Food in developed countries, is more than half a

Food borne diseases caused byparasitic organisms transmitted by fish pose major public health problems, andworldwide the number of people at risk, including those in developed countries,is more than half a billion.

Some of these parasites are highlypathogenic, and human infection is a result of the consumption of raw orundercooked fish infected by the parasites. Gizan region lies in the southwestcorner of Saudi Arabia. Gazan City is situated on the coast of the Red Seawhich represents a main source for most of the commercial fisheries of SaudiArabia. This area suffered from various sources of heavy metal pollution whichmay affect productivity of commercial fish. Since no data was available aboutzoonotic parasites infecting fishes in this area, the present study representsthe first parasitological report for different parasitic organisms inhabitingdifferent organs of fish which are considered as a major source of pathogenicdiseases after their infection to human. Taxonomy of these parasites will becarried out morphologically by light and scanning electron microscopy and bymolecular phylogeny. The distribution of pollutants and theirsubsequent effects and their health risks on marine organisms will be assessedby measuring the heavy metal concentrations in water, sediments and edibletissues of fish and detecting if these pollutants influence directly orindirectly the prevalence, intensity and susceptibility of fish for parasiticinfection. Since fish parasites closelyinteract with the metabolisms of the host and thoseparasites infra-populations can beaffected by changes of the host physiology and substancesaccumulated with the host´sfood.

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So, the present study aimed also for measuring the heavy metalsconcentrations in tissues of the recovered parasites to elucidate if theseparasites may serve as accumulation or biological indicators for waterpollution or not. This study will documented most of zoonotic parasites thatthreaten fish productivity in KSA and further studies will be supported tosolve this problem and to introduce the best way to the best health for fish consumers.Parasitic zoonosesFish as well as other aquatic and terrestrial animals may beaffected by infectious diseases and parasitization (Morsy et al.

2013;Tepe et al.2013; Shamsi andSuthar 2016; Cleveland et al.2017). Parasitic diseases are caused by animalparasites – protozoans, worms and crustaceans.

There arerelatively few studies on fish parasites inhabiting the Red Sea water at thesouthern part of Saudi Arabia, especially with regard to the parasite-hostrelationship. Previous studies focused primarily on classifying the parasitesof fish in the eastern Gulf (Saoud, 1986, 1988; Elnaffar et al. 1992; Al-Mathal, 2001; Al-Zubaidy and Mhaisen, 2011; Amin et al. 2015). The only study regardingfish zoonotic parasites in the southern region of Saudi Arabia was at Najranarea for metacercariae and nematode larvae with incomplete description.

No datawas available regarding fish parasites and their zoonoses from Gizan coasts.People handling andconsuming meat from fishes, such as fishermen and sellers, or people who comeinto contact with fish for research or during daily and monthly care ofaquarium and cage fishes are also at risk of contracting these zoonoticdiseases. Well-known zoonotic infectious agents transmitted to humans fromcaptive fishes are protozoon such asCryptosporidium and Giardia; Nematodes such as Capillariaphilippinensis, Dioctophymiasis renalae, Eustrongyloides, Gnathostomahispidum, G.

spinigerum, G. doloresi and G. nipponicum, Anisakis simplex,A.

typical, A. physeteris, Pseudoterranova decipiens and Contracaecumosculatum; Cestodes such as Diphyllobothrium latum, D. cordatum, D.pacifica, D. dendriticum, D. alascense, D.

lanceolatum, D. ursi, D. dalliae, D.nihonkaiense, D.

hians, D. cameroni, D. yonagoense, D. scoticum; Trematodessuch as Clonorchis sinensis, Opisthorchis viverrini, O. felineus,Heterophyes nocens, H. continua, H. heterophyes, H. dispar, Heterophyopiscontinua, Haplorchis taichui, H.

pumilo, Metagonimus yokogawai, M. takashii,Pyidiopsis summa, Diorchitrema falcatus, Stictodora fascatum, Centrocestusarmatus, Echinostoma hortense, E. cinetorchis, Echinochasmus japonicus,Paragonimus westermani, Nanophyetus salmincola (Brockelman et al. 1987,Campbell et al.

1988, Kuhn et al. 2017).  More than 1 billion people worldwide are infectedwith one or more species of gastrointestinal nematode parasites (WHO,2004) which cause a wide range of conditions from themild to the lethal. Humans can also be as accidental hosts for nematodeparasites that can’t progress their life cycles in humans but nevertheless cancause debilitating diseases directly or initiating immune hypersensitivitystates (Audicana and Kennedy, 2008).There is a tight connection between parasites and host, basedmainly on the fact that parasites cause the increase of fatalities in hosts(Lester, 2010; Kuhn et al.

2017). There are, however, cases in which abalance sets between parasites and hosts, but it depends on the intensity ofparasitization, immunity of the hosts to parasites and the general living conditionsof the hosts (Zaharia et al. 2012). In fish infested mainly by nematodelarvae, severe liver illnesses were reported, as well as the significantreduction of size, hemorrhages, reduction of the fat content in the liver,weight loss and reduction of the weight gain coefficient of fish (?o?oiu et al. 2013;Abdel-Gaber et al. 2016).

Aquatic pollution andits impact on parasitic infectionsOver the last few decades; aquaticpollution is still a problem in many freshwater and marine environments as itcauses negative effects for the health of the respective organisms (Nachevet al. 2015). Aquatic organisms such as fish accumulate metals toconcentrations many times higher than those in water (Al-Sultany, 2014).Heavymetal contamination may have devastating effects on the ecological balance ofthe recipient environment and a diversity of aquatic organisms. The dischargeof large amounts of metal-contaminated waste water, industries bearing heavymetals, such as cadmium, chromium, copper, nickal, arsenic, lead and zinc isconsidered the most hazardous among the chemical-intensive industries (Mason etal. 2002; Awadallahand Salem 2015). Because of their high solubility in the aquaticenvironments, heavy metals can be absorbed by living organisms (Saeed and Shaker,2008). Once they enter the food chain, large concentrations of heavy metals mayaccumulate in the human body (Mustapha and Lawal, 2014).

If the metals areingested above the permitted concentration, they can cause serious healthdisorders or reduce mental and central nervous function, lower energy levelsand damage to blood composition, lung, kidney, liver and other vital organs(Tunali et al. 2006). The presence of these metals in water streams andmarine water systems causes a significant health threat to the aquaticcommunity the most common being damage of the gill of the fish (Tunali etal. 2006, Awadallahand Salem 2015).Mason et al. (2002) stated that metal ions can beincorporated into food chains and concentrated in aquatic organisms to a levelthat affects their physiological state. Of the effective toxic pollutants arethe heavy metals which have drastic environmental impact on all organisms(Saeed and Shaker, 2008).

Trace metals such as Zn, Cu and Fe play a biochemicalrole in the life processes of all aquatic plants and animals but in traceamount, their increase and accumulation over the permissive limits may produceadverse and hazardous effects on organism’s health (Ba?yi?it and Tekin-Özan2013; Mariné Oliveira et al. 2016).It is also possible that environmental toxicants mayincrease the susceptibility of aquatic animals to various diseases byinterfering with the normal functioning of their immune, reproductive anddevelopmental processes (Al-Sultany 2014).The frequency of life threatening infections caused by consumption of untreatedwater has increased worldwide and is becoming an important cause of mortalityin developing countries (Dans et al.

2014).Oceans are largely contaminated with industrial pollutants like Hg, Pb, As, Cd,Zn and Cu, which become concentrated in the flesh of the fish (Otachi et al.,2014). These pollutants might promote increased parasitism in aquatic animals,especially in fish by impairing the host’s immune response (Khan and Thulin,1991; Paller et al. 2016).

It is clear that several fish diseases and abnormalities occur in highlypolluted areas. Pollutants might influence, directly or indirectly, theprevalence, intensity and pathogencity of parasites. There is a developinginformation about the possible relationship of parasitism and pollution (Morsy etal. 2012).Parasitesas biological indicators of pollutionSeveral metazoan fish parasites have been successfullyapplied as biological indicators for pollution. Nematodes of the Ascaridoidea(families: Anisakidae) have been recorded worldwide naturally parasitizingapproximately 200 fish species (Morsy etal. 2015), 25 cephalopod species (Mazhar et al.

2014; Gilbert and  Avenant-Oldewage, 2017),marine mammals, and humans can also become accidental hosts by ingesting fishinfected with third-stage larvae (Javed and Usmani 2014; Paller et al. 2016).Sures et al. (2004) stated that certain parasites,particularly intestinal parasites of fish can accumulate heavy metals atconcentrations that are orders of magnitude higher than those in the hosttissues or the environment. Anisakis simplex can accumulate heavy metalas pb and cu by atomic adsorption spectrometry to a level far in excess tothose in their host tissues.

Such phenomenon of conspicuous metal accumulation makes fishparasites could be applied to environmental monitoring. So, intestinalparasites have thus gained attention from ecologists and environmentaltoxicologists within the last decade (Dans etal. 2014; Shamsi andSuthar, 2015).Fish parasites closelyinteract with the metabolisms of the host. Thus, parasiteinfra-populations can be affectedby changes of the host physiology and substances accumulated with the host´s food. In such cases, somefish parasites can accumulate pollutants in a much higher concentration astheir host organisms, and serve as accumulation indicators (Nachev et al. 2013; Lacerda et al. 2017).

For example, some acanthocephalans specifically accumulatecertain heavy metals in greater amounts than their host, and can be used asaccumulation indicators of heavy metal pollution (Javed and Usmani, 2014; Gilbert and  Avenant-Oldewage, 2017).Adults of the acanthocephalans Pomphorhynchus laevis and Paratenuisentisambiguus accumulate lead and cadmium in a greater amount than theirhosts (Anguilla anguilla, Leuciscus cephalus, Perca fluviatilis).However, there is some dependence on the parasitic life cycle stage.Sures and Reimann (2003) compared the heavy metalconcentration of the acanthocephalan Aspersentis megarhynchus with themuscle of the antarctic rock cod Notothenia coriiceps. Most of theelements were found in significantly higher concentrations in theacanthocephalan than in the muscle of its host. Levels of Ag, Co and Ni in the muscleof N. coriiceps were even below the detection limit, and were only foundin the worm.

Othermetals commonly associated with human activities (e.g. Pb, Cd, Cu) wereaccumulated to a high degree in the parasite, demonstrating that pollutants ofanthropogenic origin are dispersed within this remote, fairly unpollutedenvironment. Cestodes can also be used as accumulation indicators (Sures et al.2017).

For example, lead and cadmium is found in significantlyhigher concentrations in the tissues of the cestode Monobothrium wagenerithan in their fish host tissues (Tinca tinca from Ruhr River).The marine cestode Bothriocephalus scorpii (Cestoda) from Scophthalmusmaximus (Gdansk Bay) was found to accumulate these heavy metals especiallyin the posterior part of the proglottids, while the anterior part revealed thesame amounts of the heavy metals as the fish host tissues.) Fish collection Fish samples were collected from fishermen at the fishlanding center of Red Sea, Jizan or from a fish market nearby. They Will betransferred in a cooler packed with ice blocks in order to maintain thefreshness and later brought to the Parasitology laboratory, Biology department,College of Science, King Khalid University where Fish will be identified andclassified according to Froese and Pauly (2017). Each fish specimen will beopened up dorso-ventrally and the internal organs will be examined. The entiredigestive system will be removed and placed in a Petri dish filled with aphysiological saline 0.65%. The gut will be divided into sections and eachsection will be examined for cestode, nematode and acanthocephalan parasites.

Gonads, liver, heart and gall bladder and the pericardial cavity were examinedfor parasites. The stomach and intestine were dissected, opened longitudinallyand examined for the presence of any trematode parasites under a stereomicroscope III) Morphologicalstudya.Light microscopyRelaxationis the first important step during examination of trematodes, and nematodeparasites, without relaxation these parasites become strongly contracted andcoiled thus making subsequent examination more difficult (Li et al. 2011; Smales, 2014). Cestodes willbe placed in between two glass slides with a drop of 70% alcohol and subjectedto a pressure until relaxed. Care is usually taken to avoid the use of strongpressure particularly on delicate parasites.

Nematodes will be transferred intoa clean saline solution or warm 70% ethyl alcohol for few minutes in therefrigerator till relaxation. Cestodes will be fixed in 4% formalin and thetime of fixation depends on the size and thickness of parasites, being 2-4hours for small parasites and 12–24 hours for larger ones. After fixation, thecollected samples will be washed in distilled water for 15 minutes to removethe excess fixative and then processed to staining which is carried out byusing acetic acid alum carmine for 5-10 minutes (Artis et al.,2012).  The specimens were then clearedin xylene, then mounted in Canada balsam, covered with cover glass and left todry in an oven at 40C.

Hot70% ethyl alcohol was the best fixative for nematodes. Following 12 hours, thefixative will be replaced by Lactophenol which is poured to cover the preservednematodes and left for at least 24 hours to allow cleaning.b.Scanning electronmicroscopyTo reveal the ultrastructures ofthe recovered parasites, the specimens will be fixed in 3 % bufferedgluteraldehyde, washed in cacodylate buffer, and dehydrated in a graded seriesof ethanol alcohol (10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90% and 100 (%. After passing through an ascending seriesof Genosolv-D, they were processed in a critical point drier “Bomer -900” with freon 13, and sputter -coated with gold – palladium in aTechnics Hummer V, and examined with an EtecAutoscan at 20 KV Jeol scanning EM.IV) Determination of residual heavy metals in water, fishand parasites tissues:a.

WaterTrace elements inwater were measured using flame atomic absorption spectrophotometer (Thermo ScientificICE 3300, UK) with double beam and deuterium background corrector according toAPHA (2005).b.TissuesSamples from parasites and infected fish were dried separately at 105oCfor 12 hours, burned in a muffle furnace at 550oC for 16 hours,acid-digested (HNO3, H2SO4) and diluted withde-ionized water to known volume (25ml) using the dry-ashing procedure proposedby Hseu (2004) and Gilbertand  Avenant-Oldewage (2017). Analytical blanks were run in thesame way as the samples, and concentrations were determined using standardsolutions prepared in the same acid matrix.

Standards for instrument calibration were prepared on the basis ofmono-element certified reference solution inductively coupled plasma standard (Merck).Standard reference material (National Institute of Standards and TechnologyNIST, USA) was used to validate analysis, and the metal recoveries rangedbetween 90 and 110 %.c.Water andsediment samples Water and sediment samples will be collected from the fishlanding spot of Red sea at Jizan coast. Water samples will be collected incleaned plastic bottles following filtration through Whatmann filter paper andkept in a refrigerator at 4°C with addition of 2 mg/l HNO3 before laboratoryanalysis  (Mastoi et al.

1997). Sediment samples will be collected with astainless steel Ekman grab sampler, which allows free water through the samplerduring descent penetration. The sediment samples will be collected from thesame sampling spot. They will be air dried for several days over Pyrex petridishes and then samples will be dried in an oven at 105°C in laboratory.V)Molecularanalysisa.DNAextraction AND PCR amplification:Total genomic DNA (gDNA) was extracted from ethanolpreserved specimensusing the DNeasy tissue kit (QIAGEN) following the manufacturer’s instructions.

PCR was performed with a totalvolume of 20 ?l consisting of approximately 10 ng of DNA, 5 ?l of 5x MyTaqReaction Buffer (Bioline), 0.75 ?l of each primer (10 pmols) and 0.25 ?l of TaqDNA polymerase (Bioline MyTaq™ DNA Polymerase), made up to 20 ?l withInvitrogen™ ultraPURE™ distilled water. Amplification was carried out on a MJResearch PTC-150 thermocycler.PCR amplicons were either gel-excised using a QIAquickTM GelExtraction Kit (QIAGEN) orpurified directly using QIAquickTM PCR Purification Kit (QIAGEN)following the standard manufacturer-recommended protocol.

Cycle-sequencing fromboth strands was carriedout on an ABI 3730 DNA Analyser, Big Dye version 1.1. using ABI BigDyeTMchemistry.b.Phylogenetic analysisSequence identity for the recovered data was checked usingthe Basic Local Alignment Search Tool (BLAST) (www. Thesequence trimming for the congeneric species recovered was carried out by Bioeditv 7.2.5, sequence alignment was done by CLUSTAL W v2 and the phylogenetic treeswere construced using MEGA 6 program.

Polystoma was employed as an out-group.VI)Histopathological StudyForhistopathological study, liver, gills, intestinal tissues from infected and noninfected fish  will be preserved in 10%formal saline, dehydrated in aseries of alcohols, cleared in xylol, embedded in paraffin wax and sectioned bya microtome at 6 µm thick were made using Leitz Wetzlar® basesledge microtome. Tissue sections will be stained using Haematoxylin and Eosin(H&E) stain as described by Bancroft and Cook (1984). They will beplaced haematoxylin stain for 10 minutes and washed in tap water. Sections willbe placed in 1% acid alcohol for a minute and washed by tap water. “The blue”sections will be washed by tap water and then placed in eosin for 2 minutes andwashed by tap water, dehydrated through grades of alcohol (70%, 90% and 100%),cleared with xylene and mounted using DPX mountant then examined and photographed by a Zeiss research photomicroscope(Corleton, 1967). VII)Statistical AnalysesStatistical analyses will be carried out using SPSS v.

15 software. All data will be expressed as means ± standard error of mean(SEM).