Methods for infertility and the generation of unisexual offspring

Abstract

This disclosure presents a method for producing sterile, sex-determined fish, crustaceans, or mollusks. [Solution] This method involves the breeding of (i) female fish, crustaceans, or mollusks with fertile homozygous mutations having at least a first and a second mutation, and (ii) male fish, crustaceans, or mollusks with fertile homozygous mutations having at least a first and a second mutation, for the production of sterile sex-determined fish, crustaceans, or mollusks. The first mutation inhibits one or more genes that determine sex differentiation, and the second mutation inhibits one or more genes that determine gamete function, thereby restoring the reproductive capacity of the fertile homozygous female fish, crustaceans, or mollusks and the fertile homozygous male fish, crustaceans, or mollusks. Furthermore, this disclosure presents not only the breeding stock itself, but also a method for generating breeding stock in the production of sterile sex-determined fish, crustaceans, or mollusks.

Description

[Technical Field]

[0001] Statement on the Rights of the Government The methods of work described herein were supported by grant # 2018-33522-28745 from the United States Department of Agriculture and the National Food and Agricultural Research Institute. The U.S. Government has certain rights to these inventions.

[0002] field This disclosure primarily relates to methods for sterilizing and sexing freshwater and marine organisms. [Background technology]

[0003] background The following sections do not acknowledge that the matters discussed herein constitute prior art or the knowledge of those skilled in the art.

[0004] Fish species have been genetically modified (GE) to produce valuable medical proteins or to incorporate traits advantageous for aquaculture. Growth rates, feed conversion ratios, disease resistance, and nutritional benefits have been improved in various fish species to meet future demand for seafood and the need for improved sustainability in the aquaculture industry. However, the global adoption of these GE fish is hindered by concerns about their accidental release into natural ecosystems. Farmed fish have proven to reproduce and survive in natural environments. Similarly, GE fish may have natural relatives, and genetic modification increases the likelihood of spreading to entire natural populations and altering the natural gene pool. Therefore, commercial GE fish pose a potential environmental threat and present a challenge for policymakers and regulatory bodies to conduct risk-benefit assessments.

[0005] One way to address one or more of the aforementioned problems is to sterilize fish. Induction of triploidy is the most commonly used and well-studied method for producing infertile fish. Triploid fish are typically produced by subjecting fertilized eggs to temperature or pressure shocks, forcing the incorporation of a second polar body and producing cells with three pairs of chromosomes (3N). Because triploid fish have an extra pair of chromosomes, meiosis is inhibited, and the gonads do not develop normally. On an industrial scale, reliably subjecting egg masses to pressure or temperature shocks is complex and extremely costly. In addition to triploidy induced by physical treatment, there are also genetically induced triploids, which are produced by crossing tetraploid fish with diploid fish. However, tetraploid fish are difficult to produce because of their low embryonic viability and slow growth. Triploid males produce normal haploid spermatozoa, making it possible for males to fertilize eggs, although there are some examples where this is less efficient. Furthermore, some species exhibited characteristics related to the triploid phenotype, such as stunted growth and susceptibility to disease, resulting in lower performance.

[0006] Another method involves sterilizing fish through hormonal treatment over several weeks. However, such processes, often involving prolonged and intensive stimulation, are not highly effective in sterilization and may even be associated with reduced fish growth. Furthermore, treatment with synthetic steroids can lead to higher mortality rates.

[0007] Transgenic-based technologies also exist for sterilizing fish, involving procedures to insert transgenes that eliminate developing embryos by inducing germ cell lethality or inhibiting migration patterns. However, the effectiveness of these transgenes depends on silencing and positional effects. Therefore, such methods are subject to an extensive regulatory review process before they can be accepted for commercial use.

[0008] One method of sterilizing fish involves knocking down or knocking out genes that control primordial germ cell (PGC) development. Such methods have been reported to induce PGC deletion and infertility. However, the infertility trait in these fish is not hereditary. Therefore, utilizing knockdown or knockout methods for genes controlling PGC development presents logistical and cost challenges, making the efficient mass production of sterile fish on a commercial scale difficult.

[0009] The mechanisms governing sexual or gonadal differentiation in bony fish are complex processes influenced by internal factors (genetic and endocrine factors) and external factors such as interactions and environmental conditions (water temperature, pH, and oxygen), and the relative contributions of these factors vary greatly among species.

[0010] Improvements in the production of sterile, sex-determined fish, crustaceans, or mollusks are desirable. [Overview of the Initiative] [Problems that the invention aims to solve]

[0011] introduction The purpose of this introduction is to present this specification to the reader and not to define any invention. One or more inventions may be described below or in other parts of this document in the form of a combination or partial combination of apparatus, method, or process. The inventors do not waive or deny any rights to any other inventor or any invention disclosed herein by not describing such invention or inventor in these claims.

[0012] One or more of the aforementioned methods used to sterilize freshwater and marine organisms may result in: (1) insufficient effectiveness; (2) difficulty in transmitting the sterile trait, for example, by performing genetic selection to identify subpopulations of sterile individuals and / or by repeating the treatment in each generation; (3) increased operating costs resulting from, for example, significant changes in livestock practices, inability to transfer in multiple species, increased production time, increased proportion of low-growth sterile organisms and increased susceptibility to disease, increased mortality rates of sterile organisms, or a combination thereof; (4) gene influx into wild populations and aquaculture, colonization of new habitats by introduced species; or (5) a combination thereof.

[0013] This disclosure presents methods for producing sex-determined, sterile freshwater and marine organisms by inhibiting sex differentiation and gamete formation pathways. One or more examples of this disclosure may, compared to one or more of the aforementioned methods used to sterilize freshwater and marine organisms,: (1) enhance the effectiveness of sterilization by enabling mass production of sterile individuals and ensuring that all individuals are completely sterile; (2) reduce operating costs by reducing the amount of costly processing equipment, making it feasible on a large scale, making it transferable across multiple species, reducing feed, shortening production time, reducing the proportion of sexually mature organisms, increasing the physical size of sexually mature organisms, or a combination thereof; (3) reduce gene influx into wild populations and colonization of new habitats by aquaculture and introduced species; (4) improve aquaculture performance by reducing energy loss for gonadal development; or (5) a combination thereof.

[0014] One or more examples in this disclosure may improve the feed conversion ratio (FCR = weight gained per unit of feed given) by at least 10% and accelerate the growth rate by 20% compared to other methods currently used in production systems (methyltestosterone treatment). These have the advantage of potentially affecting only feed costs (direct reduction in feed costs) and labor (reduction in labor due to shorter farming times). Based on the average itemized costs of a US domestic tilapia farm producing 1,000 pounds, savings of $0.23 may be realized for each market-sized fish (1.5 pounds) using all male sterile tilapia, suggesting that maintaining production cost savings could lead to an increase in profit margins of approximately 130%.

[0015] Furthermore, this disclosure discusses not only the breeding stock itself, but also methods for generating breeding stock of freshwater and marine organisms used to produce sex-determined, sterile freshwater and marine organisms.

[0016] This disclosure presents a method for producing sterile sex-determined fish, crustaceans, and mollusks, including the following steps: (i) breeding a fertile hemizygous mutant female fish, crustacean, or mollusk having at least the first and second mutations with a fertile hemizygous mutant male fish, crustacean, or mollusk having at least the first and second mutations; and selecting homozygous primitives, which are homozygous mutant primitives that are sterile sex-determined fish, crustaceans, or mollusks, in which the first mutation inhibits one or more genes that determine sex differentiation, and the second mutation inhibits one or more genes that determine gamete function.

[0017] The disclosure also presents a method for producing sterile sex-determined fish, crustaceans, or mollusks, including the following steps: to produce sterile sex-determined fish, crustaceans, or mollusks, (i) breed fertile homozygous mutant female fish, crustaceans, or mollusks having at least the first and second mutations; and (ii) breed fertile homozygous mutant male fish, crustaceans, or mollusks having at least the first and second mutations; the first mutation inhibits one or more genes that determine sex differentiation, the second mutation inhibits one or more genes that determine gamete function, and the reproductive capacity of the fertile homozygous female fish, crustaceans, or mollusks and the fertile homozygous mutant male fish, crustaceans, or mollusks is restored.

[0018] Restoring fertility may involve germline stem cell transplantation. Further restoration of fertility may involve sex steroid changes. These sex steroid changes may be estrogen changes or aromatase inhibitor changes.

[0019] This germline stem cell transplantation may include the following steps: obtaining germline stem cells from a sterile homozygous male fish, crustacean, or mollusk having at least the first and second mutations, or from a sterile homozygous female fish, crustacean, or mollusk; and transplanting the germline stem cells into a germline-free male fish, crustacean, or mollusk graft. This germline-free male fish, crustacean, or mollusk graft and germline-free female fish, crustacean, or mollusk graft may be homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes. This germline-free male fish, crustacean, or mollusk graft and germline-free female fish, crustacean, or mollusk graft may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0020] This germline stem cell transplantation may include the following steps: obtaining spermatogonial stem cells from a sterile homozygous male fish, crustacean, or mollusk having at least the first and second mutations, or oogonial stem cells from a sterile homozygous female fish, crustacean, or mollusk having at least the first and second mutations; and transplanting the spermatogonial stem cells into the testes of a germ-free male fish, crustacean, or mollusk, or the oogonial stem cells into the ovaries of a germ-free female fish, crustacean, or mollusk. These germ-free, fertile male fish, crustacean, or mollusk and germ-free, fertile female fish, crustacean, or mollusk may be homozygous for mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes. These germ-free male fish, crustacean, or mollusk grafts and germ-free female fish, crustacean, or mollusk grafts may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0021] This sterilized sex-determined fish, crustacean, or mollusk may be a sterilized male fish, crustacean, or mollusk. The first mutation may involve mutations in one or more genes that regulate androgen and / or estrogen synthesis. The first mutation may involve mutations in one or more genes that regulate the expression of aromatase Cyp19a1a, Cyp17, or a combination thereof. One or more genes that regulate the expression of aromatase Cyp19a1a may be one or more genes selected from the group consisting of cyp19a1a, FoxL2, and orthologs. The second mutation may involve mutations in one or more genes that regulate spermatogenesis, and the second mutation may involve mutations in one or more genes that develop macrocephaly spermatozoa. The second mutation in one or more genes that develop macrocephaly spermatozoa may cause sperm with a round head, round nucleus, detached midpiece, partially coiled tail, or a combination thereof. This second mutation may involve mutations in one or more genes selected from the group consisting of Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, and their orthologs.

[0022] This sterile sex-determined fish, crustacean, or mollusk may be a sterile female fish, crustacean, or mollusk. The first mutation may include mutations in one or more genes that regulate the expression of aromatase Cyp19a1a inhibitor. One or more genes that regulate the expression of aromatase Cyp19a1a inhibitor may include mutations in one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologs. The second mutation may include mutations in one or more genes that regulate oogenesis, folliculogenesis, or a combination thereof. One or more genes that regulate oogenesis may regulate the synthesis of estrogen. One or more genes that regulate the synthesis of estrogen may be FSHR or its ortholog. One or more genes that regulate folliculogenesis may regulate the expression of vitellogenin. One or more genes that regulate the expression of vitellogenin may be vtgs or its ortholog. One or more genes that regulate the expression of vitellogenin may be mutations in genes encoding or controlling vitellogenin; estrogen receptor 1; Cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; Choriogenin H; peroxisome proliferator-activated receptor; steroidogenic acute regulatory protein, or their orthologs.

[0023] The present disclosure also presents a method for producing a sterile sex-determined fish, crustacean, or mollusk, including the following steps: To produce a sterile sex-determined fish, crustacean, or mollusk, (i) a fertile female fish, crustacean, or mollusk with a homozygous mutation is (ii) bred with a fertile male fish, crustacean, or mollusk with a homozygous mutation, the mutation directly or indirectly inhibits spermatogenesis, and / or directly inhibits vitellogenesis, and the fertility of the fertile female fish, crustacean, or mollusk and the fertile male fish, crustacean, or mollusk is restored.

[0024] Mutations that directly or indirectly inhibit spermatogenesis may be mutations in Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, or their orthologs. Mutations that directly inhibit yolk formation may be mutations in genes encoding or controlling vitellogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein, or their orthologs. These fertile female fish, crustaceans, or mollusks and fertile male fish, crustaceans, or mollusks may have multiple homozygous mutants that directly or indirectly inhibit spermatogenesis; directly inhibit yolk formation; or a combination of both.

[0025] This restoration of fertility may include germline stem cell transplantation. This restoration of fertility may further include sex steroid changes. These sex steroid changes may be estrogen changes or aromatase inhibitor changes.

[0026] This germline stem cell transplantation may include the following steps: obtaining germline stem cells from infertile homozygous male fish, crustaceans, or mollusks with at least homozygous mutations, or from infertile homozygous females with at least homozygous mutations; and transplanting the germline stem cells into germline-free male fish, crustaceans, or mollusks, or germline-free female fish, crustaceans, or mollusks. Germline-free male fish, crustaceans, or mollusks and germline-free female fish, crustaceans, or mollusks may be homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes. These germline-free male fish, crustaceans, or mollusks and germline-free female fish, crustaceans, or mollusks may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0027] These fertile female fish, crustaceans, or mollusks, and fertile male fish, crustaceans, or mollusks, may have additional homozygous mutations that determine sex differentiation. These sex differentiation-determining mutations may modulate the expression of aromatase Cyp19a1a, Cyp17, an inhibitor of aromatase Cyp19a1a, or a combination thereof. Mutations that modulate Cyp17 expression may be mutations in cyp17I or its ortholog. Mutations that modulate aromatase Cyp19a1a inhibitors may be mutations in Gsdf, dmrt1, Amh, Amhr, or their orthologs.

[0028] The breeding steps of the methods disclosed herein may include crossbreeding or hormonal treatment and breeding methods for determining sexual differentiation.

[0029] The fish, crustaceans, or mollusks of the methods disclosed herein may be fish.

[0030] The disclosure also presents sterile, sex-determined fish, crustaceans, or mollusks having at least the first and second mutations, and for producing fertile, homozygous mutant fish, crustaceans, or mollusks in which the first mutation inhibits one or more genes determining sex differentiation, the second mutation inhibits one or more genes determining gamete function, and the fertile homozygous mutant fish, crustaceans, or mollusks have restored fertile homozygous mutant fish, crustaceans, or mollusks. This restoration of fertile capacity may include germline stem cells. This restoration of fertile capacity may further include sex steroid changes. These sex steroid changes may be estrogen changes or aromatase inhibitor changes.

[0031] This germline stem cell transplantation may include the following steps: obtaining germline stem cells from a sterile homozygous male fish, crustacean, or mollusk with at least the first and second mutations; and transplanting the germline stem cells into a germline-free male fish, crustacean, or mollusk transplant, or a germline-free female fish, crustacean, or mollusk transplant. This germline-free male fish, crustacean, or mollusk transplant and germline-free female fish, crustacean, or mollusk transplant may be homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes. This germline-free male fish, crustacean, or mollusk transplant and germline-free female fish, crustacean, or mollusk transplant may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0032] This germline stem cell transplantation may include the following steps: obtaining spermatogonial stem cells from a sterile homozygous male fish, crustacean, or mollusk having at least the first and second mutations, or oogonial stem cells from a sterile homozygous female fish, crustacean, or mollusk having at least the first and second mutations; and transplanting the spermatogonial stem cells into the testes of a germ-free, fertile male fish, crustacean, or mollusk, or the oogonial stem cells into the ovaries of a germ-free, fertile female fish, crustacean, or mollusk. This germ-free, fertile male fish, crustacean, or mollusk and germ-free, fertile female fish, crustacean, or mollusk may be homozygous for mutations in the dnd, Elavl2, vasa, nanos3, or piwi-like genes. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0033] This sterile, sex-determined fish, crustacean, or mollusk may be a sterile male fish, crustacean, or mollusk. The first mutation may involve mutations in one or more genes that regulate the synthesis of androgens and / or estrogens. The first mutation may involve mutations in one or more genes that regulate the expression of aromatase Cyp19a1a, Cyp17, or a combination thereof. The one or more genes that regulate the expression of aromatase Cyp19a1a may be one or more genes selected from the group consisting of cyp19a1a, FoxL2, and their orthologues. The one or more genes that regulate the expression of Cyp17 may be cyp17I or its orthologue. This second mutation may involve mutations in one or more genes that regulate spermatogenesis. This second mutation may involve mutations in one or more genes that cause megazoospermia. A second mutation in one or more genes that causes megaheaded spermatozoospermia can result in sperm with a round head, round nucleus, truncated mid-section, partially coiled tail, or a combination thereof. This second mutation may include mutations in one or more genes selected from the group consisting of Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, and their orthologues.

[0034] This sterile, sex-determined fish, crustacean, or mollusk may be a sterile female fish, crustacean, or mollusk. The first mutation may involve a mutation in one or more genes that modulate the expression of the aromatase Cyp19a1a inhibitor. The one or more genes that modulate the expression of the aromatase Cyp19a1a inhibitor may involve a mutation in one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologues. The second mutation may involve a mutation in one or more genes that modulate oogenesis, follicular formation, or combination. The one or more genes that modulate oogenesis may modulate estrogen synthesis. The one or more genes that modulate estrogen synthesis may be FSHR or its orthologue. The one or more genes that modulate follicular formation may modulate vitelogenin expression. The one or more genes that modulate vitelogenin expression may be vtgs or its orthologue. One or more genes that regulate vitelogenin expression may be mutations in genes encoding or controlling vitelogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein; or their orthologs.

[0035] The disclosure also presents fertile fish, crustaceans, or mollusks with homozygous mutations that directly or indirectly inhibit spermatogenesis and / or directly inhibit yolk formation, and restore the fertility of fertile male fish, crustaceans, or mollusks.

[0036] Mutations that directly or indirectly inhibit spermatogenesis may be mutations in Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, or their orthologs. Mutations that directly inhibit yolk formation may be mutations in genes encoding or controlling vitellogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein, or their orthologs. These fertile fish, crustaceans, or mollusks may have multiple homozygous mutants that directly or indirectly inhibit spermatogenesis; directly inhibit yolk formation; or a combination of both. This restoration of fertility may involve germline stem cells. This restoration of fertility may further involve sex steroid changes. These sex steroid changes may be estrogen changes or aromatase inhibitor changes.

[0037] This germline stem cell transplantation may include the following steps: obtaining germline stem cells from sterile homozygous male fish, crustaceans, or mollusks with at least homozygous mutations, or from sterile homozygous female fish, crustaceans, or mollusks with at least homozygous mutations; and transplanting the germline stem cells into germline-free male fish, crustaceans, or mollusks or germline-free female fish, crustaceans, or mollusks. These germline-free male fish, crustaceans, or mollusks and germline-free female fish, crustaceans, or mollusks may be homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes. These germline-free male fish, crustaceans, or mollusks and germline-free female fish, crustaceans, or mollusks may be created using ploidy manipulation. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by hybridization. These germ-free male fish, crustaceans, or mollusks and germ-free female fish, crustaceans, or mollusks may be created by exposure to high levels of sex hormones.

[0038] These fertile fish, crustaceans, or mollusks may have additional homozygous mutations that determine sex differentiation. These sex differentiation-determining mutations may modulate the expression of aromatase Cyp19a1a, Cyp17, an inhibitor of aromatase Cyp19a1a, or a combination thereof. The mutation that modulates the expression of aromatase Cyp19a1a may be one or more genes selected from the group consisting of cyp19a1a, FoxL2, and their orthologues. The one or more genes that modulate the expression of aromatase Cyp19a1a inhibitors may be one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologues.

[0039] The production of sterile, sex-determined fish, crustaceans, or mollusks may involve a breeding step that includes hybridization or hormonal treatment and breeding methods to determine sex differentiation.

[0040] The fertile fish, crustaceans, or mollusks disclosed herein may be fish.

[0041] The disclosure also presents a method for producing fertile homozygous mutant fish, crustaceans, and mollusks that produce sterile sex-determined fish, crustaceans, and mollusks, including the following steps: (i) breeding a fertile hemizygous mutant female fish, crustacean, or mollusk having at least the first and second mutations with a fertile hemizygous mutant male fish, crustacean, or mollusk having at least the first and second mutations; selecting a homozygous primitive by genotype selection; and restoring the fertile capacity of the homozygous primitive, wherein the first mutation inhibits one or more genes that determine sex differentiation, and the second mutation inhibits one or more genes that determine gamete function.

[0042] Further embodiments and functions described herein will become apparent to those skilled in the art upon reviewing the following descriptions of specific examples in conjunction with the accompanying drawings. The methods and organisms disclosed to date are described only through examples, with reference to the attached figures. [Brief explanation of the drawing]

[0043] [Figure 1] Figure 1 is a flowchart illustrating an example of a method for generating and propagating a sterile, sex-determined fish, crustacean, or mollusk-like organism. [Figure 2]Figure 2 shows an example and graph illustrating the identification and selection methods for F0 mosaic founder variants. Mutant alleles were identified by fluorescence PCR using gene-specific primers designed to amplify the peripheral region of the target locus (120–300 bp). For fluorescence PCR, combinations of both gene-specific primers and two forward oligonucleotides fitted with the fluorescent dye 6-FAM or NED were added to the reaction. A control reaction using wild-type DNA was used to confirm the presence of single-peak amplification at individual loci. The resulting amplicons were degraded by capillary electrophoresis (CE) with the addition of a LIZ-labeled size criterion to determine amplicon size appropriate to base pair resolution (Retrogen). Raw trace files were analyzed with Peak Scanner software (Thermo Fisher). Peak size compared to wild-type peak control determined the nature (insertion or deletion) and length of the mutation. The number of peaks indicates the level of mosaicism. The inventors selected F0 mosaic founders with the fewest mutant alleles (selectively 2-4 peaks). [Figure 3] Figure 3 is a graph visualizing heterozygous genotypes, homozygous mutants, and wild-type samples using melting curve plots. It can be seen that fluorescence changes negatively with temperature (-dF / dT). Each trace represents a sample. In this example, the melting temperature of the wild-type allele is ~81°C (wild-type peak), and the melting temperature of the homozygous mutant product (homozygous deletion peak) is ~79°C. The other traces represent heterozygous types. [Figure 4] Figure 4, panels A to D, shows photographs of the growth process of tilapia F0 generation compared to bi-allele knockout of the pigment-forming gene. [Figure 5]Figure 5, panels A and B, are photographs of tilapia after multi-gene targeting, including dead end1 (dnd) and tyrosinase (Tyr). Figure 5, panel A, shows an albino with a deficiency of F0 Tyr. Figure 5, panel B, shows dissected testes of control (WT) and sterile (F0 dnd KO) tilapia. [Figure 6] Figure 6, panels A and B, show photographs of the testes and ovaries of Elavl2-knockout tilapia (Elavl2△8 / △8) lacking germ cells (arrows point to the gonads). The small photographs show the urogenital process. The Elavl2 mutant was generated by microinjecting an artificial nuclease targeting the Elavl2 coding sequence into one-cell stage tilapia embryos before cell cleavage. One of the resulting founder males mated with a wild-type female and produced heterozygous mutants in the F1 generation. By mating with the F1 mutant Elavl2△8 / +, approximately 25% of the eggs produced the F2 generation of bisexual homozygous mutants. [Figure 7] Panels A through C of Figure 7 show the selected mutant alleles at the tilapia cyp17 locus. Panel A of Figure 7 is a schematic of the cyp17 gene. Exons (E1-8) are shown in shaded boxes; the translation start and stop sites are ATG and TAA, respectively. The arrows point to the target position in the first exon. Panel B of Figure 7 is the wild-type reference sequence (SEQ ID NO: 60) with the germline mutant allele (SEQ ID NO: 61) of a Cyp17 F0 mutant tilapia offspring. This 11nt+5nt deletion is expected to produce a truncated protein that cleaves at amino acid 44 instead of position 521. Panel C of Figure 7 is the mutant cyp17 allele (SEQ ID NO: 63) with the expected WT protein sequence (SEQ ID NO: 62) and the first 16 amino acids matching those of the wild-type Cyp17 protein, with 44 amino acids misencoded. [Figure 8]Figure 8 panels A to C are graphs and photographs showing that cyp17 deficiency produces only male offspring lacking secondary sexual characteristics. Figure 8 panel A is a graph showing fish of the Cyp17 mutant that are completely male-biased. Founder males with germline mutations at the cyp17 locus were bred with wild-type females, and F1 offspring of these males and females carrying the null Δ16-cyp17 allele were selected and mated to produce F2 generations of wild-type (WT) homozygous (- / -) and hemizygous mutants (+ / -). The graph shows the number of males and females of these genotypes. Figure 8 panel B shows undetectable levels of testosterone in the cyp17 deficiency mutant. Blood was collected from the tail vein and centrifuged at 3000 rpm for 10 minutes. Plasma was separated, frozen at -80°C, and plasma free testosterone levels were measured by enzyme-linked immunosorbent assay (ELISA) (Cayman Chemical, Michigan, USA). Plasma samples were analyzed three times. Figure 8, panel C, shows a photograph of a cyp17 F0 KO (- / -) male with underdeveloped UGP compared to an age-matched, untreated male (right). [Figure 9] Panels A through E of Figure 9 show that Cyp17-deficient mutants are sexually delayed, exhibiting small testes and oligospermia. F2 offspring of hemizygous cyp17 mutants were raised to 5 months of age, weighed (Panel C of Figure 9), and genotyped. Panel A of Figure 9 shows exposed testes (Panel A) and dissected testes (Panel B) of euthanized males, illustrating the color and size gradient of homozygous WT testes, which appear to be sexually delayed and possess the most mature gonads. Panel E of Figure 9 shows the size of exfoliated fish sperm from eight homozygous and WT males, and Panel F of Figure 9 shows a comparison of sperm concentrations by spectrophotometric analysis (absorbance at 600 nm). [Figure 10]Panels A through C of Figure 10 represent the mutant alleles selected at the tilapia tight junction constituent protein 1 (Tjp1a) locus. Panel A of Figure 10 is a schematic of the Tjp1a gene. Exons (E1-32) are shown in shaded boxes; the translation start and stop sites are ATG and TAA, respectively. Arrows point to target exons 15 and 17. Panel B of Figure 10 is the wild-type reference sequence (SEQ ID NO: 71) with the selected germline mutant allele (SEQ ID NO: 72) of a Tjp1a F0 mutant tilapia offspring. This 7 nt deletion is expected to produce a truncated protein that cleaves at amino acid 439 instead of position 1652. Panel C of Figure 10 is the expected WT protein sequence (SEQ ID NO: 73) and the mutant Tjp1a allele (SEQ ID NO: 74) where the first 439 amino acids match those of the wild-type Tjp1a protein. [Figure 11] Figure 11 panels A to C show selected mutations at the Hippocampus abundant transcript 1a (Hiat1) locus. Figure 11 panel A is a schematic of the tilapia Hiat1 gene. Exons (E1-12) are shown in shaded boxes; the 5' and 3' untranslated regions are shown in open boxes. Arrows point to target exons 4 and 6. Figure 11 panel B is the wild-type reference sequence (SEQ ID NO: 75) with the selected germline mutant allele (SEQ ID NO: 76) of a Hiat1 F0 mutant tilapia offspring. The location of the 17-nucleotide deletion is indicated by a dash. This frameshift mutation is expected to produce a truncated protein that cleaves at amino acid 234 instead of position 491. Figure 11 Panel C shows the expected WT protein sequence (SEQ ID NO: 77) and the deletion mutant Hiat1 protein (SEQ ID NO: 78), in which the first 218 amino acids match those of the wild type, with the following 16 amino acids being misencoded. [Figure 12]Figure 12 panels A to C show selected mutations at the tilapia Small ArfGAP2 (Smap2) locus. Figure 12 panel A is a schematic of the tilapia Smap2 gene. Exons (E1-12) are shown in shaded boxes, and the 3' untranslated region is shown in an open box. Arrows point to target exons 2 and 9. Figure 12 panel B is the wild-type reference sequence (SEQ ID NO: 79) with the selected germline mutant allele (SEQ ID NO: 80) of a Smap2 F0 mutant tilapia offspring. The location of the 17-nucleotide deletion is indicated by a dash. This frameshift mutation is expected to produce a truncated protein that cleaves at amino acid 118 instead of position 429. Figure 12 Panel C shows the expected WT protein sequence (SEQ ID NO: 81) and the deletion mutant Smap2 protein (SEQ ID NO: 82), in which the first 53 amino acids match those of the wild type, while the following 63 amino acids are misencoded. [Figure 13] Figure 13 panels A to C show the selected mutant alleles at the tilapia casein kinase 2, alpha-prime polypeptide (Csnk2a2) gene locus. Figure 13 panel A is a schema of the Csnk2a2 gene. Exons (E1-11) are shown in shaded boxes; the translation start and stop sites are ATG and TGA, respectively. Arrows point to target exons 1 and 2. Figure 13 panel B is the wild-type reference sequence (SEQ ID NO: 83) with the selected germline mutant allele (SEQ ID NO: 84) of a Csnk2a2 F0 mutant tilapia offspring. This 22 nt deletion is expected to produce a truncated protein that cleaves at amino acid 31 instead of position 350. Figure 13 panel C shows the expected WT protein sequence (SEQ ID NO: 85) and the mutant Csnk2a2 allele (SEQ ID NO: 86) with the first 31 amino acids incorrectly encoded. [Figure 14]Figure 14 panels A to C show selected mutant alleles at the tilapia Golgi-associated PDZ and coiled-coil motif (Gopc) loci. Figure 14 panel A is a schematic of the Gopc gene. Exons (E1-9) are shown in shaded boxes; the translation start and stop sites are ATG and TAA, respectively. Arrows point to target exons 1 and 2. Figure 14 panel B is the wild-type reference sequence (SEQ ID NO: 87) with a selected germline mutant allele (SEQ ID NO: 88) from a Gopc F0 mutant tilapia offspring. This 8 nt deletion is expected to produce a truncated protein that cleaves at amino acid 30 instead of position 444. Figure 14 Panel C shows the expected WT protein sequence (SEQ ID NO: 89) and the mutant Gopc allele (SEQ ID NO: 90), in which the first 9 amino acids match those of the wild-type Gopc protein, while the following 21 amino acids are incorrectly encoded. [Figure 15] Figure 15, panels A and B, show photographs and graphs of tilapia testis formation-specific genes mimicking human and mouse deletion knockout phenotypes. Figure 15, panel A, shows sperm malformations in F0 deletion tilapia for five candidate genes. Microscopic images of sperm were collected from wild-type (WT) and Tjp1a, Gopc, Smap2, Hiat1, and Csnk2a2 F0 mutant fish, respectively. Black arrows indicate the size of WT sperm heads, and yellow arrows indicate large, round sperm heads. The scale bar is 100 μm. Figure 15, panel B, shows the fertilization success rate of hand-detached gametes, which were mixed with dried gametes (200 oocytes and detached fish sperm), immediately activated with 2 mL of incubation water, and fertilized in vitro. Data are mean + / - SD, n=3 replicates. [Figure 16]Figure 16 panels A to C show images and graphs illustrating the expression levels of SMS genes in fertile, germ-free testes. Figure 16 panel A shows dissected testes from 4-month-old dnd1 knockout and age-matched wild-type controls. Figure 16 panel B shows the relative expression levels of vasa, while the Sertoli-specific gene Dmrt1 is expressed at the same level in the testes of wild-type and infertile tilapia, while germ-specific genes are reduced to undetectable levels in the testes of dnd1 KO fish. Beta-actin was used as a reference gene to normalize the expression levels of Vasa and Dmrt1. Figure 16 panel C shows the relative expression levels of the SMS genes Tjp1a, Hiat1, Gopc, and Csnk2a2 in the testes of wild-type and infertile tilapia. Dmrt1 was used as a reference gene to normalize the expression levels of SMS genes. In all cases, the values ​​represent a mean of 3 biological replicates, + / - SD. [Figure 17] Panels A through C of Figure 17 show the selected mutations at the Cyp9a1a locus. Panel A of Figure 17 is a schematic of the Cyp9a1a gene. Exons (E1-9) are shown in shaded boxes. Arrows point to target exons 1 and 9. Panel B of Figure 17 is the wild-type reference sequence (SEQ ID NO: 65) with selected germline mutant alleles (SEQ ID NOs: 66 and 67) from offspring of Cyp19a1a F0 mutant tilapia. These 7 nt (deletion 8 and insertion 1) and 10 nt deletions are indicated by dashes. These frameshifts are expected to produce truncated proteins that cleave at amino acids 12 and 11 instead of positions 511. Figure 17, panel C, shows the expected WT protein sequence (SEQ ID NO: 68) and deletion mutant proteins (SEQ ID NO: 69 and 70), where the first 7 and 5 amino acids match those of the wild-type Cyp19a1a protein, and the following 5 and 6 amino acids are incorrectly encoded. The altered amino acids are highlighted. [Figure 18]Figure 18 shows the breeding schema of double heterozygous parents and a description and table of the expected genotypes of mutant offspring. m1, 2, and 3 show different mutations at the Tjp1a locus in F0 mosaic females. The columns in the table show the expected sex ratio and fertility status, and the expected frequency of F2 offspring for each combination of cyp17 and Tjp1a alleles. Offspring expected to be all male and infertile are enclosed in a box. [Figure 19] Figure 19 panels A to C show the selected mutations at the Dmrt1 locus. Figure 19 panel A is a schematic of the Dmrt1 gene. Exons (E1-9) are shown in shaded boxes. Arrows point to target exons 1 and 3. Figure 19 panel B is the wild-type reference sequence (SEQ ID NO: 91) with the selected germline mutant alleles (SEQ ID NOs: 92 and 93) from the Dmrt1 F0 mutant tilapia. These 7 nt and 13 nt deletions are indicated by dashes. These frameshift mutations are expected to produce a truncated protein that cleaves at amino acids 40 and 38 instead of position 293. Figure 19 panel C shows the expected WT protein sequence (SEQ ID NO: 94) and the deletion mutant protein (SEQ ID NO: 95 and 96), where the first 16 amino acids match those of the wild-type Dmrt1 protein, and the following 24 and 22 amino acids are misencoded. The changed amino acids are highlighted. [Figure 20]Panels A through C of Figure 20 show selected mutations at the growth / differentiation factor 6-B-like locus (Gsdf). Panel A of Figure 20 is a schematic of the tilapia Gsdf gene. Exons (E1-5) are shown in shaded boxes. Arrows point to target exons 2 and 4. Panel B of Figure 20 is the wild-type reference sequence (SEQ ID NO: 97) with the selected germline mutant alleles (SEQ ID NOs: 98 and 99) of the Gsdf F0 mutant tilapia. These 5 nt and 22 nt deletions are indicated by dashes. These frameshift mutations are expected to produce truncated proteins that cleave at amino acids 56 and 46 instead of positions 213. Figure 20, panel C, shows the expected WT protein sequence (SEQ ID NO: 100) and deletion mutant proteins (SEQ ID NO: 101 and 102), where the first 52 and 46 amino acids match those of the wild-type Gsdf protein, with the following 4 and 0 amino acids being incorrectly encoded. The altered amino acids are highlighted. [Figure 21] Panels A through C of Figure 21 show the selected mutations at the tilapia follicle-stimulating hormone receptor (FSHR) gene locus. Panel A of Figure 21 is a schematic of the tilapia FSHR gene. Exons (E1-15) are shown in shaded boxes; the 5' and 3' untranslated regions are shown in open boxes. Arrows point to target exons 11 and 15. Panel B of Figure 21 is the wild-type reference sequence (SEQ ID NO: 103) with the sequence (SEQ ID NO: 104) of the selected germline mutation allele of an FSHR F0 mutant tilapia offspring. The 5-nucleotide position is indicated by a dash. This frameshift mutation is expected to produce a truncated protein that cleaves at amino acid 264 instead of position 689. Figure 21 Panel C shows the expected WT protein sequence (SEQ ID NO: 105) and the deletion mutant FSHR protein (SEQ ID NO: 106), in which the first 258 amino acids match those of the wild type, with the following 6 amino acids being misencoded. [Figure 22]Figure 22 panels A to C show selected mutations at the vitelogenin Aa (VtgAa) locus. Figure 22 panel A is a schematic of the tilapia VtgAa gene. Exons (E1-35) are shown in shaded boxes. Arrows point to target exons 7 and 22. Figure 22 panel B is the wild-type reference sequence (SEQ ID NO: 107) with the selected germline mutant alleles (SEQ ID NOs: 108 and 109) from the VtgAa F0 mutant tilapia. These 5 nt and 25 nt deletions are indicated by dashes. These frameshift mutations are expected to produce truncated proteins that cleave at amino acids 279 and 301 instead of positions 1657. Figure 22, panel C, shows the expected WT protein sequence (SEQ ID NO: 110) and deletion mutant proteins (SEQ ID NO: 111 and 112), where the first 278 and 269 amino acids match those of the wild-type VtgAa protein, with the following 1 and 32 amino acids incorrectly encoded. The altered amino acids are highlighted. [Figure 23] Figure 23 panels A to C show selected mutations at the vitelogenin Ab (VtgAb) locus. Figure 23 panel A is a schematic of the tilapia VtgAb gene. Exons (E1-35) are shown in shaded boxes; the 5' untranslated region is shown in an open box. Arrows point to target exons 5 and 22. Figure 23 panel B is the wild-type reference sequence (SEQ ID NO: 113) with the sequence (SEQ ID NOs: 114) of the selected germline mutant allele of a VtgAb F0 mutant tilapia offspring. The location of the 8-nucleotide deletion is indicated by a dash. This frameshift mutation is expected to produce a truncated protein that cleaves at amino acid 202 instead of position 1747. Figure 23, Panel C, shows the expected WT protein sequence (SEQ ID NO: 115) and the deletion mutant VtgAb protein (SEQ ID NO: 116), where the first 270 amino acids match those of the wild-type VtgAb protein, and the following 32 amino acids are incorrectly encoded. The altered amino acids are highlighted. [Figure 24] Figures 24, panels A and B, are photographs and graphs showing the tails of females lacking VtgAa for viable offspring. Figure 24, panel A, is a photograph of embryos in methylene blue-containing (Roth, 0.01% of the incubation solution) incubation water 8 hours after fertilization. Blue staining indicates unfertilized eggs and dead embryos. Embryos were examined daily under a lighted stereomicroscope, and the number of dead embryos was counted and removed. Figure 24, panel B, shows the survival rates in F0 VtgAa male and female offspring crossed with wild-type fish. Data are mean + / - SD, n=2x3 replicates. [Figure 25] Figure 25 shows the breeding method and the genotypes of mutant offspring of double hemizygous mutant parents. m1-n and m1 represent mosaic mutations in F0 and specific mutations selected for their respective target loci. The F1 genotypes shown correspond to one of the four alleles that the inventors are trying to establish. The columns in the table show the predicted sex ratio and fertility status, and the expected relative frequency of F2 offspring for each allele combination. Offspring that are expected to be all female and infertile are enclosed in a red box. [Figure 26] Figure 26 shows photographs illustrating the effects of FSHR deficiency on ovarian development. The gonads of siblings from 12-month-old female controls (WT body color - lower panel) and albino F0 FSHR mutant females (FSHR - / -, tyr- / -; upper panel), which were similar in body size, were dissected for morphological analysis. The left image shows the dissected intraperitoneal ovaries of the control and mutant females. White arrows point to the gonads, and black arrows point to the urogenital process. The FSHR mutation resulted in complete cessation of follicular formation, leading to atrophic, string-like gonads. Wild-type females had large, protruding urogenital processes, while albino F0 FSHR - / - females had much smaller processes. [Figure 27]Figure 27 shows a germ cell transplantation method that enables the mass production of donors derived from gametes containing the FEM (cyp17, Cyp19a1a), SMS (Tjp1a, Csnk2a2, Gopc, Smap2, Hiat1), MA (Dmrt1, Gsdf), and FLS genes (Vtgs, FSHR). In mutant donors, these defective genes result in an increased number of monosexual males (FEM gene) or females (MA gene) individuals, or the presence of non-functional sperm (SMS gene) or oocytes (FLS gene). Therefore, mass production of these homozygous mutants is impossible. To circumvent this limitation, the inventors targeted only genes whose mutant phenotypes result from dysfunction in somatic cells, not germ cells, and produced chimeric embryos using a "germ cell transplantation" technique. To produce chimeras, ovarian or testicular cell suspensions from homozygous mutant juvenile fish were transplanted into the peritoneal cavity of germ-less host organisms that were wild-type relative to the target gene. This method results in chimeric embryos from the wild-type host that possess normal somatic cells but a mutant reproductive system. These chimeric host organisms, by possessing functional somatic cells, restore a normal sex ratio and / or infertility. These host fish can be used as commercial breeding stock to mass-produce unisexual and / or infertile fish. [Figure 28]Figure 28 shows a germ cell transplantation method for mass-producing functional sperm with the spermatogenesis defect gene (SMS (-)). No defects are observed in primordial germ cells (PGCs) and spermatogonial cell generation in SMS nulls obtained from heterozygous SMS mutant parents. However, at maturity, SMS mutant males produce only immotile sperm with rounded heads that are not fertile. Female SMS mutants are fertile. This SMS gene is expressed in somatic cells surrounding active germ cells (Sertoli cells and Leydig cells). A deficiency in the SMS protein causes a defective microenvironment that impairs sperm maturation. To restore spermatogenesis, germline stem cells can be isolated from SMS mutant juvenile fish and transplanted into recipient embryos that have a functional SMS gene, although their own PGC levels are drastically reduced. The transplanted SMS - / - spermatogonial stem cells form the recipient's gonads, and since SMS is essential for their continued development, the recipient's somatic cells nourish the transplanted germ cells, restoring spermatogenesis and enabling the production of functional sperm. All of these cells possess the mutant SMS gene. [Figure 29]Figure 29 shows a germ cell transplantation method to produce functional eggs with the vitelogenin-deficient gene (Vtg (-)). No defects are observed in primordial germ cells (PGCs) and oogonia generation in Vtg-null fish from heterozygous Vtg mutant parents. However, at maturity, Vtg mutant females produce only oocytes lacking the Vtg protein, leading to female infertility. Vtg-deficient males develop normally and are fertile. The Vtg gene is normally expressed in the Vtg protein, which is transported to oocytes via hepatocytes and bloodstream. The deficiency of the Vtg protein causes the egg to lack essential nutrients necessary for early embryonic or larval development, leading to developmental arrest. Therefore, Vtg- / - females are prolific. To restore yolk formation, germline stem cells are isolated from Vtg null mutant juvenile fish. Although their own PGCs are drastically reduced, they can be transplanted into embryos of recipients that possess the functional Vtg gene. The transplanted Vtg - / - germline stem cells form the recipient's gonads, and the surrogate mother's hepatocytes ensure that nutrients supporting early development are properly delivered to the egg. Female recipients mated with Vtg - / - males produce viable Vtg - / - offspring. [Figure 30] Figure 30 shows a germ cell transplantation method for producing viable FSHR-mutant oocytes (FSHR (-)). No defects were observed in primordial germ cells (PGCs) and oogonia generation in FSHR-null oocytes obtained from heterozygous FSHR mutant parents. However, at maturity, FSHR mutant females do not respond to FSHR-mediated signals, leading to the cessation of follicular formation and resulting in female development. FSHR knockout males develop normally and are fertile. Because FSHR is expressed alone in the follicular cells of the body, the amount of PGCs in FSHR null mutant juvenile fish is drastically reduced. However, by transplanting germline stem cells into the embryo of a recipient organism with a functional FSHR gene, normal oocyte development can be restored, making it possible to produce viable oocytes. Female recipient organisms mated with FSHR (- / -) males produce only FSHR (- / -) offspring. [Figure 31]Figure 31 shows a germ cell transplantation method for producing functional FEM-mutant oocytes (FEM: Cyp19a1a and cyp17). The inventors found no defects in primordial germ cell (PGC) and oogonia generation in FEM-null fish primitives obtained from heterozygous FEM mutant parents. However, at maturity, FEM mutant females do not convert androgens to estrogen, thereby reprogramming ovarian somatic cell-instructing cells (follicular membrane cells and granulosa cells) and testicular somatic cell-instructing cells (Leydig cells and Sertoli cells), reverting genetically female to phenotypic male. They do not respond to FSHR-mediated signals, leading to cessation of follicular formation and resulting in femaleness. FSHR knockout males develop normally and are fertile. FEM-deficient males develop normally and are fertile. The FEM gene is normally expressed in ovarian somatic cells. To enable the mass production of oocytes with the FEM-deficient gene, germline stem cells are isolated from FEM null mutant juvenile fish. Although their own PGCs are drastically reduced, they can be transplanted into embryos of recipients that possess functional FEM genes. The transplanted FEM - / - germline cells form the recipient's gonads. These somatic cells surrounding the donor oocyte produce normal amounts of estrogen, enabling the progression of follicular formation and the maintenance of female fate. These recipient females, when mated with FEM (- / -) males, produce only FEM - / - offspring. [Figure 32] Figure 32 is a diagram illustrating a method for mass-producing all-male, infertile fish. Double KO parents (e.g., SMS and cyp17) can be propagated using the germ cell transplantation techniques shown in Figures 27-32. These parent stocks produce only donors derived from gametes carrying the mutated gene. Natural and artificial mating of these parent stocks produces only all-male, infertile populations. [Figure 33]Panels A and B of Figure 33 illustrate germ cell transplantation experiments that successfully produced and generated donors derived from tilapia gametes. Panel A of Figure 33 shows germ cell transplantation into newly hatched tilapia larvae lacking germ cells. Mutant donor spermatogonial stem cells (SSCs) were transplanted into the peritoneal cavity of larvae with drastically reduced endogenous germ cells. Two groups of SSCs were transplanted simultaneously: one group with an in-frame △3nt deletion in the reference gene and a 6nt insertion in the pigment gene (tyri6 / i6), and another group with an out-of-frame 4nt deletion in the reference gene and a 22 deletion in the pigment gene (tyr△22 / △22). This 3nt deletion was not expected to alter gene function and therefore served as a positive control. These transplanted cells migrated and formed the genital ridge of the transplantee. After sexual maturity was achieved, gametes were collected from the transplanted fish, and their DNA was analyzed by a PCR fragment size assay using PCR primers adjacent to the mutant region of the donor derived from the gametes. The amplified products were classified and detected by capillary electrophoresis. After spermatogonial stem cell transplantation, the percentage of female and male transplants producing functional oocytes and sperm derived from donor cells was determined. Figure 33 Panel B shows the analysis of the capillary fragment lengths of sperm DNA collected from wild-type controls and transplanted fertile tilapia. The traces below show only donors derived from △3nt and △4nt deletion fragments of the reference gene, along with 6nt insertions and △22nt deletion fragments in the pigment gene. A negative control with a gene-specific fragment (268bp) for the wild-type-sized test gene and 467nt for the tyr gene is shown as a reference. [Figure 34]Figure 34 panels A to D show different methods for breeding a number of unisexual infertile individuals. FEM- / - and MA- / - represent female and male null genes. SMS- / - and FLS- / - represent spermatogenesis and follicularogenesis null genes. Male and female breeding stock were produced only by germ cell transplantation using steroid hormone manipulation and germ cell transplantation (Figure 34 panels A and B) (Figure 34 panels C and D). It is possible to cross a limited number of breeding stock to mass-produce millions of all-male infertile embryos (Figure 34 panels A and C) or all-female infertile embryos (Figure 34 panels B and D) for use in aquaculture systems. [Modes for carrying out the invention]

[0044] Overall, this disclosure provides a method for generating sterile, sex-determined fish, crustaceans, or mollusks. This method includes: (i) breeding a fertile hemizygous mutant female fish, crustacean, or mollusk with at least the first and second mutations; and breeding a fertile hemizygous mutant male fish, crustacean, or mollusk with at least the first and second mutations; and selecting homozygous mutant primitives, which are sterile, sex-determined fish, crustaceans, or mollusks, by genotype selection. The first mutation inhibits one or more genes that determine sex differentiation. The second mutation inhibits one or more genes that determine gamete function.

[0045] This disclosure also describes a method for producing sterile sex-determined fish, crustaceans, or mollusks. This method includes the following steps: to produce sterile sex-determined fish, crustaceans, or mollusks, (i) breed a fertile homozygous mutant female fish, crustacean, or mollusk having at least the first and second mutations with (ii) a fertile homozygous mutant male fish, crustacean, or mollusk having at least the first and second mutations. The first mutation inhibits one or more genes that determine sex differentiation. The second mutation inhibits one or more genes that determine gamete function. The fertile capacity of the fertile homozygous female fish, crustacean, or mollusk and the fertile homozygous mutant male fish, crustacean, or mollusk is restored.

[0046] This disclosure also describes a method for producing sterile sex-determined fish, crustaceans, or mollusks. This method includes the following steps: to produce sterile sex-determined fish, crustaceans, or mollusks, (i) a fertile homozygous mutant female fish, crustacean, or mollusk with a homozygous mutation is (ii) bred with a fertile homozygous mutant male fish, crustacean, or mollusk with a homozygous mutation. This mutation directly or indirectly inhibits spermatogenesis and / or directly inhibits yolk formation. The fertility of the fertile female fish, crustacean, or mollusk and the fertile male fish, crustacean, or mollusk is restored.

[0047] This disclosure also describes a method for producing fertile homozygous mutant fish, crustaceans, or mollusks that produce sterile sex-determined fish, crustaceans, or mollusks. This method includes the following steps: to produce sterile sex-determined fish, crustaceans, or mollusks, (i) breed fertile hemizygous mutant female fish, crustaceans, or mollusks having at least the first and second mutations with (ii) hemizygous mutant male fish, crustaceans, or mollusks having at least the first and second mutations; select a homozygous primitive by genotype selection; and restore the fertility of this homozygous primitive. The first mutation inhibits one or more genes that determine sex differentiation. The second mutation inhibits one or more genes that determine gamete function.

[0048] This disclosure also presents fertile homozygous mutant fish, crustaceans, or mollusks for producing sterile, sex-determined fish, crustaceans, or mollusks. In these fertile homozygous mutant fish, crustaceans, or mollusks having at least the first and second mutations, the first mutation inhibits one or more genes that determine sex differentiation, and the second mutation inhibits one or more genes that determine gamete function. The fertile homozygous mutant fish, crustaceans, or mollusks regain their reproductive capacity.

[0049] This disclosure also presents fertile homozygous mutant fish, crustaceans, or mollusks for producing sterile, sex-determined fish, crustaceans, or mollusks. This mutation directly or indirectly inhibits spermatogenesis and / or directly inhibits yolk formation, restoring the fertile capacity of these fertile fish, crustaceans, or mollusks.

[0050] In the context of this disclosure, "fish" refers to any cephalate animal that has gills and lacks fingers or limbs. Examples of fish include carp, tilapia, salmon, trout, and catfish. In the context of this disclosure, "crustaceans" refers to all arthropod taxa. Examples of crustaceans include crabs, lobsters, crayfish, and shrimp. In the context of this disclosure, "mollusks" refers to any invertebrate animal that has a soft, unsegmented body and is usually covered by a calcareous shell. Examples of mollusks include bivalves, scallops, oysters, octopuses, squid, and chitons.

[0051] A sterile fish, crustacean, or mollusk is any fish, crustacean, or mollusk whose ability to reproduce or mate to produce offspring is reduced compared to a wild-type control; for example, a sterile fish, crustacean, or mollusk has a reduced probability of producing viable offspring by approximately 50%, 75%, 90%, 95%, or 100%. In contrast, a fertile fish, crustacean, or mollusk is any fish, crustacean, or mollusk that is capable of reproducing or mate to produce offspring. Reproduction and mating are all processes in the mating of a male and female species to produce offspring or offspring.

[0052] Sex-determined fish, crustaceans, or mollusks are the primitive forms of all fish, crustaceans, or mollusks, whose primitive sex is predetermined by inhibiting the primitive sex differentiation pathway. There are also some examples where sex-determined primitives of the same generation are monosex.

[0053] The function of gametes is the process by which gametes combine with other gametes during fertilization.

[0054] Mutations that inhibit one or more genes determining sex differentiation are any gene mutations that directly or indirectly modulate gonadal function. Direct or indirect action on gonadal function means (1) mutating the coding sequence of one or more gonadal cell genes; (2) mutating the non-coding sequence that moderately controls the transcription of one or more gonadal cells; (3) mutating the coding sequence of one or more other genes involved in the post-translational regulation of gonadal cells; or (4) a combination thereof. Modulation of gonadal function means determining whether the gonads produce female gametes or male gametes. Examples of when masculinization is preferred include, for example, the modification of one or more genes that regulate androgen and / or estrogen synthesis, such as the modification of the expression of aromatases Cyp19a1a, Cyp17, or a combination thereof. Genes involved in regulating aromatase Cyp19a1an expression include cyp19a1a, FoxL2, sf1 (steroid-producing factor 1), and their orthologues. Genes involved in regulating Cyp17 expression include cyp17I or its orthologue. Examples of when feminization is preferred include the regulation of one or more genes that regulate the expression of aromatase Cyp19a1a inhibitors. Genes that regulate the expression of aromatase Cyp19a1a inhibitors include Gsdf, dmrt1, Amh, Amhr, and their orthologues.

[0055] Alternatively, sex differentiation may be determined without one or more gene mutations. Examples of non-genetic mutation methods that determine sex differentiation include methods that utilize sex reversal (hormone manipulation) and breeding, progeny testing, androgenesis, and gynogenesis, which can produce monosexual male or female populations that are homozygous XX, YY, or ZZ (see example

[21] ; Dunham 2004, as included by reference). In some examples of this disclosure, breeding (i) homozygous mutant fertile female fish, crustaceans, or mollusks with (ii) homozygous mutant fertile male fish, crustaceans, or mollusks to produce sterile sex-determined fish, crustaceans, or mollusks includes a non-genetic mutation method for determining sex differentiation. In some examples of this disclosure using salmon, the creation of sex-reversed males (XX) and breeding with females produces monosexual female primitives. In other examples provided herein, the determination of sex differentiation can be achieved through interspecific hybridization (see, as indicated by reference, Pruginin, Rothbard et al. 1975, Wolters and DeMay 1996).

[0056] Mutations that inhibit one or more genes that determine gamete function are any gene mutations that directly or indirectly regulate spermatogenesis, oogenesis, and / or follicular formation in order to produce infertile fish, crustaceans, or mollusks. Direct or indirect regulation of spermatogenesis, oogenesis, and / or follicular formation means (1) mutating the coding sequence of one or more gamete genes; (2) mutating the non-coding sequence that partially controls the transcription of one or more gamete genes; (3) mutating the coding sequence of other genes that are involved in the post-translational regulation of one or more gamete genes; or (4) a combination thereof.

[0057] Mutations that directly or indirectly inhibit spermatogenesis and / or directly inhibit yolk formation are any gene mutations that directly or indirectly regulate spermatogenesis and / or directly inhibit yolk formation in order to produce infertile fish, crustaceans, or mollusks. Direct or indirect regulation of spermatogenesis means (1) mutating the coding sequence of one or more gamete genes involved in spermatogenesis; (2) mutating non-coding sequences that partially control the transcription of one or more gamete genes involved in spermatogenesis; (3) mutating the coding sequence of other genes involved in the post-translational regulation of one or more gamete genes involved in spermatogenesis; or (4) a combination thereof. Direct regulation of yolk formation means (1) mutating the coding sequence of one or more gamete genes involved in yolk formation; (2) mutating non-coding sequences that partially control the transcription of one or more gamete genes involved in yolk formation; or (3) a combination thereof, in order to produce infertile fish, crustaceans, or mollusks.

[0058] Examples of situations where the production of infertile male fish, crustaceans, or mollusks is preferred include the modification of one or more genes that regulate spermatogenesis. Examples of one or more genes that regulate spermatogenesis may cause megahead spermatosis, sperm with a rounded head, rounded nucleus, truncated midsection, partially coiled tail, or a combination thereof. Examples of genes that cause megahead spermatosis include Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, and their orthologues. Examples of situations where the production of infertile female fish, crustaceans, or mollusks is preferred include the modification of one or more genes that regulate oogenesis, follicular formation, or combinations thereof. Examples of one or more genes that regulate oogenesis include one or more genes that regulate estrogen synthesis. Examples of one or more genes that regulate estrogen synthesis include FSHR or its orthologue. Examples of one or more genes that regulate follicular formation include one or more genes that regulate vitellogenesis expression. Examples of one or more genes that regulate vitellogenesis expression include vtgs or its orthologue. Examples of mutations that directly or indirectly inhibit spermatogenesis include those in Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, or their orthologs. Examples of mutations that directly inhibit yolk formation include those in genes encoding or controlling vitellogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein, or their orthologs.

[0059] A mutation can be any type of change in the nucleotide sequence in question, such as nucleotide insertions, nucleotide deletions, and nucleotide substitutions.

[0060] Restoring fertility or reproductive capacity refers to any process by which infertile fish, crustaceans, or mollusks are converted into fertile fish, crustaceans, or mollusks. In some cases, aromatase inhibitors are administered to infertile fish, crustaceans, or mollusks to restore reproductive capacity. In other cases, reproductive capacity is restored through germline stem cell transplantation of infertile fish, crustaceans, or mollusks. Germline stem cell transplantation refers to any process by which germline stem cells from infertile fish, crustaceans, or mollusks are transplanted into fertile fish, crustaceans, or mollusks to restore reproductive capacity. In some examples provided herein, germline stem cell transplantation is a process comprising: obtaining germline stem cells from a sterile homozygous male fish, crustacean, or mollusk having at least the first and second mutations, or from a sterile homozygous female fish, crustacean, or mollusk having at least the first and second mutations; and transplanting these germline stem cells into a germline-depleted male fish, crustacean, or mollusk or a germline-depleted female fish, crustacean, or mollusk. The germline-depleted male or female fish, crustacean, or mollusk is any embryo that has a drastically reduced number of its own germ cells but possesses a functional copy of the target gene that determines sex differentiation and gamete function. Alternatively, the germline-depleted The transplanted host organism is a chimeric fish, crustacean, or mollusk with normal somatic cells but a mutant germline. Transplants lacking germ cells can sometimes be created through ploidy manipulation, hybridization, or exposure to high levels of sex hormones. Exposure of aquatic juvenile fish to high levels of sex hormones can lead to sterilization of the exposed animals. While this method has been proven (Hunter et al, 1982; Solar et al, 1984; Piferrer et al, 1994), it is not used on a commercial scale.While this method may be effective in creating infertile fish, it has not been proven to be effective in inducing infertility in 100% treated fish. While treated fish may be ideal for research or as recipients for germ cell transplantation, this method is not sufficient to produce sterile fish for commercial aquaculture (Hunter, GA, EM Donaldson, FW Goetz, and PR Edgell. 1982. Production of all-female and sterile coho salmon and experimental demonstration of male heteromorphism, Journal of the American Society of Fisheries Science 111: 367-372; Piferrer, F, M Carillo, S. Zanuy, II Solar, and EM Donaldson. 1994. Induction of sterilization in coho salmon (Oncorhynchus kisutch) by androgen immersion before initial feeding, Aquaculture 119: 409-423; and Solar, I., EM Donaldson, and GA Hunter. 1984.). Optimization of treatment methods for controlled sex differentiation by oral administration of 17α-methyltestosterone and sterilization in wild rainbow trout (Salmo gairdeneri Richardson), see Aquaculture 42: 129-139).

[0061] In some cases, this germline stem cell transplantation is a process that includes: obtaining spermatogonial stem cells from infertile homozygous male fish, crustaceans, or mollusks, or oogonial stem cells from infertile homozygous female fish, crustaceans, or mollusks; and transplanting the spermatogonial stem cells into the peritoneal cavity of germline-free embryos, or into germline-free differentiated testes or ovaries of fish, crustaceans, or mollusks. If necessary, in addition to germline stem cell transplantation, exogenous steroids, such as estrogen, are provided to the infertile fish, crustaceans, or mollusks to restore fertility. In other cases, aromatase inhibitors are provided to the infertile fish, crustaceans, or mollusks to restore fertility.

[0062] Figure 1 is a flowchart according to this disclosure showing a method for creating male and female breeding stock, such as fertile homozygous mutant male and female fish, crustaceans, or mollusks, for use in producing, for example, sterile sex-determined fish, crustaceans, or mollusks.

[0063] Figure 1 shows the genetic pathways that govern sex differentiation and gametification, as well as the knockout (KO) method for producing monosexual, infertile populations.

[0064] One or more mutations in the genes cyp19a1a, FoxI2, or a combination thereof result in reduced or decreased estrogen expression, which is responsible for testis formation and the production of male fish, crustaceans, or mollusks. Similarly, one or more mutations in the gene cyp17 result in reduced or decreased estrogen and androgen, which are responsible for the production of male fish, crustaceans, or mollusks. One or more additional mutations in genes that inhibit spermatogenesis (SMS) result in male fish, crustaceans, or mollusks. Thus, infertile homozygous mutant male fish, crustaceans, or mollusks are produced.

[0065] In an additional step used to breed this strain, the reproductive capacity of the infertile homozygous mutant male fish, crustacean, or mollusk may be restored by estrogen treatment. The following treatment produces a reproductive homozygous mutant female fish, crustacean, or mollusk. In this sex reversal process, the phenotypic female has one or more mutations that inhibit spermatogenesis, and produces fertile oocytes with one or more further spermatogenic mutations, enabling breeding of this strain. Alternatively, as described in Example 10, the reproductive capacity of the infertile homozygous mutant male fish, crustacean, or mollusk may be restored by incorporating the germ cells of the infertile homozygous mutant male fish, crustacean, or mollusk into the testicular cells of a reproductive wild-type male, thereby enabling breeding of this strain.

[0066] In contrast to Figure 1, one or more mutations in the genes Gsdf, Dmrt1, or a combination thereof lead to inactivation of the Cyp19a1a inhibitor, resulting in high or increased estrogen expression, which is responsible for ovarian development and the production of female fish, crustaceans, or mollusks. One or more additional mutations in genes that regulate oogenesis, follicular development (FLS), or a combination thereof result in infertile female fish, crustaceans, or mollusks. Thus, infertile homozygous mutant female fish, crustaceans, or mollusks are produced.

[0067] In an additional step used to breed this strain, the reproductive capacity of the infertile homozygous mutant female fish, crustacean, or mollusk may be restored by treatment with an aromatase inhibitor. The following treatment produces a reproductive homozygous mutant male fish, crustacean, or mollusk. In this sex reversal process, the phenotypic male has one or more mutations that inhibit oogenesis, follicular formation, or a combination thereof, and is reproductive, producing sperm with one or more mutations that further inhibit oogenesis, follicular formation, or a combination thereof, enabling breeding of this strain. Alternatively, as described in Example 10, the reproductive capacity of the infertile homozygous mutant female fish, crustacean, or mollusk may be restored by incorporating the germ cells of the infertile homozygous mutant female fish, crustacean, or mollusk into the ovarian cells of a reproductive wild-type female in order to produce a reproductive homozygous mutant female fish, crustacean, or mollusk, thereby enabling breeding of this strain. [Examples]

[0068] Example 1 - Materials and Methods Statement regarding the animals used and ethical considerations: All experiments, in compliance with U.S. regulations to ensure animal welfare and livestock practices, were conducted in accordance with CAT-004, an IACUC-approved animal protocol. The tilapia (Oreochromis niloticus) strain used in this study originated from a Brazilian strain obtained from a U.S. commercial producer.

[0069] Nuclease production and methods:Generation of F0 mutants: Tilapia orthologs of cyp17, Cyp19a1a, Tjp1a, Csnk2a2, Hiat1, Smap2, Gopc, Gsdf, Dmrt1, FSHR, and vitelogenin genes (VtgAa and VtgAb) were identified in silico from genome databases.

[0070] To create DNA double-strand breaks (DSBs) at specific gene locations, the inventors used artificial nucleases. In most applications, the absence of repair genes resulted in the production of a single DSB, leading to non-homologous inactivation and binding to the repair pathway (NHEJ). This NHEJ can result in an incomplete repair pathway, generating an insertion or deletion (indel) at the target site. The introduction of an indel can create a frameshift within the gene's coding region, resulting in an abnormal protein product with an incorrect amino acid sequence. To increase the frequency of null mutations in the target gene, the inventors targeted two different exons simultaneously in addition to the target cyp17. In parallel with this target gene, the inventors co-targeted a chromogenic gene that functions as a mutagenic color selection marker. Typically, there is a correlation between the mutation frequencies of chromogenic genes and the target gene. Therefore, embryos with complete pigment deficiency (albino phenotype) were preferentially selected compared to mosaic pigment phenotypes (partial gene inactivation). To confirm the function of the newly designed nuclease, five albino embryos were selected from each treated group, and gene modifications at the gene locus were quantitatively analyzed by PCR fragment analysis. If all five tested embryos did not show indels at the target gene locus, the treated embryos from the same g-loop were removed. Furthermore, the inventors preferentially grew embryo groups in which mutations occurred at the 1- or 2-cell stage (e.g., detecting 2 or 4 mutant alleles at the target gene locus by fragment analysis).

[0071] The template DNA encoding the artificial nuclease was linearized and purified using a DNA Clean & Concentrator-5 column (Zymo Research). Using the mMESSAGE mMACHINE T3 kit (Invitrogen), 1 microgram of the linearized template was used to synthesize cap RNA, which was purified using Qiaquick (Qiagen) and stored in RNase-free water at a final concentration of 800 ng / μl at -80°C.

[0072] Injection into the embryo: Embryos were produced via in vitro fertilization. A solution containing an artificial nuclease, totaling approximately 10 nL in volume, was simultaneously injected into the cytoplasm of one-cell stage embryos. Typically, injection into 200 embryos yields 10 to 60 embryos completely lacking pigment (albino phenotype). The viability of the embryos / larvae was monitored for 10 to 12 days after injection.

[0073] Founder's Choice: At least 10 albino embryos were incubated until 3 months of age, and gene modifications were quantitatively analyzed by fluorescence PCR fragment analysis (see Table 1, columns 8 and 11 for gene-specific phenotyping primers). The inventors preferred to select founders in which mutations occurred at the 1- or 2-cell stage (e.g., 2 or 4 mutant alleles were detected at the target gene locus by fragment analysis (Figure 2)).

[0074] F1 genotyping:These selected founders were crossbred with wild-type strains. These F1 offspring were raised to 2 months of age, anesthetized in MS-222 (tricaine) 200 mg / L, and transferred to a clean surface using a plastic spoon. The fins were cut off with a razor blade and placed on a well (a 96-well plate with caps). The fin DNA was then analyzed by fluorescence PCR, and the fish from which the fins were removed were placed in individual containers. Briefly, to digest the tissue overnight and extract gDNA, 60 μl of a solution containing 9.4% Kirex and 0.625 mg / ml of proteinase K was added to each well in a 55°C incubator. The plate was then agitated and centrifuged. Finally, to remove all PCR inhibitors from the mixture, the gDNA extract was diluted 10-fold with ultra-clean water. The inventors selected approximately 20 juvenile fish with the same size variant and analyzed mainly 80 juvenile / founder fish for rearing purposes.

[0075] Fluorescence PCR (see Figure 2): For the PCR reaction, 5 μL of a PCR master mix (Quiagen Multiplex PCR) was used, which contained 3.8 μL of water, 0.2 μL of fin DNA, and three primers: a tailed primer labeled with a fluorescent tag (6-FAM, NED), an amplicon-specific forward primer with a forward tail (SEQ ID NO: 117: 5′ -TGTAAAACGACGGCCAGT-3′ and SEQ ID NO: 118: 5′ -TAGGAGTGCAGCAAGCAT-3′), and an amplicon-specific reverse primer (fluorescent PCR gene-specific primers are listed in Table 1). The PCR conditions were as follows: denaturation at 95°C for 15 minutes, followed by 30 amplification cycles (30 seconds at 94°C, 45 seconds at 57°C, and 45 seconds at 72°C), then 8 amplification cycles (30 seconds at 94°C, 45 seconds at 53°C, and 45 seconds at 72°C), and a final extension reaction at 72°C for 10 minutes, followed by indefinite holding at 4°C. The resulting amplicons were digested by capillary electrophoresis (CE) using a 1:10 dilution (1-2 microliters) with added LIZ labeling size criteria to determine amplicon size accurately at base pair resolution (Retrogen Inc., San Diego). Raw trace files were analyzed using Peak Scanner software (Thermo Fisher). Peak size compared to wild-type peak control determined the nature (insertion or deletion) and length of the mutation. The number of peaks indicates the level of mosaicism. The inventors selected F0 mosaic founders with the fewest mutant alleles (selectively 2-4 peaks).

[0076] The size of this allele was used to refine the indel mutation under observation. For further confirmation by sequencing, mutants expected to be frameshift mutations were selected, rather than multiples of 3 bp. To facilitate genotyping by QPCR thawing analysis for the next generation, mutations larger than 8 bp but smaller than 30 bp were preferred. For sequence confirmation, the PCR product of this selected indel was further sent to gene sequencing. Sequencing chromatography of PCR with two reads simultaneously indicates the presence of an indel. The onset of deletion or insertion usually begins when the sequence reads branch. This dual sequence is then carefully analyzed to detect unique nucleotide reads. Subsequently, the pattern of unique nucleotide reads is analyzed against a series of artificial single read patterns generated by gradually shifting the wild-type sequence itself.

[0077] QPCR genotyping of F1 and F2 generationsReal-time qPCR was performed using a ROTOR-GENE RG-3000 real-time PCR system (Corbett Research). A total of 10 μL of 1-μL gene DNA (gDNA) template (diluted at 5-20 ng / μl) was used, containing 0.15 μM concentrates of forward and reverse primers and 5 μL of QPCR 2x Master Mix (Apex Bio). The qPCR primers used are listed in Table 2 (genotyping RT-PCR primers are in columns 11-14). This qPCR was performed for 40 cycles at 95°C for 15 seconds and 60°C for 60 seconds, and then melting curve analysis was performed to confirm the specificity of the assay (67°C to 97°C). In this method, a short PCR amplicon (approximately 120-200 bp) containing the target region was generated from the gDNA sample, and this depends on temperature-dependent dissociation (melting curve). When indel induction occurs within hemizygous gDNA, heteroduplex molecules and different homoduplex molecules are formed. Multiple double-strand molecule formation is detected by the melting profile, where double dissolution indicates whether it functions as a single species or as one or more species. Generally, the symmetry of the melting curve and melting temperature suggests uniformity of the dsDNA sequence and its length. Therefore, homozygous and wild-type (WT) gDNAs exhibit symmetrical melting curves distinguishable by different melting temperatures. This melting analysis was performed in comparison to a reference DNA sample (from control wild-type DNA) amplified simultaneously with the same mastermix reaction. In short, the changes in the melting profile differ depending on the amplicons generated from homozygous, hemizygous, and WT gDNA (see Figure 3).

[0078] Evaluation of sterilization in malesFor each genotype, the amount and concentration of exfoliable sperm collected from five males (5 months old) were measured. Sperm were counted using spectrophotometry (optical density (OD) 600 nm) and Neubauer hemocytometers of serially diluted samples. Sperm motility was measured in terms of sperm viability in the field of view [4]. Morphology of spermatids stained with nigrosine and eosin was analyzed at 400x magnification using a light microscope. Wild-type eggs collected from three different females were fertilized in vitro to determine the optimal sperm-to-egg ratio (sperm 5.10). 6 The fertilization capacity of sperm in 100 eggs was analyzed. Sperm collected from wild-type males was used, and the quality of wild-type eggs was also investigated. The fertilization rate was expressed as the percentage of surviving embryos relative to the total number of eggs collected 24 hours after fertilization. The mean values ​​obtained from these investigations were compared across all mutant genotypes using independent t-tests.

[0079] Evaluation of sterilization in females The inventors recorded the weight of all sampled fish. For each genotype, at least six 4- and 6-month-old females were dissected, and in situ images were taken before dissection of the gonads. The mean total gonad weight index was statistically compared across all genotypes (independent t-test). The survival rates of at least six eggs, embryos, and larvae from outcrosses with wild-type males were statistically analyzed and compared to controls (wild-type females mated with mutant males).

[0080] Isolation of donor cells and transplantation of gonadal cells:As described by Lacerda, germline stem cells were collected from the gonads of 3-4 month old fish (~50-70g) by enzymatic digestion [5]. Briefly, fresh isolated gonads were finely chopped and cultured for 3-4 hours at 25°C in PBS (pH 8.2) dissolved in 1 ml of 0.5% trypsin (Worthington Biochemical Corp., Lakewood, New Jersey) containing 5% fetal bovine serum (Gibco Invitrogen Co., Grand Island, New York) and 0.05% DNase I (Roche Diagnostics, Mannheim, Germany). During culture, gentle pipetting was performed to physically inhibit any intact remaining parts of the gonad. Prior to storage on ice until transplantation, any unseparated cell clumps were removed. The resulting cell suspension was filtered through a 42 μm nylon screen (N-No.330T; Tokyo Screen Co., Ltd., Tokyo, Japan), and then resuspended in L-15 medium (Gibco Invitrogen Co.).

[0081] Germ cell-free transplant larvae (5-7 dpf) were anesthetized with 0.0075% ethylmethanesulfonate 3-aminobenzoate (Sigma-Aldrich Inc.) and transferred to petri dishes covered with 2% agar. Approximately 15,000 testicular cells were injected into the peritoneal cavity of about 80 larval offspring of Elavl2 hemizygous mutant parents to perform cell transplantation. Alternatively, MSC homozygous females and wild-type males were mated, and embryos without PGCs were collected [6]. After transplantation, the transplant larvae were returned to aerated embryo culture medium and grown to adulthood.

[0082] [Table 1]

[0083] [Table 2]

[0084] Example 2 - Use of gene editing tools for inducing bi-allele knockout in tilapia F0 generation The inventors separately targeted two genes involved in pigment formation: the genes encoding tyrosinase (tyr) [2] and the mitochondrial inner membrane protein MpV17 (mpv17) (Krauss, Astrinides et al. 2013) [8]. The inventors found that 50% and 46% of the embryos injected at the Tyr and mpv17 loci, respectively, showed high mutability (Figure 4). The loss-of-function alleles became cell-autonomously pigmentless melanocytes in the embryonic body (Figure 4 panel B) and retinal pigment epithelium (Figure 4 panel C), producing embryonic phenotypes ranging from complete to partial loss of melanin and iridophoric pigments, which were easily identifiable compared to the wild-type phenotype (Figure 4 panels A and C). Embryos showing complete pigment loss (10-30% of the treated fish) were raised to 3 months of age, and all of them lacked wild-type tyr and mpv17 sequences. These fish exhibit translucent and albino phenotypes (Figure 4, Panel D), suggesting that functional studies can be conducted in F0 tilapia.

[0085] Example 3 - Multiple gene targeting in tilapia The inventors investigated whether multiple genome loci can be targeted simultaneously and whether mutagenicity efficiency measured at one locus can predict mutations at another locus in the tilapia genome. To test the hypothesis of the present invention, the inventors co-targeted tyr and Dead-end1 (dnd). Dnd is a PGC-specific RNA-binding protein (RBP) that maintains germ cell fate and motility [3]. After injection of programmed nucleases, the inventors found that mutations in both the tyr and dnd gene targets were highly correlated. Approximately 95% of albino (tyr) mutants also had mutations at the dnd locus, demonstrating the suitability of pigment deficiency as a selection marker (Figure 5 Panel A). Further analysis of the gonads of 10 albino fish revealed that the testes of 6 were translucent and lacked germ cells (Figure 5 Panel B). The expression of vasa, a germ cell-specific marker strongly expressed in wild-type testes, was not detected at all in the testes of dnd mutants. This result suggests that the expression of zygosity-dependent dnd is necessary for the maintenance of germ cells, and furthermore, dnd mRNA and / or protein contributed from the mother cannot restore the loss of zygosity of this gene.

[0086] Example 4 - Production of gonads without germ cells The inventors produced infertile tilapia by injecting antisense-modified oligonucleotides (dnd-morpholino and dnd-AUM oligo) to transiently silence the dnd gene in the embryo. The inventors also produced infertile tilapia after exposing embryos to the small molecule first detected on screen to remove PGCs from zebrafish

[10] . The inventors further produced infertile tilapia using a gene knockout method, as described for dnd in the preceding section (Example 3). The inventors also found that it is possible to produce hermaphroditic adults without germ cells by crossing an Elavl2 heterozygous mutant system with selected homozygous mutant offspring (Figure 6). These gene KO methods, in combination with the other methods described above, produce infertile tilapia, which exhibit either a female urogenital process (UGP) and string-like gonads or a male UGP and translucent tube-like gonads (Figure 6). However, these methods result in only 25% infertility of offspring from heterozygous mutant parents, and other knockdown methods are not stable or reliable enough to ensure complete sterilization of all fish in each batch, making them unsuitable solutions for the commercial production of infertile fish. In the present invention, the mass production of infertile fish relies on surrogate parents that are born without germ cells, undergo germline stem cell transplantation, and ultimately produce donors derived from sperm or eggs. In the methods of the present invention, knockout methods are preferred for sterilizing these surrogate parents (e.g., elavl2-null offspring of heterozygous parents; see Example 11). Other knockout methods besides Elavl2 may be used to produce infertile surrogates, such as null mutants for dead-end1, vasa, nanos3, or piwi-like genes. Such knockout surrogates ensure that, after transplantation, only donors derived from gametes are produced. Depending on the species of fish, crustaceans, or mollusks, alternative methods such as hybridization and triploidization can be used to produce sterile transplants (Benfey et al., 1984; Felip et al., 2001).

[0087] Example 5 - Cyp17I is necessary for the development of female Nile tilapia. Because estrogen plays a crucial role in female differentiation, the balance of steroid-producing hormones can govern sexual differentiation and gonadal mutations in bony fish. However, gonadal differentiation and gamete formation in the absence of both androgens and estrogen remain uninvestigated. To achieve this goal, we have produced an in vitro fertilized tilapia model lacking the cyp17I gene.

[0088] In Nile tilapia, this enzyme is expressed only in theca cells and responds to luteinizing hormone to produce androgens (LH)

[13] . The androgens are then converted to estrogen by follicle-stimulating hormone (FSH)-induced aromatase (cyp19a1a) in the granulosa cells of adjacent developing follicles. Therefore, functional deletion of cyp17 (by gene editing knockout) should simultaneously block the synthesis of both androgens and estrogens. Consistent with this model, the inventors found that 20 out of 22 selected F0 albino / cyp17 mutants developed as phenotypic males and all had extremely small UGPs (Figure 8 Panel C). Given the relationship between androgens and urogenital processes, this genital atrophy is not unexpected

[14] . The reproductive capacity of these F0 males remained unchanged, which may be due to a partial functional deficit phenotype in the mosaic F0 context. For a complete phenotypic analysis, the inventors generated individuals with the same null Δ16-cyp17 mutation in all cells of each individual through selective breeding of F1 offspring (Figure 7). Crosses between F1 heterozygotes (cyp17+ / -) produced ~360 F2 offspring, including Mendelian segregation of normal wild-type (n = 110; cyp17+ / +), hemizygotes (n = 159; cyp17+ / -), and homozygotes (n = 91; cyp17- / -). A total of 155 F2 offspring were sexed at 6 months of age based on morphological characteristics of the urogenital process (UGP). The inventors found that all 33 homozygous fish developed as phenotypic males with atrophied UGP (Figure 8, Panel A). The results of this invention suggest that Cyp17 is essential for female development.

[0089] Subsequently, the inventors measured the amount of free plasma testosterone in wild-type and cyp17 mutant tilapia using ELISA. While the mean testosterone level of 86 pg / mL was measured in wild-type (cyp17+ / +) and heterozygous mutant tilapia (cyp17+ / -), no detectable levels of testosterone were found in homozygous mutants (cyp17- / -) (Figure 8, Panel B). This supports the idea that this enzyme is essential for androgen production.

[0090] The inventors further investigated the morphology and function of the gonads in fish deficient in Cyp17. Simultaneous-sized 5-month-old male siblings cyp17+ / +, cyp17+ / -, and cyp17- / - were dissected, and all organs other than the gonads were removed from the body cavity (Figure 9 Panel A). The wild-type (WT) and hemizygous mutants had pink testes, as is typically seen in sexually mature males, while the homozygous mutants had translucent testes (Figure 9 Panels A and B). Furthermore, the mutant testes were 50% smaller than those of the control (Figure 9 Panel D), and the amount of exfoliable sperm was 20% less than that of the WT (Figure 9 Panel E). In addition, sperm concentration in the homozygous cyp17 mutants decreased 20-fold and 6-fold at 5 and 6 months of age, respectively (Figure 9 Panel F). As evidenced by the inventors' successful fertilization of WT eggs with fish sperm collected from 10 null mutants, no defects were found in the morphology, motility, or function of the sperm.

[0091] The fact that cyp17 null mutants are capable of spermatogenesis suggests that androgens are not essential for this process in Nile tilapia. Therefore, mutants lacking function in this gene may not be sufficient to produce populations of entirely infertile males. To identify the regulatory mechanisms responsible for functional spermatogenesis, the inventors further investigated genes associated with infertile males in mammals.

[0092] Example 6 - Gene candidate targeting spermatogenesis There are significant differences in the morphology and function of mammalian and fish sperm. In particular, mammalian sperm possess acrosomes (important organelles necessary for penetration into the chorionic membrane of the egg) and are mobile in semen, while fish sperm lack acrosomes and are immobile in semen. Megacephalic spermatopathy is rare and a severe form of human infertility characterized by sperm defects in both morphology and function. However, a fish model of this disease likely failed to develop because fish sperm lacked acrosomes. Using genome databases, the inventors identified tilapia orthologs in silico of the following mammalian genes: Csnk2a2

[15] , Gopc [16, 17], Hiat1

[18] , Tjp1a, and Smap2

[21] . To explore the function of these genes in tilapia, the inventors targeted two different exons for each gene (see Figures 10–14). The pigment-forming gene (tyrosinase) was used as a co-target and as a mutagenic color selection marker.

[0093] Along with untreated controls, approximately 20 embryos were grown to adulthood for each candidate gene exhibiting pigmentation abnormalities. At 5 months of age, sperm were stripped from F0 males and WT controls, and sperm concentration, motility, and morphology were analyzed. Compared to controls, all F0 mutant males produced thin sperm. Microscopic examination revealed that the majority of mutant sperm exhibited only trembling motion, and the inventors found widespread (25%–95%) abnormal sperm head morphology, a defect feature seen in megacephalis spermatopathy in humans and mice (Figure 15 Panel A). These mutations resulted in a significant decrease in fertilization rates (Figure 15 Panel B). Furthermore, the inventors found a positive correlation between the severe infertility phenotype and the lowest fertilization rate seen in the Tjp1a mutant, where 95% of sperm are deformed (Figure 15 Panels A and B). The inventors found that all of these F0 mutant females are capable of reproduction.

[0094] The results of this invention demonstrate the existence of evolutionarily conserved pathways that control spermatogenesis in fish and mammals. These results support the concept that targeted inhibition of these corresponding genes would similarly induce infertility phenotypes in many other bony fish, or in other broader taxa.

[0095] Example 7 - All sterile male fish in a cyp17 KO background To manipulate male sterilization, the inventors first controlled a critical branching point in steroid hormone synthesis, regulating the production of both androgens and estrogens. They evaluated the effect of null mutations in the cyp17 gene. The inventors found that all cyp17- / - fish developed as males. Surprisingly, the fish sperm produced by cyp17- / - contained a small number of mature sperm capable of fertilizing oocytes by in vitro fertilization. The inventors then investigated the possibility of blocking spermatogenesis. In a preliminary screening of the invention, the inventors focused on five genes (collectively referred to as spermatogenesis-specific genes or SMS genes: Smap2, Cnsk2a2, Gopc, Hiat1, and Tjp1a) whose mutations cause low fertility in F0 males with severe oligoastheno teratozoospermia, while F0 mutant females are fully fertile. Previous genetic characteristics of F0 KO fish typically indicate mosaic mutations at the corresponding target loci, often in-frame mutations leading to partial phenotypic recovery. Therefore, to measure complete loss-of-function phenotypes, the inventors also performed additional phenotypic characterization on homozygous SMS null mutants. Furthermore, to explore the effect of simultaneously impairing spermatogenesis and steroid hormone synthesis, the inventors established a tilapia strain with double homozygous mutants.

[0096] experiment: To analyze the in vivo function of dual gene knockout in cyp17 and one of these five SMS genes, the inventors used F0 SMS mutant females and cyp17 Δ16 / +Males were crossbred. The genotypes of the offspring (120-180 individuals) were determined for each target gene by PCR fragment analysis (described in Figure 2)

[22] . The inventors raised only individuals with the same mutant allele, hereafter referred to as m1, at the selected SMS locus (typically 12-50% of the F1 offspring population share the same genotype) (Figure 18). Double heterozygotes (e.g., cyp17) Δ16 / + ; SMS m1 / + At least 10 individuals were raised to adulthood. These double heterozygotes were crossbred, and the genotype of the offspring at 1 month of age was analyzed by QPCR thawing. For each of the 9 individuals (see Figure 9) selected to secure potential F2 genotypes, at least 30 fish are currently being raised to adulthood, and their reproductive capacity will be analyzed. Females with cyp17+ / +; SMS+ / m1 (e.g., cyp17+ / +; Tjp1a+ / m1) were reserved for the studies described in the following two sections. Figure 9 summarizes this experimental scheme using Tjp1a as an example of an SMS gene target.

[0097] Without being constrained by theory, the inventors hypothesize that, in finned fish, as seen in mammals, null mutants in all five conserved spermatogenesis-specific genes result in oligoastheno teratospermia, causing infertility. The inventors predict that all double homozygous mutants (cyp17- / -; SMS- / -) will develop into infertile males with even fewer sperm than any single KO male (SMS- / -) with spermatogenesis defects. Indeed, cyp17- / - fish should be deficient in 11-ketotestosterone, a positive regulator of spermatogenesis. This is consistent with the concept that androgens play a paracrine role in spermatogenesis, and the low sperm count of cyp17- / - tilapia has been previously shown. Figure 9 shows nine genotypes, along with four different corresponding phenotypes, with expected proportions: 1) fertile hermaphrodites ~56%, 2) fertile females and infertile males ~19%, 3) all infertile females ~19%, and 4) all infertile males ~6%. By examining each trait individually, the inventors predict that 62% of the offspring population will be male, and 25% of these males will be infertile.

[0098] Example 8 - All sterile male fish in a cyp19a1a KO background An alternative method for generating an all-male population is the inactivation of Cyp19a1a aromatase (hereinafter referred to as Cyp19). The inventors created an out-of-frame mutation in the coding sequence of the tilapia cyp19 gene (Figure 17). This enzyme is produced by gonadal somatic cells and converts testosterone to estrogen. Consistent with this model, the inventors found that among 25 selected F0 Cyp19 mutants, 20 mutants developed as phenotypic males, showing a strong bias toward males (Table 3). In particular, these mutant males exhibited what appeared to be normal male urogenital tracts, unimpaired androgen production, and normal development of secondary sexual characteristics. This is in contrast to cyp17 KO males, which are deficient in androgens and therefore develop atrophied urogenital tracts. By generating all-male, sterile tilapia populations that either express or do not express androgens (as seen in cyp19 KO and cyp17 KO backgrounds, respectively), we were able to explore the effect of male steroid hormones on the growth capacity of tilapia. The testosterone-stimulating effect and responsiveness of GH secretion in mammals are well-established. For complete phenotypic analysis, we generated populations with the same null mutant in all cells of the individual. To breed the F2 generation, we selected heterozygous cyp19 F1 offspring with a Δ10-cyp19 deletion in the first exon. This frameshift mutation was expected to create a truncated protein lacking >98% of the wild-type amino acid sequence (Figure 17). This F2 generation was genotyped and sex-determined. As expected, the inventors found that while the hemizygous (n=97) and wild-type (n=40) tilapia had a normal sex ratio, all homozygous Δ10-cyp19 tilapia developed as males (n=38). Furthermore, to investigate the effect of simultaneously impairing spermatogenesis and steroid hormone synthesis, the inventors established a tilapia strain with a double homozygous mutation.

[0099] experiment: The inventors crossed the heterozygous F1 male Δ10-cyp19a1a with the heterozygous mutant female (Gopc △8 / + ; Smap2 △17 / + ; Tjp1a △7 / + ; Csnk2a2 △22 / + ; Hiat1 △17 / + ) of Example 7. When inhibited in the Cyp17 null background (results of Example 7), only the SMS gene that causes male infertility was selected. The offspring were genotyped, and at least 10 heterozygotes were raised to adulthood, sex-determined, and interbred. As described in Example 7, the reproductive ability of the resulting offspring was analyzed. At least 5 different double KO males were generated. Without being bound by theory, the inventors expect that the double KO cyp19- / -; SMS- / - fish will develop as infertile males, and 62% of the offspring population will be male and 25% of them will be infertile.

Table 3

[0100] Example 9 - Evaluation of two genes targeting male differentiation and two other genes controlling oogenesis to generate a population of all infertile males The transcriptional inhibitor gonadal somatic cell-derived factor (Gsdf) is a TGF-β superfamily member that is expressed only in the gonads of fish, mainly in Sertoli cells. Similarly, this transcription factor Dmrt1 is preferentially expressed in the epithelial cells of the testis and in pre-Sertoli and Sertoli cells. Both genes are required for normal testis development ([23, 24]).

[0101] To produce an all-female tilapia population, the inventors generated null mutations in either the Dmrt1 or Gsdf gene (male gene or MA) (Figures 19 and 20). The inventors found that 19 out of 20 Gsdf mutant albino tilapia developed as females (Table 3). In contrast, F0 mutants exhibiting mosaic pigment deficiency had a normal sex ratio. Assuming a positive correlation in mutagenicity frequencies between the co-targeted tyrosinase and Gsdf genes, the results of the present invention suggest that a high mutation rate in Gsdf causes XY males to undergo sex reversal to female. Surprisingly, the inventors did not observe a female bias in the selected F0 Dmrt1 mutants (Table 3).

[0102] To manipulate female sterilization, the inventors targeted genes involved in follicular maturation. They identified two genes in the molecular pathway that controls follicular formation: 1) FSHR, which functions upstream of ovarian estrogen synthesis, and 2) vitelogenin (Vtgs), which functions downstream of ovarian estrogen synthesis. Viterogenin is preferentially produced by the liver, and the follicle-stimulating hormone (FSH) receptor FSHR is expressed in the theca cells surrounding developing oocytes. To verify the necessity of FSHR and Vtgs in normal ovarian development (follicular formation-specific genes or FLS), the inventors produced loss-of-function mutations in these genes in each F0 lineage (Figures 22-24).

[0103] The inventors found that FSHR is essential for follicular formation, and that inhibiting the FSHR gene resulted in complete failure of follicular activation and female sterilization (Figure 26 and Table 3). As previously described for zebrafish

[29] , in tilapia, genetic masculinization from female to male did not occur after FSHR mutation. However, the inventors found that F0 FSHR mutant females had significantly smaller urogenital projections compared to control females. This observation appears to reflect reduced estrogen levels in FSHR mutants, which is consistent with the role of FSHR in locally increasing aromatase expression and estrogen production. The inventors did not find any significant reproductive phenotypes in F0 FSHR mutant males.

[0104] Nile tilapia possesses three Vtg genes

[25] , two of which are complete Vtgs types (VtgAa and VtgAb) and one of the incomplete C-type bony fish vitellogenin types, and is deficient in three protein domains (VtgC). Since VtgAa and VtgAb are expressed at higher levels than VtgC and are presumed to be important for early embryonic development, we targeted these two genes both together and separately (Figures 22, 23 and Table 3). Consistent with function in oocyte mutations and nutritional maintenance for embryogenesis, we found that of the four females examined, three F0 females with mutations in VtgAa repeatedly failed to produce viable offspring (Figure 24). The inventors also found that one F0 female out of five, which had a mutation in VtgAb, produced embryonic offspring that died before hatching (data not shown).

[0105] For the complete characterization of the phenotype, it is important to generate the same mutation in all cells of the animal. Therefore, the inventors established and characterized four tilapia strains with insufficient masculinization and yolk formation.

[0106] At 6 months of age, double heterozygous mutants (e.g., Dmrt1) possess the same gene-specific indel at each gene locus (Table 3). △7 / + - FSHR △5 / + To identify the mosaic F0 XX MA m 1-n Female (e.g., Dmrt1 m) 1-n or Gsdf m 1-n ) Mosaic F0 FLS m 1-n The male and its genotyped F1 offspring were crossbred.

[0107] experiment: We are currently raising at least 10 double heterozygous individuals (for each of the four gene combinations) to adulthood. This WT allele should ensure that these F1 fish develop as both males and females. These double heterozygous mutants will then be incrossed, and their offspring will be genetically determined by QPCR thawing analysis at 1 month of age. For each of the 9 individuals (see Figure 25) that represent potential genotypes, we are currently raising at least 30 fish to adulthood, and we plan to analyze their reproductive capacity.

[0108] Figure 25 shows nine genotypes and four corresponding phenotypes, which the inventors predict will result in: 1) ~56% fertile hermaphrodites, 2) ~19% fertile females and infertile individuals, 3) ~19% all fertile males, and 4) ~6% all infertile females. By examining each trait individually, the inventors predict that 62% of the offspring will be female, and of these, 25% will be infertile.

[0109] Phenotypic investigations of the present invention in F0 mutant lines (Table 3) are largely consistent with the initial hypothesis of the present invention, and the inventors fully anticipate the support for the genotype-phenotype relationship in the next generation. The inventors found that inactivation of FSHR and Vtgs causes female infertility, while a deficiency of Gsdf causes feminization. This result strongly suggests that dual FSHR-Gsdf KOs develop into a population of monosexually infertile females characterized by atrophied ovaries with follicles arrested at the pre-vitellation stage. The lack of sex-differentiated phenotypes in F0 Dmrt1 mutants appears to reflect incomplete editing, local mosaicism, and correction by non-mutant genes. However, without being constrained by theory, the inventors believe that all dual FSHR-Dmrt1 KOs that inherit mutations from the reproductive system will become a population of female infertility. In the F0 mutagenesis screen of the present invention, the inventors observed that blocking of essential yolk protein precursors (as seen in Vtg KO) impairs oocyte quality and reduces embryonic development and viability. Therefore, the inventors predicted that double KO Gsdf-Vtgs and Dmrt1-Vtgs would result in a population of monosexually infertile females.

[0110] Example 10 - Propagation of all-male and all-female sterile lines by reproductive system transplantation into sterile adult surrogates Examples 8 and 9 described above demonstrate a method for producing parthenogenetically infertile fish by breeding double hemizygous mutants and individually selecting subpopulations of double KO offspring. However, this method may not be sufficiently efficient and may be too expensive for use in industrial settings. Intracytoplasmic sperm injection in assisted reproductive technology is a solution for breeding male breeding stock with spermatogenesis defects. However, this method is also not scalable for mass production of commercial breeding stock (as it requires a one-at-a-time approach). The key to large-scale production is to generate male and female breeding stock that produce only mutant gametes, since there is no need to select for the identification of double KO offspring. Importantly, these mutant gametes must be functional so that the natural mating of these breeding stocks can be used to produce viable individuals of parthenogenetically infertile offspring. This is only possible if the sex ratio and gamete function are restored in the breeding stock. The inventors hypothesized that this could be achieved by transplanting germline stem cells from double KO mutant fish into germ cell-less transplants that do not have mutations in the same gene. These transplanted parent plants possess normal somatic cells but are mutant germlines (see Figures 27–32). These chimeric transplants are assumed to have functional MA or FEM somatic cells (Figure 34 panels C and D) or functional SMS or FLS somatic cells that ensure a normal sex ratio in order to restore spermatogenesis (Figure 28) or oogenesis (Figures 29 and 30), and that the mutated genes are non-functional in the germ cells.

[0111] Since spermatogenesis defects result from defects in the germ cell or somatic cell environment, the inventors analyzed SMS gene expression to preferentially identify genes that are not expressed within germ cells (Figure 16). In infertility testing, the SMS gene expression test of the present invention refers to the role of gonadal somatic cells in supporting germ cell development. For example, the inventors found that while Hiat1 and Gopc expression levels were only slightly lower compared to fertile testes, Tjp1a was strongly expressed in infertile testes at higher levels than in wild-type testes (Figure 16).

[0112] These results indicate that mutants of these genes construct the testicular microenvironment, where spermatogenesis is impaired due to Sertoli and / or Leydig-specific defects (Figure 28). Therefore, we anticipate that transplantation of spermatogonial stem cells from male knockout infertile donors into a tolerant wild-type testicular environment will restore sperm function and reproductive capacity (Figure 28).

[0113] Similarly, FSHR and Vtgs are expressed only in somatic cells (theca cells and hepatocytes, respectively). Thus, oocytes with null alleles of these genes should retain intrinsic capacity for proliferation and differentiation, ensuring that transplantation into a WT / acceptable recipient allows oogonial stem cells from a sterile female mutant donor to rearrange the ovary and differentiate into functional oocytes (Figures 29 and 30). For this reason, we believe that a male or female recipient can produce gametes with the donor genotype.

[0114] Example 11 - Elavl2 KO implants are capable of producing functional gametes. To confirm that infertile Elavl2 KO transplants can produce functional gametes from donor-derived germ cells, the inventors collected spermatogonial stem cells from albino (tyr- / -) male tilapia with mutations (in-frame and out-frame) in the reference gene (Figure 33 Panel A). The inventors transplanted testicular cell suspensions from both mutant lines into transplant embryo offspring with drastically reduced germ cells from Elavl2 - / + and tyr+ / + parents. To select homozygous Elavl2- / - mutants, the inventors genotyped the transplanted fish and raised them to adulthood. At 5 months of age, when crossbred with albino males and females, 31-50% of the transplanted Elavl2- / - males and 40% of the transplanted Elavl2- / - females at 6 months of age produced only albino offspring. The Elavl2- / - control, which did not receive transplantation, was infertile. Therefore, Elavl2- / - transplantees are capable of producing donor-derived gametes after germline stem cell transplantation, demonstrating the feasibility of creating tilapia that produce only donor-derived gametes. To analyze gametes containing the Tyr allele, albinism was used, and a high-throughput assay that allows for easy quantification of the effectiveness of germline transplantation of the mutant allele was performed. However, these experiments do not indicate successful reproduction of the null mutation. To achieve this goal, the inventors extracted and analyzed sperm DNA from one infertile transplantee by PCR fragment sizing analysis. The amplified products were classified using capillary electrophoresis (Figure 33, Panel B). The results revealed that the transplanted fish produced only sperm containing intra- and extra-frame (3 nt and 4 nt) deletion fragments derived from the donor, suggesting that this null allele (4 nt deletion) can form gonads and proliferate as efficiently as the positive control mutation (Figure 33, Panel B).

[0115] experiment:Spermatogonial stem cells and oogonial stem cells (SSCs, OSCs) are isolated from tilapia strains where the juvenile fish are all male and all female, respectively (developed according to Examples 7, 8, and 9). After collection, these germline stem cells are transplanted into Elavl2 KO transplant juvenile fish as described above. Without being constrained by theory, the inventors anticipate the production of functional sperm and oocytes with this donor genotype. In vitro fertilization assays are performed to evaluate the function of the donor-derived gametes produced after transplantation. Furthermore, the inventors anticipate that albino offspring result from the mating of naturally pigment-deficient transplants with albino strains that possess albino donor gametes. The inventors determine the genotypes of 10 offspring based on mutations in spermatogenesis and yolk formation-specific genes derived from the donor.

[0116] As shown in Figure 34 Panel B, a surrogate mother is mated with a double KO male whose sex has been reversed by treatment with an aromatase inhibitor, producing all female infertile offspring. Alternatively, a surrogate father is mated with a double KO female mutant whose sex has been reversed after estrogen treatment, producing an all male infertile population (Figure 34 Panel A). Sex reversal of double KOs using estrogen (as shown in Figure 34 Panel A) or androgen inhibitors (as shown in Figure 34 Panel B) can be replaced by germination transplantation methods, as they produce female breeding stock (Figure 34 Panel C) or male breeding stock (Figure 34 Panel D).

[0117] Example 12 - Aquarium Growth Test Energy directed towards the gonads impairs body growth, resulting in a direct trade-off between growth and reproduction. Due to a physiological process that requires rapid yolk formation

[26] and a high metabolic rate, Nile tilapia mature early and are capable of breeding throughout the year. Furthermore, tilapia species can suppress growth to maintain fertility

[27] , and in other fish species, puberty onset can have a significant impact on important production parameters in aquaculture, such as appetite, growth rate, feed conversion efficiency, meat traits, appearance, health, welfare, and survival rate. Therefore, delaying or blocking sexual maturation could offer significant benefits to the commercial aquaculture industry. As part of efforts to increase the number of sterile, unisexual populations, we have targeted genes in which mutations block or delay puberty onset. However, for these effects, targeted genes may have multifaceted effects that are detrimental to the strain, acting through unknown hormones, physiological or behavioral changes.

[0118] experiment To generate groups for growth capacity testing, produce single pairs of embryos (at least three separate crosses) from each target strain. Rear the treated and control embryos separately according to the established incubation procedure. At the feeding stage, sex change half of the control animals using an appropriate exogenous hormone protocol (e.g., administer methyltestosterone or DES). When the fish in the (treated and control) groups reach an average weight of 60g, attach PIT tags and divide into six 1000L tanks (three control tanks and three treated tanks, 50 fish / tank). Feed all fish satiety three times a day.

[0119] Each fish is individually weighed and measured in length every four weeks until it reaches a commercially viable size (680g Sdv:77g, 8 months old). At the end of the experiment, these fish are euthanized and sex is determined by the structure of the urogenital opening. The inventors individually record the weight of the dissected gonads and carcasses to calculate the gonadal weight index (GSI) and carcass index (n=60 per group). The specific growth rate (G) is calculated according to the formula of Houde and Scheckter

[28] .

[0120] Without being constrained by theory, the inventors believe that almost all double KO fish produced in Examples 7, 8, and 9 will develop as unisexual and become sterile without other biological processes being impaired. Therefore, the selected mutations should not negatively affect the overall fish capacity. In contrast, the inventors expect an increase in growth rate and feed conversion ratio related to gonad weight. The mutant lines should be sexually delayed (male sterile) or immature (females stopped at pre-vitellation eggs). If the inventors were to sterilize only a portion of the unisexual population, the inventors expect that the increase in productivity in the tilapia, as a result of reduced energy consumption, would be proportional to the proportion of sterile fish in the population. In all cases, the inventors expect that sterile fish and fish with atrophied gonads will outperform fully fertile partners (e.g., unisexual populations derived from exogenous hormone treatment) in terms of growth characteristics.

[0121] reference 1. Dunham, R., Aquaculture and Fisheries Biotechnology, CABI Publishing. 2004. 2. Pruginin, Y., et al., All-male broods of Tilapia nilotica× T. aurea hybrids. Aquaculture, 1975. 6(1): p. 11-21. 3. Wolters, WR and R. DeMay, Production characteristics of striped bass× white bass and striped bass× yellow bass hybrids. Journal of the World Aquaculture Society, 1996. 27(2): p. 202-207. 4. McMaster, M., et al., Milt characteristics, reproductive performance, and larval survival and development of white sucker exposed to bleached kraft mill effluent. Ecotoxicology and environmental safety, 1992. 23(1): p. 103-117. 5. Lacerda, S.M., et al., A new and fast technique to generate offspring after germ cells transplantation in adult fish: the Nile tilapia (Oreochromis niloticus) model. PLoS One, 2010. 5(5): p. e10740. 6. Lauth, X. and J.T. Buchanan, Maternally induced sterility in animals. 2015, Google Patents. 7. Koga, A., et al., Insertion of a novel transposable element in the tyrosinase gene is responsible for an albino mutation in the medaka fish, Oryzias latipes. Molecular and General Genetics MGG, 1995. 249(4): p. 400-405. 8. Krauss, J., et al., transparent, a gene affecting stripe formation in Zebrafish, encodes the mitochondrial protein Mpv17 that is required for iridophore survival. Biology open, 2013. 2(7): p. 703-710. 9. Weidinger, G., et al., dead end, a Novel Vertebrate Germ Plasm Component, Is Required for Zebrafish Primordial Germ Cell Migration and Survival. Current Biology, 2003. 13(16): p. 1429-1434. 10. Peterson, R.T. and P.J. Schlueter, Germ cell ablation compounds and uses thereof. 2017, Google Patents. 11. Benfey, T.J. and A.M. Sutterlin, Growth and gonadal development in triploid landlocked Atlantic salmon (Salmo salar). Canadian Journal of Fisheries and Aquatic Sciences, 1984. 41(9): p. 1387-1392. 12. Felip, A., et al., Induction of triploidy and gynogenesis in teleost fish with emphasis on marine species. Genetica, 2001. 111(1-3): p. 175-195. 13. Zhou, L.-Y., et al., A novel type of P450c17 lacking the lyase activity is responsible for C21-steroid biosynthesis in the fish ovary and head kidney. Endocrinology, 2007. 148(9): p. 4282-4291. 14. Carlisle, S., et al., Carneiro, L. & Grober, MS (2000). Effects of 11-ketotestosterone on genital papilla morphology in the sex changing fish Lythrypnus dalli. Journal of Fish Biology 57, 445-456. The resolution of Fig. 2 when originally printed was unsatisfactory. The correct. Journal of Fish Biology, 2001. 58: p. 299. 15. Xu, X., et al., Globozoospermia in mice lacking the casein kinase II α′ catalytic subunit. Nature genetics, 1999. 23(1): p. 118. 16. Yao, R., et al., Lack of acrosome formation in mice lacking a Golgi protein, GOPC. Proceedings of the National Academy of Sciences, 2002. 99(17): p. 11211-11216. 17. Suzuki-Toyota, F., et al., Factors maintaining normal sperm tail structure during epididymal maturation studied in Gopc- / - mice. Biology of reproduction, 2007. 77(): p. 71-82. 18. Doran, J., et al., Mfsd14a (Hiat1) gene disruption causes globozoospermia and infertility in male mice. Reproduction, 2016. 152(1): p. 91-99. 19. Rocha, D. and N. Affara, The genetic basis of impaired spermatogenesis and male infertility. Current Obstetrics & Gynaecology, 2000. 10(3): p. 139-145. 20. Truong, B., et al., Searching for candidate genes for male infertility. Asian journal of andrology, 2003. 5(2): p. 137-147. 21. Funaki, T., et al., The Arf GAP SMAP2 is necessary for organized vesicle budding from the trans-Golgi network and subsequent acrosome formation in spermiogenesis. Molecular biology of the cell, 2013. 24(17): p. 2633-2644. 22. Oka, K., et al., Genotyping of 38 insertion / deletion polymorphisms for human identification using universal fluorescent PCR. Molecular and cellular probes, 2014. 28(1): p. 13-18. 23. Jiang, D.N., et al., gsdf is a downstream gene of dmrt1 that functions in the male sex determination pathway of the Nile tilapia. Molecular reproduction and development, 2016. 83(6): p. 497-508. 24. Li, M., et al., Efficient and heritable gene targeting in tilapia by CRISPR / Cas9. Genetics, 2014. 197(2): p. 591-599. 25. Davis, L.K., et al., Gender-specific expression of multiple estrogen receptors, growth hormone receptors, insulin-like growth factors and vitellogenins, and effects of 17β-estradiol in the male tilapia (Oreochromis mossambicus). General and comparative endocrinology, 2008. 156(3): p. 544-551. 26. Naylor, R.L., et al., Effect of aquaculture on world fish supplies. Nature, 2000. 405(6790): p. 1017. 27. Coward, K. and NR Bromage, Spawning frequency, fecundity, egg size and ovarian histology in groups of Tilapia zillii maintained upon two distinct food ration sizes from first-feeding to sexual maturity. Aquatic Living Resources, 1999. 12(1): p. 11-22. 28. Houde, ED, Growth rates, rations and cohort consumption of marine fish larvae in relation to prey concentrations. Rapp. P.-V. Reun. Cons. Int. Explor. Mer, 1981. 178: p. 441-453. 29. Zhang, Z., et al., Disruption of zebrafish follicle-stimulating hormone receptor (fshr) but not luteinizing hormone receptor (lhcgr) gene by TALEN leads to failed follicle activation in females followed by sexual reversal to males. Endocrinology, 2015. 156(10): p. 3747-3762.

[0122] List of sequences

[0123] SEQ ID NO 1 Length: 38 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (FAM) Array: 1 TGTAAAACGACGGCCAGTttgaagttgctacataaaag

[0124] SEQ ID NO 2 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 2 TGGTTGATGACAATCACACTGT

[0125] SEQ ID NO 3 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( NED ) Array: 3 TAGGAGTGCAGCAAGCATtgttctacatcatcacccttctc

[0126] SEQ ID NO 4 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 4 AGCAGACAGACGAGCAGTATCAG

[0127] SEQ ID NO 5 Length: 41 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (FAM) Array: 5 TGTAAAACGACGGCCAGT TGATGGAGAGCTTCATCTACGAA

[0128] SEQ ID NO 6 Length: 20 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 6 GTTCCAGGTTAAATTGATTG

[0129] SEQ ID NO 7 Length: 41 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 7 TAGGAGTGCAGCAAGCAT gcgtgatttgctgacctttttac

[0130] SEQ ID NO 8 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 8 acacttacCCTGAGAATCTGG

[0131] SEQ ID NO 9 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 9 TGTAAAACGACGGCCAGTGAAAAAGGATGgtgagggatgac

[0132] SEQ ID NO 10 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 10 GAGTGTGTCTACCACACGGAAAA

[0133] SEQ ID NO 11 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 11 TGTAAAACGACGGCCAGTgtatttagaaggcggtgaaggtc

[0134] SEQ ID NO 12 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 12 CAGTTTGGCACATGAGCATCGTA

[0135] SEQ ID NO 13 Length: 37 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 13 TAGGAGTGCAGCAAGCAT ATGCTCATGTGCCAAACTG

[0136] SEQ ID NO 14 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 14 cCTTCAGGATTTTCACCACCACT

[0137] SEQ ID NO 15 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 15 TGTAAAACGACGGCCAGTtactgacacatccagcagcgtct

[0138] SEQ ID NO 16 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 16 cagcactgagccgtcagtattct

[0139] SEQ ID NO 17 Length: 37 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 17 TAGGAGTGCAGCAAGCAT TGGAGCCTACCTGTCTGAG

[0140] SEQ ID NO 18 Length: 20 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 18 tactcacAGCGAAGGGGTCT

[0141] SEQ ID NO 19 Length: 38 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 19 TAGGAGTGCAGCAAGCAT gctcctctgcgaagactctc

[0142] SEQ ID NO 20 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 20 aagacctccgacCTGGACTTGCT

[0143] SEQ ID NO 21 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 21 TGTAAAACGACGGCCAGT AGAGGAGGGCACAGTCAAGAAAC

[0144] SEQ ID NO 22 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 22 TTGGATATCCCATTTGGTTCAT

[0145] SEQ ID NO 23 Length: 40 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 23 TAGGAGTGCAGCAAGCAT tttaacggtgttggcagagatt

[0146] SEQ ID NO 24 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 24 AGATCCACATCCACGAAAGCCT

[0147] SEQ ID NO 25 Length: 37 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 25 TGTAAAACGACGGCCAGT tgcccctttaaaccaccta

[0148] SEQ ID NO 26 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 26 CTCAGCTTGGCCTTGCTTGACAT

[0149] SEQ ID NO 27 Length: 39 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 27 TAGGAGTGCAGCAAGCAT ttgccaggacccATGAGCCAG

[0150] SEQ ID NO 28 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 28 AGACACGTATCCGTGATTTCTAC

[0151] SEQ ID NO 29 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 29 TGTAAAACGACGGCCAGT ctcttcatcctctgtgtctcatc

[0152] SEQ ID NO 30 Length: 22 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Primer Array: 30 GGGTTTCCAGCAGGAGGTCAGA

[0153] SEQ ID NO 31 Length: 39 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Forward-tailed primer (NED) Array: 31 TAGGAGTGCAGCAAGCAT ttatgttcagGTGCCAAGGTG

[0154] SEQ ID NO 32 Length: 23 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Primer Array: 32 TGGCTGTGTGAGAAACGATGCTG

[0155] SEQ ID NO 33 Length: 35 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Forward-tailed primer ( FAM ) Array: 33 TGTAAAACGACGGCCAGT agATCTGGGCTGGGACA

[0156] SEQ ID NO 34 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 34 tgttaactatacCTGTGTGTTGG

[0157] SEQ ID NO 35 Length: 38 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 35 TAGGAGTGCAGCAAGCAT ttttctccgcttgcttctgc

[0158] SEQ ID NO 36 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 36 AAAGAGCTGAATAGGAGGAAGTT

[0159] SEQ ID NO 37 Length: 39 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 37 TGTAAAACGACGGCCAGT CATCTTGGCGTTCTTCTGTGT

[0160] SEQ ID NO 38 Length: 21 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 38 CTTGAGGGCAGCTGAGATGGC

[0161] SEQ ID NO 39 Length: 40 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Forward tailed primer (NED) Sequence: 39 TAGGAGTGCAGCAAGCAT GCAATCCTTGATGCTCCTTGAC

[0162] SEQ ID NO 40 Length: 22 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Primer Sequence: 40 CTGAGACTCTATGTCGTTGATA

[0163] SEQ ID NO 41 Length: 41 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Forward tailed primer ( FAM ) Sequence: 41 TGTAAAACGACGGCCAGT AGAAGATCATCAAACACATCACG

[0164] SEQ ID NO 42 Length: 23 Type: DNA Organism: Artificial sequence Other information: Details of artificial sequence: Primer Sequence: 42 GACTTGTTGAGCAGTTGCATCAA

[0165] SEQ ID NO 43 Length: 38 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: Forward-tailed primer (NED) Array: 43 TAGGAGTGCAGCAAGCAT ttttgtgatctagTCTGGAG

[0166] SEQ ID NO 44 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 44 gctcttacAGCTTCACAATCAT

[0167] SEQ ID NO 45 Length: 41 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: Forward-tailed primer ( FAM ) Array: 45 TGTAAAACGACGGCCAGT AGAAG ATCATCAAACACATCACG

[0168] SEQ ID NO 46 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 46 GACTTGTTGAGCAGTTGCATCAA

[0169] SEQ ID NO 47 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 47 GAACCAAACCCCTCTGTCACTG

[0170] SEQ ID NO 48 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 48 GTAATTCACTCCGCAGGCTCAG

[0171] SEQ ID NO 49 Length: 17 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 49 ggcgATGAATCCTGTAG

[0172] SEQ ID NO 50 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 50 ATGGCATTTGAGGTCACAGAGA

[0173] SEQ ID NO 51 Length: 19 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 51 GTTCAAGAAGGGAGAGAGT

[0174] SEQ ID NO 52 Length: 18 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 52 AAAAATTCCCACATCGTT

[0175] SEQ ID NO 53 Length: 20 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 53 tgctttggcttcagTGTATC

[0176] SEQ ID NO 54 Length: 19 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 54 AATGCGTTCGAATGTAGAA

[0177] Saturation ID NO 55 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 55 CATCTGCTTCATCCTGGTGGCTG

[0178] SEQ ID NO 56 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 56 AATTTGGGCATCTTCATCTGTAT

[0179] SEQ ID NO 57 Length: 22 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 57 GACAGACTTGACCTTGGAGATG

[0180] SEQ ID NO 58 Length: 21 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 58 ATGTCTGCTTCGACTGGATGC

[0181] SEQ ID NO 59 Length: 23 Type: DNA Biology: Artificial sequences Other information: Details of artificial sequences: primers Array: 59 GCCATCGAAACATGGACATACTG

[0182] SEQ ID NOs 60 and 62 (wild-type Cyp17a1) Length: 1563bp and 521aa Type: cDNA (SEQ ID NO: 60) and protein (SEQ ID NO: 62) Organism: Nile tilapia [ka] [ka]

[0183] SEQ ID NOs 61 and 63 (Cyp17a1 mutant allele - 16nt deletion) Length: 1563 bp and 44 aa Type: cDNA (SEQ ID NO: 61) and protein (SEQ ID NO: 63) Organism: Nile tilapia [ka]

[0184] SEQ ID NOs 65 and 68 (wild-type Cyp19a1a) Length: 1707bp and 511aa Type: cDNA (SEQ ID NO: 65) and protein (SEQ ID NO: 68) Organism: Nile tilapia [ka] [ka]

[0185] SEQ ID NOs 66 and 69 (Cyp19a1a mutant allele - 7nt deletion) Length: 1707bp and 12aa Type: cDNA (SEQ ID NO: 66) and protein (SEQ ID NO: 69) Organism: Nile tilapia [ka]

[0186] SEQ ID NOs 67 and 70 (Cyp19a1a mutant allele - 10nt deletion) Length: 1707bp and 11aa Type: cDNA (SEQ ID NO: 67) and protein (SEQ ID NO: 70) Organism: Nile tilapia [ka]

[0187] SEQ ID NOs 71 and 73 (wild-type Tjp1a) Length: 6674bp and 1652aa Type: cDNA (SEQ ID NO: 71) and protein (SEQ ID NO: 73) Organism: Nile tilapia [ka] [ka] [ka] [ka] [ka] [ka]

[0188] SEQ ID NOs 72 and 74 (Tjp1a mutant allele - 7nt deletion) Length: 6674bp and 439aa Type: cDNA (SEQ ID NO: 72) and protein (SEQ ID NO: 74) Organism: Nile tilapia [ka] [ka]

[0189] SEQ ID NOs 75 and 77 (wild-type Hiat1a) Length: 5281bp and 491aa Type: cDNA (SEQ ID NO: 75) and protein (SEQ ID NO: 77) Organism: Nile tilapia [ka] [ka] [ka] [ka]

[0190] SEQ ID NOs 76 and 78 (Hiat1a mutant allele - 17nt deletion) Length: 5281bp and 234aa Type: cDNA (SEQ ID NO: 76) and protein (SEQ ID NO: 78) Organism: Nile tilapia [ka]

[0191] SEQ ID NOs 79 and 81 (wild-type Smap2) Length: 4207bp and 429aa Type: cDNA (SEQ ID NO: 79) and protein (SEQ ID NO: 81) Organism: Nile tilapia [ka] [ka] [ka]

[0192] SEQ ID NOs 80 and 82 (Smap2 mutant allele - 17nt deletion) Length: 4207bp and 118aa Type: cDNA (SEQ ID NO: 80) and protein (SEQ ID NO: 82) Organism: Nile tilapia [ka]

[0193] SEQ ID NOs 83 and 85 (wild-type Csnk2a2) Length: 1053 bp and 350 aa Type: cDNA (SEQ ID NO: 83) and protein (SEQ ID NO: 85) Organism: Nile tilapia [ka]

[0194] SEQ ID NOs 84 and 86 (Csnk2a2 mutant allele - 22nt deletion) Length: 1053bp and 31aa Type: cDNA (SEQ ID NO: 84) and protein (SEQ ID NO: 86) Organism: Nile tilapia [ka]

[0195] SEQ ID NOs 87 and 89 (Wild-type Gope) Length: 1335bp and 444aa Type: cDNA (SEQ ID NO: 87) and protein (SEQ ID NO: 89) Organism: Nile tilapia [ka] [ka]

[0196] SEQ ID NOs 88 and 90 (Gope variant allele - 8nt deletion) Length: 1335bp and 30aa Type: cDNA (SEQ ID NO: 88) and protein (SEQ ID NO: 90) Organism: Nile tilapia [ka]

[0197] SEQ ID NOs 91 and 94 (wild-type DMRT-1) Length: 882bp and 293aa Type: cDNA (SEQ ID NO: 91) and protein (SEQ ID NO: 94) Organism: Nile tilapia [ka]

[0198] SEQ ID NOs 92 and 95 (DMRT-1 mutant allele - 7nt deletion) Length: 882bp and 40aa Type: cDNA (SEQ ID NO: 92) and protein (SEQ ID NO: 95) Organism: Nile tilapia [ka]

[0199] SEQ ID NOs 93 and 96 (DMRT-1 mutant allele - 13nt deletion) Length: 882bp and 38aa Type: cDNA (SEQ ID NO: 93) and protein (SEQ ID NO: 96) Organism: Nile tilapia [ka]

[0200] SEQ ID NOs 97 and 100 (Wild-type GSDF) Length: 840bp and 213aa Type: cDNA (SEQ ID NO: 97) and protein (SEQ ID NO: 100) Organism: Nile tilapia [ka]

[0201] SEQ ID NOs 98 and 101 (GSDF variant allele - 5nt deletion) Length: 840bp and 56aa Type: cDNA (SEQ ID NO: 98) and protein (SEQ ID NO: 101) Organism: Nile tilapia [ka]

[0202] SEQ ID NOs 99 and 102 (GSDF variant allele - 22nt deletion) Length: 840bp and 46aa Type: cDNA (SEQ ID NO: 99) and protein (SEQ ID NO: 102) Organism: Nile tilapia [ka]

[0203] SEQ ID NOs 103 and 105 (wild-type FSHR) Length: 5853bp and 689aa Type: cDNA (SEQ ID NO: 103) and protein (SEQ ID NO: 105) Organism: Nile tilapia [ka] [ka] [ka] [ka] [ka]

[0204] SEQ ID NOs 104 and 106 (FSHR mutant allele - 5nt deletion) Length: 5853bp and 264aa Type: cDNA (SEQ ID NO: 104) and protein (SEQ ID NO: 106) Organism: Nile tilapia [ka]

[0205] SEQ ID NOs 107 and 110 (wild-type VtgAa) Length: 4974bp and 1657aa Type: cDNA (SEQ ID NO: 107) and protein (SEQ ID NO: 110) Organism: Nile tilapia [ka] [ka] [ka] [ka] [ka]

[0206] SEQ ID NOs 108 and 111 (VtgAa mutant allele - 5nt deletion) Length: 4974bp and 279aa Type: cDNA (SEQ ID NO: 108) and protein (SEQ ID NO: 111) Organism: Nile tilapia [ka]

[0207] SEQ ID NOs 109 and 112 (VtgAa mutant allele - 25nt deletion) Length: 4974bp and 301aa Type: cDNA (SEQ ID NO: 109) and protein (SEQ ID NO: 112) Organism: Nile tilapia [ka]

[0208] SEQ ID NOs 113 and 115 (wild-type VtgAb) Length: 5339bp and 1747aa Type: cDNA (SEQ ID NO: 113) and protein (SEQ ID NO: 115) Organism: Nile tilapia [ka] [ka] [ka] [ka] [ka]

[0209] SEQ ID NOs 114 and 116 (VtgAb mutant allele - 8nt deletion) Length: 5339bp and 202aa Type: cDNA (SEQ ID NO: 114) and protein (SEQ ID NO: 116) Organism: Nile tilapia [ka]

[0210] SEQ ID NO 117 Length: 18 Type: DNA Biology: Artificial sequences Other information: Artificial sequence details: 5' tailed primer extended sequence (FAM) Array: 1 TGTAAAACGACGGCCAGT

[0211] SEQ ID NO 118 Length: 18 Type: DNA Biology: Artificial sequences Other information: Details of the artificial sequence: 5' tailed primer extended sequence ( NED ) Array: 3 TAGGAGTGCAGCAAGCAT

[0212] In the above explanation, numerous details are presented for the purpose of providing a thorough understanding of the examples. However, it will be obvious to those skilled in the art that these specific details are not mandatory.

[0213] The above embodiments are for illustrative purposes only. Modifications, alterations, and variations of specific embodiments are possible for those skilled in the art. The claims should not be limited to the specific embodiments shown herein, but should be interpreted in accordance with the entire specification.

Claims

1. A method for producing sterile, sex-determined fish, crustaceans, or mollusks, including the following steps: (i) breeding fertile hemizygous mutant female fish, crustaceans, or mollusks having at least the first and second mutations, and (ii) breeding fertile hemizygous mutant male fish, crustaceans, or mollusks having at least the first and second mutations; and Selection of homozygous primitive, sterile, sex-determined fish, crustaceans, or mollusks with homozygous mutations through genotypic selection. The initial mutation inhibits one or more genes that determine sex differentiation, and The second mutation inhibits one or more genes that determine gamete function.

2. A method for producing sterile, sex-determined fish, crustaceans, or mollusks, including the following steps: To produce sterile sex-determined fish, crustaceans, or mollusks, (i) breed fertile homozygous mutant female fish, crustaceans, or mollusks having at least the first and second mutations, and (ii) breed fertile homozygous mutant male fish, crustaceans, or mollusks having at least the first and second mutations, The initial mutation inhibits one or more genes that determine sex differentiation. The second mutation inhibits one or more genes that determine gamete function, and The reproductive capacity of homozygous female fish, crustaceans, or mollusks with reproductive capacity, and homozygous mutant male fish, crustaceans, or mollusks with reproductive capacity, was restored.

3. The method according to claim 2, wherein the restoration of reproductive capacity includes germline stem cell transplantation.

4. The method according to claim 3, wherein the restoration of reproductive capacity further includes a change in sex steroids.

5. The method according to claim 4, wherein the change in sex steroids is a change in estrogen or a change in aromatase inhibitors.

6. The method according to any one of claims 3 to 5, wherein the germline stem cell transplantation includes the following steps: Obtain germline stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least the first and second mutations, or from sterile homozygous female fish, crustaceans, or mollusks having at least the first and second mutations; and These germline stem cells are transplanted into male fish, crustaceans, or mollusks that lack germ cells, or into female fish, crustaceans, or mollusks that lack germ cells.

7. The method according to claim 6, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and the germ cell-less transplanted female fish, crustacean, or mollusk are homozygous for null mutations of dnd, Elavl2, vasa, nanos3, or piwi-like genes.

8. The method according to claim 6, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are produced by a ploidy operation.

9. The method according to claim 6, wherein the germ cell-less male transplant fish, crustacean, or mollusk and the germ cell-less female transplant fish, crustacean, or mollusk are produced by crossbreeding.

10. The method according to claim 6, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are prepared by exposure to high levels of sex hormones.

11. The method according to any one of claims 3 to 5, wherein the germline stem cell transplantation includes the following steps: Obtain spermatogonial stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least the first and second mutations, or obtain oogonial stem cells from sterile homozygous female fish, crustaceans, or mollusks having at least the first and second mutations; and These spermatogonial stem cells are transplanted into the testes of reproductively capable male fish, crustaceans, or mollusks that lack germ cells, or these oogonial stem cells are transplanted into the ovaries of reproductively capable female fish, crustaceans, or mollusks that lack germ cells.

12. The method according to claim 11, wherein the germ-free, fertile male fish, crustacean, or mollusk and the germ-free, fertile female fish, crustacean, or mollusk are homozygous for mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes.

13. The method according to claim 11, wherein the germ cell-free transplanted male fish, crustacean, or mollusk and germ cell-free transplanted female fish, crustacean, or mollusk are produced by polyploidy.

14. The method according to claim 11, wherein the germ cell-less male transplant fish, crustacean, or mollusk and the germ cell-less female transplant fish, crustacean, or mollusk are produced by crossbreeding.

15. The method according to claim 11, wherein the germ-free male fish, crustacean, or mollusk and germ-free female fish, crustacean, or mollusk are created by exposing them to high levels of sex hormones.

16. The method according to any one of claims 1 to 15, wherein the sterile sex-determined fish, crustacean, or mollusk is a sterile male fish, crustacean, or mollusk.

17. The method according to any one of claims 1 to 16, wherein the initial mutation comprises a mutation in one or more genes that regulate the synthesis of androgens and / or estrogens.

18. The method according to claim 17, wherein the initial mutation comprises a mutation in one or more genes that modulate the expression of aromatase Cyp19a1a, Cyp17, or a combination thereof.

19. The method according to claim 18, wherein one or more genes that regulate the expression of aromatase Cyp19a1a are one or more genes selected from the group consisting of cyp19a1a, FoxL2, and their orthologues.

20. The method according to claim 17, wherein one abnormal sound gene that regulates the expression of Cyp17 is cyp17I or its ortholog.

21. The method according to any one of claims 1 to 20, wherein the second mutation comprises a mutation in one or more genes that regulate spermatogenesis.

22. The method according to claim 21, wherein the second mutation includes a mutation in one or more genes that cause megacephalic spermatopathy.

23. The method according to claim 22, wherein a second mutation in one or more genes causing megahead spermatophobia results in a round head, a round nucleus, a truncated mid-section, a partially coiled tail, or a combination thereof.

24. The method according to claim 23, wherein the second mutation comprises a mutation in one or more genes selected from the group consisting of Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, and their orthologues.

25. The method according to any one of claims 1 to 15, wherein the sterile sex-determined sterile fish, crustacean, or mollusk is a sterile female fish, crustacean, or mollusk.

26. The method according to any one of claims 1 to 15 and 25, wherein the initial mutation comprises a mutation in one or more genes that modulate the expression of an aromatase Cyp19a1a inhibitor.

27. The method according to claim 26, wherein one or more genes for preparing the expression of an aromatase Cyp19a1a inhibitor are one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologues.

28. The method according to any one of claims 1 to 15 and 25 to 27, wherein the second mutation comprises one or more genes that modulate oogenesis, follicular formation, or a combination thereof.

29. The method according to claim 28, wherein one or more genes that regulate oogenesis regulate estrogen synthesis.

30. The method according to claim 29, wherein one or more genes that regulate estrogen synthesis are FSHR or an ortholog thereof.

31. The method according to claim 28, wherein one or more genes that regulate follicular formation regulate the expression of vitellogenin.

32. The method according to claim 31, wherein one or more genes that regulate vitelogenin expression are vtgs or their orthologs.

33. The method according to claim 31, wherein one or more genes that regulate vitelogenin expression are mutations in genes encoding or controlling vitelogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein; or orthologs thereof.

34. Methods for producing sterile, sex-determined fish, crustaceans, or mollusks, including the following steps: To produce sterile, sex-determined fish, crustaceans, or mollusks, (i) a fertile female fish, crustacean, or mollusk with a homozygous mutation is (ii) bred with a fish, crustacean, or mollusk with a homozygous mutation. This mutation directly or indirectly inhibits spermatogenesis, and / or directly inhibits yolk formation, The reproductive capacity of these fertile female fish, crustaceans, or mollusks, and fertile male fish, crustaceans, or mollusks, has been restored.

35. The method according to claim 34, wherein the mutation that directly or indirectly inhibits spermatogenesis is a mutation in Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, or an ortholog thereof.

36. The method according to claim 34 or 35, wherein the mutation that directly inhibits yolk formation is a mutation in a gene encoding or controlling vitellogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein; or an ortholog thereof.

37. The method according to claim 34, 35, or 36, wherein the fertile female fish, crustacean, or mollusk and the fertile male fish, crustacean, or mollusk have multiple homozygous mutations that directly or indirectly inhibit spermatogenesis; directly inhibit yolk formation; or a combination of both.

38. The method according to any one of claims 34 to 37, wherein the restoration of reproductive capacity includes the transplantation of germline stem cells.

39. The method according to claim 38, wherein the restoration of reproductive capacity further includes a change in sex steroids.

40. The method according to claim 39, wherein the change in sex steroids is a change in estrogen or a change in aromatase inhibitors.

41. The method according to claims 38-40, wherein the germline stem cell transplantation includes the following steps: To obtain germline stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least homozygous mutations, and from sterile homozygous female fish, crustaceans, or mollusks having at least homozygous mutations; and These germline stem cells are transplanted into male fish, crustaceans, or mollusks that lack germ cells, or into female fish, crustaceans, or mollusks that lack germ cells.

42. The method according to claim 41, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and the germ cell-less transplanted female fish, crustacean, or mollusk are homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes.

43. The method according to claim 41, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are produced by a ploidy operation.

44. The method according to claim 41, wherein the germ cell-less male transplant fish, crustacean, or mollusk and the germ cell-less female transplant fish, crustacean, or mollusk are produced by crossbreeding.

45. The method according to claim 41, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are prepared by exposure to high levels of sex hormones.

46. The method according to any one of claims 34 to 45, wherein the fertile female fish, crustacean, or mollusk and the fertile male fish, crustacean, or mollusk have additional homozygous mutations that determine sex differentiation.

47. The method according to claim 46, wherein the mutation determining sex differentiation modulates the expression of aromatase Cyp19a1a, Cyp17, an inhibitor of aromatase Cyp19a1a, or a combination thereof.

48. The method according to claim 47, wherein the mutation that modulates the expression of Cyp17 is a mutation in cyp17I or its ortholog.

49. The method according to claim 47 or 48, wherein the mutation modulating the aromatase Cyp19a1a inhibitor is a mutation in Gsdf, dmrt1, Amh, Amhr, or its ortholog.

50. The method according to claims 34 to 45, wherein this breeding step includes crossbreeding or hormonal manipulation and breeding methods to determine sexual differentiation.

51. The method according to any one of claims 1 to 50, wherein the fish, crustacean, or mollusk is a fish.

52. A sterile, sex-determined fish, crustacean, or mollusk whose reproductive capacity is restored, in which the first mutation inhibits one or more genes that determine sex differentiation, the second mutation inhibits one or more genes that determine gamete function, and in which the reproductive capacity of the fertile, homozygous mutant fish, crustacean, or mollusk is restored, or a fertile, homozygous mutant fish, crustacean, or mollusk having at least the first and second mutations.

53. The restoration of this reproductive capacity includes the transplantation of germline stem cells in the fertile, homozygous mutant fish, crustacean, or mollusk according to claim 52.

54. The recovery of this reproductive capacity further includes sex steroid changes, wherein the reproductive capacity is homozygous mutation of the fish, crustacean, or mollusk according to claim 53.

55. The fertile, homozygous mutant fish, crustacean, or mollusk according to claim 54, wherein the sex steroid change is an estrogen change or an aromatase inhibitor change.

56. This germline stem cell transplantation is the next step for this fertile, homozygous mutant fish, crustacean, or mollusk: Obtain germline stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least the first and second mutations, or from sterile homozygous female fish, crustaceans, or mollusks having at least the first and second mutations; and Germline stem cells are transplanted into male fish, crustaceans, or mollusks that lack germ cells, or into female fish, crustaceans, or mollusks that lack germ cells.

57. The reproductively capable homozygous mutant fish, crustacean, or mollusk according to claim 56, wherein the germ-less male transplanted fish, crustacean, or mollusk and the germ-less female transplanted fish, crustacean, or mollusk are homozygous for a null mutation in the dnd, Elavl2, vasa, nanos3, or piwi-like gene.

58. The germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are created by polyploidy to produce the fertile, homozygous mutant fish, crustacean, or mollusk according to claim 56.

59. A reproductively capable homozygous mutant fish, crustacean, or mollusk according to claim 56, created by crossbreeding a germ-cell-less male transplant fish, crustacean, or mollusk and a germ-cell-less female transplant fish, crustacean, or mollusk.

60. The reproductively capable homozygous mutant fish, crustacean, or mollusk according to claim 56, which is created by exposing the germ-free male transplanted fish, crustacean, or mollusk and the germ-free female transplanted fish, crustacean, or mollusk to high levels of sex hormones.

61. This germline stem cell transplantation includes the following steps in the fertile homozygous mutant fish, crustacean, or mollusk according to any one of claims 53 to 55: Obtain spermatogonial stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least the first and second mutations, and obtain oogonial stem cells from sterile homozygous female fish, crustaceans, or mollusks having at least the first and second mutations; and These spermatogonial stem cells are transplanted into the testes of reproductively capable male fish, crustaceans, or mollusks that lack germ cells, or these oogonial stem cells are transplanted into the ovaries of reproductively capable female fish, crustaceans, or mollusks that lack germ cells.

62. The fertile homozygous mutant fish, crustacean, or mollusk according to claim 61, wherein the germ-free, fertile male fish, crustacean, or mollusk and the germ-free, fertile female fish, crustacean, or mollusk are homozygous for mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes.

63. The germ cell-less transplanted male fish, crustacean, or mollusk and germ cell-less transplanted female fish, crustacean, or mollusk are created by polyploidy to produce the fertile, homozygous mutant fish, crustacean, or mollusk described in claim 61.

64. A reproductively capable homozygous mutant fish, crustacean, or mollusk according to claim 56, created by crossbreeding a germ-cell-less male transplant fish, crustacean, or mollusk and a germ-cell-less female transplant fish, crustacean, or mollusk.

65. The reproductively capable homozygous mutant fish, crustacean, or mollusk according to claim 61, which is created by exposing the germ-free male transplanted fish, crustacean, or mollusk and the germ-free female transplanted fish, crustacean, or mollusk to high levels of sex hormones.

66. The fertile, homozygous variant of a fish, crustacean, or mollusk according to any one of claims 52 to 65, wherein the sterile, sex-determined sterile fish, crustacean, or mollusk is a sterile male fish, crustacean, or mollusk.

67. A fertile, homozygous mutant fish, crustacean, or mollusk according to any one of claims 52 to 66, wherein the initial mutation comprises a mutation in one or more genes that regulate the synthesis of androgens and / or estrogens.

68. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 67, wherein the initial mutation comprises a mutation in one or more genes that modulate the expression of aromatase Cyp19a1a, Cyp17, or a combination thereof.

69. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 68, wherein one or more genes that regulate the expression of aromatase Cyp19a1a are one or more genes selected from the group consisting of cyp19a1a, FoxL2, and its orthologues.

70. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 68, wherein one or more genes that regulate the expression of Cyp17 are cyp17I or an ortholog thereof.

71. A fish, crustacean, or mollusk having a fertile, homozygous mutation according to any one of claims 52 to 70, wherein the second mutation comprises a mutation in one or more genes that regulate spermatogenesis.

72. The fertile, homozygous mutant fish, crustacean, or mollusk according to claim 71, wherein the second mutation includes a mutation in one or more genes that cause megacephalic spermatopathy.

73. A fertile, homozygous mutant fish, crustacean, or mollusk of claim 72, in which a second mutation in one or more genes causing megacephalic spermatozoa results in sperm having a round head, a round nucleus, a truncated midsection, a partially coiled tail, or a combination thereof.

74. The fertile, homozygous mutant fish, crustacean, or mollusk according to claim 73, wherein the second mutation comprises a mutation in one or more genes selected from the group consisting of Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, and their orthologues.

75. The reproductively capable homozygous mutant fish, crustacean, or mollusk according to any one of claims 52 to 65, wherein the sterile sex-determined sterile fish, crustacean, or mollusk is a sterile female fish, crustacean, or mollusk.

76. A fertile, homozygous mutant fish, crustacean, or mollusk according to any one of claims 52-65 and 75, wherein the initial mutation comprises a mutation in one or more genes that modulate the expression of an aromatase Cyp19a1a inhibitor.

77. The fertile, homozygous mutant fish, crustacean, or mollusk according to claim 76, wherein one or more genes that modulate the expression of the aromatase Cyp19a1a inhibitor are one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologues.

78. A fertile, homozygous mutant fish, crustacean, or mollusk according to any one of claims 52-65 and 75-77, wherein the second mutation comprises a mutation in one or more genes that modulate oogenesis, follicular formation, or combination.

79. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 78, wherein one or more genes that regulate oogenesis regulate estrogen synthesis.

80. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 79, wherein one or more genes that regulate estrogen synthesis are FSHR or its ortholog.

81. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 80, wherein one or more genes that regulate follicular formation regulate the expression of vitellogenin.

82. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 80, wherein one or more genes that regulate vitelogenin expression are vtgs or their orthologs.

83. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 82, wherein one or more genes that regulate vitelogenin expression are mutations in genes encoding or controlling vitelogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein; or an ortholog thereof.

84. A mutation that directly or indirectly inhibits spermatogenesis and / or directly inhibits follicular formation, and a fertile fish, crustacean, or mollusk having a homozygous mutation that restores the fertility of the fertile fish, crustacean, or mollusk, resulting in the production of infertile, sex-determined fish, crustacean, or mollusks.

85. A fertile, homozygous mutant fish, crustacean, or mollusk according to claim 84, wherein the mutation that directly or indirectly inhibits spermatogenesis is a mutation in Gopc, Hiat1, Tjp1a, Smap2, Csnk2a2, or its ortholog.

86. The fertile fish, crustacean, or mollusk according to claim 84 or 85, wherein the mutation that directly inhibits yolk formation is a mutation in a gene encoding or controlling vitellogenin; estrogen receptor 1; cytochrome p450, family 1, subfamily a; zona pellucida glycoprotein; choriogenin H; peroxisome proliferator-activated receptor; steroid-producing acute regulatory protein, or an ortholog thereof.

87. The fertile female fish, crustacean, or mollusk according to claim 84, 85, or 86, having multiple homozygous mutations that directly or indirectly inhibit spermatogenesis; directly inhibit yolk formation; or a combination of both.

88. A fertile fish, crustacean, or mollusk according to any one of claims 84 to 87, wherein the restoration of fertility includes the transplantation of germline stem cells.

89. The reproductive capacity of the fish, crustacean, or mollusk according to claim 88, further comprising a sex steroid alteration.

90. The fertile fish, crustacean, or mollusk according to claim 89, wherein the change in sex steroids is a change in estrogen or a change in aromatase inhibitors.

91. The reproductively capable fish, crustacean, or mollusk according to any one of claims 88 to 90, wherein the reproductive stem cell transplantation includes the following steps: Obtaining germline stem cells from sterile homozygous male fish, crustaceans, or mollusks having at least homozygous mutations, or from sterile homozygous female fish, crustaceans, or mollusks having at least homozygous mutations; and These germline stem cells are transplanted into male fish, crustaceans, or mollusks that lack germ cells, or into female fish, crustaceans, or mollusks that lack germ cells.

92. The reproductive fish, crustacean, or mollusk according to claim 91, wherein the germ-free male transplanted fish, crustacean, or mollusk and the germ-free female transplanted fish, crustacean, or mollusk are homozygous for null mutations in dnd, Elavl2, vasa, nanos3, or piwi-like genes.

93. The reproductively capable fish, crustacean, or mollusk according to claim 91, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and the germ cell-less transplanted female fish, crustacean, or mollusk are created by polyploidy manipulation.

94. The reproductively capable fish, crustacean, or mollusk according to claim 91, which is produced by crossbreeding a germ-free male transplant fish, crustacean, or mollusk and a germ-free female transplant fish, crustacean, or mollusk.

95. The reproductively capable fish, crustacean, or mollusk according to claim 91, wherein the germ cell-less transplanted male fish, crustacean, or mollusk and the germ cell-less transplanted female fish, crustacean, or mollusk are created by exposing them to high levels of sex hormones.

96. The reproductively capable fish, crustacean, or mollusk according to any one of claims 84 to 95, wherein the reproductively capable fish, crustacean, or mollusk has additional homozygous mutations that determine sex differentiation.

97. The fertile fish, crustacean, or mollusk according to claim 96, wherein the mutation determining sex differentiation modulates the expression of aromatase Cyp19a1a, Cyp17, an inhibitor of aromatase Cyp19a1a, or a combination thereof.

98. The fertile fish, crustacean, or mollusk according to claim 97, wherein one or more genes that regulate the expression of aromatase Cyp19a1a are one or more genes selected from the group consisting of cyp19a1a, FoxL2, and their orthologues.

99. The fertile fish, crustacean, or mollusk according to claim 97, wherein one or more genes that modulate the expression of the aromatase Cyp19a1a inhibitor are one or more genes selected from the group consisting of Gsdf, dmrt1, Amh, Amhr, and their orthologues.

100. A fertile fish, crustacean, or mollusk according to claims 84 to 95, wherein the production of a sterile, sex-determined fish, crustacean, or mollusk includes a breeding step comprising hybridization or hormonal manipulation and a breeding method for determining sex differentiation.

101. The reproductively capable fish, crustacean, or mollusk according to any one of claims 52 to 100, wherein the reproductively capable fish, crustacean, or mollusk is a fish.

102. The following steps involve producing fertile, homozygous mutant fish, crustaceans, or mollusks that produce sterile, sex-determined fish, crustaceans, or mollusks: (i) breeding fertile hemizygous female fish, crustaceans, or mollusks having at least the first and second mutations; (ii) breeding fertile hemizygous male fish, crustaceans, or mollusks having at least the first and second mutations; Genotype selection is used to select homozygous atoms; and To restore the reproductive capacity of this homozygous primitive, The initial mutation inhibits one or more genes that determine sex differentiation, and The second mutation inhibits one or more genes that determine gamete function.

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