roland Billard, Bernard Breton

► To cite this version:

Roland Billard, Bernard Breton. Control of reproduction and fish farming. Current trends in comparative endocrinology, Hong-Kong University Press, 1985, 962-209-039-7. hal-01600838

HAL Id: hal-01600838

https://hal.science/hal-01600838

Submitted on 8 Jun 2020

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_2.png" /></figure>

Distributed under a Creative Commons Attribution - ShareAlike 4.0 International License
# Control of Reproduction and Fish Farming

R. Billard and B. Breton

*Laboratoire de Physiologie des Poissons, Institut National de la Recherche Agronomique, Campus de Beaulieu, Rennes Cedex, France*

In fish farming one of the critical steps is the quantity and quality of available juveniles; they must have a good survival rate and a good growth ability. Juveniles may be taken from the wild, a technique that has been successfully used for the yellowtail (*Seriola quinqueradiate*) and elvers (*Anguilla*), but the availability of many species varies greatly, and fish breeders are often faced with a shortage of fry. Furthermore, the availability may also be seasonal. A better method is by controlling reproduction and maintaining a brood stock. The stock may be taken from the wild with the same limitations outlined above, but a more preferable solution is to maintain a permanent stock of males and females so that fry may be produced by placing both sexes in a pond where natural reproduction can occur (e.g. carp, *Cyprinus carpio*, in a Dubicz pond). However, the success of this approach depends on the climatic conditions and a good knowledge of the optimal environmental conditions for spawning.

To ensure the availability of fry when desired, the tendency now is to control each step of reproduction, that is gametogenesis, ovulation, spermiation and also artificial insemination and egg incubation. For such control, it is necessary to know the physiological factors regulating the process of reproduction, and our knowledge in this field has increased considerably during the last decade.¹,² Our understanding of the role played by environmental factors at each stage of reproduction has also increased substantially;³⁻⁸ some are physical or chemical (photoperiod, temperature, water quality, substrate, current), others are biological (social, nutritional, pathological). These factors do not act independently, but rather interact and undergo substitution.

The management of gametes also requires an understanding of the physiology of the sperm and ova, and the effects that environmental factors such as temperature and water composition have

on their viability. The process of obtaining gametes can be controlled by hormones or by changes in the external factors, but to be utilizable, both the eggs and spermatozoa, when they are stored or being used for insemination, must be maintained in the proper media under the correct physical conditions.

## Control of gametogenesis

### Hormonal treatment

This technique consists of continuously supplying exogenous, gonadotropin or cholesterol implants, and has been attempted in juvenile salmonids and successfully used on males;⁹,¹⁰ vitellogenesis, however, is only slightly stimulated in the females.¹⁰,¹¹ In prepubertal and adult fish a continuous supply of pituitary extracts or GtH initiates a normal spermatogenetic cycle, including spermiation, both in salmonids¹² and eels.¹³ Injec- tion of LHRH twice a week for 6 weeks also ad- vances vitellogenesis and ovulation in the medaka (*Oryzias latipes*).¹⁴ This technique is not very useful in fish farming, however, because it is necessary to handle the fish frequently and the hormone is costly and not always available. It is feasible when native strains are stocked in rivers, e.g. the Fraser, where the pink salmon only spawn in alternate years.¹⁵

### Manipulation of environmental factors

Modification in the timing of gametogenesis has been achieved by changing a few environmental factors and some biological factors.

*Photoperiod.* The reproductive cycle of adult salmonids can be advanced or delayed by chang- ing the photoperiodic regime.¹⁶,¹⁷ Gametogenesis
usually occurs during decreasing photoperiods, and by artificially advancing the normal decreasing regime reproduction is also advanced. This technique is currently used in fish farming,¹⁸ but may entail problems of temperature interference; while the gametogenesis of rainbow trout occurs at temperatures around 18-20°C, lower temperatures are necessary for incubation and, if the farmer wants eggs out-of-season, cool water is necessary. Other species also have been shown to respond to changes in the photoperiodic regime by a modification of the time of maturation, thus mullet (*Mugil cephalus*), kept at a constant temperature of 21°C, mature out-of-season when kept under short days (6L:18D),¹⁹ whereas long photoperiods associated with high temperatures stimulates gametogenesis in *Lepomis cyanellus*²⁰ and *Notemigonus crysoleucas*.²¹ In the sea bass, *Dicentrarchus labrax* the time of reproduction is advanced by advancing the normal decreasing photoperiod (Barnabé, unpublished) or by dephasing the photo-thermoperiodic regime.²²

*Temperature.* In some species the influence of temperature on gametogenesis appears to be more important than the photoperiod. This is the case in carp. These fish are seasonal breeders in the temperate zone but reproduce several times a year under a constant temperature (20°C).²³ The thermal requirements are different for gametogenesis and spawning in species which are usually spring or early summer spawners. In some cases, these requirements vary according to the stage of gametogenesis. In *Microgrex terrae sanctae*, vitellogenesis occurs only at low temperatures but oogonial proliferation occurs at high temperatures.²⁴ A minimal temperature is also necessary to initiate vitellogenesis in the tench.²⁵ The optimal temperature for the metabolization of the various hormones may also be different. Many other physico-chemical factors, such as salinity, pollution, physically-induced stress and chemically-induced sterilization, are also important and require further investigation.

*Nutrition.* Food requirements for brood stock are very crucial since gonadal weight can reach 30% of body weight. The quality of the food is important in relation to egg quality.²⁶ Lack of food may interfere with the pattern of GtH secretion; for in-

stance, the starvation of goldfish, *Carassius auratus*, kept at 30°C, results in changes or suppression of the daily pattern of plasma GtH, and thus inhibits reproduction.²⁷ This suggests that a daily cycle of GtH is of major importance in gonadal activity.⁸

*Other biological factors.* Overcrowding inhibits gametogenesis,²⁸,²⁹ and parasites may also prevent gonadal development.³⁰

*Inhibition of gametogenesis*

Another aspect of the control of gametogenesis is the prevention of gonadal development. Sterile fish are expected to grow faster with a better conversion rate because they do not have to develop gonads and the quality of the flesh is higher than in mature fish. This problem has been reviewed recently;¹⁷ there are several possible strategies but none are yet available to the fish farmer. Gametogenesis may be inhibited by supplying androgens via the diet during the period of gonadal activity, or further gonadal development may be prevented by giving sex hormones to sterilize juvenile fish at the time of sexual differentiation. However, using steroid hormones implies the possible toxicity of some residues and its psychological effect on the consumer. Other methods of sterilization, such as irradiation, triploidy and interspecific hybridization, have been explored.¹⁷

*Control of ovulation and spermination*

The final event of the reproductive cycle corresponding to gamete release (ovulation and spermiation) can be controlled by placing fish in the appropriate environment or by changing some internal regulatory factors such as the supply of hormones or neurohormones. Endocrine controls may be by a direct stimulation on the gonads using pituitary extracts and/or steroids, the so-called hypophysation technique,³¹,³² or by stimulation of the pituitary to induce endogenous GtH release, using mammalian LHRH or an analog LHRHa, or by influencing the gonadal feedback on the pituitary with substances such as the antiestrogens, chromophene citrate or tamoxifen.
## Induction of ovulation

In the actual practice of fish farming, ovulation is induced in two types of fish, namely, species which do not spawn spontaneously in fish-farm conditions or which would spawn in such conditions that most of the eggs would be lost, and species such as salmonids, in which ovulation, occurring spontaneously, can be advanced or synchronized for practical purposes by hormonal treatment.

**Induced ovulation.** In the case of direct stimulation of the ovaries, the most common way is to inject crude pituitary extracts or partially purified gonadotropins such as SGG 100³³ and PPSG.³⁴,³⁵ Among the mammalian gonadotropins, only HCG in very high doses (up to 50,000 IU/kg in mullet³⁸) has been shown to be potent in a limited number of fish species, such as sea beam (*Sparus aurata*)³⁶ and sea bass.³⁷ Very high doses of various steriods have been used with varying success,³⁹ and, so far, they are not widely used. A combination of pituitary extracts and 17α-hydroxy-20β-dihydroprogesterone (17α-20βP) has been successfully used in carp, but 17α-20βP given alone does not induce follicular rupture.⁴⁰

The induction of ovulation by stimulating the pituitary secretion of endogenous GtH, has been used only recently in a few species of fish. The successful use of LHRHa injections to phytophagous fish has been reported in China, and this method is now currently used in fish farming there.⁴¹ Attempts to use LHRH on other species has not been successful, and very high does must be used for limited success; 100 µg of LHRH/kg in tench,⁴² 10 mg/kg in goldfish⁴³ and 50 µg of LHRHa/kg in carp (Crim et al., unpublished). In pike, another LHRHa, (Gly 10-[D-try⁶] LHRH ethylanide), has no effect on ovulation.³⁵ Some reasons for this limited success have been discussed;³⁵ among them are possible species specificity and also methods of delivery – the pituitary portal system is not well-developed in some fish species. The number and timing of injections are also important; in carp the overall amount of GtH released increases after repeated injections (Fig. 1). It should be noted that the pituitary response changes according to the stage of gametogenesis.⁴⁴ More generally, basic information is lacking on endogenous

<figure><img src="image_3.png" /><figcaption>**Fig. 1.** Changes in plasma c-GtH in carp after 3 successive LHRHa injections given 3h apart. Rearing temperature: 18°C. The treatment was not followed by ovulation but germinal vesicle breakdown (GVBD) occurred in some females (n = 5 per group).</figcaption></figure>

LHRH levels and the half-life of injected compounds.

Antiestrogens, such as cis-chlomiphene citrate, induce the release of GtH in carp⁴⁵ and goldfish,⁴⁶ and in the latter, ovulation has also been induced by a lesion of the preoptic region.⁴⁷ This has also been reported in common carp by Peter et al. (unpublished), and strongly suggests a central inhibition of GtH secretion by the pituitary, and the existence of a possible gonadotropin release-inhibitory factor, at least in cyprinids.

**Advancement and synchronization of ovulation in salmonids.** Ovulation is usually stimulated in this group of fish by exogenous hormones. In rainbow trout the normal spawning time is advanced by 4 to 6 weeks after combined injections of trout pituitary extract (3.25 µg t-GtH/kg), followed 2 days later by 17α-20βP (3 mg/kg).⁴⁸ In species sensitive to captivity, such as the salmon, this method reduces the period of acclimatization and limits mortality.⁴⁹,⁵⁰ A combination of pituitary extracts and LHRHa has recently been used for the same purpose.⁵¹ This method has also been used in brown trout (*Salmo trutta*) to advance and
stimulate ovulation which is considerably retarded when the temperature drops to less then 2 to 3°C (Fig. 2). When reproduction is advanced by photoperiodic change to mid-summer, ovulation can be synchronized, making it possible to accurately predict ovulation and to take the eggs quickly, thus avoiding the rapid aging of the ova which occurs at high summer temperatures. Spawning has been advanced in rainbow trout by combining the use of warm water (to stimulate body growth at early stages), changing the photoperiod and hypophysation.⁵²

*Manipulation of external factors.* Some environmental factors, such as dissolved oxygen in carp⁵³ and the presence of spawning substrate in goldfish,⁵⁴ enhance or trigger ovulation. In these species the timing of the ovulatory GtH surge is synchronized by the photoperiod.⁸,⁵⁴ Nutritional factors are also important, and a protein-rich diet should be given to carp just before spawning.⁵⁵ After spawning, heavy feeding reduces female mortality. The social environment may interfere with ovulation and spawning, as well as with gametogenesis. Spawning and successful reproduction may be inhibited when fish are kept at high densities.²⁸,⁵⁶ Spawning may be stimulated by sex pheromones,⁵⁷ and these have been identified and isolated in some fish species.⁵⁸ Other factors, such as changes in the chemical composition of the water⁵⁹ and changes in temperature and the physical environment in 'ovulatory' ponds or

<figure><img src="image_4.png" /></figure>

**Fig. 2.** Advancement of synchronization of ovulation in brown trout. The normal spawning season peaks in early December. Each female received one pituitary taken from a male of the same age. Ovulation was advanced to the end of October in one group and late spawning females, in which ovulation was delayed due to low temperature, were induced to ovulate at the beginning of January. Numbers in each group in brackets.

<figure><img src="image_5.png" /></figure>

**Fig. 3.** Stimulation of GtH secretion and spermiation in carp after 5 daily injections of LHRHa (n = 7 per group) (C. Weil and F. Takashima, unpublished).

aquaria, are also probable factors influencing gametogenesis. More investigation is needed in this field; the idea is to provide the optimal environment for brood stock during spawning, thus potentiating the effects of the various treatments and, in some cases, ovulation without hypophysation.

*Stimulation of spermiation*

The spermatogenetic production of teleost fish is very high but the spermiation yield is poor and sometimes only a low percentage of the spermatozoa present in the testes are released during the period of spermiation.⁶⁰ In some species, such as pike (*Esox lucius*), the volume of milt collected is very low and stimulation is necessary.³⁵ In other species, the amount of milt produced must be increased for better use of the brood stock; pituitary hormones, sex steroids³⁴ and LHRHa, which raise plasma GtH and sperm volume (Fig. 3),⁶¹ have all been used to stimulate spermiation.

*Conclusions*

The technique of hypophysation has allowed many species to be developed for aquaculture.⁶²,⁶³ However, this technique is primitive, and its efficiency (½ of females selected and ovulated over
the total number of the brood stock) is not clearly established, nor its mortality after spawning determined. A mortality rate of 57% has been reported in China after HCG or pituitary extract treatment, but this rate dropped to 3-12% after the use of LHRHa.³² The biological activity of crude pituitary extracts, which are very widely used, is usually not known, although routine bioassays are available. The dose may be too low (and not efficient) or too high; even if ovulation occurs, data on the consequences on the following reproductive cycle are not available. In addition, crude extracts include several hormones and factors which may not be at all desirable.

Species specificity of gonadotropin⁶⁴ is another problem; only salmonid and carp pituitaries are now commercially available and, if they are to be used on distantly-related species, higher doses must be given. Moreover, the formation of antibodies, shown for HCG⁴¹ (Avtalion, personal communication), may cause immunological problems.

The timing of hormone injection is also a factor since plasma GtH varies daily when females reach full maturity,⁸,⁶⁵ and the GtH receptors show daily cyclicity.⁶⁶ Furthermore, the metabolic clearance rate varies.⁸

Another serious problem concerning induced ovulation is the definition of the optimal stage of ovarian development in order to apply the treatment.³⁹ The size of the abdomen is measured in phytophagous fish⁶⁷ and a corresponding hormone dose is given. Follicular diameter has been used in mullet⁶⁸ and the position of the germinal vesicle in carp,⁶⁹ pike⁷⁰,³⁴ and salmonids.⁴⁸ However, there is still great individual variability in response to the treatment and more precis criteria are needed.

The possibilities of stimulating endogenous pituitary GtH secretion are interesting. This would solve the problem of the availability of pituitary extracts and the species specificity of gonadotropin. The use of LHRHa, a standard product of known biological activity, is still far from being applicable to all species; some species are obviously easier to spawn than others. More research is necessary to determine the level of the ovulatory GtH surge and the structure of some fish gonadotropin-releasing hormones which are probably different from LHRH and should be

more potent. A better understanding of the endocrinology of fish reproduction is absolutely crucial to all these problems. Spawning may, in the future, be partly induced by hormone supply and a more adapted environment, but more indirect ways, such as changing the endocrine balance (inhibitory or stimulatory feed back) must be considered.

## Gamete management

The management of gametes in the fish farm remains primitive. The gametes are collected and mixed together with water. Efficiency is very poor since one male is used to fertilize one or only a few females. Several points can be improved: (i) collection of the gametes at the right time after ovulation or during the period of spermiation. The milt cannot be stripped in species like the catfish (*Silurus glanis*) and other techniques of sperm release are required; (ii) more efficient use of gametes via an effective technique of artificial insemination; and (iii), incubation in optimal environmental conditions.

## Optimal collection time

The gametes undergo aging processes after they are released from the follicles or the testicular lobules. The *in vivo* survival of the gametes in the genital tract depends mostly on the sex, the species, and the temperature. Post-ovulatory survival of rainbow trout eggs varies from 6-10 to 36 days.⁵⁷,⁷¹,⁷² The percentage of deformed larvae increases with the aging of the ova (Fig. 4).⁵⁴ In trout, a high level of GtH remains in the plasma for some time when the ova are left in the body cavity,⁷³ but this does not seem to be related to ovum survival (Fig. 4). It has been suggested that factors originating from post-ovulatory corpora lutea may interfere with ovum survival.⁷⁴ The survival of pike ova at 10°C does not exceed 2 days.³⁴ In cyprinids, survival is much shorter, a few hours in common carp and about one hour in phytophagous fish.⁵⁵ In these species, the eggs should be collected very quickly after ovulation, the time of ovulation, sometimes being identified by signs of spawning behavior. As the hypophysiation-ovulation interval depends on the temperature, an
<figure><img src="image_6.png" /><figcaption>Fig. 4. Post-ovulatory survival of ova left in the body cavity of female rainbow trout. Survival was measured by the percentage of total embryonated eggs over the total number of eggs and the percentage of abnormal embryos. Plasma s-GtH was measured during the same period.</figcaption></figure>

exact knowledge of the thermal regime helps in predicting the time of ovulation. Sometimes to avoid overripening, male and female phytophagous fish are stored in circular tanks where they spawn spontaneously and the eggs are collected at the tank outlet.

Concerning sperm survival, its quality tends to decrease during the spawning season; in sea bass the motility decreases,⁷⁵ in pike the duration and intensity of motility slightly decrease at the end of a 2-month sampling period.⁷⁶ In rainbow trout, the ability for cryopreservation tends to decline towards the end of the spawning season.⁷⁷ It should be kept in mind that the quality of the sperm is not constant throughout the spawning season and that the amount of sperm used in artificial insemination varies accordingly. Moreover, the short duration of ovum survival after ovulation in some species limits considerably the possibility of collecting ovulated females in the wild. Induced breeding is therefore more convenient.

Fertilization in fish usually takes place externally and the operation of artificial insemination is very simple. The object is to make efficient use of the gametes available, but several problems in this domain are lacking a solution.

## Technique of artificial insemination

In the actual practice of artificial insemination, water is added to the gametes. Neither fresh water nor sea water is the best medium for gamete survival.⁷⁸ For instance, the sperm of many species do not survive for more than 1 min in fresh water or the ova for more than 4 min. It is therefore more practical to use a diluent which improves gamete survival and fertilization yield. Such diluents have been developed but their composition varies according to the gamete physiology of the individual species. Good results have been obtained in salmonids and pike with a saline solution (osmotic pressure 250 mOsm/kg and pH 9);¹⁷ the optimal osmotic pressure for carp is 120 mOsm with pH 8-9. For sea bass and sea bream, optimal salinity is about 20 ppm.⁷⁹ Many other factors, such as temperature at the time of insemination and mechanical and thermal shock, are also implicated.

### Minimum number of spermatozoa needed.
The number of spermatozoa needed to fertilize one ovum is a basic factor in any technique of artificial insemination. Sperm volumes vary from 10,000 – 300,000 for trout,⁸⁰ 400,000 – 700,000 for pike, and 13,000 – 30,000 for carp (Marcel *et al.*, unpublished).

The optimal ratio of spermatozoa to ova should be determined in a well-defined volume of diluent to ensure a good mixture of gametes. Sperm dilution can be as high as 1/10,000, and for carp and pike approximately one litre of eggs is diluted in one litre of diluent, then 1 ml of sperm added immediately afterwards. The amount of sperm for trout is 1 ml at the onset of the spawning season and 3 ml at the end.⁸⁰ After mixing by pouring back and forth twice, the eggs are left for 5 (pike) or 15 (trout) min then transferred to incubators.

### Sticky eggs.
The eggs of some species are very sticky and may aggregate and die from anoxia in conditions of artificial incubation. It is therefore essential to remove the sticky layer surrounding them before they are transferred to the incubator. In pike and *Thymallus* this layer is simply removed in the insemination diluent, while *Corregonus* eggs are washed in an acid solution (pH 6.2-6.9).⁸¹ The removal of carp sticky layer is more complex,
and in addition the eggs must be treated for 45 min
in a solution containing 4 g of NaCl + 20 g of
urea/1.⁸²

Incubation

Various devices are used for incubating fish eggs. Salmonid eggs are usually kept immobile on a tray. Carp, pike and most marine fish eggs are incubated in small or large conical jars where they float and are constantly stirred at a slow rate. Many environmental factors, such as dissolved oxygen, water temperature, thermic and mechanical shock, light intensity, salinity, pH, disease and even prophylactic treatment,¹⁷ may interfere with the quality of embryonic development. Endogenous egg quality is important too.⁸³ These factors vary according to the species and the stage of development. The incubation conditions are relatively well-defined for some old cultured species (cyprinids, salmonids) but they still need to be thoroughly investigated for new cultured species. It is better to incubate the eggs from each female separately and eliminate the females which produce poor-looking eggs.

Discussion

We have noted that the management of gametes is
strongly linked to brood stock management. The
control of the reproductive cycle and gamete
release should yield sperm and ova of very good
quality which must be efficiently used. The hat-
chery is the key unit in fish farming, integrating all
aspects of reproductive control. However, at pre-
sent, the control of reproduction is in a very early
stage of development; only a few species are really
domesticated (and therefore used for experimenta-
tion), and only some of the various steps of
reproduction can be controlled. With the excep-
tion of the wide use of LHRHa in China, progress
in fish reproductive endocrinology during the last
decade has not yet had a noticeable impact on fish
farming. There is no standard gonadotropin pre-
paration available to the fish farmer who, instead,
often uses the wrong hormone at the wrong dose
at the wrong time; the technique of artificial in-
semination and the concept of gamete manage-
ment discussed in the present paper, are not yet

widely used in the practice of fish farming.
Modern techniques have obviously not progressed
from the experimental stage in the laboratory to
practical use on the fish farm, even in developed
countries, and programs for their development are
therefore necessary.

There is a great need for controlling the
reproduction of new species that are candidates
for aquaculture, or species in which induced
breeding is attempted. Such techniques should in-
tegrate the various aspects of reproductive
physiology and internal and external factors of
regulation. Hormone supply, although very
popular, is not the only approach. The choice of
the right technique should be based on a good
knowledge of the reproductive physiology of each
species. The techniques must also be simple and
easy to apply in fish farms. The number of species
may be limited. Even then it will not be possible to
carry out intensive fundamental research in all of
them, but the fish farmer should be able to benefit
from basic information gathered on some model
species and adapted through studies of com-
parative physiology and endocrinology. Since the
FAO workshop in 1976,⁸⁴ some methods, such as
the use of mammalian gonadotropins, have not
turned out to be widely applicable; others
(hypophysation and incubation) have had a very
limited development. However, some new per-
spectives, e.g. the use of LHRH or a combination
of hormones, the choice of the environment and
good management of brood stock and gametes,
have opened up.

Acknowledgements

We wish to thank Ms A. Daifuku for editing the
English manuscript and Drs A. Fostier and B.
Jalabert for their fruitful criticism.

References

¹ Billard, R. & Breton, B. (1978). In *Rhythmic Activity of Fishes* (eds. Thorpe, J.E.), pp. 31–45. Academy Press, New York and London.

² Peter, R.E., & Crim, L.W. (1978). *Ann. Rev. Physiol.* **41**, 323–35.
³ De Vlaming, V.L. (1974). In *Control of Sex in Fishes* (ed. Shreck, C.B.), pp. 13–63. Virginia Polytechnic Institute and State University, Blackburg, Virginia.

⁴ Billard, R., Breton, B., Fostier, A., Jalabert, B. & Weil, C. (1978). In *Comparative Endocrinology* (eds. Gaillard, P.J. & Boer, H.H.), pp. 37–48. Elsevier/ North Holland Biomedical Press, Amsterdam.

⁵ Breton, B., Fostier, A., Jalabert, B. & Weil, C. (1980). In *La Pisciculture en Etang* (ed. Billard, R.), pp. 149–61. INRA Paris.

⁶ Peter, R.E. & Hontela, A. (1978). In *Environmental Endocrinology* (eds. Assenmacher, I. & Farner, D.S.), pp. 20–25. Springer Verlag, Berlin and Heidelberg.

⁷ Larsen, L.O. (1981). In *3th Congr. Europ. Soc. Comp. Physiol. Biochem., Holland, Sept 1981*. Pergamon Press, Oxford.

⁸ Peter, R.E. (1981). *Gen. Comp. Endocrinol.* **45**, 294–305.

⁹ Robertson, O.H. & Rinfret, A.P. (1957). *Endocrinology* **60**, 559–62.

¹⁰ Magri, M.H., Billard, R., Reinaud, P. & Fostier, A. (1981). In *11th Conf. Europ. Soc. Comp. Endocrinol., Jerusalem*.

¹¹ Funk, J.D., Donaldson, E.M. & Dye, H.M. (1972). *Can. J. Zool.* **51**, 493–500.

¹² Funk, J.D. & Donaldson, E.M. (1972). *Can. J. Zool.* **50**, 1413–19.

¹³ Fontaine, M. (1936). *C.R. Acad. Sci. (Paris)* **202**, 1312–14.

¹⁴ Chan, K.K. (1977). *Can. J. Zool.* **55**, 155–60.

¹⁵ Mackinnon, C.N. & Donaldson, E.M. (1978). *Can. J. Zool.* **56**, 86–89.

¹⁶ Htun-Han, M. (1977). *Library Inf. Leaflet, MAAF Lowestoft*, **6**, p. 30.

¹⁷ Billard, R. (1981). *World Conf. Aquacult. Vencie*. In press.

¹⁸ Nomura, M. (1982). *Bull. Jap. Soc. Sci. Fish.* **28**, 1070–76.

¹⁹ Kuo, C.M., Nash, C.E. & Shehaded, Z.H. (1974). *Aquaculture* **3**, 25–43.

²⁰ Kaya, C.M. & Hasler, A.D. (1972). *Trans. Am. Fish. Soc.* **101**, 270–75.

²¹ De Vlaming, V.L. (1975). *Biol. Bull.* **148**, 402–15.

²² Girin, M. & Devauchelle, N. (1978). *Ann. Biol. Anim. Bioch. Biophys.* **18**, 1059–65.

²³ Kossmann, H. (1976). *EIFAC Tech. Pap.* **25**, 122–26.

²⁴ Yaron, Z., Cocos, M. & Salzer, H. (1980). *J. Fish. Biol.* **16**, 371–82.

²⁵ Breton, B., Horoszewicz, L., Bieniarz, K. & Epler, P. (1980). *Reprod. Nutr. Develop.* **20**, 1011–24.

²⁶ Omae, H., Suzuki, S.R. & Shimma, Y. (1978). *EIFAC Tech. Pap.* F-14, 2–20.

²⁷ Gillet, C., Billard, R. & Breton, B. (1981). *Cahier Lab. Montereau* **11**, 49–56.

²⁸ Greene, G.N. (1970). Ph.D. Diss. Auburn University.

²⁹ Yu, M. & Perlemutter, A. (1970). *Growth* **34**, 153–75.

³⁰ Hoffman, G.L. & Meyer, F.P. (1974). T.F.H. Publ. Inc., Neptune, N.J.

³¹ Pickford, G.E. & Atz, J.W. (1957). *New York Zool. logical Society*, New York.

³² Harvey, B.J. & Hoar, W.S. (1979). Intern. Devel. Res. Center, Ottawa, Ontario.

³³ Donaldson, E.M., Yamasaki, F., Dye, H.M. & Philleo, W.W. (1972). *Gen. Comp. Endocrinol.* **18**, 469–81.

³⁴ De Montalembert, G., Bry, C. & Billard, R. (1978). *Am. Fish. Soc. Spec. Publ.* **1**, 217–25.

³⁵ Billard, R. & Marcel, J. (1980). *Aquaculture* **21**, 181–95.

³⁶ Gordin, H. & Zohar, Y. (1978). *Ann. Biol. Anim. Bioch. Biophys.* **18**, 985–90.

³⁷ Barnabé, G. & René, F. (1972). *C.R. Acad Sci. (Paris)*, sér. D **275**, 2741–44.

³⁸ Kuo, C.M., Shehadeh, Z.H. & Nash, C.E. (1973). *Aquaculture* **1**, 429–32.

³⁹ Shedhadeh, Z.H. (1976). *EIFAC Tech. Pap.* **25**, 72–89.

⁴⁰ Jalabert, B., Breton, B., Brzuska, E., Fostier, A. & Wieniawaski, J. (1977). *Aquaculture* **10**, 353–64.

⁴¹ Coopertive Team for Hormonal Application in Pisciculture (1977). *Scientia Sinica* **20**, 469–74.

⁴² Kouril, J. & Barth, T. (1981). *Bull. VURH Vodnany* **9**, 13–18.

⁴³ Lam, T.J., Pandey, S. & Hoar, W.S. (1975). *Can. J. Zool.* **53**, 1189–92.

⁴⁴ Weil, C. (1981). Thèse Doctorat, Univ. P. et M. Curie Paris, p. 250.

⁴⁵ Breton, B., Jalabert, B. & Fostier, A. (1975). *Gen. Comp. Endocrinol.* **25**, 400–404.

⁴⁶ Pandey, S., Stacey, N. & Hoar, W.S. (1973). *Can. J. Zool.* **51**, 1315–16.

⁴⁷ Peter, R.E. & Paulencu, C.R. (1980). *Neuroendocrinology* **31**, 133–41.

⁴⁸ Jalabert, B., Breton, B. & Fostier, A. (1978). *Ann. Biol. Anim. Bioch. Biophys.* **18**, 977–84.

⁴⁹ Jalabert, B., Goetz, F.W., Breton, B., Fostier, A. & Donaldson, E.M. (1978). *J. Fish. Res. Board Can.* **35**, 1423–29.

⁵⁰ Jalabert, B., Le Bail, P.Y. & Cuinat, R. (1980). *Bull. Fr. Pisc.* **279**, 57–64.

⁵¹ Donaldson, E.M., Hunter, G. & Dye, H.M. (1981). *Aquaculture* **26**, 129–41.

⁵² Jennerich, S. (1978). *Fischerei-Forschung* **16**, 7–19.

⁵³ Horvath, J. & Peteri, A. (1980). *Aquacultura Hungarica* (Szarvas) **11**, 15–18.

⁵⁴ Stacey, N.E., Cook, A.F. & Peter, R.E. (1979). *J. Fish. Biol.* **15**, 349–61.

⁵⁵ Horvath, L. (1978). *Aquaculture Hungarica* (Szarvas) **1**, 58–65.

⁵⁶ Fitzgerald, G.J. (1976). Ph.D. Thesis. Univ. West Ontario, Canada.

⁵⁷ Kausch, H. (1976). *EIFAC Tech. Pap.* **25**, 43–52.

⁵⁸ Algranati, F.D. & Perlmutter, A. (1981). *Env. Biol. Fish.* **6**, 31–38.

⁵⁹ Lake, J.S. (1967). *Austr. J. Mar. Freshwater Res.* **18**, 137–53.

⁶⁰ Billard, R., Fostier, A., Weil, C. & Breton, B. (1982). *Can. J. Aqua. Sci. Fish.* **39**, In press.

⁶¹ Peter, R.E. (1980). *Can. J. Zool.* **58**, 1100–1104.
62 Fontaine, M. (1976). *J. Fish. Res. Board Can.* **33**, 922-39.

63 Caudhuri, H. (1976). *J. Fish. Res. Board Can.* **33**, 940-47.

64 Breton, B., Billard, R. & Jalabert, B. (1973). *Ann. Biol. Anim. Bioch. Biophys.* **13**, 347-62.

65 Zohar, Y., Breton, B. & Billard, R. (1981). In *The II Conf. Europ. Soc. Comp. Endocrinol.*, Jerusalem.

66 Kuo, C.M., Watanabe, W.O. (1978). *Ann. Biol. Anim. Bioch. Biophys.* **18**, 946-65.

67 Antalfi, A. & Tölg, E. (1976). *EIFAC Tech. Pap.* **25**, 111-21.

68 Shedhadeh, Z.H., Kuo, C.M. & Milisen, K.K. (1973). *J. Fish. Biol.* **5**, 489-96.

69 Brzuska, E. & Bieniarz, K. (1977). *I.R.S.* **105**, 1-26.

70 De Montalembert, G., Jalabert, B. & Bry, C. (1978). *Ann. Biol. Anim. Bioch. Biophys.* **18**, 969-75.

71 Bry, C. (1981). *Aquaculture* **24**, 153-60.

72 Escaffre, A.M. & Billard, R. (1979). *Bull. Fr. Pisc.* **272**, 56-70.

73 Jalabert, B. & Breton, B. (1980). *C.R. Acad. Sci. (Paris,)* sér. D. **290**, 799-801.

74 Lam, T.J., Nagahama, Y., Chan, K. & Hoar, W.S. (1978). *Can. J. Zool.* **56**, 2029-36.

75 Billard, R., Dupont, J. & Barnabé, G. (1977). *Aquaculture* **11**, 363-67.

76 De Montalembert, G., Marcel, J. & Billard, R. (1980). *Bull. Fr. Pisc.* **276**, 90-103.

77 Legendre, M. & Billard, R. (1980). *Reprod. Nutr. Develop.* **20**, 1859-68.

78 Billard, R. (1978). *Aquaculture* **14**, 187-98.

79 Billard, R. (1978). *Actes de colloques du CNEXO* **8**, 59-73.

80 Billard, R. (1975). *Bull. Fr. Pisc.* **257**, 121-35.

81 Prokes, H. (1975). *Zool. Listy* **24**, 185-96.

82 Woynarovich, E. & Woynarovich, A. (1980). *Aquacultura Hungarica* (Szarvas) **11**, 19-21.

83 Zhukinskiy, B.N. & Nedyalkov, G.F. (1980). *Hydrobiol. J.* **16**, 51-65.

84 EIFAC (1976). *EIFAC Tech. Pap.* **25**, p. 180.
Current Trends in
Comparative Endocrinology

Editors

B. Lofts and W. N. Holmes

Volume Two

<figure><img src="image_7.png" /></figure>

Hong Kong University Press

<figure><img src="image_8.png" /></figure>