Survey
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
RH: GRIFFIN ET AL. – EYEFLUKE INFECTION IN MOSQUITOFISH MICROHABITAT SELECTION AND EYEFLUKE INFECTION LEVELS IN THE WESTERN MOSQUITOFISH (GAMBUSIA AFFINIS) Shane L. Griffin, Nichole Carpenter, Autumn Smith-Herron*, and Kristin K. Herrmann Department of Biological Sciences, Tarleton State University, Box T-0100, Stephenville, Texas 76402. Correspondence should be sent to Shane L. Griffin at: [email protected] ABSTRACT: A variety of trematode species infect the eyes of fish as second intermediate hosts. In most cases the definitive host is a piscivorous bird. Studies of a few species have shown an increase in transmission due to decreased visual acuity of the fish host. However, this may vary depending on trematode microhabitat choice within the eye. Some trematode species are found in the lens, some in the vitreous humor and others have been reported from the retina. Here we report 3 genera of eyeflukes in 3 locations of the eye in the intermediate fish host, Gambusia affinis. Clinostomum metacercariae were found attached to the outer sclera within the eye orbit, and Diplostomum metacercariae were found in the lens. Posthodiplostomum metacercariae were confirmed by histology to reside between the choroid and pigmented retina. Posthodiplostomum metacercariae were found in both eyes of all 20 fish examined and in high intensities (up to 27 metacercariae per eye). High trematode intensities between the choroid and pigmented retina found in this study may disrupt vision in this fish host. Our study is the first to document the microhabitat of all 3 trematode metacercariae within the eye of G. affinis. Eyeflukes are parasitic trematodes that commonly parasitize freshwater fish species (Chappell, 1995). Eyefluke infections can result in deleterious effects on the second intermediate fish host’s fitness and overall survivability. Infections can result in lens opacity or dislocation, cataracts, detached retina or choroid, capsular rupture, and exophthalmia, which can lead to 1 reduced vision or blindness in some cases (Rushton, 1937; Shariff et al., 1980; Grobbelaar et al., 2015). Impaired vision may result in emaciation (Shariff et al., 1980) from a decreased ability of a host to forage, as well as hinder reproduction and predator avoidance (Seppälä et al., 2004, 2005; Barber, 2007). The complex life cycles of eyeflukes involve multiple stages, which incorporate both sexual and asexual reproduction. Multiple transmission events take place throughout the life cycle typically involving 2 intermediate hosts and a definitive host. Transmission to the first intermediate snail host occurs by ingestion of eggs or penetration by the free-swimming miracidia. In the snail host, miracidia undergo asexual reproduction and produce cercariae, which eventually penetrate the epithelium of the second intermediate host, commonly a fish. In some cases, as here, cercariae migrate to the eye and develop into metacercariae. Transmission to the definitive host occurs via ingestion by a bird or predatory fish, depending on the trematode species (Shoop, 1988). Preliminary parasite data collected from local Texas river systems in 2013 indicated high intensity eyefluke infections in several local genera of fishes including the Western mosquitofish, Gambusia affinis (Poeciliidae) (K. K. Herrmann, pers. obs.). This common fish species is favorable as the second intermediate host of various trematode species (Davis and Huffman, 1977), which are found as metacercariae throughout the organs, tissues, and eyes. Some eyefluke studies document microhabitat locations within the eyes of many species of fish (Ferguson, 1943). In his synopsis of strigeid metacercariae of fishes, Hoffman (1960) recounts studies documenting multiple species of eyefluke infecting various locations within the eyes. He notes specifically, Diplostomum flexicaudum, Diplostomum spathaceum, and Diplostomum lenticola infections of the lens, with D. spathaceum, Diplostomum scheuringi, and Tylodelphis 2 clavata infecting the vitreous humor of various species of fish. Hoffman (1999) notes studies reporting Diplostomum huronense infections of the lens and vitreous humor, as well as Diplostomum pungiti infections of the retina and retinal area. Posthodiplostomum brevicaudatum infection of the retina and choroid coat has also been reported (Wisniewski, 1958). However, numerous studies on eyeflukes do not report a specific microhabitat. The objective of this study was to investigate eyefluke infections within a single population of G. affinis. In doing so, we aim to document infection intensities and morphologically identify the genera of the metacercariae constituting these infections. Information regarding eyefluke microhabitat commonly lacks specificity in many studies of various host species including G. affinis. Therefore, we further examined the location of each taxa to increase specificity in assessing potential microhabitat selection. MATERIALS AND METHODS Twenty G. affinis individuals were collected via seine net from the Paluxy River (32.230460°N, -97.776282°W) in Glen Rose, Texas in June 2014. Individuals were identified to species level (Thomas et al., 2007). Live fish were transported to the lab in an aerated container. Individuals were kept in an aerated aquarium and fed ad lib. Upon processing, individuals were euthanized in a Tricaine-Mesylate (MS-222) solution. Length, weight, and sex were recorded for each individual prior to eye dissection. Eyes were carefully removed by severing all tissue and the optic nerve to prevent damage. Left and right eyes were differentiated to determine potential eye preference. Eyes were dissected in 0.075% saline solution. The eye layers visible using a dissection microscope were completely separated, and the number of metacercariae and their relative locations were recorded (Fig. 1). Eyeflukes were fixed and stained for identification. 3 Specimens were deposited in the Sam Houston State University Parasitology Collection (SHSUP001580- SHSUP001586). Statistical analyses were conducted using SPSS (v21). An additional 7 G. affinis eyes were fixed in HistoTainer 10% neutral buffered formalin (VWR) for histological analysis to confirm metacercariae location (Fig. 2). Of these, 4 eyes were from individuals collected from the original study site on the Paluxy River that harbored high infection intensities. The other 3 eyes were from individuals collected from the Bosque River (31.976781°N, -98.031078°W) in Hico, Texas to serve as a control. According to preliminary data from 2013, this site harbored 0% eyefluke prevalence in 48 fishes including G. affinis, Cyprinella venusta, and Cyprinella lutrensis. Histology was conducted according to standard histology procedures (Humason, 1962). Eyes were embedded in paraffin wax and sliced into 7 µm sections using a microtome. Tissue sections were floated in water and placed on microscope slides coated in a protein adhesive. Sections were then stained in Hematoxylin & Eosin and examined using an Olympus BX53 microscope paired with an Olympus DP72 camera (Olympus Corporation, Tokyo, Japan). RESULTS All 20 fish were mature adult females ranging from 35.75 to 52.56 mm in total length with a mean of 41.77 mm. Mean weight was 0.91 g and ranged from 0.60 to 1.84 g. From the 40 G. affinis eyes dissected, a total of 466 metacercariae were recovered. Of these, 10 encysted metacercariae were found attached to the outer surface of the sclera within the eye orbit, 446 encysted metacercariae were found between the choroid and pigmented retina (Figs. 1, 2), and 10 unencysted metacercariae were found in the lens. Sclera metacercariae were identified as Clinostomum sp. (Caffara et al., 2011). Choroid metacercariae were identified as Posthodiplostomum sp. (Hughes, 1928; Hunter and Hunter, 1940; Hoffman, 1999). Unencysted 4 lens metacercariae were identified as Diplostomum sp. (Palmieri et al., 1976; McKeown and Irwin, 1995). Infection intensities of Clinostomum and Diplostomum metacercariae each ranged from 0 to 2 individuals per eye, whereas Posthodiplostomum metacercariae ranged from 3 to 27 individuals per eye (Table I). There was no significant difference between the left and right eyes in abundance of total eye parasites (t = -1.89, p = 0.075), Clinostomum, Posthodiplostomum or Diplostomum metacercariae (t = 0.00, p = 1.0; t = -1.90, p = 0.072; t = 0.62, p = 0.54; respectively). Data from both eyes were pooled to test for relationships between total length of fish and intensity. Infection intensity was not correlated with fish length for total eye parasites combined (r = 0.366, p = 0.056), or for Clinostomum or Diplostomum metacercariae (r = 0.301, p = 0.098; r = -0.069, p = 0.386; respectively). However, total fish length is correlated with combined intensity of Posthodiplostomum metacercariae in both eyes (r = 0.408, p = 0.037). DISCUSSION We examined a single population of G. affinis individuals, and documented 3 genera of metacercariae inhabiting 3 locations within the eye. Posthodiplostomum metacercariae were reported between the choroid and pigmented retina of both eyes in all 20 fish. Wisniewski (1958) also documented Posthodiplostomum metacercariae embedded in the choroid coat of the eye, as well as cercarial migration to that location, however this was in unspecified fish host species. Studies by Janovy and Hardin (1987, 1988), and Janovy et al. (1997) report Posthodiplostomum metacercariae in the eye and body cavity, but do not go on to specify which microhabitat within the eye. A study on fathead minnows, Pimephales promelas, also documents Posthodiplostomum infections in the eyes, as well as in the head, liver, and body cavity, but again does not specify the location within the eye (Mitchell et al., 1982). Hunter and Hunter (1940) reported 5 Posthodiplostomum minimum infections occurring in all visceral organs of various fish species, and Rakauskas and Blaževičius (2009) documented Posthodiplostomum infections occurring on the skin, fins, and gills of roach. Brock and Font (2009) reported Posthodiplostomum metacercariae in G. affinis, however the metacercariae were found in the body cavity only. Our study is the first to specifically document high intensities of Posthodiplostomum metacercariae in the choroid of G. affinis. Such high intensity, up to 27 metacercariae per eye, most likely disrupts the vision of the fish and could lead to lowered fitness as shown in experiments with other fish host species (Rushton, 1937; Shariff et al., 1980; Grobbelaar et al., 2015). We found Clinostomum metacercariae attached to the outer sclera within the eye orbit of 8 individuals. In his classification of the genus, Hoffman (1999) documents the occurrence of Clinostomum metacercariae in various fishes, with a range broad enough that he hypothesized members of this genus could likely infect most species of fish. Other studies on Clinostomum reported metacercariae in multiple fishes, being found encysted in the musculature (Cort, 1913; Meade and Bedinger, 1972; Dias et al., 2003), mesentery and viscera (Dias et al., 2003), and subcutaneously (Hopkins, 1933; Al-Awadi et al., 2010). Mitchell et al. (1982) also documented ocular infections of Clinostomum metacercariae in P. promelas. Unfortunately, the study only generalized metacercariae location as being in the eye and did not report a specific microhabitat. Overall, we found a lack of information in the literature regarding studies of Clinostomum metacercariae specifically being attached to the sclera or located in the eye orbit in G. affinis. In our study, Diplostomum metacercariae were documented in a single lens of 7 individuals and in both lenses of 1 individual. Infections of Diplostomum metacercariae occur in the aqueous and vitreous humor (Hoffman, 1960; Shariff et al., 1980; Chappell, 1995; Grobbelaar et al., 2015), retina (Shariff et al., 1980; Bortz et al., 1988; Chappell, 1995), brain 6 (Hoffman, 1960; Grobbelaar et al., 2015), and mesentery and coelom (Hoffman, 1960). However, Davis and Huffman (1977) and Aho et al. (1982) both document Diplostomum infections occurring in the coelom of G. affinis, but neither found ocular infections. Numerous studies have documented the lens microhabitat of Diplostomum metacercariae in various fishes (see Hoffman, 1960; Shariff et al., 1980; Bortz et al., 1988; Dwyer and Smith, 1989; Chappell, 1995), however we again encountered a lack of studies reporting Diplostomum lens infections in G. affinis. Certain trematodes such as Diplostomum spp. are unencysted and free-moving while inhabiting the eye of a teleost host (Grobbelaar et al., 2015). Diplostomum metacercariae documented in the lens during this study were all found to exhibit this. Several hypotheses exist as to why this phenomenon occurs, and generally involve the lens microhabitat serving as a protective barrier against environmental factors. One hypothesis suggests the lens may provide protection while passing through the definitive host’s stomach (Szidat, 1969). Further, trematodes that inhabit the lens tend to exhibit lower host specificity from the reduced immune response that occurs within the lens (Locke et al., 2010) and may therefore be found in a variety of fish host species. We found a significant correlation between metacercarial infection intensities and fish host size only for Posthodiplostomum metacercariae. This is likely because Posthodiplostomum metacercariae were found in high intensities in all fish sampled, whereas the sample size for Clinostomum and Diplostomum was only 10 individuals each. Correlations of parasite intensity in fishes generally constitute positive relationships with size and therefore age. For example, Janovy et al. (1997) reported significant positive correlations in terms of fish host size when analyzing ectoparasitic monogenean and larval trematode abundances. Similarly, Poulin (2000) 7 showed positive correlations in all of the parasite-host relationships tested, although these relationships were presumed to vary in their significance based on a multitude of biological factors and statistical groupings. Another potential explanation for the lack of significance with Clinostomum and Diplostomum metacercariae in terms of fish size is that the G. affinis used in the study only represent a portion of the size range of the population, which does not allow for assessing correlations between host size and infection intensity in an entire population. All individuals in this study were more than 35 mm in length. This size limit was chosen to increase the probability of obtaining infections that would allow for description of eyefluke microhabitat. Furthermore, this size limitation may have contributed to the sex bias of all females in our study. This situation poses questions regarding the sex ratio of the population sampled and the effects of infection on G. affinis males. The body size of male individuals varies depending on maturation relative to the breeding season with males maturing and achieving larger size later in the season (Hughes, 1985). Regardless, males generally tend to be smaller (18 to 30 mm) than females (30 to 65 mm; Kuntz, 1914). Thus, the size limit utilized in this study could explain the lack of male G. affinis individuals. Alternatively, the population sampled could be heavily female-biased, which can be common in poeciliid populations (Magurran, 2011). Further research would be necessary to determine the sex ratio in the sampled population, the effects of eyefluke infection on male individuals, as well as any potential differences between the sexes. Future studies may also expand on our results to reveal any spatial and temporal patterns of all helminth species parasitizing G. affinis. ACKNOWLEDGMENTS We thank C. Pyle and T.E. Barnes for their assistance in collection of G. affinis. We also thank the Office of Student Research and Creative Activities for funding this project and 8 providing an assistantship for SLG. Fish collection was conducted under Scientific Research Permit (SPR-0403-284) issued by Texas Parks & Wildlife. This project was conducted in accordance with the protocol and procedures established by the Animal Care and Use Committee at Tarleton State University (Approval #: 05-005-2014). LITERATURE CITED Aho, J. M., J. W. Camp, and G. W. Esch. 1982. Long-term studies on the population biology of Diplostomulum scheuringi in a thermally altered reservoir. Journal of Parasitology 68: 695-708. Al-Awadi, H. M., F. T. Mhaisen, and F. F. Al-Joborae. 2010. Parasitic fauna of fishes in Bahr Al-Najaf depression, mid Iraq. Bulletin of the Iraq National History Museum 11: 1-9. Barber, I. 2007. Parasites, behaviour and welfare in fish. Applied Animal Behaviour Science 104: 251-264. Bortz, B. M., G. E. Kenny, G. B. Pauley, and A. H. Bunt‐Milam. 1988. Prevalence of two site‐specific populations of Diplostomum spp. in eye infections of rainbow trout, Salmo gairdneri Richardson, from lakes in Washington State, USA. Journal of Fish Biology 33: 31-43. Brock, S., and W. F. Font. 2009. Helminths of the Western mosquitofish (Gambusia affinis) in Bayou Traverse, Louisiana, U.S.A. Comparative Parasitology 76: 210-221. Caffara, M., S. A. Locke, A. Gustinelli, D. J. Marcogliese, and M. L. Fioravanti. 2011. Morphological and molecular differentiation of Clinostomum complanatum and Clinostomum marginatum (Digenea: Clinostomidae) metacercariae and adults. Journal of Parasitology 97: 884-891. Chappell, L. H. 1995. The biology of diplostomatid eyeflukes of fishes. Journal of Helminthology 69: 97-101. 9 Cort, W. W. 1913. Notes on the trematode genus Clinostomum. Transactions of the American Microscopical Society 32: 169–182. Davis, J. R., and D. G. Huffman. 1977. A comparison of the helminth parasites of Gambusia affinis and Gambusia geiseri (Osteichthyes: Poeciliidae) from the upper San Marcos River. Southwestern Naturalist 22: 359–366. Dias, M. L. G. G., J. C. Eiras, M. H. Machado, G. T. R. Souza, and G. C. Pavanelli. 2003. The life cycle of Clinostomum complanatum Rudolphi, 1814 (Digenea, Clinostomidae) on the floodplain of the high Paraná River, Brazil. Parasitology Research 89: 506-508. Dwyer, W. P., and C. E. Smith. 1989. Metacercariae of Diplostomum spathaceum in the eyes of fishes from Yellowstone Lake, Wyoming. Journal of Wildlife Diseases 25: 126-129. Ferguson, M. S. 1943. Development of eye flukes of fishes in the lenses of frogs, turtles, birds, and mammals. Journal of Parasitology 29: 136-142. Grobbelaar, A., L. L. van As, J. G. van As, and H. J. B. Butler. 2015. Pathology of eyes and brain of fish infected with diplostomids, southern Africa. African Zoology 50: 181-186. Hoffman, G. L. 1960. Synopsis of Strigeoidea (Trematoda) of fishes and their life cycles. US Fish & Wildlife Publications 60: 439-466. Hoffman, G. L. 1999. Parasites of North American freshwater fishes, 2nd ed. Cornell University Press, Ithaca, 576 p. Hopkins, S. H. 1933. Note on the life history of Clinostomum marginatum (Trematoda). Transactions of the American Microscopical Society 52: 147–149. Hughes, A. L. 1985. Male size, mating success, and mating strategy in the mosquitofish Gambusia affinis (Poeciliidae). Behavioral Ecology and Sociobiology 17: 271-278. 10 Hughes, R. C. 1928. Studies on the trematode family Strigeidae (Holostomidae) No. IX. Neascus Van Cleavei (Agersborg). Transactions of the American Microscopical Society 47: 320-341. Humason, G. L. 1962. Animal tissue techniques, 2nd ed. W.H. Freeman & Company, San Francisco, California, 469 p. Hunter, G. W., and W. S. Hunter. 1940. Studies on the development of the metacercaria and the nature of the cyst of Posthodiplostomum minimum (MacCallum 1921) (Trematoda; Strigeata). Transactions of the American Microscopical Society 59: 52-63. Janovy Jr., J., and E. L. Hardin. 1987. Population dynamics of the parasites in Fundulus zebrinus in the Platte River of Nebraska. Journal of Parasitology 73: 689–696. Janovy Jr., J., and E. L. Hardin. 1988. Diversity of the parasite assemblage of Fundulus zebrinus in the Platte River of Nebraska. Journal of Parasitology 74: 207–213. Janovy Jr., J., S. D. Snyder, and R. E. Clopton. 1997. Evolutionary constraints on population structure: the parasites of Fundulus zebrinus (Pisces: Cyprinodontidae) in the south Platte River of Nebraska. Journal of Parasitology 83: 584–592. Kuntz, A. 1914. Notes on the habits, morphology of the reproductive organs, and embryology of the viviparous fish (Gambusia affinis). Bulletin of the United States Bureau of Fisheries 33: 177190. Locke, S. A., D. J. McLaughlin, and D. J. Marcogliese. 2010. DNA barcodes show cryptic diversity and a potential physiological basis for host specificity among Diplostomoidea (Platyhelminthes: Digenea) parasitizing freshwater fishes in the St. Lawrence River, Canada. Molecular Ecology 19: 2813-2827. Magurran, A. E. 2011. Sexual coercion. In Ecology and evolution of poeciliid fishes, J. P. Evans, A. Pilastro, and I. Schlupp (eds.). University of Chicago Press, Chicago, Illinois, p. 214-215. 11 McKeown, C. A., and S. W. B. Irwin. 1995. The life cycle stages of three Diplostomum species maintained in the laboratory. International Journal for Parasitology 25: 897-906. Meade, T. G., and C. A. Bedinger. 1972. Helminth parasitism in some species of fresh water fishes of eastern Texas. Southwestern Naturalist 16: 281–295. Mitchell, A. J., C. E., Smith, and G. L. Hoffman. 1982. Pathogenicity and histopathology of an unusually intense infection of white grubs (Posthodiplostomum m. minimum) in the fathead minnow (Pimephales promelas). Journal of Wildlife Diseases 18: 51-57. Palmieri, J. R., R. A. Heckmann, and R. S. Evans. 1976. Life cycle and incidence of Diplostomum spathaceum Rudolphi (1819) (Trematoda: Diplostomatidae) in Utah. Western North American Naturalist 36: 86-96. Poulin, R. 2000. Variation in the intraspecific relationship between fish length and intensity of parasitic infection: Biological and statistical causes. Journal of Fish Biology 56: 123-137. Rakauskas, V., and Č. Blaževičius. 2009. Distribution, prevalence and intensity of roach (Rutilus rutilus (Linnaeus, 1758)) parasites in inland waters of Lithuania in 2005–2008. Acta Zoologica Lituanica 19: 99-108. Rushton, W. 1937. Blindness in freshwater fish. Nature 140: 1014. Seppälä, O., A. Karvonen, and E. T. Valtonen. 2004. Parasite-induced change in host behaviour and susceptibility to predation in an eye fluke–fish interaction. Animal Behaviour 68: 257-263. Seppälä, O., A. Karvonen, and E. T. Valtonen. 2005. Manipulation of fish host by eye flukes in relation to cataract formation and parasite infectivity. Animal Behaviour 70: 889-894. Shariff, M., R. H. Richards, and C. Sommerville. 1980. The histopathology of acute and chronic infections of rainbow trout Salmo gairdneri Richardson with eye flukes, Diplostomum spp. Journal of Fish Diseases 3: 455–465. 12 Shoop, W. L. 1988. Trematode transmission patterns. Journal of Parasitology 74: 46-59. Szidat, L. 1969. Structure, development, and behaviour of new strigeatoid metacercariae from subtropical fishes of South America. Journal of the Fisheries Board of Canada 26: 753-786. Thomas, C., T. H. Bonner, and B. G. Whiteside. 2007. Freshwater fishes of Texas, 1st ed. Texas A&M University Press, College Station, Texas, 175 p. Wisniewski, W. L. 1958. The development cycle of Posthodiplostomum brevicaudatum (v. Nordmann, 1832) Kozicka, 1958. Acta Parasitologica Polonica 6: 251-272. Figure 1. Dissection of a Gambusia affinis eye infected with Posthodiplostomum metacercariae showing (A) posterior and (B) lateral views. Figure 2. Histological sections of Gambusia affinis: (A) an uninfected eye and (B) an infected eye showing an encysted Posthodiplostomum metacercaria in the choroid (S = sclera, C = choroid, PR = pigmented retina, M = metacercaria, CW = cyst wall). *Texas Invasive Species Institute, Sam Houston State University, Box 2506, Huntsville, Texas 77340. 13