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Yavari, M. Environmental Contaminants-Related Fertility Threat in Male Fishes. Encyclopedia. Available online: (accessed on 11 December 2023).
Yavari M. Environmental Contaminants-Related Fertility Threat in Male Fishes. Encyclopedia. Available at: Accessed December 11, 2023.
Yavari, Mina. "Environmental Contaminants-Related Fertility Threat in Male Fishes" Encyclopedia, (accessed December 11, 2023).
Yavari, M.(2021, October 26). Environmental Contaminants-Related Fertility Threat in Male Fishes. In Encyclopedia.
Yavari, Mina. "Environmental Contaminants-Related Fertility Threat in Male Fishes." Encyclopedia. Web. 26 October, 2021.
Environmental Contaminants-Related Fertility Threat in Male Fishes

Public concern regarding environmental contaminants (ECs)-related reproductive disorders has increased due to increasing global rates of infertility. All kinds of ECs are on rise rapidly in developing and industrializing low- and middle-income countries. The aquatic environments throughout the world are repositories for enormous amounts of ECs. As the biology of the reproductive system is highly conserved in vertebrates, wildlife or laboratory studies on fish provide significant information to establish a detailed risk assessment, and to identify novel or more sensitive endpoints for ECs-related reproductive disorders. The adverse effects of ECs on endocrine regulation of reproduction in male fishes have been extensively studied and reviewed; however, our knowledge on the effects and mechanisms of action of ECs on determinants of male fertility is limited.

fertility endpoints industrial pollutants pesticides pharmaceuticals sperm quality

1. Introduction

Global rates of environmental contaminants (ECs)-related reproductive disorders have been increasing over the past 50 years. In human beings, the incidences of testicular dysgenesis syndrome, including hypospadias (urethra opens on the underside of the penis instead of the tip), cryptorchidism (one or both testes not descended into the scrotum), testicular cancer, low semen quality, and infertile men, show global increases associated with ECs [1][2][3][4][5][6][7][8][9]. Landrigan et al. [10] reported that all kinds of ECs are all on the rise rapidly in developing and industrializing low-income and middle-income countries. The public concern regarding ECs-related reproductive disorders was originally linked to observations of reduced fertility, birth defects, and sexual developmental disorders in wildlife [11]. For over 30 years, the World Health Organization (WHO), National Institute of Health (NIH, USA), European Food Safety Authority, and the other organizations composed of working groups of experts in endocrinology, risk assessment, and toxicology, have conducted studies to examine the adverse effects of ECs on reproduction in humans and wildlife. These studies have shown that there are about 800 natural and man-made chemicals known or suspected to interfere with physiological and endocrinological regulation of reproduction [12]. However, our knowledge on the ECs-related hormonal dysfunctions that cause diminished fertility is limited to a small fraction of these chemicals. To reduce ECs-related fertility threat in males, it is critical to identify the contaminants that interfere with determinants of fertility, including sperm production, morphology, genome, and motility, and to characterize their modes of action on reproductive endocrine system. In this regard, interdisciplinary efforts that combine knowledge from wildlife, experimental animals, and human infertility clinics are needed to provide a more holistic approach for ECs-related reproductive disorders and fertility threat.
The aquatic environment is at greatest risk from pollutants since all chemicals will eventually find themselves in the rivers, lakes, and oceans as the final repository [13]. As biology of reproduction is highly conserved in vertebrates [14][15][16], studies on fishes as model organisms provide significant information to establish a detailed risk assessment and to establish novel or more sensitive endpoints for ECs-related fertility threat. Frequent clear evidences show reproductive disorders in fishes from polluted aquatic environments (see Section 4). The adverse effects of ECs on endocrine regulation of reproduction in male fishes have been extensively studied and reviewed in laboratory studies [13][17][18][19][20][21][22][23][24][25]. In contrast, our knowledge to understand whether ECs-disrupted hormonal functions result in diminished fertility is poor. To answer, it is critical to uncover the adverse effects of ECs on sperm production, morphology, genome, and motility kinetics as key determinants of fertility.
We have recently reviewed the toxicity of ECs on sperm morphology and motility in fishes, in vitro [26]. The review showed that ECs, in a dose-dependent manner, cause damage to sperm morphology and interfere with sperm energetics and motility kinetics, and thus affect male fertility. However, significant decreases or complete suppression of sperm motility and fertilizing ability occurred mostly at concentrations considerably higher than those reported in the aquatic environment or exceeding the WHO recommended limits for surface waters. Recently, Carnevali et al. [27] and Golshan and Alavi [25] suggested that ECs are capable of affecting sperm quality in fishes associated with alternations in hormonal functions of hypothalamus–pituitary–testis (HPT).

2. An Introduction to Reproductive Biology in Male Fishes

It is essential to review reproductive biology, fertility indicators, and determinants of fertility in male fishes before delving into the ECs-related fertility threat. These provide the basic information to better understand multiplicity of sites through which ECs interfere with fertility. To clarify the terminology, “semen” refers to seminal plasma and sperm and “sperm” refers to sperm cells in the present review.

2.1. Anatomy of Reproductive Organ

In general, the male reproductive organ consists of a paired testes, the testicular duct, and the sperm duct in fishes [28][29] (Figure 1). In some primitive fishes (such as sturgeons), the testes release sperm into the testicular ducts, which pass the kidneys. At spawning, semen is released into the aquatic environment through the urinary ducts opened into the urogenital opening (Figure 1A,C). In most bony fishes (teleosts), neither testicular ducts nor sperm ducts attach to the kidneys. The sperm is released from the testes into sperm ducts where seminal plasma is secreted. At spawning, semen is released into the aquatic environment through the sperm ducts opened in the urogenital opening (Figure 1B, bookmark0D).
Figure 1. Reproductive system in male fishes. Panels (A,C) show the anatomy of the reproductive system in primitive fishes (sturgeons). Sperm is released from the testes into the testicular ducts, which pass the kidney. At spawning, semen is released into the aquatic environment through the urinary ducts opened into the urogenital opening (UO). Panels (B,D) show the anatomy of the reproductive system in bony fishes (teleosts). Sperm is released from the testes into the testicular ducts. At spawning, semen is released into the aquatic environment through the sperm ducts opened into the UO. Panels (E,F) are schematic of the tubular testis and lobular testis, respectively. Panel G shows testicular compartments in fishes. K, kidneys; SD, sperm duct; SC-I, primary spermatocyte, SC-II, secondary spermatocyte; SG, spermatogonia; SP, spermatid; SZ, sperm; T, testis; TD, testicular duct; UB, Urinary bladder; UD (WD), urinary duct (Wolffian duct). The panels are modified from Grier [30], Nagahama [31]; Alavi et al. [28] and Dzyuba et al. [29]. The photo of panel A is courtesy of Associate Professor Borys Dzyuba from the sterlet (Acipenser ruthenus). The photo of panel B is from S. M. H. Alavi from the Northern pike (Esox Lucius). Panels C-G credits: © S. Barzegar-Fallah.
The testes are divided into the “tubular type” and the “lobular type” according to the distributions of spermatogonia in the seminiferous region [30][32][31][33] (Figure 1E, bookmark0F).
In the tubular type, as spermatogonia divide and enter in meiosis, the cysts migrate towards the region of the spermatic ducts located in the central region of the testis, where the cysts open to release sperm (Figure 1E). This type of testicular arrangement is found in zebrafish (Danio rerio) and guppy (Poecilia reticulate).
In the lobular type, the testis is composed of numerous lobules that are separated from each other by a thin layer of fibrous connective tissue, and spermatogonia are spread along the germinal compartment throughout the testis. The cysts do not migrate or become displaced during their development, and sperm is released into the lobular lumen (Figure 1F). This type of testicular arrangement is found in Japanese medaka (Oryzias latipes), common carp (Cyprinus carpio), goldfish (Carassius auratus), and rainbow trout (Oncorhynchus mykiss).
The testicular compartment contains Sertoli cells, Leydig cells, blood/lymphatic vessels, macrophages and mast cells, and neural and connective tissue cells (Figure 1G). The Leydig and Sertoli cells are involved in biosynthesis of steroid hormones to regulate sperm production and maturation.
The testicular ducts are located adjacent to the testes, which continue into the sperm ducts on the ventral sides. Testicular and sperm ducts possess very similar structural and enzyme-histochemical characteristics, and play key roles in nutrition of sperm, storage of sperm, synthesis of steroids, secretion of proteins and enzymes, and formation of the seminal plasma [34][35]. Maturation of sperm to acquire potential for motility and fertilizing ability occurs in the sperm ducts [36][37].

2.2. Spermatogenesis

Sperm is produced from spermatogonia following divisions [15][38][39]. During the process of spermatogenesis, diploid spermatogonia type A divides mitotically to produce diploid spermatogonia type B. The final mitotic division of spermatogonia type B produces diploid primary spermatocytes that undergo the first meiotic division to form haploid secondary spermatocytes. The second meiotic division produces haploid spermatids that transform into the flagellated sperm.

2.3. Sperm Morphology

Sperm is differentiated into a head, midpiece, and flagellum in fishes [40][41][42] (Figure 2). The head of sperm contains DNA for transferring a haploid set of the chromosomes into the oocyte upon fertilization. Mitochondria and proximal and distal centrioles are located in the midpiece. Mitochondria deliver energy that is required for the beating of the sperm motility apparatus with a “9 + 2” structure called the “axoneme” [43][44]. Both proximal and distal centrioles consist of nine peripheral triplets of microtubules. The distal centriole forms the basal body of the axoneme. Sperm is acrosomeless in teleostean fishes, while it possesses acrosome in primitive fishes, including hagfish and sturgeons [45][46].
Figure 2. Sperm morphology in primitive (chondrostei) and bony (teleostei) fishes. Sperm is composed of a head (nucleus, N), midpiece (M) and flagellum (F). In chondrostei fishes (such as sturgeons), there is an acrosome (A) at the top of the head of sperm. The ultrastructure compartments of sperm are similar between chondrostei and teleostei fishes: DC, distal centriole; PC, proximal centriole; Mt, mitochondria. The structure of the motility apparatus called “axoneme” is highly conserved, and possesses the typical 9 + 2 microtubule structure of cilia surrounded by plasma membrane. The electron micrographs are selected from the Russian sturgeon (Acipenser gueldenstaedtii) [47], and Atlantic halibut (Hippoglossus hippoglossus) sperm [48]. The schematic of the axoneme is from Inaba [49].

2.4. Sperm Physiology

The seminal plasma is a product of Sertoli cells, testicular ducts, and sperm ducts, and its composition is different among fishes that may reflect species variations. The main role of seminal plasma is to create an optimal environment for the storage of sperm during maturation in the sperm ducts. Seminal plasma maintains sperm viability, motility, and fertilizing ability, and protects sperm against damage caused by proteolytic or oxidative attacks [50][51].

2.5. Sperm Motility

Sperm is generally immotile in the seminal plasma and the sperm ducts of fishes, and motility is triggered upon discharge into the aquatic environment (Figure 3A). In most freshwater and marine fishes, osmolality of the seminal plasma is the key factor to maintain sperm in the quiescent state in the sperm ducts [52]. In some freshwater fishes, including Salmonidae and Acipenseridae, high concentrations of potassium (K+) ions inhibits sperm motility in the seminal plasma [53][54][55]. At spawning, a hypo-osmotic and a hyper-osmotic signal is necessary for initiation of sperm motility in freshwater and marine fishes, respectively [42][56][57][58][59]. Changes of osmolality around sperm accompanied by K+ efflux in freshwater fishes and water efflux in marine fishes trigger sperm motility signaling. Activation of sperm motility is associated with an increase in intracellular pH and calcium (Ca2+) ions in both freshwater and marine fishes, while cyclic adenosine monophosphate (cAMP) remains unchanged. However, studies show that demembranated sperm of salmonid and sturgeon fishes require cAMP for the axonemal beating [60][61][62]. In some marine fishes, it has been shown that 17,20β,21-trihydroxy-4-pregnen-3-one (17,20β-P) is capable to induce sperm hypermotility by increasing cAMP and intracellular Ca2+ through a membrane progesterone receptor [63][64].
Figure 3. Sperm motility signaling and kinetics in fishes. Panel (A) summarizes sperm motility signaling in fishes. Sperm is immotile in the sperm ducts and seminal plasma. At spawning, a hypo- osmolality accompanied by K+ efflux or hyper-osmolality accompanied by water efflux trigger sperm motility activation in freshwater and marine fish species, respectively. Activation of ATP- dependent sperm motility initiation is associated with an increase in intracellular calcium ([Ca2+]i) ions in all fish species and an increase in intracellular potassium ([K+]i) ions in marine species, while cyclic adenosine monophosphate (cAMP) remains unchanged. However, demembranated sperm of salmonids requires cAMP for axonemal beating. Panel (B) shows sperm motility kinetics in fishes. After initiation of sperm motility, percentage of motile sperm and sperm velocity decrease rapidly in both freshwater and marine fishes due to depletion of adenosine triphosphate (ATP) content. Panel (C) is a schematic representing various sperm velocity parameters analyzed by a computer-assisted sperm analysis. The curvilinear velocity (VCL) is the velocity along the trajectory of sperm head. The straight line velocity (VSL) is the straight line distance between the start and end points of the track divided by the duration of the movement. The angular path velocity (VAP) is the velocity along a derived smoothed path.
After initiation of sperm motility in the aquatic environment, duration of motility is very short in fishes from a few seconds to several minutes or hours depending on the species [28][29][51][59][65][66]. The inter-species differences probably depend on the capacity of the sperm to restore intracellular ATP and creatine phosphate concentrations [67][68]. Once sperm motility is initiated, the percentage of motile sperm and sperm velocity rapidly decrease, which are associated with a large, but not complete depletion of ATP [46][59][66] (Figure 3B). Fish sperm can regenerate ATP from phosphocreatine and ADP; however, this ATP regeneration system does not prevent the precipitous decline in ATP levels during motility [69][70][71].

3. Fertility Indicators and Assessments in Fishes

Fertilization and hatching rates are calculated to assess fertility in fishes (Figure 4A). The fertilization rate is the percentage of oocytes that become fertilized upon spawning or artificial insemination, and is calculated as number of fertilized eggs/initial number of oocytes × 100. Successful fertilization depends on onset of release of sperm from males and ova from females [28][72]. During the short period of motility, sperm must penetrate the oocyte through a funnel called the “micropyle” to fertilize it [31][73]. A fertilized egg can be easily identified by the presence of a multi-cellular blastodisc (cleavage), which occurs from several hours to a few days post fertilization and depends on fish species and environmental factors including temperature. The hatching rate is the percentage of hatched larvae, and calculated as number of hatched larvae/initial number of oocytes × 100. Once embryonic development is completed, larvae hatch [74][75].
Figure 4. Indicators and determinants of fertility in male fishes. Fertilization success is assessed by fertilization rate or hatching rate (A). Sperm production, morphology, genome and motility are key determinants of fertility in male fishes (B).

4. Determinants of Fertility in Male Fishes

Analyses of sperm production, morphology, genome, and motility kinetics are basically important to assess fertility in male fishes (Figure 4B).

4.1. Sperm Production

Frequent studies have shown that fertilization rate positively correlates with sperm volume, sperm density, number of sperm per oocyte, and density of sperm in the water during fertilization [76][77][78][79][80][81][82][83]. One can weigh semen mass, measure semen volume, or count sperm density to evaluate sperm production.

4.2. Sperm Morphology

There is a species-specific relationship between the head size of sperm and diameter of micropyle in fishes [49][73][74]. This indicates that sperm of one species can penetrate only into the oocyte of similar species. A change in the size of sperm head is a mirror of the size of nucleus [84][85][86]. It has also reported that sperm with a smaller head can move faster than those with a larger head [86][87][88]. Additionally, both positive and negative correlations have been reported between the length of flagellum and the sperm velocity [86][87][88][89][90][91]. These suggest that alternations in the size of sperm head and length of flagellum can result in diminished fertility by affecting sperm penetration into the oocytes or sperm motility performance. Various microscopic techniques including scanning and electron microscopy are valuable methods to assess sperm morphology [43][45][92].

4.3. Sperm Genome

Upon fertilization, sperm with a haploid number of chromosomes transmit a parental genome to the next generation. Alternation of chromosome material, Y chromosome deletion, and ploidy level are among factors that affect fertility. The integrity of sperm DNA correlates with fertilization and embryonic development in fishes [93][94]. Fertility threat has been frequently reported in polyploid fish, which were associated with failure of testicular development [95], enlarged head size making penetration of sperm through a normal-sized micropyle difficult [96][97], reduced sperm production [84][98], and increased abnormal sperm with malformation of the head, mitochondria, and flagellum resulting in decreasing motility and velocity [84][99]. One can assess the integrity of DNA using a comet assay, sperm chromatin structure assay, or terminal deoxynucleotidyl transferase dUTP nick end labelling assay (TUNEL) [100][101]. Chromosome number and DNA content can be counted or assessed using a flow cytometry, respectively [84][86][95].

4.4. Sperm Motility Kinetics

Duration of sperm motility, percentage of motile sperm, and sperm velocity are key determinants for fertility in male fishes [28][79][102]. It has been shown that sperm with faster movement and a longer period of motility have more chance to approach an oocyte to fertilize it [78][80][103][104]. In addition, it has been suggested that sperm velocity and the duration of motility are positively correlated with ATP content of sperm [59][66]. A computer-assisted sperm analysis (CASA) provides a valuable tool to assess sperm motility kinetics [105][106][107]. Percentage of sperm motility is evaluated by counting the number of motile sperm and total number of sperm. The sperm velocity is the distance between the starting and ending points of the motility track divided by the time spent for this movement. Based on sperm head positions during the period of motility, various sperm velocity parameters are identified, including curvilinear velocity (VCL, the velocity along the sperm head trajectory), straight line velocity (VSL, the straight line distance between the start and end points of the sperm head trajectory), and the angular path velocity (VAP, the velocity along a derived smoothed path) (Figure 3C).


  1. Richiardi, L.; Bellocco, R.; Adami, H.O.; Torrang, A.; Barlow, L.; Hakulinen, T.; Rahu, M.; Stengrevics, A.; Storm, H.; Tretli, S.; et al. Testicular cancer incidence in eight Northern European countries: Secular and recent trends. Cancer Epidemiol. Prev. Biomark. 2004, 13, 2157–2166.
  2. Andersson, A.M.; Jørgensen, N.; Main, K.M.; Toppari, J.; Meyts, E.R.D.; Leffers, H.; Juul, A.; Jensen, T.K.; Skakkebæk, N.E. Adverse trends in male reproductive health: We may have reached a crucial ‘tipping point’. Int. J. Androl. 2008, 31, 74–80.
  3. Phillips, K.P.; Tanphaichitr, N. Human exposure to endocrine disrupters and semen quality. J. Toxicol. Environ. Health B Crit. Rev. 2008, 11, 188–220.
  4. Lund, L.; Engebjerg, M.C.; Pedersen, L.; Ehrenstein, V.; Nørgaard, M.; Sørensen, H.F. Prevalence of hypospadias in Danish boys: A longitudinal study, 1977–2005. Eur. Urol. 2009, 55, 1022–1026.
  5. Toppari, J.; Virtanen, H.E.; Main, K.M.; Skakkebaek, N.E. Cryptorchidism and hypospadias as a sign of testicular dysgenesis syndrome (TDS): Environmental connection. Birth Defects Res. A Clin. Mol. Teratol. 2010, 88, 910–919.
  6. Mascarenhas, M.N.; Flaxman, S.R.; Boerma, T.; Vanderpoel, S.; Stevens, G.A. National, regional, and global trends in infertility prevalence since 1990: A systematic analysis of 277 health surveys. PLoS Med. 2012, 9, e1001356.
  7. Agarwal, A.; Mulgund, A.; Hamada, A.; Chyatte, M.R. A unique view on male infertility around the globe. Reprod. Biol. Endocrinol. 2015, 13, 37.
  8. Inhorn, M.C.; Patrizio, P. Infertility around the globe: New thinking on gender, reproductive technologies and global movements in the 21st century. Hum. Reprod. Update 2015, 21, 411–426.
  9. Skakkebaek, N.E.; Rajpert-De Meyts, E.; Buck Louis, G.M.; Toppari, J.; Andersson, A.M.; Eisenberg, M.L.; Jensen, T.K.; Jørgensen, N.; Swan, S.H.; Sapra, K.J.; et al. Male reproductive disorders and fertility trends: Influences of environment and genetic susceptibility. Physiol. Rev. 2016, 96, 55–97.
  10. Landrigan, P.J.; Fuller, R.; Acosta, N.J.R.; Adeyi, O.; Arnold, R.; Basu, N.N.; Baldé, A.B.; Bertollini, R.; Bose-O’Reilly, S.; Boufford, J.I.; et al. Lancet Commission on pollution and health. Lancet 2018, 391, 462–512.
  11. EFSA Scientific Committee. Scientific Opinion on the Hazard Assessment of Endocrine Disruptors: Scientific Criteria for Identification of Endocrine Disruptors and Appropriateness of Existing Test Methods for Assessing Effects Mediated by These Substances on Human Health and the Environment. EFSA J. 2013, 11, 3132. Available online: (accessed on 10 April 2021).
  12. Bergman, Å.; Heindel, J.J.; Jobling, S.; Kidd, K.A.; Zoeller, R.T. The State-of-the-Science of Endocrine Disrupting Chemicals; World Health Organization and United Nations Environment Programme Report–2012; WHO: Geneva, Switzerland, 2013; Available online: (accessed on 10 April 2021).
  13. Kime, D.E. Endocrine Disruptors in Fish; Kluwer Academic Publishers: Boston, MA, USA, 1998.
  14. Kah, O. Endocrine targets of the hypothalamus and pituitary. In Fish Physiology: Fish Neuroendocrinology; Bernier, N.J., Van Der Kraak, G., Farrell, A.P., Brauner, C.J., Eds.; Elsevier Inc.: Amsterdam, The Netherlands, 2009; Volume 28, pp. 75–112.
  15. Schulz, R.W.; de França, L.R.; Lareyre, J.J.; Le Gac, F.; Chiarini-Garcia, H.; Nobrega, R.H.; Miura, T. Spermatogenesis in fish. Gen. Comp. Endocrinol. 2010, 165, 390–411.
  16. Dufour, S.; Quérat, B.; Tostivint, H.; Pasqualini, C.; Vaudry, H.; Rousseau, K. Origin and evolution of the neuroendocrine control of reproduction in vertebrates, with special focus on genome and gene duplications. Physiol. Rev. 2020, 100, 869–943.
  17. Gregory, M.; Aravindakshan, J.; Nadzialek, S.; Cyr, D.G. Effects of endocrine disrupting chemicals on testicular functions. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 161–214.
  18. Ankley, G.T.; Bencic, D.C.; Breen, M.S.; Collette, T.W.; Conolly, R.B.; Denslow, N.D.; Edwards, S.W.; Ekman, D.R.; Garcia-Reyero, N.; Jensen, K.M.; et al. Endocrine disrupting chemicals in fish: Developing exposure indicators and predictive models of effects based on mechanism of action. Aquat. Toxicol. 2009, 92, 168–178.
  19. Scholz, S.; Klüver, N. Effects of endocrine disrupters on sexual, gonadal development in fish. Sex. Dev. 2009, 3, 136–151.
  20. Bosker, T.; Munkittrick, K.R.; MacLatchy, D.L. Challenges and opportunities with the use of biomarkers to predict reproductive impairment in fishes exposed to endocrine disrupting substances. Aquat. Toxicol. 2011, 100, 9–16.
  21. Leet, J.K.; Gall, H.E.; Sepúlveda, M.S. A review of studies on androgen and estrogen exposure in fish early life stages: Effects on gene and hormonal control of sexual differentiation. J. Appl. Toxicol. 2011, 31, 379–398.
  22. Mennigen, J.A.; Stroud, P.; Zamora, J.M.; Moon, T.W.; Trudeau, V.L. Pharmaceuticals as neuroendocrine disruptors: Lessons learned from fish on prozac. J. Toxicol. Environ. Health. B Crit. Rev. 2011, 14, 387–412.
  23. Hano, T. Studies on the evaluation of the effects of endocrine disrupting chemicals using transgenic see-through medaka (Oryzias latipes), olvas-GFP/STII-YI strain. Bull. Fish. Res. Agency 2012, 36, 1–56.
  24. Abdel-Moneim, A.; Coulter, D.P.; Mahapatra, C.T.; Sepúlveda, M.S. Intersex in fishes and amphibians: Population implications, prevalence, mechanisms and molecular biomarkers. J. Appl. Toxicol. 2015, 35, 1228–1240.
  25. Golshan, M.; Alavi, S.M.H. Androgen signaling in male fishes: Examples of anti-androgenic chemicals that cause reproductive disorders. Theriogenology 2019, 139, 58–71.
  26. Hatef, A.; Alavi, S.M.H.; Golshan, M.; Linhart, O. Toxicity of environmental contaminants to fish spermatozoa functions in vitro—A review. Aquat. Toxicol. 2013, 140–141, 134–144.
  27. Carnevali, O.; Santangeli, S.; Forner-Piquer, I.; Basili, D.; Maradonna, F. Endocrine disrupting chemicals in aquatic environment: What are the risks for fish gametes? Fish Physiol. Biochem. 2018, 44, 1561–1576.
  28. Alavi, S.M.H.; Linhart, O.; Coward, K.; Rodina, M. Fish spermatology: Implication for aquaculture management. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 397–460.
  29. Dzyuba, V.; Shelton, W.L.; Kholodnyy, V.; Boryshpolets, S.; Cosson, J.; Dzyuba, B. Fish sperm biology in relation to urogenital system structure. Theriogenology 2019, 132, 153–163.
  30. Grier, H.J. Cellular organization of the testis and spermatogenesis in fishes. Am. Zool. 1981, 21, 345–357.
  31. Nagahama, Y. The functional morphology of teleosts gonads. In Fish Physiology, Reproduction Part A.; Hoar, W.S., Randall, D.J., Donaldson, E.M., Eds.; Academic Press Inc.: London, UK, 1983; Volume IX, pp. 223–275.
  32. Billard, R. Spermatogenesis and spermatology of some teleost fish species. Reprod. Nutr. Dev. 1986, 2, 877–920.
  33. Parenti, L.R.; Grier, H.J. Evolution and phylogeny of gonad morphology in bony fishes. Integr. Comp. Biol. 2004, 44, 333–348.
  34. Lahnsteiner, F.; Patzner, R.A. The spermatic duct of blenniid fish (Teleostei, Blenniidae): Fine structure, histochemistry and function. Zoomorphology 1990, 110, 63–73.
  35. Lahnsteiner, F.; Patzner, R.A.; Weismann, T. The spermatic ducts of salmonid fishes (Salmonidae, Teleostei). Morphology, histochemistry and composition of the secretion. J. Fish Biol. 1993, 42, 79–93.
  36. Miura, T.; Yamauchi, K.; Takahashi, H.; Nagahama, Y. The role of hormone in the acquisition of sperm motility in salmonid fish. J. Exp. Zool. 1992, 261, 359–363.
  37. Morisawa, S.; Ishida, K.; Okuno, M.; Morisawa, M. Roles of pH and cyclic adenosine monophosphate in the acquisition of potential for sperm motility during migration from the sea to the river in chum salmon. Mol. Reprod. Dev. 1993, 34, 420–426.
  38. Vizziano, D.; Fostier, A.; Loir, M.; Le Gac, F. Testis development, its hormonal regulation and spermiation induction in teleost fish. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 103–140.
  39. Watanabe, A.; Onitake, K. The regulation of spermatogenesis in fish. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 141–160.
  40. Jamieson, B.G.M. Fish Evolution and Systematics: Evidence from Spermatozoa; Cambridge University Press: Cambridge, UK, 1991.
  41. Hara, M.; Okiyama, M. An ultrastructural review on the spermatozoa of Japanese fishes. Bull. Oce. Res. Inst. Univ. Tokyo 1998, 33, 1–138.
  42. Lahnsteiner, F.; Patzner, R.A. Sperm morphology and ultrastructure in fish. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 1–61.
  43. Ingermann, R.L. Energy metabolism and respiration in fish spermatozoa. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 241–266.
  44. Inaba, K.; Shiba, K. Microscopic analysis of sperm movement: Links to mechanisms and protein components. Microscopy 2018, 67, 144–155.
  45. Morisawa, S.; Cherr, G.N. Acrosome reaction in spermatozoa from hagfish (Agnatha) Eptatretus burgeri and Eptatretus stouti: Acrosomal exocytosis and identification of filamentous actin. Dev. Growth Differ. 2002, 44, 337–344.
  46. Alavi, S.M.H.; Hatef, A.; Pšenička, M.; Kašpar, V.; Boryshpolets, S.; Dzyuba, B.; Cosson, J.; Bondarenko, V.; Rodina, M.; Gela, D.; et al. Sperm biology and control of reproduction in sturgeon: (II) Sperm morphology, acrosome reaction, motility and cryopreservation. Rev. Fish Biol. Fish. 2012, 22, 861–886.
  47. Hatef, A.; Alavi, S.M.H.; Rodina, M.; Linhart, O. Morphology and fine structure of the Russian sturgeon, Acipenser gueldenstaedtii (Acipenseridae, Chondrostei) spermatozoa. J. Appl. Ichthyol. 2012, 28, 978–983.
  48. Alavi, S.M.H.; Butts, I.A.E.; Hatef, A.; Mommens, M.; Trippel, E.A.; Litvak, M.K.; Babiak, I. Sperm morphology, ATP content and analysis of motility in Atlantic halibut (Hippoglossus hippoglossus L.). Can. J. Zool. 2011, 89, 219–228.
  49. Inaba, K. Molecular architecture of sperm flagella: Molecules for motility and signaling. Zool. Sci. 2003, 20, 1043–1056.
  50. Ciereszko, A. Chemical composition of seminal plasma and its physiological relationship with sperm motility, fertilizing capacity and cryopreservation success in fish. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 215–240.
  51. Kowalski, R.K.; Cejko, B.I. Sperm quality in fish: Determinants and affecting factors. Theriogenology 2019, 135, 94–108.
  52. Morisawa, M.; Suzuki, K. Osmolality and potassium ions: Their roles in initiation of sperm motility in teleosts. Science 1980, 210, 1145–1147.
  53. Morisawa, M.; Suzuki, K.; Morisawa, S. Effects of potassium and osmolality on spermatozoan motility of salmonid fishes. J. Exp. Biol. 1983, 107, 105–113.
  54. Billard, R. Reproduction in rainbow trout: Sex differentiation, dynamics of gametogenesis, biology and preservation of gametes. Aquaculture 1992, 100, 263–298.
  55. Alavi, S.M.H.; Cosson, J.; Karami, M.; Amiri, B.M.; Akhoundzadeh, M.A. Spermatozoa motility in the Persian sturgeon, Acipenser persicus: Effects of pH, dilution rate, ions and osmolality. Reproduction 2004, 128, 819–828.
  56. Morisawa, M.; Oda, S.; Yoshida, M.; Takai, H. Transmembrane signal transduction for the regulation of sperm motility in fishes and ascidians. In The Male Gamete: From Basic to Clinical Applications; Gagnon, C., Ed.; Cache River Press: Vienna, IL, USA, 1999; pp. 149–160.
  57. Alavi, S.M.H.; Cosson, J. Sperm motility in fishes: (II) Effects of ions and osmotic pressure. Cell Biol. Int. 2006, 30, 1–14.
  58. Morisawa, M. Adaptation and strategy for fertilization in the sperm of teleosts fish. J. Appl. Ichthyol. 2008, 24, 362–370.
  59. Alavi, S.M.H.; Cosson, J.; Bondarenko, O.; Linhart, O. Sperm motility in fishes: (III) diversity of regulatory signals from membrane to the axoneme. Theriogenology 2019, 136, 143–165.
  60. Morisawa, M.; Okuno, M. Cyclic AMP induces maturation of trout sperm axoneme to initiate motility. Nature 1982, 295, 703–704.
  61. Cosson, M.P.; Cosson, J.; Andre, F.; Billard, R. cAMP/ATPrelationshipinthe activation of trout sperm motility: Their interaction in membrane-deprived models and in live spermatozoa. Cell Motil. Cytoskelet. 1995, 31, 159–176.
  62. Linhart, O.; Cosson, J.; Mims, S.D.; Shelton, W.L.; Rodina, M. Effects of ions on the motility of fresh and demembranated paddlefish (Polyodon spathula) spermatozoa. Reproduction 2002, 124, 713–719.
  63. Tubbs, C.; Thomas, P. Progestin signaling through an olfactory G protein and membrane progestin receptor-a in Atlantic croaker sperm: Potential role in induction of sperm hypermotility. Endocrinology 2009, 150, 473–484.
  64. Tan, W.; Aizen, J.; Thomas, P. Membrane progestin receptor-alpha mediates progestin-induced sperm hypermotility and increased fertilization success in southern flounder (Paralichthys lethostigma). Gen. Comp. Endocrinol. 2014, 200, 18–26.
  65. Alavi, S.M.H.; Cosson, J. Sperm motility in fishes: (I) Effects of pH and temperature. Cell Biol. Int. 2005, 29, 101–110.
  66. Cosson, J. Frenetic activation of fish spermatozoa flagella entails short-term motility, portending their precocious decadence. J. Fish Biol. 2010, 76, 240–279.
  67. Ziętara, M.S.; Biegniewska, A.; Rurangwa, E.; Swierczynski, J.; Ollevier, F.; Skorkowski, E.F. Bioenergetics of fish spermatozoa during semen storage. Fish Physiol. Biochem. 2009, 35, 607–614.
  68. Dzyuba, B.; Bondarenko, O.; Fedorov, P.; Gazo, I.; Prokopchuk, G.; Cosson, J. Energetics of fish spermatozoa: The proven and the possible. Aquaculture 2017, 472, 60–72.
  69. Lahnsteiner, F.; Patzner, R.A.; Weismann, T. Energy resources of spermatozoa of the rainbow trout Oncorhynchus mykiss (Pisces, Teleostei). Reprod. Nut. Dev. 1993, 33, 349–360.
  70. Saudrais, C.; Fierville, F.; Loir, M.; Le Remeur, E.; Cibert, C.; Cosson, J. The use of phosphocreatine plus ADP as energy source for motility of membrane-derived trout spermatozoa. Cell Motil. Cytoskel. 1998, 41, 91–106.
  71. Woolsey, J.; Ingermann, R.L. Acquisition of the potential for sperm motility in steelhead (Oncorhynchus mykiss): Effect of pH on dynein ATPase. Fish Physiol. Biochem. 2003, 29, 47–56.
  72. Kinsey, W.H.; Sharma, D.; Kinsey, S.C. Fertilization and egg activation in fishes. In The Fish Oocyte: From Basic Studies to Biotechnological Applications; Babin, P.J., Cerdà, J., Lubzens, E., Eds.; Springer: Dordrecht, The Netherlands, 2007; pp. 397–409.
  73. Kudo, S. Fertilization, cortical reaction, polyspermy preventing and anti-microbial mechanisms in fish eggs. Bull. Inst. Zool. Acad. Sinica 1991, 16, 313–340.
  74. Ginzburg, A.S. Fertilization in Fishes and Problem of Polyspermy; Israel Program for Scientific Translations; National Technical Information Service US Department of Commerce: Springfield, VA, USA, 1972.
  75. Iwamatsu, T. Stages of normal development in the medaka Oryzias latipes. Mech. Dev. 2004, 121, 605–618.
  76. Tvedt, H.B.; Benfey, T.J.; Martin-Robichaud, D.J.; Power, J. The relationship between sperm density, spermatocrit, sperm motility and fertilization success in Atlantic halibut, Hippoglossus hippoglossus. Aquaculture 2001, 194, 191–200.
  77. Rideout, R.M.; Trippel, E.A.; Litvak, M.K. Relationship between sperm density, spermatocrit, sperm motility and spawning date in wild and cultured haddock. J. Fish Biol. 2004, 65, 319–332.
  78. Kaspar, V.; Kohlmann, K.; Vandeputte, M.; Rodina, M.; Gela, D.; Kocour, M.; Alavi, S.M.H.; Hulak, M.; Linhart, O. Equalizing sperm concentrations in a common carp (Cyprinus carpio) sperm pool does not affect variance in proportions of larvae sired in competition. Aquaculture 2007, 272 (Suppl. S1), S204–S209.
  79. Hatef, A.; Niksirat, H.; Alavi, S.M.H. Composition of ovarian fluid in endangered Caspian brown trout, Salmo trutta caspius, and its effects on spermatozoa motility and fertilizing ability compared to freshwater and a saline medium. Fish Physiol. Biochem. 2009, 35, 695–700.
  80. Butts, I.A.E.; Trippel, E.A.; Litvak, M.K. The effect of sperm to egg ratio and gamete contact time on fertilization success in Atlantic cod Gadus morhua. Aquaculture 2009, 286, 89–94.
  81. Butts, I.A.E.; Roustaian, P.; Litvak, M.K. Fertilization strategies for winter flounder: Effects of spermatozoa density and the duration of gamete receptivity. Aquac. Biol. 2012, 16, 115–124.
  82. Linhart, O.; Cheng, Y.; Xin, M.M.; Rodina, M.; Tučková, V.; Shelton, W.L.; Kašpar, V. Standardization of egg activation and fertilization in sterlet (Acipenser ruthenus). Aquac. Rep. 2020, 17, 100381.
  83. Cheng, Y.; Franek, R.; Rodina, M.; Xin, M.; Cosson, J.; Zhang, S.; Linhart, O. Optimization of Sperm Management and Fertilization in Zebrafish (Danio rerio (Hamilton)). Animals 2021, 11, 1558.
  84. Linhart, O.; Rodina, M.; Flajšhans, M.; Mavrodiev, N.; Nebesárová, J.; Gela, D.; Kocour, M. Studies on sperm of diploid and triploid tench, Tinca tinca (L.). Aquac. Int. 2006, 14, 9–25.
  85. Piferrer, F.; Beaumont, A.; Falguière, J.C.; Flajšhans, M.; Haffray, P.; Colombo, L. Polyploid fish and shellfish: Production, biology and applications to aquaculture for performance improvement and genetic containment. Aquaculture 2009, 293, 125–156.
  86. Alavi, S.M.H.; Drozd, B.; Hatef, A.; Flajšhans, M. Sperm morphology, motility and velocity in naturally polyploid European weatherfish (Misgurnus fossilis L.). Theriogenology 2013, 80, 153–160.
  87. Pšenička, M.; Flajšhans, M.; Hulák, M.; Kašpar, V.; Rodina, M.; Borishpolets, S.; Linhart, O. The influence of ploidy level on ultrastructure and motility of tench Tinca tinca (L.) spermatozoa. Rev. Fish Biol. Fish. 2010, 20, 331–338.
  88. Pšenička, M.; Kašpar, V.; Rodina, M.; Gela, D.; Hulák, M.; Flajšhans, M. Comparative study on ultrastructure and motility parameters of spermatozoa of tetraploid and hexaploid Siberian sturgeon Acipenser baerii. J. Appl. Ichthyol. 2011, 27, 683–686.
  89. Gage, M.J.G.; MacFarlane, C.; Yeates, S.; Shackleton, R.; Parker, G.A. Relationships between sperm morphometry and sperm motility in the Atlantic salmon. J. Fish Biol. 2002, 60, 1528–1539.
  90. Alavi, S.M.H.; Psenicka, M.; Rodina, M.; Policar, T.; Linhart, O. Changes of sperm morphology, volume, density and motility and seminal plasma composition in Barbus barbus (Cyprinidae: Teleostei) during the reproductive season. Aquat. Living Resour. 2008, 21, 75–80.
  91. Alavi, S.M.H.; Rodina, M.; Viveiros, A.T.M.; Cosson, J.; Gela, D.; Boryshpolets, S.; Linhart, O. Effects of osmolality on sperm morphology, motility and flagellar wave parameters in Northern pike (Esox lucius L.). Theriogenology 2009, 72, 32–43.
  92. Ishijima, S.; Hara, M.; Okiyama, M. Comparative studies on spermatozoan motility of Japanese fishes. Bull. Oce. Res. Inst. Univ. Tokyo 1998, 33, 139–152.
  93. Beirão, J.; Boulais, M.; Gallego, V.; O’Brien, J.K.; Peixoto, S.; Robeck, T.R.; Cabrita, E. Sperm handling in aquatic animals for artificial reproduction. Theriogenology 2019, 133, 161–178.
  94. Figueroa, E.; Lee-Estévez, M.; Valdebenito, I.; Farías, J.G.; Romero, J. Potential biomarkers of DNA quality in cryopreserved fish sperm: Impact on gene expression and embryonic development. Rev. Aquac. 2020, 12, 382–391.
  95. Kim, D.S.; Jo, J.Y.; Lee, T.Y. Induction of triploidy in mud loach (Misgurnus mizolepis) and its effect on gonad development and growth. Aquaculture 1994, 120, 263–270.
  96. Chourrout, D.; Chevassus, B.; Krieg, F.; Happe, A.; Burger, G.; Renard, P. Production of second generation triploid and tetraploid rainbow trout by mating tetraploid males and diploid females-potential of tetraploid fish. Theor. Appl. Genet. 1986, 72, 193–206.
  97. Oshima, K.; Morishima, K.; Yamaha, E.; Arai, K. Reproductive capacity of triploid loaches obtained from Hokkaido Island, Japan. Ichthyol. Res. 2005, 52, 1–8.
  98. Fujimoto, T.; Yasui, G.S.; Yoshikawa, H.; Yamaha, E.; Arai, K. Genetic and reproductive potential of spermatozoa of diploid and triploid males obtained from interspecific hybridization of Misgurnus anguillicaudatus female with M. mizolepis male. J. Appl. Ichthyol. 2008, 24, 430–437.
  99. Kawamura, K.; Ueda, T.; Aoki, K.; Hosoya, K. Spermatozoa in triploids of the rosy bitterling, Rhodeus ocellatus ocellatus. J. Fish Biol. 1999, 55, 420–432.
  100. Cabrita, E.; Robles, V.; Rebordinos, L.; Sarasquete, C.; Herraez, M.P. Evaluation of DNA damage in rainbow trout (Oncorhynchus mykiss) and gilthead sea bream (Sparus aurata) cryopreserved sperm. Cryobiology 2005, 50, 144–153.
  101. Shaliutina, A.; Hulak, M.; Gazo, I.; Linhartova, P.; Linhart, O. Effect of short-term storage on quality parameters, DNA integrity, and oxidative stress in Russian (Acipenser gueldenstaedtii) and Siberian (Acipenser baerii) sturgeon sperm. Anim. Reprod. Sci. 2013, 139, 127–135.
  102. Linhart, O.; Rodina, M.; Gela, D.; Kocour, M.; Vandeputte, M. Spermatozoal competition in common carp (Cyprinus carpio): What is the primary determinant of competition success? Reproduction 2005, 130, 705–711.
  103. Gage, M.J.G.; MacFarlane, C.P.; Yeates, S.; Ward, R.G.; Searle, J.B.; Parker, G.A. Spermatozoal traits and sperm competition in Atlantic salmon: Relative sperm velocity is the primary determinant of fertilization success. Curr. Biol. 2004, 14, 44–47.
  104. Linhart, O.; Alavi, S.M.H.; Rodina, M.; Gela, D.; Cosson, J. Comparison of sperm velocity, motility and fertilizing ability between firstly and secondly activated spermatozoa of common carp (Cyprinus carpio). J. Appl. Ichthyol. 2008, 24, 386–392.
  105. Wilson-Leedy, J.G.; Ingermann, R.L. Development of a novel CASA system based on open source software for characterization of zebrafish sperm motility parameters. Theriogenology 2007, 67, 661–672.
  106. Cosson, J. Methods to analyse the movements of the spermatozoa and their flagella. In Fish Spermatology; Alavi, S.M.H., Cosson, J., Coward, K., Rafiee, G., Eds.; Alpha Science Ltd.: Oxford, UK, 2008; pp. 63–102.
  107. Amann, R.P.; Waberski, D. Computer-assisted sperm analysis (CASA): Capabilities and potential developments. Theriogenology 2014, 81, 5–17.
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