How The Understanding of Sex Determination in Fish can Help Improve the Control of Phenotypic Sex in Aquaculture

How The Understanding of Sex Determination in Fish can Help Improve the Control of Phenotypic Sex in Aquaculture

Introduction

Sex determination or control is a very important and highly targeted area of aquaculture research; due to its influences on productivity, husbandry management and economics. The ability to control sexual differentiation, reproduction and maturation, gives aquatic farmers, the control over breeding processes, both in hatcheries and throughout the grow-out period. It could be argued that in aquaculture species, which have become global merchandise, primary facilitators of their largescale industrial production, can be attributed to the control over sex and reproduction. However, in some species, where production is yet to reach industrial scale, the definition of sex differentiation and improved dependability of reproduction remains an important area of applied research (Budd et al., 2015).The need for sex control in aquaculture is evidently expressed in the desire to achieve several broad goals in the industry including: prevention of uncontrolled reproduction and precocious maturation e.g. in tilapia; the desire for monosex population as a result of sexual growth dimorphism and economic value of the sexes; to reduce the impact of phenotypic sex on product quality, as in the production of Atlantic salmon and oysters; increase the stability of mating systems in species such as groupers; and prevention of the negative impacts which result from the unintentional introduction (escapees) of genetically improved strains or non-indigenous species to the culture environment. The culture system and reproductive biology of the species concerned, has a great influence on the relative importance of each goal mentioned above.

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Precocious maturation materializes in several cultured species such as: the Nile tilapia Oreochromis niloticus, (Nash and Novotny, 1995); freshwater crayfish, Cherax destructor, (Lawrence et al., 2007); and Atlantic salmon, Salmo salar; all of which have the tendency to attain sexual maturation and reproduce before reaching the suitable size for harvest. Sexual maturation impairs growth, as energy is diverted to reproduction, creates variances in size at harvest, and results in overpopulation of the culture system; which in turn creates an inability to control feeding rates and animal density. In addition, deterioration in flesh quality, as commonly observed in female Atlantic salmon, is as a result of sexual maturity being attained through the diversion of energy, like lipids, towards reproductive processes, thereby, resulting in differences in economic values between sexes (Piferrer et al., 2009). In culture species like the Nile tilapia, sex specific growth rate provokes the desire for monosex culture as males grow faster with lower food conversion ratios than females (Al-Hafedh and Alam, 2007). Also, female kuruma prawns (Penaeus japonicus) tend to be larger in size at harvest, than males (Coman et al., 2008). As a result, aquatic farmers have adopted several approaches to produce monosex populations for culture, both manually by hand sexing or selective removal, and/or the use of technologies like: exogenous hormones treatment e.g. 17-estradiol in Atlantic cod, Gadus morhua, (Lin, Benfey and Martin-Robichaud, 2012), or 17 ⍺-methyltestosterone in Nile tilapia, Oreochromis niloticus, (Kwon et al., 2000); chromosome ploidy manipulation in rainbow trout, Oncorhynchus mykiss; hybridization between, Oreochromis aurea and  Oreochromis niloticus; environment manipulation i.e. manipulation of social factors in orange-spotted grouper, Epinnephelus coiodes, for the production of male broodstock(Liu and Sadovy de Mitcheson, 2010) or temperature treatment during gonadal differentiation in European sea bass, Dicentrarchus labrax, (Navarro-Martín et al., 2011); and the use of maker assisted selection e.g. in the production of monosex populations in Turbot, Scophthalmus maximus, (Martínez et al., 2009). In hatcheries, sex control is also important for the production of seedstock, particularly if the aim is to produce specific family combinations for selective breeding (Budd et al., 2015).

Sex Determination and Differentiation in Fish

Evolutionarily, sex determination and differentiation in fish, is a very diverse and highly plastic developmental mechanism (Kobayashi, Nagahama and Nakamura, 2013). Such diversity makes it challenging when trying to cultivate a general understanding of sex in fish. However, at the individual or species level, profound plasticity of sexual phenotype, can present increased opportunity for sex control, which is very important for cultured fish. Sex determination can be described as the mechanism, genetic or environmental cues, which ultimately controls the sex of an individual (Devlin and Nagahama, 2002). For example, inheritance of the Y chromosome in mammals, determines that an individual will develop as male. This type of sex determining mechanism, is referred to as genotypic sex determination (GSD), where sex is determined at conception, and there is the expectation of genetic differences between sexes. However, during embryonic development in reptiles, the temperature experienced, rather than genetics factors, provides the cue for sex determination, a sex determining mechanism referred to as environmental sex determination (ESD), where there are inconsistences in the genetic differences between sexes and sex is determined after fertilization, in response to environmental cues. However, sex differentiation refers to the physical materialization resulting from sex determining cues. It largely pertains to the transformation of an undifferentiated primordium into testicular or ovarian tissues, which follows on from a sex determining cue (Devlin and Nagahama, 2002). In some cases, there is a partial overlap between the processes of sex determination and gonadal differentiation, resulting in the terms being used interchangeably (Penman and Piferrer, 2008; Heule, Salzburger and Bohne, 2014).

Techniques Used to Manipulate Sex in Aquaculture

Given the immense variability and complexity of sex determining mechanisms in fish, no single approach has been proven effective for sex manipulation in all cultured fish species. Rather, as mentioned above, several major approaches including:

Utilisation of Exogenous Hormones to Control Sex

 

Due to the ease of application, on a commercial scale, and its consistency in producing monosex population, administration of exogenous hormones to control sex, is considered as the most frequently used and reliable technique of all the external factors know to control sexual development in fish. Sex steroids, steroid hormones known to interact with androgen and oestrogen receptors, are critically important to the natural processes of phenotypic sex determination, therefore, providing the basis for utilisation of exogenous sex steroids to manipulate sex ratios in cultured fish. This technique was first successfully demonstrated in medaka, Oryzias latipes, were androgen and oestrogens were administered to sexually undifferentiated fish, resulting in functional males and females respectively (Yamamoto, 1953 & 1958). Onwards from these early experiments, similar treatments have also been administered to several fish species, demonstrating the possibility that sex steroid therapy, can alter the natural course of sex differentiation in fish, towards a desired gonadal phenotype (Pandian and Sheela, 1995). For example, the use of 17 ⍺-methyltestosterone, an exogenous androgen, has proven effective in 35 fish species for the masculinisation of genetically female fish. However, despite the success of this technique, with many aquaculture species, the use of hormones for sex reversal in fish should be undertaken with extreme caution, in other to prevent any adverse effects to the animal, the farmers, consumers of the end product and the culture environment. For instance, an overdose or prolonged treatment with hormones may lead to deformities in the animal or skew the sex ratio towards the undesired sex (Beardmore, Mair and Lewis, 2001). This use of hormones in hatcheries, require careful handling to prevent adverse effects on human health and limit its impact on the environment. This is facilitated by proper and safe disposal of hormone laden waters. Within European Union (EU) and its member states, the direct use of hormones to commercial grow outs food fish, is banned (EU Directive 96/22/EC and 2003/74/EC). However, the use of hormonal sex reversal on broodfish, is legal. Outside the EU, the use of hormonal sex reversal is still practiced, particularly for sex determination for Nile tilapia, in countries where this species is produced on a commercial scale. Most of these countries, like China, Egypt and India, have prohibited this practice, however, ineffective enforcement of the ban still permits the continued practice of this technique in these countries.

Chromosome-set Ploidy Manipulation and Sex Control

 

Chromosome-set ploidy manipulation or whole genome manipulation is a highly researched technique with regards to sex control in aquaculture. It is routinely adopted for the commercial production of several aquaculture species (Piferrer et al., 2009; Tiwary, Kirubagaran and Ray, 2004). This technique relies on the application of physical or chemical shocks to alter the normal developmental processes of gametogenesis. Depending on the timing of the applied shock, individuals with a haploid (n); triploid (3n); and tetraploid (4n) cells can be produced; compared to the natural diploid (2n) somatic cell chromosome set. In aquaculture, however, triploids are mostly desired. This is mostly due to the gonadal sterility of the organism, and the advantages resulting in this. During meiosis the trivalent homologous chromosomes of triploids, are unable to pair correctly during prophase I; rendering the organism sterile (Cal et al., 2010). This chromosomal sterility severely impairs pubertal gonad development and subsequently, reproductive maturation. Because of the post-meiotic process of vitellogenesis in oocytes, the retardation or delay of ovarian development and maturation is higher than in male testis.

Consequently, ovaries in triploids individuals, are generally smaller and remain inactive (Benfey, 1999). The production of triploids has been demonstrated in about 49 commercial aquaculture species (Piferrer et al., 2009). However, despite this number, the routine use of this technique for sex control on a commercial scale is a lot lower. In large-scale aquaculture productions, triploidy is mostly restricted to use in various salmonids (Salmo salar, Oncorhynchus masou, Oncorhynchus rhodurus, Oncorhynchus mykiss, Salmo trutta, Salvelinus fontinalis), grass carp (Ctenopharyngodon idella), Arctic char (Salvelinus alpinus), loach (Misgurnus anguillicaudatus), oysters (Crassostrea gigas, Crassostrea virginica, Saccostrea commercialis), ayu (Plecoglossus altivelis), and tilapia (Piferrer, 2001).

Aside from the fact that triploid culture is viewed as a panacea for sex control in fish, there are several challenges with the commercial application of this technique on many aquaculture species. An obvious example is the acquisition of newly fertilised eggs within the timeframe required for the efficient extrusion of the second polar body. In mass spawning species like snappers and barramundi (Sparus spp.), 100 per cent triploidy is crucial for the interference of polar body extrusion in all oocytes. Commercially, this is difficult to achieve, as there are intraspecific family variabilities with regards to the timing of polar body extrusion and oocyte sensitivity to the duration and intensity of the environmental shock applied (Felip et al., 1997). Another challenge limiting the commercial use of this technique for sex control is that triploids often exhibit a higher rate of deformities, lower survival rates, and reduce prepubertal growth compared to diploids. However, this is species specific (Budd et al., 2015).

The Use of Temperature for Sex Ratio Manipulation   

Several pieces of research have documented the effect of temperature on sex ratio, in over 60 fish species; with many of commercial importance to aquaculture (Devlin and Nagahama, 2002; Baroiller and D’Cotta, 2001; Ospina-Álvarez and Piferrer, 2008). A well-studied example regarding the impact of temperature on sex ratio is the European seabass (Dicentrarchus labrax). Cultured populations of this species exhibit a strong male-bias sex ratio, whereas, the total opposite of this occurs in the wild (Piferrer et al., 2005). Between 0 – 60 days post fertilisation, the thermosensitive period, exposure to high temperature has been shown to produce male-biased sex ratio, through an epigenetic modification referred to as DNA methylation. DNA methylation of the gene, cyp19a, coding for gonadal aromatase, is linked to the suppression or downregulation of the gene expression and the subsequent masculinization of genetically female fish (Navarro-Martín et al., 2011). Experimental treatments of 15 oC and 20 oC have been demonstrated to achieve 77% female and 73% male populations respectively (Socorro et al., 2007). This knowledge is invaluable to seabass farmers and offers them the practical tool for sex control of this species.

Next Generation Sequencing and Sex control 

Genomic approaches have significantly contributed to the rapid discovery of genes, gene functions and pathways associated with sex differentiation and gonadal maintenance in several commercially important fish species (Heule, Salzburger and Bohne, 2014). Next-generation sequencing and high throughput single-nucleotide polymorphism (SNP) genotyping technologies have paved the way for the use quantitative trait loci (QTL) studies and high-resolution linkage mapping for marker-assisted selection (MAS) breeding schemes (Yue, 2013). In species with genetically determined systems like the Nile tilapia, high-density marker genome screening, using restriction site associated DNA (RAD) markers, are revolutionising the process of identifying sex associated loci and the development of sex-specific markers in these species. For example, the X-linked Oni23063 and Oni28137 SNP markers in Nile tilapia (Palaiokostas et al., 2013). In populations were variable sex ratios within a species exist, due to both environmental and genetic effects, sex can be viewed as a quantitative threshold trait (Grossen, Neuenschwander and Perrin, 2010; and Vandeputte et al., 2006). In such cases, QTL mapping can help determine the number, and chromosomal regions responsible for sex determination. It can also assist in the locations of significant sex-determining genes within the genome.

In over 40 aquaculture species, QTLs have identified several phenotypic traits, including sex traits (Yue, 2013). These efforts are mostly directed towards locating sex, or early maturation QTLs in species like tilapia, gilt-head sea bream, Sparus aurata, rainbow trout, turbot and several other species (Martínez et al., 2009; Palaiokostas et al., 2013; Cnaani, 2013; Loukovitis et al., 2011).

Conclusion

Sex determination or sex control is a very complex and mostly species-specific process in fish. As a result, the successful application of a sex-determining approach in one species is certainly not guaranteed to work for another. However, there is a wealth of knowledge on several techniques (with more being discovered), available to aquatic farmers to enable them to influence the sex of their culture species. Many of these techniques can now be applied on a commercial scale. Next-generation sequencing is rapidly expanding our knowledge of the genes involved in sexual development, and the processes by which environmental factors can induce phenotypic changes. A collaboration of our current knowledge on sex determination in fish, together with new genetic and epigenetic understanding, will undoubtedly result in further advances in sex control in fish. It will act as a significant catalyst for selective breeding and the culture more efficient/productive fish populations in future.   

Bibliography

Al-Hafedh, Y. and Alam, A. (2007). Design and Performance of an Indigenous Water Recirculating Aquaculture System for Intensive Production of Nile Tilapia, Oreochromis niloticus (L.), in Saudi Arabia. International Journal of Recirculating Aquaculture, 8(1).

Baroiller, J. and D’Cotta, H. (2001). Environment and sex determination in farmed fish. Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology, 130(4), pp.399-409.

Beardmore, J., Mair, G. and Lewis, R. (2001). Monosex male production in finfish as exemplified by tilapia: applications, problems, and prospects. Aquaculture, 197(1-4), pp.283-301.

Benfey, T. (1999). The Physiology and Behavior of Triploid Fishes. Reviews in Fisheries Science, 7(1), pp.39-67.

Budd, A., Banh, Q., Domingos, J. and Jerry, D. (2015). Sex Control in Fish: Approaches, Challenges and Opportunities for Aquaculture. Journal of Marine Science and Engineering, 3(2), pp.329-355.

Cal, R., Terrones, J., Vidal, S., Martínez, P. and Piferrer, F. (2010). Differential incidence of gonadal apoptosis in triploid-induced male and female turbot (Scophthalmus maximus). Aquaculture, 307(3-4), pp.193-200.

Cnaani, A. (2013). The Tilapias’ Chromosomes Influencing Sex Determination. Cytogenetic and Genome Research, 141(2-3), pp.195-205.

Coman, F., Sellars, M., Norris, B., Coman, G. and Preston, N. (2008). The effects of triploidy on Penaeus (Marsupenaeus) japonicus (Bate) survival, growth and gender when compared to diploid siblings. Aquaculture, 276(1-4), pp.50-59.

Devlin, R. and Nagahama, Y. (2002). Sex determination and sex differentiation in fish: an overview of genetic, physiological, and environmental influences. Aquaculture, 208(3-4), pp.191-364.

Felip, A., Zanuy, S., Carrillo, M., Martínez, G., Ramos, J. and Piferrer, F. (1997). Optimal conditions for the induction of triploidy in the sea bass (Dicentrarchus labrax L.). Aquaculture, 152(1-4), pp.287-298.

Grossen, C., Neuenschwander, S. and Perrin, N. (2010). TEMPERATURE-DEPENDENT TURNOVERS IN SEX-DETERMINATION MECHANISMS: A QUANTITATIVE MODEL. Evolution, 65(1), pp.64-78.

Heule, C., Salzburger, W. and Bohne, A. (2014). Genetics of Sexual Development: An Evolutionary Playground for Fish. Genetics, 196(3), pp.579-591.

Kobayashi, Y., Nagahama, Y. and Nakamura, M. (2013). Diversity and Plasticity of Sex Determination and Differentiation in Fishes. Sexual Development, 7(1-3), pp.115-125.

Kwon, J., Haghpanah, V., Kogson‐Hurtado, L., McAndrew, B. and Penman, D. (2000). Masculinization of genetic female Nile tilapia (Oreochromis niloticus) by dietary administration of an aromatase inhibitor during sexual differentiation. Journal of Experimental Zoology, 287(1), pp.46-53.

Lawrence, C., Morrissy, N., Vercoe, P. and Williams, I. (2007). Hybridization in Australian Freshwater Crayfish-Production of All-Male Progeny. Journal of the World Aquaculture Society, 31(4), pp.651-658.

Lin, S., Benfey, T. and Martin-Robichaud, D. (2012). Hormonal sex reversal in Atlantic cod, Gadus morhua. Aquaculture, 364-365, pp.192-197.

Liu, M. and Sadovy de Mitcheson, Y. (2010). The influence of social factors on juvenile sexual differentiation in a diandric, protogynous grouper Epinephelus coioides. Ichthyological Research, 58(1), pp.84-89.

Loukovitis, D., Sarropoulou, E., Tsigenopoulos, C., Batargias, C., Magoulas, A., Apostolidis, A., Chatziplis, D. and Kotoulas, G. (2011). Quantitative Trait Loci Involved in Sex Determination and Body Growth in the Gilthead Sea Bream (Sparus aurata L.) through Targeted Genome Scan. PLoS ONE, 6(1), p.e16599.

Martínez, P., Bouza, C., Hermida, M., Fernández, J., Toro, M., Vera, M., Pardo, B., Millán, A., Fernández, C., Vilas, R., Viñas, A., Sánchez, L., Felip, A., Piferrer, F., Ferreiro, I. and Cabaleiro, S. (2009). Identification of the Major Sex-Determining Region of Turbot (Scophthalmus maximus). Genetics, 183(4), pp.1443-1452.

Nash, C. and Novotny, A. (1995). Production of aquatic animals. Amsterdam: Elsevier.

Navarro-Martín, L., Viñas, J., Ribas, L., Díaz, N., Gutiérrez, A., Di Croce, L. and Piferrer, F. (2011). DNA Methylation of the Gonadal Aromatase (cyp19a) Promoter Is Involved in Temperature-Dependent Sex Ratio Shifts in the European Sea Bass. PLoS Genetics, 7(12), pe1002447.

Palaiokostas, C., Bekaert, M., Khan, M., Taggart, J., Gharbi, K., McAndrew, B. and Penman, D. (2013). Mapping and Validation of the Major Sex-Determining Region in Nile Tilapia (Oreochromis niloticus L.) Using RAD Sequencing. PLoS ONE, 8(7), p.e68389.

Pandian, T. and Sheela, S. (1995). Hormonal induction of sex reversal in fish. Aquaculture, 138(1-4), pp.1-22.

Penman, D. and Piferrer, F. (2008). Fish Gonadogenesis. Part I: Genetic and Environmental Mechanisms of Sex Determination. Reviews in Fisheries Science, 16(sup1), pp.16-34.

Piferrer, F. (2001). Endocrine sex control strategies for the feminization of teleost fish. Aquaculture, 197(1-4), pp.229-281.

Piferrer, F., Beaumont, A., Falguière, J., Flajšhans, M., Haffray, P. and Colombo, L. (2009). Polyploid fish and shellfish: Production, biology and applications to aquaculture for performance improvement and genetic containment. Aquaculture, 293(3-4), pp.125-156.

Piferrer, F., Blázquez, M., Navarro, L. and González, A. (2005). Genetic, endocrine, and environmental components of sex determination and differentiation in the European sea bass (Dicentrarchus labrax L.). General and Comparative Endocrinology, 142(1-2), pp.102-110.

Tiwary, B., Kirubagaran, R. and Ray, A. (2004). The biology of triploid fish. Reviews in Fish Biology and Fisheries, 14(4), pp.391-402.

Yamamoto, T. (1953). Artificially induced sex-reversal in genotypic males of the medaka (Oryzias latipes). Journal of Experimental Zoology, 123(3), pp.571-594.

Yamamoto, T. (1958). Artificial induction of functional sex-reversal in genotypic females of the medaka (Oryzias latipes). Journal of Experimental Zoology, 137(2), pp.227-263.

Yue, G. (2013). Recent advances of genome mapping and marker-assisted selection in aquaculture. Fish and Fisheries, 15(3), pp.376-396.