Food borne diseases caused by
parasitic organisms transmitted by fish pose major public health problems, and
worldwide the number of people at risk, including those in developed countries,
is more than half a billion. Some of these parasites are highly
pathogenic, and human infection is a result of the consumption of raw or
undercooked fish infected by the parasites. Gizan region lies in the southwest
corner of Saudi Arabia. Gazan City is situated on the coast of the Red Sea
which represents a main source for most of the commercial fisheries of Saudi
Arabia. This area suffered from various sources of heavy metal pollution which
may affect productivity of commercial fish. Since no data was available about
zoonotic parasites infecting fishes in this area, the present study represents
the first parasitological report for different parasitic organisms inhabiting
different organs of fish which are considered as a major source of pathogenic
diseases after their infection to human. Taxonomy of these parasites will be
carried out morphologically by light and scanning electron microscopy and by
molecular phylogeny. The distribution of pollutants and their
subsequent effects and their health risks on marine organisms will be assessed
by measuring the heavy metal concentrations in water, sediments and edible
tissues of fish and detecting if these pollutants influence directly or
indirectly the prevalence, intensity and susceptibility of fish for parasitic
infection. Since fish parasites closely
interact with the metabolisms of the host and those
parasites infra-populations can be
affected by changes of the host physiology and substances
accumulated with the host´s
food. So, the present study aimed also for measuring the heavy metals
concentrations in tissues of the recovered parasites to elucidate if these
parasites may serve as accumulation or biological indicators for water
pollution or not. This study will documented most of zoonotic parasites that
threaten fish productivity in KSA and further studies will be supported to
solve this problem and to introduce the best way to the best health for fish consumers.
Fish as well as other aquatic and terrestrial animals may be
affected by infectious diseases and parasitization (Morsy et al. 2013;
Tepe et al.
2013; Shamsi and
Suthar 2016; Cleveland et al.
2017). Parasitic diseases are caused by animal
parasites – protozoans, worms and crustaceans. There are
relatively few studies on fish parasites inhabiting the Red Sea water at the
southern part of Saudi Arabia, especially with regard to the parasite-host
relationship. Previous studies focused primarily on classifying the parasites
of 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 regarding
fish zoonotic parasites in the southern region of Saudi Arabia was at Najran
area for metacercariae and nematode larvae with incomplete description. No data
was available regarding fish parasites and their zoonoses from Gizan coasts.
People handling and
consuming meat from fishes, such as fishermen and sellers, or people who come
into contact with fish for research or during daily and monthly care of
aquarium and cage fishes are also at risk of contracting these zoonotic
diseases. Well-known zoonotic infectious agents transmitted to humans from
captive fishes are protozoon such as
Cryptosporidium and Giardia; Nematodes such as Capillaria
philippinensis, Dioctophymiasis renalae, Eustrongyloides, Gnathostoma
hispidum, G. spinigerum, G. doloresi and G. nipponicum, Anisakis simplex,
A. typical, A. physeteris, Pseudoterranova decipiens and Contracaecum
osculatum; 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; Trematodes
such as Clonorchis sinensis, Opisthorchis viverrini, O. felineus,
Heterophyes nocens, H. continua, H. heterophyes, H. dispar, Heterophyopis
continua, Haplorchis taichui, H. pumilo, Metagonimus yokogawai, M. takashii,
Pyidiopsis summa, Diorchitrema falcatus, Stictodora fascatum, Centrocestus
armatus, 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 infected
with one or more species of gastrointestinal nematode parasites (WHO,
2004) which cause a wide range of conditions from the
mild to the lethal. Humans can also be as accidental hosts for nematode
parasites that can’t progress their life cycles in humans but nevertheless can
cause debilitating diseases directly or initiating immune hypersensitivity
states (Audicana and Kennedy, 2008).
There is a tight connection between parasites and host, based
mainly on the fact that parasites cause the increase of fatalities in hosts
(Lester, 2010; Kuhn et al. 2017). There are, however, cases in which a
balance sets between parasites and hosts, but it depends on the intensity of
parasitization, immunity of the hosts to parasites and the general living conditions
of the hosts (Zaharia et al. 2012). In fish infested mainly by nematode
larvae, severe liver illnesses were reported, as well as the significant
reduction 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 and
its impact on parasitic infections
Over the last few decades; aquatic
pollution is still a problem in many freshwater and marine environments as it
causes negative effects for the health of the respective organisms (Nachev
et al. 2015). Aquatic organisms such as fish accumulate metals to
concentrations many times higher than those in water (Al-Sultany, 2014).Heavy
metal contamination may have devastating effects on the ecological balance of
the recipient environment and a diversity of aquatic organisms. The discharge
of large amounts of metal-contaminated waste water, industries bearing heavy
metals, such as cadmium, chromium, copper, nickal, arsenic, lead and zinc is
considered the most hazardous among the chemical-intensive industries (Mason et
al. 2002; Awadallah
and Salem 2015). Because of their high solubility in the aquatic
environments, heavy metals can be absorbed by living organisms (Saeed and Shaker,
2008). Once they enter the food chain, large concentrations of heavy metals may
accumulate in the human body (Mustapha and Lawal, 2014). If the metals are
ingested above the permitted concentration, they can cause serious health
disorders or reduce mental and central nervous function, lower energy levels
and damage to blood composition, lung, kidney, liver and other vital organs
(Tunali et al. 2006). The presence of these metals in water streams and
marine water systems causes a significant health threat to the aquatic
community the most common being damage of the gill of the fish (Tunali et
al. 2006, Awadallah
and Salem 2015).
Mason et al. (2002) stated that metal ions can be
incorporated into food chains and concentrated in aquatic organisms to a level
that affects their physiological state. Of the effective toxic pollutants are
the heavy metals which have drastic environmental impact on all organisms
(Saeed and Shaker, 2008). Trace metals such as Zn, Cu and Fe play a biochemical
role in the life processes of all aquatic plants and animals but in trace
amount, their increase and accumulation over the permissive limits may produce
adverse and hazardous effects on organism’s health (Ba?yi?it and Tekin-Özan
2013; Mariné Oliveira et al. 2016).
It is also possible that environmental toxicants may
increase the susceptibility of aquatic animals to various diseases by
interfering with the normal functioning of their immune, reproductive and
developmental processes (Al-Sultany 2014).
The frequency of life threatening infections caused by consumption of untreated
water has increased worldwide and is becoming an important cause of mortality
in 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 highly
polluted areas. Pollutants might influence, directly or indirectly, the
prevalence, intensity and pathogencity of parasites. There is a developing
information about the possible relationship of parasitism and pollution (Morsy et
as biological indicators of pollution
Several metazoan fish parasites have been successfully
applied as biological indicators for pollution. Nematodes of the Ascaridoidea
(families: Anisakidae) have been recorded worldwide naturally parasitizing
approximately 200 fish species (Morsy et
al. 2015), 25 cephalopod species (Mazhar et al. 2014; Gilbert and Avenant-Oldewage, 2017),
marine mammals, and humans can also become accidental hosts by ingesting fish
infected 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 at
concentrations that are orders of magnitude higher than those in the host
tissues or the environment. Anisakis simplex can accumulate heavy metal
as pb and cu by atomic adsorption spectrometry to a level far in excess to
those in their host tissues.
Such phenomenon of conspicuous metal accumulation makes fish
parasites could be applied to environmental monitoring. So, intestinal
parasites have thus gained attention from ecologists and environmental
toxicologists within the last decade (Dans et
al. 2014; Shamsi and
Fish parasites closely
interact with the metabolisms of the host. Thus, parasite
infra-populations can be affected
by changes of the host physiology and substances accumulated with the host´s food. In such cases, some
fish parasites can accumulate pollutants in a much higher concentration as
their host organisms, and serve as accumulation indicators (Nachev et al. 2013; Lacerda et al. 2017).
For example, some acanthocephalans specifically accumulate
certain heavy metals in greater amounts than their host, and can be used as
accumulation indicators of heavy metal pollution (Javed and Usmani, 2014; Gilbert and Avenant-Oldewage, 2017).
Adults of the acanthocephalans Pomphorhynchus laevis and Paratenuisentis
ambiguus accumulate lead and cadmium in a greater amount than their
hosts (Anguilla anguilla, Leuciscus cephalus, Perca fluviatilis).
However, there is some dependence on the parasitic life cycle stage.
Sures and Reimann (2003) compared the heavy metal
concentration of the acanthocephalan Aspersentis megarhynchus with the
muscle of the antarctic rock cod Notothenia coriiceps. Most of the
elements were found in significantly higher concentrations in the
acanthocephalan than in the muscle of its host. Levels of Ag, Co and Ni in the muscle
of N. coriiceps were even below the detection limit, and were only found
in the worm.
metals commonly associated with human activities (e.g. Pb, Cd, Cu) were
accumulated to a high degree in the parasite, demonstrating that pollutants of
anthropogenic origin are dispersed within this remote, fairly unpolluted
environment. Cestodes can also be used as accumulation indicators (Sures et al.
2017). For example, lead and cadmium is found in significantly
higher concentrations in the tissues of the cestode Monobothrium wageneri
than in their fish host tissues (Tinca tinca from Ruhr River).
The marine cestode Bothriocephalus scorpii (Cestoda) from Scophthalmus
maximus (Gdansk Bay) was found to accumulate these heavy metals especially
in the posterior part of the proglottids, while the anterior part revealed the
same amounts of the heavy metals as the fish host tissues.
) Fish collection
Fish samples were collected from fishermen at the fish
landing center of Red Sea, Jizan or from a fish market nearby. They Will be
transferred in a cooler packed with ice blocks in order to maintain the
freshness and later brought to the Parasitology laboratory, Biology department,
College of Science, King Khalid University where Fish will be identified and
classified according to Froese and Pauly (2017). Each fish specimen will be
opened up dorso-ventrally and the internal organs will be examined. The entire
digestive system will be removed and placed in a Petri dish filled with a
physiological saline 0.65%. The gut will be divided into sections and each
section will be examined for cestode, nematode and acanthocephalan parasites.
Gonads, liver, heart and gall bladder and the pericardial cavity were examined
for parasites. The stomach and intestine were dissected, opened longitudinally
and examined for the presence of any trematode parasites under a stereo
is the first important step during examination of trematodes, and nematode
parasites, without relaxation these parasites become strongly contracted and
coiled thus making subsequent examination more difficult (Li et al. 2011; Smales, 2014). Cestodes will
be placed in between two glass slides with a drop of 70% alcohol and subjected
to a pressure until relaxed. Care is usually taken to avoid the use of strong
pressure particularly on delicate parasites. Nematodes will be transferred into
a clean saline solution or warm 70% ethyl alcohol for few minutes in the
refrigerator till relaxation. Cestodes will be fixed in 4% formalin and the
time of fixation depends on the size and thickness of parasites, being 2-4
hours for small parasites and 12–24 hours for larger ones. After fixation, the
collected samples will be washed in distilled water for 15 minutes to remove
the excess fixative and then processed to staining which is carried out by
using acetic acid alum carmine for 5-10 minutes (Artis et al.,
2012). The specimens were then cleared
in xylene, then mounted in Canada balsam, covered with cover glass and left to
dry in an oven at 40C.
70% ethyl alcohol was the best fixative for nematodes. Following 12 hours, the
fixative will be replaced by Lactophenol which is poured to cover the preserved
nematodes and left for at least 24 hours to allow cleaning.
To reveal the ultrastructures of
the recovered parasites, the specimens will be fixed in 3 % buffered
gluteraldehyde, washed in cacodylate buffer, and dehydrated in a graded series
of ethanol alcohol (10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90% and 100 (%. After passing through an ascending series
of Genosolv-D, they were processed in a critical point drier “Bomer –
900” with freon 13, and sputter -coated with gold – palladium in a
Technics Hummer V, and examined with an EtecAutoscan at 20 KV Jeol scanning EM.
IV) Determination of residual heavy metals in water, fish
and parasites tissues:
Trace elements in
water were measured using flame atomic absorption spectrophotometer (Thermo Scientific
ICE 3300, UK) with double beam and deuterium background corrector according to
Samples from parasites and infected fish were dried separately at 105oC
for 12 hours, burned in a muffle furnace at 550oC for 16 hours,
acid-digested (HNO3, H2SO4) and diluted with
de-ionized water to known volume (25ml) using the dry-ashing procedure proposed
by Hseu (2004) and Gilbert
and Avenant-Oldewage (2017). Analytical blanks were run in the
same way as the samples, and concentrations were determined using standard
solutions prepared in the same acid matrix.
Standards for instrument calibration were prepared on the basis of
mono-element certified reference solution inductively coupled plasma standard (Merck).
Standard reference material (National Institute of Standards and Technology
NIST, USA) was used to validate analysis, and the metal recoveries ranged
between 90 and 110 %.
Water and sediment samples will be collected from the fish
landing spot of Red sea at Jizan coast. Water samples will be collected in
cleaned plastic bottles following filtration through Whatmann filter paper and
kept in a refrigerator at 4°C with addition of 2 mg/l HNO3 before laboratory
analysis (Mastoi et al. 1997). Sediment samples will be collected with a
stainless steel Ekman grab sampler, which allows free water through the sampler
during descent penetration. The sediment samples will be collected from the
same sampling spot. They will be air dried for several days over Pyrex petri
dishes and then samples will be dried in an oven at 105°C in laboratory.
extraction AND PCR amplification:
Total genomic DNA (gDNA) was extracted from ethanol
using the DNeasy tissue kit (QIAGEN) following the manufacturer’s instructions. PCR was performed with a total
volume of 20 ?l consisting of approximately 10 ng of DNA, 5 ?l of 5x MyTaq
Reaction Buffer (Bioline), 0.75 ?l of each primer (10 pmols) and 0.25 ?l of Taq
DNA polymerase (Bioline MyTaq™ DNA Polymerase), made up to 20 ?l with
Invitrogen™ ultraPURE™ distilled water. Amplification was carried out on a MJ
Research PTC-150 thermocycler.
PCR amplicons were either gel-excised using a QIAquickTM Gel
Extraction Kit (QIAGEN) or
purified directly using QIAquickTM PCR Purification Kit (QIAGEN)
following the standard manufacturer-recommended protocol. Cycle-sequencing from
both strands was carried
out on an ABI 3730 DNA Analyser, Big Dye version 1.1. using ABI BigDyeTM
Sequence identity for the recovered data was checked using
the Basic Local Alignment Search Tool (BLAST) (www.ncbi.nih.gov/BLAST/). The
sequence trimming for the congeneric species recovered was carried out by Bioedit
v 7.2.5, sequence alignment was done by CLUSTAL W v2 and the phylogenetic trees
were construced using MEGA 6 program. Polystoma was employed as an out-group.
histopathological study, liver, gills, intestinal tissues from infected and non
infected fish will be preserved in 10%
formal saline, dehydrated in a
series of alcohols, cleared in xylol, embedded in paraffin wax and sectioned by
a microtome at 6 µm thick were made using Leitz Wetzlar® base
sledge microtome. Tissue sections will be stained using Haematoxylin and Eosin
(H&E) stain as described by Bancroft and Cook (1984). They will be
placed haematoxylin stain for 10 minutes and washed in tap water. Sections will
be 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 and
washed 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
Statistical analyses will be carried out using SPSS v.
15 software. All data will be expressed as means ± standard error of mean