Oocyte distribution within and between ovary lobes is largely homogeneous in Lachnolaimus maximus (Perciformes: Labridae).
A number of fish reproduction studies have addressed oocyte distribution pattern in the ovary. Most of these studies have focused on a species' reproductive biology, defining spawning season, and characterizing sexual differentiation and gonad development. To identify the homogeneity degree in ovary oogenesis, analyses have been commonly done along three regions (anterior, middle and posterior) of each ovary lobe (right and left). Generally, three parameters have been used in most of these studies: average oocyte diameter, oocyte density per unit of weight; and oocyte density per unit of ovary tissue surface. Studies have addressed females in different maturation phases and all oocyte stages present in the ovary (Tricas & Hiramoto, 1989; Lee, Liu, Su, & Wu, 2005; Weng, Liu, Lee, & Tsai, 2005; Hainfellner et al., 2011; Liao & Chang, 2011, Wu, Su, Liu, Weng, & Wu, 2012), as well as only vitellogenic or mature oocytes (Witthames & Greer Walker, 1995; Ma et al., 1998; Nichol & Acuna, 2001; McElroy et al., 2013). Some do not include exact data on female maturation phases and/or the oocyte stages analyzed (Matic-Skoko, Kraljevic, & Dulcic, 2004; Kennedy et al., 2007; Renones, Grau, Mas, Riera, & Saborido-Rey, 2010; Sequeira, Neves, Paiva, Vieira, & Gordo, 2012; Yang, Chen, & Hu, 2013).
Gonadal development studies have focused in standard histological methods, and very few studies have used digital image analysis, to calculate average oocyte diameter and estimate oocyte density (Nichol & Acuna, 2001; McElroy et al., 2013); although this type of analysis lias been in use in the biological sciences for over a decade (Klibansky & Juanes, 2008). The application of digital image processing techniques may generate data on structure from images normally intended to improve image quality and in the search of data (Pertusa, 2003). They can also be used to count and measure cells or other small particles in an image, and to make precise measurements; besides, the use of these techniques has reduced the processing time, health risks and costs of standard histological methods.
Hogfish Lachnolaimus maximus (Walbaum, 1792) is a labrid fish species distributed from the mid-Atlantic seaboard of the United States (North Carolina) to the North coast of South America (Venezuela and Guyana), including the Bermudas, the Gulf of Mexico and the Caribbean Sea (Carpenter, 2002). It is important in commercial and recreational fisheries throughout its range, and presently, the IUCN (2016) has considered the species to be at high risk of extinction in the wild (Vulnerable A2bd). Growth and recruitment stage overfishing has been reported in a L. maximus population in South Florida (US) (McBride & Murphy, 2003; McBride & Richardson, 2007; McBride, Thurman, & Bullock, 2008). The L. maximus population in Southern Gulf of Mexico, on the continental shelf of the Yucatan Peninsula (known as Campeche Bank), is considered a high value alternative target fishing resource. To date, no studies have been done on the biology, exploitation level and conservation status of L. maximus population in Campeche Bank.
The reproductive biology of this species' lias been studied in populations of Puerto Rico (Colin, 1982), Cuba (Claro, Garcia-Cagide, & Fernandez de Alaiza, 1989), North Carolina (Parker, 2000), the Northeast Gulf of Mexico, and South Florida (Davis, 1976; McBride, Johnson, Bullock, & Stengard, 2001; Robinson & Prince, 2003; McBride & Johnson, 2007; McBride et al., 2008; Munoz, Burton, Brennan, & Parker, 2010; Collins & McBride, 2015). Of these studies, six have included histological analysis of gonads to identify species sexuality (protogynous hermaphrodite) and reproductive cycle, and to estimate fecundity. Only two (Davis, 1976; McBride & Johnson, 2007) have addressed spatial development of oocytes in the ovaries. Neither of these reported studies have evidence of heterogeneous oocyte development between lobes or within sections of the same lobe. But Davis (1976) only analyzed two females, while McBride and Johnson (2007) made a cursory examination of a small number of slides (3 locations) from a limited number of females (n= 14) to ensure that no clear difference in oocyte distribution patterns occurred between locations (McBride, personal communication, May 19, 2016).
The objective of the present study was to characterize oocyte distribution patterns in different developmental stages and different sections of the ovary of L. maximus, by the use digital images taken from histological sections of female gonads, in different development stages. The resulting oocyte distribution patterns will determine the sampling protocol for histological sampling from gonads of this species, thus providing more accuracy to analyses of main reproductive parameters (i.e. maturity, reproductive cycle and fecundity) of L. maximus in Campeche Bank.
MATERIALS AND METHODS
Specimen collection: The data analyzed here were collected as part of a project studying L. maximus reproduction in Campeche Bank. Specimens were collected between May 2013 and January 2014 in three coastal marine zones, corresponding to areas reported to have substantial abundance of L. maximus along the coast of the state of Yucatan (MexicanoCintora, Leonce-Valencia, Salas, & Vega-Cendejas, 2007): Celestun (west zone: 20[grados]51'33" N - 90[grados]24'0" W); Dzilam de Bravo (central zone: 21[grados]23'33" N - 88[grados]53'29" W); and Rio Lagartos (East zone: 21[grados]35'51" N - 88[grados]9'28" W). Specimens were caught between six and 22 m depth by professional fishers using harpoons and compressor diving gears. Sex identification was done based on sexual dimorphism traits described by Davis (1976), Claro et al. (1989) and McBride and Johnson (2007). For each specimen we recorded fork length (FL; nearest 0.1 cm) with a graduated ictiometer and whole weight (WW; with an electric scale Sartorius model TE2101 with a 2 100 ([+ or -] 0.1) g capacity). A total of 850 specimens were collected, of which 646 were female (size range: 13.9-39.4 cm FL; 135-1 132 g WW) and 204 were male (26.3-47.4 cm FL; 344-1 885 g WW). Based on previous data on its reproductive cycle of populations in Florida and Cuba (Davis, 1976; Claro et al., 1989; McBride et al., 2008; Collins & McBride, 2015), the ninemonth collection period considered, included the probable maturation season of L. maximus in Campeche Bank.
Female selection and ovary histology:
From the total sample of females, a pre-selection was made of those exhibiting visible oocytes in the ovaries (N = 47; 20.2-35.3 cm FL; 171-939 g WW), taking care to identify mature adult individuals with secondary growth oocytes. Six samples were extracted from each preselected ovary for histological analysis: one from the anterior, middle and posterior sections of the left and right ovarian lobes. These were fixed in Bouin's solution for four to five days, embedded in paraplast, sectioned at a 6 [micro]m thickness (4 to five sections per sampled region), and stained with Gabe and Martoja's one-step trichrome stain (Gabe, 1968). Based on the histological analyses, the preselected females were classified by their gonad development phase into one of the following reproductive phases or sub-phases (Brown-Peterson, Wyansky, Saborido-Rey, Macewicz, & Lowerre-Barbieri, 2011): early developing (ED), developing (D), spawning capable (SC) and actively spawning (AS).
The histological sections of only 23 females (20.3-34.0 cm FL; 191-814 g WW) were of sufficient quality (i.e. sections with no broken or incomplete oocyte and adequate color contrast) to allow digital image processing.
Image analysis: Digital images were taken from three microscopic fields (each field = 3.8 [mm.sup.2]) of each ovary region section using a camera (Axiocam MRc) attached to a microscope (Axioscop; 25X). Each image was analyzed with the Image-Pro Plus 6.0 program (Media Cybernetics, Inc.).
Before beginning the oocyte count, all oocytes (that did not appear complete in the right and upper portions of each field) were excluded to prevent recounting of the same oocyte. Each image was processed with spectral analysis and a binary mask generated within which the selected oocytes were counted. Oocyte stages were identified based on the criteria developed by Wallace and Selman (1981), Brown-Peterson, Thomas and Arnold (1988) and Lowerre-Barbieri et al. (2009): primary growth (PG) oocyte; cortical alveolar (CA) oocyte; primary, secondary and tertiary vitellogenic (Vtgl, 2 and 3, respectively) oocytes; and oocyte maturation (OM). The oocyte maturation stage included: germinal vesicle migration, germinal vesicle breakdown, yolk coalescence, and hydration events.
Contingency tables (rows x columns) were applied to each group of females which had been classified into the four reproductive phases or sub-phases to analyze the frequency distributions of oocyte stage (columns) at two levels (rows): 1) anterior, middle and posterior regions of each ovary lobe, and 2) right and left lobes of the ovary. A Pearson's Chi-squared ([chi square]) was used to determine if oocyte frequencies were uniform between rows and columns. When distributions were not uniform, a replicated goodness-of-fit test (heterogeneity G-test) (Sokal & Rohlf, 1995) was applied to identify which rows differed in terms of oocyte frequency between columns. All statistical analyses ([alpha] = 0.05) were run with the InfoStat program (Di Rienzo et al., 2013).
Following the degree of ovary development, the 23 selected females analyzed were classified in four reproductive phases or subphases: eight were in the ED sub-phase; four in the D phase; five in the SC phase; and six in the AS sub-phase (Fig. 1). The latter did not exhibit newly collapsed post-ovulatory follicles (POFs) in the ovary. The different oocyte stages characteristic of each female reproductive phases and sub-phases were observed in all the examined ovary sections and lobes (Table 1). In addition, the oocyte density typical of each phase and sub-phase was generally very similar between ovary regions and lobes; the one exception being Vtg3 oocytes in the right ovary of SC phase females.
The oocyte frequency analysis showed that the oocyte distribution was uniform in all three regions of the left lobe ([chi square], 0.1423 < P < 0.3858), no matter the phase or sub-phase (Table 2). In the right lobe, oocyte frequencies were also uniform in females at the ED subphase ([chi square], P = 0.2683), the D phase ([chi square], P = 0.4006) and the AS sub-phase ([chi square], P = 0.6852). However, females in the SC phase exhibited significant differences in oocyte frequencies in this lobe ([chi square], P = 0.0146). This discrepancy in frequencies in the SC phase females was confirmed by the G test (P = 0.0086; Table 3). Significant differences were present between any combination of the three regions in the right lobe of these females, the highest being between the middle and posterior lobes. This difference was probably caused by variability in Vtg3 oocyte frequency between the three regions of the right lobe (Table 1). The SC phase females analyzed here were likely just entering that phase. Nonetheless, if this oocyte category is excluded from the statistical analysis, distribution of all the other oocyte categories (PG, CA, Vtgl and 2 oocytes) was homogeneous throughout the three regions of the right lobe ([chi square] = 7.83; d.f. = 6; P = 0.2510). Oocyte frequency distribution between lobes was essentially uniform, no matter the reproductive phase or sub-phase (y2. 0.1459 [menor que o igual a] P [menor que o igual a] 0.7094; Table 4).
Most of females analyzed in the present study exhibited a high degree of homogeneity in oocyte distribution within and between the ovary lobes. The only observed difference in distribution was between different regions of the right lobe in SC phase females. These females had low Vtg3 oocyte densities in both lobes, as well as notable variation in the frequencies of this oocyte stage among the regions of the right lobe. This variation was apparently one of the possible factors causing the heterogeneity observed in the SC phase females. During the spawning season of L. maximus from the Eastern Gulf of Mexico, Collins and McBride (2015) observed a diel pattern of oocyte maturation in females: germinal vesicle migration of the vitellogenic oocytes occurs in the late morning-early afternoon preceding spawning day. As a consequence, few vitellogenic oocytes without any evidence of maturation were present in the ovaries at mid-day. Notwithstanding, SC phase females analyzed in the present study presented neither oocytes undergoing early nor late germinal vesicle migration. These females were not undergoing OM and probably were barely entering the SC phase and therefore displaying very few oocytes progressing towards the final vitellogenesis stage.
In this latter case, then females classified as being in the SC phase, may not necessarily be representative of this stage, and data from them must be used with caution. Larger sample sizes of SC phase females collected throughout this species' reproductive season would help to eliminate any uncertainty about Vtg3 oocyte distribution during this phase. However, even if Vtg3 oocyte distribution is heterogeneous in the right lobe during this phase, it did not affect the protocol for estimating fecundity. Female L. maximus have asynchronous oocyte development and engage in successive batch spawning (Claro et al., 1989; McBride & Johnson, 2007). This means that only females in the AS subphase, not exhibiting newly collapsed POFs in the ovary, should be selected to estimate batch fecundity (Murua et al., 2003). In the present data, AS sub-phase females exhibited a homogeneous distribution of all oocyte stages, including OM, which are those considered when estimating batch fecundity.
Two previous studies have been done on oocyte development homogeneity in L. maximus, both with a Florida population (Davis, 1976; McBride & Johnson, 2007). Over 40 years ago, Davis (1976) has addressed oocyte development in the anterior, middle and posterior regions of the same lobe and between right and left lobes. Using contingency tables based on the frequency distributions and supported with [chi square] data, this author analyzed if the distribution of oocytes in three diameter ranges differed between regions of the same lobe and/or between lobes. No differences were observed in oocyte counts in any of the three diameter classes between regions or lobes. However, this analysis was not particularly robust since the sample size was small (N = 2; 29.6 and 30.4 cm FL), and the oocyte diameter ranges were unrelated to reproductive phase, being essentially random. McBride and Johnson (2007) also studied histological sections from three sections of ovary lobes in L. maximus, and found no differences in oocyte distribution. However, these authors reached their conclusion by analyzing the oocytes stages present, tunica appearance, and presence or absence of atretic, rather than attempting to count cell numbers per stage, as was done in the present study (McBride, personal communication, May 19, 2016).
Oocyte distribution patterns in teleosts vary by species, and appear unrelated to taxonomic level or any other ecological or biogeographic characteristic. The pattern in L. maximus of largely homogeneous distribution of oocyte stages within and between ovary lobes coincides with reports for other females teleosts. It has been observed in species from both temperate waters [e.g. Clupea harengus (Linnaeus, 1758) (Ma et al., 1998), Spicara maena (Linnaeus, 1758) (Matic-Skoko et al., 2004), Gadus morhua and Merluccius merluccius (Linnaeus, 1758) (Witthames et al., 2009), Epinephelus marginatus (Lowe, 1834) (Renones et al., 2010), andPseudopleuronectes americanus (Walbaum, 1792) (McElroy et al., 2013)]; tropical and subtropical waters [e.g. Chaetodon multicinctus (Garrett, 1863) (Tricas & Hiramoto, 1989), Leiognathus equulus (Forsskal, 1775) (Lee et al., 2005), Spratelloides gracilis (Temminck, & Schlegel, 1846) (Weng et al., 2005), Tylosurus acus melanotus (Bleeker, 1850) (Liao & Chang, 2011), and Psenopsis anomala (Temminck, & Schlegel, 1844) (Wu et al., 2012)]; and temperate and tropical waters [e.g. Helicolenus dactylopterus (Delaroche, 1809) (Sequeira et al., 2012) and Trachinocephalus myops (Forster, 1801) (Yang et al., 2013)]. In contrast, other teleosts exhibit a heterogeneous oocyte distribution pattern in the ovary, as in the temperate marine flatfish species Solea solea (Linnaeus, 1758) (Witthames & Greer Walker, 1995), Limando aspera (Pallas, 1814) (Nichol & Acuna, 2001) and Pleuronectes platessa (Linnaeus, 1758) (Kennedy et al., 2007), and the freshwater species Prochilodus lineatus (Valenciennes, 1837) from Brazil (Hainfellner et al., 2011). In another example, significant differences in average oocyte size were observed along the anterior-posterior axis and between the center and periphery of ovary lobes in bigeye tuna Parathunnus sibi (= Thunnus obesus: Lowe, 1839) (Yuen, 1955) and bluefin tuna Thunnus thynnus (Linnaeus, 1758) (Baglin, 1982).
Given the homogeneous oocyte distribution pattern within and between the ovary lobes in L. maximus females in the AS sub-phase, no systematization is required of the gonad histological sampling protocol to estimate species batch fecundity. Stereological methods in tandem with digital image processing can therefore be used to make accurate measurements of oocyte size, shape, count, area and volume in the ovaries of L. maximus, and generate unbiased fecundity estimates in this species. However, ovary sampling for characterizing the reproductive cycle in this species, based on analysis of the seasonal development of reproductive phase and sub-phase frequency, needs to be done with caution due to the potential for heterogeneity among Vtg3 oocytes in the right ovary lobe of females in the SC phase. If this is a possible risk in a given sample, the present results suggest it is best to systematically take sections of any region in the left ovary when conducting a study encompassing all of a species' reproductive aspects.
Received 02-11-2016. Corrected 12-VII-2016. Accepted 11-VIII-2016.
Collections were authorized by fishing licenses: PPF/DGOPA-037/14 and PPF/ DGOPA-080/15, from Secretaria de Agricultura, Ganaderia, Desarrollo Rural, Pesca y Alimentacion/Comision Nacional de Acuacultura y Pesca. The authors thank the professional fishers Leonardo Pech, Santos Efrain Sosa Pech, Jose Nicolas Flores Aceves, Luis Emilio Aceves Nadal and Cesar Alexander Tun Pacheco for their assistance with sample collection. We also thank the fishing cooperatives Nohoch Cuch S.C.L., S.C.P.P., and Pescadores de Rio Lagartos S.C.D.R.L, S.C.P.P. The Consejo Nacional de Ciencia y Tecnologia (CONACYT) financed a doctoral research stay in La Paz, Baja California Sur through the 2013 PNPC RM program. Technical assistance with gonad histological processing was provided by Teresa Colas-Marrufo, and the digital images were taken by Ximena Renan. We thank R. S. McBride, one anonymous reviewer and the editor for providing insightful comments that improved the quality of the manuscript.
Baglin, Jr., R. E. (1982). Reproductive biology of western Atlantic bluefin tuna. Fishery Bulletin, 80, 121-134.
Brown-Peterson. N., Tilomas. P., & Arnold. C. (1988). Reproductive biology of the spotted seatrout. Cynoscion nebulosas, in south Texas. Fishery Bulletin, 86, 373-388.
Brown-Peterson. N., Wyanski. D., Saborido-Rey. F., Macewicz. B., & Lowerre-Barbieri. S. (2011). A standardized terminology for describing reproductive development in fishes. Marine and Coastal Fisheries: Dynamics. Management, and Ecosystem Science. J(l)' 52-70.
Carpenter. K. E. (2002). The living marine resources of the Western Central Atlantic. Volume 3: Bony fishes part 2 (Opistognathidae to Molidae). sea turtles and marine mammals. FAO Species Identification Guide for Fishery Purposes and American Society of ichthyologists and Herpetologists Special Publication No. 5. FAO, Rome.
Claro, R., Garcia-Cagide, A., & Fernandez de Alaiza, R. (1989). Caracteristicas biologicas del pez perro, Lachnolaimus maximus (Walbaum), en el Golfo de Batabano, Cuba. Revista Investigaciones del Mar, Cuba, 10(3), 239-252.
Colin, P. L. (1982). Spawning and larval development of the hogfish, Lachnolaimus maximus (Pisces: Labridae). Fishery Bulletin, 4, 853-862.
Collins. A. B., & McBride. R. S. (2015). Variations in reproductive potential between nearshore and offshore spawning contingents of hogfish in the eastern Gulf of Mexico. Fisheries Management and Ecology, 22, 113-124.
Davis. J. C. (1976). Biology of the hogfish. Lachnolaimus maximus (Walbaum). in the Florida Keys (Master dissertation). University of Miami. Coral Gables. FL.
Di Rienzo, J. A., Casanoves, F., Balzarini, M. G., Gonzalez. L., Tablada. M., & Robledo. C. W. (2013). Infostat (Version 2013). Grupo InfoStat. FCA. Universidad Nacional de Cordoba. Argentina. Retrieved from http://www.infostat.com.ar
Gabe. M. (1968). Techniques histologiques. Paris: Masson & Cie.
Hainfellner. P., de Souza. T. G., Nascimento. T. S. R., Freitas. G. A., & Batlouni. S. R. (2011). Heterogeneous distribution of oocytes in the ovaries of Prochilodus lineatus. Indian Journal of Science and Technology. Proceedings of 9"' International Symposium on Reproductive Physiology of Fish. Cochin India, 4, 111-112.
IUCN (International LTnion for Conservation of Nature and Natural Resources). (2016). The IUCN Red List of Threatened Species. Version 2015.4. Retrieved from dhttp://www.iucnredlist. org.
Kennedy. J., Witthames. P. R., & Nash. R. D. M. (2007). The concept of fecundity regulation in plaice (Pleuronectes platessa) tested on three Irish Sea spawning populations. Canadian Journal of Fisheries and Aquatic Sciences. 64, 587-601.
Klibansky. N., & Juanes. F. (2008). Procedures for efficiently producing high-quality fecundity data on a small budget. Fisheries Research. 89, 84-89.
Lee. C. F., Liu. K. M., Su. W. C., & Wu. C. C. (2005). Reproductive biology of the common ponyfish
Leiognathus equulus in the south-western waters off Taiwan. Fisheries Science. 71, 551-562.
Liao. Y. Y. & Chang. Y. H. (2011). Reproductive Biology of Needlefish Tylosurus acus melanotus in waters around Hsiao-Liu-Chiu Island. Southwestern Taiwan. Zoological Studies. 50(3), 296-308.
Lowerre-Barbieri. S. K. Henderson. N., Llopiz. J., Walters. S., Bickford. J., & Muller. R. (2009). Defining a spawning population (spotted seatrout Cynoscion nebulosas) over temporal, spatial, and demographic scales. Marine Ecology Progress Series, 394. 231-245.
Ma. Y. Kjesbu. O. S., & Jirgensen. T. (1998). Effects of ration on the maturation and fecundity in captive Atlantic herring (Clupea harengus). Canadian Journal of Fisheries and Aquatic Sciences. 55, 900-908.
Matic-Skoko. S., Kraljevic. M., & Dulcic. J. (2004). Fecundity of blotched picarel. Spicara maena L. (Teleostei: Centracanthidae). in the Eastern central Adriatic Sea. Acta Adraiatica. 45(2), 155-162.
McBride. R. S., & Johnson. M. R. (2007). Sexual development and reproductive seasonality of hogfish (Labridae: Lachnolaimus maximus), an hermaphroditic reef fish. Journal of Fish Biology 71, 1270-1292.
McBride. R. S., Johnson. M., Bullock, L., & Stengard, F. (2001). Preliminary observations on the sexual development of hogfish, (Pisces: Labridae). Gulf and Caribbean Fishery Institute Proceedings. 52, 98-102.
McBride. R. S., & Murphy. M. D. (2003). Current and potential yield per recruit of hogfish. Lachnolaimus maximus, in Florida. Gulf and Caribbean Fishery Institute Proceedings. 54, 513-525.
McBride. R. S., & Richardson. A. K. (2007). Evidence of size-selective fishing mortality from an age and growth study of hogfish (Labridae: Lachnolaimus maximus), a hermaphroditic reef fish. Bulletin of Marine Science. 80, 401-417.
McBride. R. S., Thurman. P. E., & Bullock. L. H. (2008). Regional variations of hogfish (Lachnolaimus maximus) life history: consequences for spawning biomass and egg production models. Journal of Northwest Atlantic Fishery Science. 41, 1-12.
McElroy. D. W., Wuenschel. M. J., Press. Y. K. Towle. E. K. & McBride. R. S. (2013). Differences in female individual reproductive potential among three stocks of winter flounder. Pseudopleuronectes americanas. Journal of Sea Research. 75, 52-61.
Mexicano-Cintora. G., Leonce-Valencia. C. O., Salas. S., & Vega-Cendejas. M. E. (2007). Recursos pesqueros de Yucatan: fichas tecnicas y referencias bibliograficas. Merida. Yucatan. Mexico': CINVESTAV-IPN.
Munoz. R. C., Burton. M. L., Brennan. K. J., & Parker. O. (2010). Reproduction, habitat utilization, and movements of hogfish (Lachnolaimus maximus) in the Florida keys. LI.S.A.: Comparisons from fished versus unfished habitats. Bulletin of Marine Science. 86(1), 93-116.
Muraa. H., Kraus. G., Saborido-Rey. F., Witthames. P., Thorsen. A., & Junquera. S. (2003). Procedures to estimate fecundity of marine fish species in relation to their reproductive strategy. Journal of Northwest Atlantic Fishery Science, 33, 33-54.
Nichol, D. G., & Acuna, E. I. (2001). Annual and batch fecundities of yellowfin sole, Limanda aspera, in the Eastern Bering Sea. Fishery Bulletin, 99, 108-122.
Parker, R. O. (2000). Courtship in hogfish, Lachnolaimus maximus, and other behavior of reef fishes off Beaufort, North Carolina. The Journal of the Elisha Mitchell Scientific Society, 116(3), 260-261.
Pertusa, J. F. (2003). Tecnicas de analisis de imagen: Aplicaciones en biologia. Universidat de Valencia, Aldaia, Spain.
Renones, O., Grau, A., Mas, X., Riera, F., & SaboridoRey, F. (2010). Reproductive pattern of an exploited dusky grouper Epinephelus marginatus (Lowe 1834) (Pisces: Serranidae) population in the western Mediterranean. Scientia Marina, 74(3), 5 23-5 3 7.
Robinson, M. P., & Prince, J. S. (2003). Morphology of the sperm of two wrasses, Thalassoma Bifasciatum and Lachnolaimus Maximus (Labridae, Perciformes). Bulletin of Marine Science, 72(1), 247-252.
Sequeira, V., Neves, A., Paiva, R. B., Vieira, A. R., & Gordo, L. S. (2012). Is the fecundity type of the zygoparous fish species Helicolenus Dactylopterus determinate or indeterminate? Vie et Milieu, 62(1), 37-42.
Sokal, R. R., & Rohlf, J. (1995). Biometry. The principles and practices of statistics in biological research (3rded.). New York: W. H. Freeman and Company.
Tricas, T. C., & Hiramoto, J. T. (1989). Sexual differentiation, gonad development, and spawning seasonality of the Hawaiian butterflyfish, Chaetodon multicinctus. Environmental Biology of Fishes, 25, 111-124.
Wallace, R. A., & Selman, K. (1981). Cellular and dynamic aspects of oocyte growth in teleosts. American Zoologist. Tampa Florida, 21, 325-343.
Weng, J. S., Liu, K. M., Lee, S. C., & Tsai, W. S. (2005). Reproductive Biology of the Blue Sprat Spratelloides gracilis in the waters around Penghu, Central Taiwan Strait. Zoological Studies, 44(4), 475-486.
Witthames, P. R., & Greer Walker, M. (1995). Determinacy of fecundity and oocyte atresia in sole (Solea solea) from the Channel, the North Sea and the Irish Sea. Aquatic Living Resources, 8, 91-109.
Witthames, P. R., Thorsen, A., Murua, H., Saborido-Rey, F., Greenwood, L. N., Dominguez, R., & Kjesbu, O. S. (2009). Advances in methods for determining fecundity: application of the new methods to some marine fishes. Fishery Bulletin, 107, 148-164.
Wootton, J. R., & Smith, C. (2015). Reproductive biology of teleost fishes. Oxford: Wiley Blackwell.
Wu, C. C., Su, W. C., Liu, K. M., Weng, J. S., & Wu, L. J. (2012). Reproductive Biology of the Japanese butterfish, Psenopsis anomala, in the waters off southwestern Taiwan. Journal of Applied Ichthyology, 28, 209-216.
Yang, J. L., Chen, L. H., & Hu, T. J. (2013). Maturity and spawning of painted lizardfish, Trachinocephalus myops (Bloch and Schneider, 1801) in the southeastern China Sea. Journal of Applied Ichthyology, 29, 1050-1055.
Yuen, H. S. H. (1955). Maturity and fecundity of bigeye tuna in the Pacific. Special Scientific Report: Fisheries (No. 150, pp. 1-30). Washington: United States of Department of the Interior, Fish and Widlife Service.
Virginia Elena Noh Quinones (1), J. Rene Torres-Villegas (2), Thierry Brule (1), Jorge L. MonteroMunoz (1), Uriel Fernando Valdez-Montiel (2)
(1.) Centro de Investigacion y de Estudios Avanzados del Instituto Politecnico Nacional, Departamento de Recursos del Mar, Unidad Merida, Ant. Carr. a Progreso Km. 6, A.P. 73 Cordemex, 97310 Merida, Yucatan, Mexico; email@example.com, tbruleMcinvestav.mx, firstname.lastname@example.org
(2.) Centro Interdisciplinario de Ciencias Marinas del Instituto Politecnico Nacional, Departamento de Pesquerias y Biologia Marina, Av. Instituto Politecnico Nacional s/n, Col. Playa Palo de Santa Rita, 23096 La Paz, Baja California Sur, Mexico; email@example.com, firstname.lastname@example.org
Leyenda: Fig. 1. Histological sections of ovaries from Lachnolaimus maximus collected on Campeche Bank (Gabe and Martoja's onestep trichrome stain; scale bar = 200 microns). A) female in early developing reproductive sub-phase (30.6 cm FL), collected 16 Oct. 2013; B) female in developing reproductive phase (31.2 cm FL), collected 16 Oct. 2013; C) female in spawning capable reproductive sub-phase (27.8 cm FL), collected 15 Oct. 2013; and D) female in actively spawning reproductive sub-phase (20.6 cm FL), collected 22 May 2013. CA = cortical alveolar oocyte; PG = primary growth oocyte; OM = oocyte maturation (germinal vesicle breakdown and yolk coalescence); Vtg1 = primary vitellogenic oocyte; Vtg2 = secondary vitellogenic oocyte; Vtg3 = tertiary vitellogenic oocyte.
TABLE 1 Oocyte density (number per unit area) frequencies observed in anterior, middle and posterior regions of left and right ovary lobes of Lachnolaimus maximus females in different reproductive phases and sub--phases Female reproductive Ovary Total oocyte counts phase Lobe Region PG CA Vtg1 Early developinga Left A 1 833 321 -- (n = 8) M 1 918 302 -- P 1 815 336 -- Right A 1 913 316 -- M 1 833 283 -- P 1 774 316 -- Developing Left A 695 142 81 (n = 4) M 728 146 81 P 816 154 63 Right A 766 129 94 M 729 150 92 P 788 127 96 Spawning capable Left A 900 131 61 (n = 5) M 971 158 70 P 930 148 91 Right A 894 124 84 M 942 138 81 P 970 131 92 Actively spawning (a) Left A 627 112 66 (n = 6) M 696 104 83 P 720 94 70 Right A 638 93 69 M 653 104 64 P 654 107 65 Female reproductive Total oocyte countsTotal area phase Vtg2 Vtg3 OM analyzed (b) ([mm.sup.2]) Early developinga -- -- -- 91.2 (n = 8) -- -- -- -- -- -- -- -- -- -- -- -- -- -- -- Developing -- -- -- 45.6 (n = 4) -- -- -- -- -- -- -- -- -- -- -- -- -- -- -- Spawning capable 27 9 -- 57.0 (n = 5) 23 5 -- 17 4 -- 32 12 -- 16 5 -- 33 1 -- Actively spawning (a) 48 30 49 68.4 (n = 6) 46 27 65 36 26 59 40 35 56 45 40 49 41 25 66 n = number of females analyzed; (a) = reproductive sub--phases; (b) = three 3.8 [mm.sup.2] microscopic fields x n; A = anterior, M = middle; P = posterior; PG = primary growth oocyte; CA = cortical alveolar oocyte; Vtgl = primary vitellogenic oocyte; Vtg2 = secondary vitellogenic oocyte; Vtg3 = tertiary vitellogenic oocyte; OM = oocyte maturation. TABLE 2 Chi-square goodness-of-fit test results for oocyte density frequencies in anterior, middle and posterior regions of left and right ovary lobes of Lachnolaimus maximus females in four reproductive phases and sub-phases Female reproductive n Ovary f d.f. P phase lobe Early developing (a) 8 Left 3.65 2 0.1610 Right 2.63 2 0.2683 Developing 4 Left 6.57 4 0.1602 Right 4.04 4 0.4006 Spawning capable 5 Left 12.2 8 0.1423 Right 19.04 8 0.0146* Actively spawning (a) 6 Left 10.65 10 0.3858 Right 7.42 10 0.6852 Female reproductive Total Total area phase oocyte analyzed (b) count ([mm.sup.2]) Early developing (a) 6 525 273.6 6 435 Developing 2 906 136.8 2 971 Spawning capable 3 545 171.0 3 555 Actively spawning (a) 2 958 205.2 2 844 (a) = reproductive sub-phases; n = number of females analyzed; [chi square] = chi-square goodness-of-fit statistic; d.f. = degrees of freedom (r-1)(c-1), with r=anterior, middle and posterior regions of each right and left lobes of ovaries, and c=oocyte stages (see Tablel), for a chi-square calculated from contingency table data; p = probability value; (b) = three 3.8 [mm.sup.2] microscopic fields x three ovary regions x n. * = significant statistical difference (P < 0.05). TABLE 3 Replicated goodness-of-fit test (heterogeneity G-test) results for oocyte density frequencies in anterior, middle and posterior regions of the right ovary lobe of five Lachnolaimus maximus females in spawning capable (SC) reproductive phase. Total area analyzed = 57.0 [mm.sup.2] Total oocyte counts Ovary PG CA Vtg1 Vtg2 Vtg3 OM Test region Anterior 894 124 84 32 12 -- Middle 942 138 81 16 5 -- Posterior 970 131 92 33 1 -- Total G Pooled 2 806 393 257 81 18 -- Pooled G Heterogeneity G Ovary G d.f. P region Anterior 1 915.88 4 0.0001* Middle 2 157.78 4 0.0001* Posterior 2 177.94 4 0.0001* 6 251.60 12 < 0.05 Pooled 6 231.11 4 < 0.05 20.49 8 0.0086 PG = primary growth oocyte; CA = cortical alveolar oocyte; Vtgl = primary vitellogenic oocyte; Vtg2 = secondary vitellogenic oocyte; Vtg3 = tertiary vitellogenic oocyte; OM = oocyte maturation; G = G statistic; d.f. = degrees of freedom; p = probability value; * = highly significant statistical difference (P < 0.05). TABLE 4 Chi-square goodness-of-fit test results for oocyte density frequencies in left and right ovary lobes of Lachnolaimus maximus females in four reproductive phases and sub-phases Female n [chi d.f. P Total Total reproductive square] oocyte area phase counts analyzed (b) ([mm. sup.2]) Early developing 8 0.60 1 0.4390 12 960 547.2 (a) Developing 4 3.85 2 0.1459 5 877 273.6 Spawning capable 5 6.20 4 0.1844 7 100 342.0 Actively spawning 6 2.94 5 0.7094 5 802 410.4 (a) (a) = reproductive sub-phases; n =number of females analyzed; [chi square] = chi-square goodness-of-fit statistic; d.f. = degrees of freedom (r-1)(c-1), with r=left or right lobes of the ovaries, and c=oocytes stages (see Table 1), p = probability value; b = three 3.8 nun2 microscopic fields x three ovary regions xtwo lobes x n.
|Printer friendly Cite/link Email Feedback|
|Author:||Quinones, Virginia Elena Noh; Torres-Villegas, J. Rene; Brule, Thierry; Montero-Munoz, Jorge L.; Mon|
|Publication:||Revista de Biologia Tropical|
|Date:||Mar 1, 2017|
|Previous Article:||Taxones superiores de hormigas como sustitutos de la riqueza de especies, en una cronosecuencia de bosques secundarios, bosque primario y sistemas...|
|Next Article:||Estimation of genetic variation in closely related cycad species in Ceratozamia (Zamiaceae: Cycadales) using RAPDs markers.|