**About the Editor**

#### **Snjezana Zrnˇ ˇ ci´c**

Snjezana Zrn ˇ ciˇ c, PhD, Doctor of Veterinary Medicine, works in the National Reference ´ Laboratory for Fish and Shellfish Diseases and has more than 30 years of experience in the pathology of aquatic organisms. Her general interests lie in the study of diseases among farmed marine fish and bivalve mollusks. She has participated in several research projects on the subject. Currently, she is serving as the General Secretary of the European Association of Fish Pathologists (EAFP). Furthermore, she participated in organization of conferences organized by the EAFP and also as a program co-chair of the European Aquaculture Society conference. She served as external expert for the UN FAO in the field of diagnostics. During her career she published more than 50 peer reviewed papers and more than 60 conferences paper, chapters in five books and edited three thematic manuals. She mentored master and doctoral thesis and participated in several working groups for advices to decision making bodies nationally and on European level.

## *Editorial* **Microbial Diseases of Marine Organisms**

**Snježana Zrnˇci´c**

Laboratory for Fish Pathology, Croatian Veterinary Institute, 10000 Zagreb, Croatia; zrncic@veinst.hr

Healthy oceans and marine environments provide critical life support functions upon which human health and well-being depend [1]. Multiple benefits are derived from marine and coastal ecosystem at local, regional and global scales, ranging from pollution control, storm protection, shoreline stabilization and habitats for species to climate mitigation and food provisioning.

At present, we are facing increasing threats to the sustainability of the marine environment caused by industrialization, tourism, marine traffic and global warming. Marine organisms, whether they are prey or predators in the food chain, are very important members of the marine ecosystem. They form associations with microorganisms, including protists, bacteria, fungi and viruses, and their relationships are mostly mutually beneficial symbiont systems. However, environmental changes induced by anthropogenic impact and climate changes may alter the symbiont relationship, and microorganisms could influence the health, physiology, behaviour and ecology of marine animals [2]. Over time, many different pathogenic microorganisms have been reported as causes of mortality of fish, molluscs, crustaceans and other marine organisms. These disease outbreaks may lead to a large decline in the host population, resulting in the endangerment of the affected species and causing an imbalance in the marine environment.

This Special Issue, titled "Microbial Diseases of Marine Organisms", is conceived as a contribution to the knowledge of the deleterious impacts of microorganisms on marine fish, molluscs, crustaceans, cetaceans or other organisms. Data presented in this Special Issue provide valuable information which could be used for establishing strategies for disease mitigation and control and consequently contribute to preserving the sustainability of the marine ecosystem. A variety of hosts such as corals [3]; different species of endemic, farmed or commercial bivalve mollusc species [4–6]; decapods [7]; and wild fish in public aquaria [8] and aquaculture facilities [9] endangered by different microorganisms is presented in this issue.

We have learnt that coral reefs are among the most biodiverse biological systems on Earth [3]. They are classified as marine invertebrates and filter the surrounding food and other particles in seawater, including pathogens such as viruses. Viruses act as both pathogens and symbionts for metazoans. Marine viruses, which are abundant in the ocean, are mostly single- or double-stranded DNA and single- or double-stranded RNA viruses. These findings were obtained using advanced identification methodologies to detect the presence of viruses in coral reefs; PCR analyses, metagenomic analyses, transcriptomic analyses; and electron microscopy. The review paper in this issue discusses and presents the discovery of different viruses in the marine environment and their hosts, the viral diversity in coral and also the presence of viruses in corallivorous fish communities in reef ecosystems. The detection methods were described, as well as the occurrence of marine viral communities in marine sponges. It was concluded that marine viral communities play a crucial role in biogeochemical cycles in the ocean, indirectly and directly. These viral communities induce mortalities and diseases in the reef ecosystem through abiotic and biotic factors, which cause disturbances in the symbiotic relationship between the coral and their surrounding hosts. The review emphasizes marine viruses from the ocean, coralassociated viruses and marine sponge and coral fish viruses in reef ecosystems, examining

**Citation:** Zrnˇci´c, S. Microbial Diseases of Marine Organisms. *J. Mar. Sci. Eng.* **2022**, *10*, 1682. https:// doi.org/10.3390/jmse10111682

Received: 1 November 2022 Accepted: 2 November 2022 Published: 7 November 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

previous research via traditional methods to modern advanced approaches. The findings of this review have enhanced our understanding of coral–virus interactions and enriched our understanding of reef-associated virus interactions and the diversity of viral communities in marine environments.

Other studied species were red claw crayfish (*Cherax quadricarinatus*) and red swamp crayfish (*Procambarus clarkia*), important for food and ornamental purposes, imported to South Korea from China and Indonesia [7]. The research comprised PCR testing to detect infectious hypodermal and hematopoietic necrosis virus (IHHNV or Decapod penstylhamaparvovirus 1). IHHNV was detected in tissue samples pooled from nine out of ten batches of red claw crayfish imported from Indonesia. Phylogenetic analysis of PCR amplicons from representative pools clustered the IHHNV strain with infectious-type II sequences commonly detected in Southeast Asian countries, rather than with type III strains detected previously in white leg shrimp (*Penaeus vannamei*) cultured in South Korea. IHHNV DNA was detected most frequently in the muscle, followed by hepatopancreas and gill tissues, suggesting that red claw crayfish could be a potential carrier of the virus. It was concluded that transboundary movements can cause significant environmental disturbance and provide opportunities for the inadvertent translocation of disease-causing pathogens to new locations. The detection of IHHNV infection in *C. quadricarinatus* by identifying the viral DNA in tissue samples of the commodity imported into South Korea as the IHHNV type II strain indicates that red claw crayfish could be a potential carrier of the infectious IHHNV, posing a threat to the cultured and wild population of crustaceans in South Korea.

The most represented marine species of this issue were bivalve molluscs: from farmed European flat oysters (*Ostrea edulis*) [5], followed by farmed and free-living Portuguese oysters (*Crassostrea angulata*) and Pacific oysters (*Crassostrea gigas*) [6], to the critically endangered noble pen shell (*Pinna nobilis*) endemic to the Mediterranean area [4]. Research on flat oysters describes the first occurrence and molecular identification and epidemiology of parasites from the genera *Bonamia*, phylogenetically positioned into a clade microcell within the genus *Haplosporidia* in a farming area along the Croatian Adriatic Coast [5]. PCR analysis and sequencing for SSU rDNA gene and BLAST analysis confirmed infection with *Bonamia exitiosa*. Although prevalence in a five-year period ranged from 3.3 to 20% at the different sites, there were no mortalities reported from the infected sites, and it seemed that infection of flat oysters with *B. exitiosa* did not affect their health. Attempt to prove the Pacific oyster as a putative vector of the parasite failed. The phylogenetic analysis did not disclose any information on the source of *B. exitiosa* origin. Since the Croatian isolate showed 100% similarity to previously sequenced isolates from Chile or Australia based on the SSU rDNA gene, the sequencing of additional genes or the whole genome should be carried out to provide us with more details on the phylogeny of the Croatian isolates. More comprehensive molecular studies of the *B. exitiosa*, together with an investigation of the natural population of *O. stentina*, which are susceptible species from production areas and natural beds along the Eastern Adriatic coast, could confirm the natural-historical origin of the parasite *B. exitiosa.* Other studied bivalve molluscs species are species nationally important for Portugal; Portuguese oyster (*Crassostrea angulata*) and Pacific oyster (*Crassostrea gigas*) from four distinctive areas in Portugal were studied to evaluate their sanitary status [6]. Collected Pacific oyster populations were cultivated in a strong ocean-influenced environment, and Portuguese oyster populations were cultivated in wild beds. The histopathological examination of both oyster species revealed the presence of parasites in gills, mantle epithelium, digestive gland tubules and connective tissue, with a moderate prevalence. In both populations, hemocytosis was observed in the connective tissue, oedema and metaplasia in the digestive gland and necrosis in the tissues. In wild populations from the Sado and Mira estuaries, the prevalence of mud blisters and gill lesions was higher than from populations produced on 0.50 m tables from mudflats. It was concluded that diseases are important risk factors which are caused, in many cases, by non-compliance with basic management rules, namely, the level of animal load in production areas, the length of time the bivalve molluscs remain in these areas, and

the introduction of seeds of unknown origin. Effective biosecurity measures and correct and early diagnostic techniques are essential to control pathogenic agents. In the production areas of the Aveiro and Alvor lagoons, these measures are implemented by producers and by the authorities. Producers understand that it is essential to prevent mortalities in bivalve mollusc populations, namely, avoiding overcrowding and preventing diseases while in Sado and Mira estuaries, wild populations were proven to be more susceptible to lesions than oysters produced on tables under the supervision of producers. Differently from previous research, another study is dedicated to the noble pen shell (*Pinna nobilis*), the largest bivalve (60–120 cm), endemic to the Mediterranean Sea [4]. It is an inhabitant of shallow waters along the Croatian Adriatic coastline and suffered high mortalities similar to other parts of the Mediterranean. The results of the study presented in this Special Issue contribute a description of the diagnostics of causative agents of mortalities and epidemiology in Mljet National Park and the Northern Adriatic. It seems that mortalities were caused by infection with the haplosporidian parasite *Haplosporidium pinnae* and bacterium *Mycobacterium* sp. The spreading pattern of the mortalities based on a pilot study undertaken in Mljet National Park, an area with a dense population of noble pen shells, was evaluated. The results of the study support the hypothesis that the increase in mortalities was influenced by high temperatures, as peak mortalities in the studied area occurred in August with a sea temperature of more than 26 ◦C. In addition, multifactorial causality was proven, as the presence of *Mycobacterium* sp. alone was detected a long time before mortalities occurred, but coinfection of *Haplosporidium pinnae* was also detected after mortality. Still, a multidisciplinary experts' approach is needed to explain the phenomenon and to set up an efficient program for the protection of noble pen shell from extinction.

The focus of the next two papers are marine fish: one studied lesser-spotted dogfish (*Scyliorhinus canicula*) juveniles reared in public aquaria which suffered from infection with two different species from the genus *Vibrio* [8], and the other studied the skin microbiota of farmed fish correlated to the use of antibiotics [9]. Although elasmobranchs are endangered species in the Mediterranean Sea, classified as on the decline due to habitat degradation and consequent to the direct impacts of fishing, lesser-spotted dogfish, a small demersal shark, is classified as being of least concern (LC) by the IUCN. Its diet and habitat requirements, as well as easy reproduction in captivity, make them favourable species in public aquaria. Reports on infectious diseases affecting sharks are scarce, and therefore, research on their susceptibility to different infectious agents may contribute to the mitigation of elasmobranchs' decline. This research proved susceptibility to two *Vibrio* species: *V. crassostreae*, previously described as a pathogen of molluscs and fish, and *V. cyclotrophicus*, reported in molluscs and as a member of the microbiome of marine copepods [8]. This study described suitable diagnostic methodology to identify specifically *V. crassostreae* and *V. cyclitrophicus*, underlining the criticalities in the identification process concerning the techniques adopted (API® 20E, MALDI-TOF MS, molecular biology) and the need for developing specific guidelines for the identification of non-major pathogenic *Vibrio* species. In addition, it highlighted the need for in-depth studies of the pathogenic mechanism of these bacteria in sharks. Bacteria from the genus *Vibrio*, together with those from the genus *Pseudomonas*, were prevalent in the study of skin microbiota of farmed European seabass (*Dicentrarchus labrax*) [9]. Some of the microbiota that were identified are known to be pathogenic to fish: *V.alginolyticus*, *V. anguillarum* and *V. harveyi*. *Vibrio* strains showed higher resistance to studied antibiotics compared to previous studies. This study provides, for the first time, information on the cultivable skin bacteria that were associated with healthy European seabass under culture conditions with and without the use of antibiotics. The obtained information will be useful in assessing how changes in cultivable microbiota may affect the health of farmed European seabass, indicating a potential problem for fish health management during disease outbreaks. Interestingly, some resistant bacteria were detected among isolated bacteria, a component of skin microbiota from the farm without the use of antibiotics that raised many different questions on the influence of the fish farms on the environment but also the influence of the environment and anthropogenic activity

on fish farming. The presence of resistant microorganisms could potentially endanger consumers' health and contribute to horizontal gene transmission. However, there is little data on the antimicrobial resistance gene transmission (AGR) in the marine environment and future studies should put a lot of effort into elucidating pathways and possibilities of AGR spreading.

Each of the published articles in this Special Issue tackled one of the niches within the marine environment and fulfilled a strong need for further and more detailed research on different marine organisms and their interactions with different microbes and the outcomes of these interactions.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


## *Review* **A Review of Marine Viruses in Coral Ecosystem**

**Logajothiswaran Ambalavanan 1, Shumpei Iehata 1, Rosanne Fletcher 1, Emylia H. Stevens <sup>1</sup> and Sandra C. Zainathan 1,2,\***


**Abstract:** Coral reefs are among the most biodiverse biological systems on earth. Corals are classified as marine invertebrates and filter the surrounding food and other particles in seawater, including pathogens such as viruses. Viruses act as both pathogen and symbiont for metazoans. Marine viruses that are abundant in the ocean are mostly single-, double stranded DNA and single-, double stranded RNA viruses. These discoveries were made via advanced identification methods which have detected their presence in coral reef ecosystems including PCR analyses, metagenomic analyses, transcriptomic analyses and electron microscopy. This review discusses the discovery of viruses in the marine environment and their hosts, viral diversity in corals, presence of virus in corallivorous fish communities in reef ecosystems, detection methods, and occurrence of marine viral communities in marine sponges.

**Keywords:** coral ecosystem; viral communities; corals; corallivorous fish; marine sponges; detection method

#### **1. Coral Ecosystem**

The earth is covered by an ocean that contains about 97% of the planet's water [1]. Mora et al. [2] have estimated that about 2.2 ± 0.18 million species are found in the marine environment but only 91% of the ocean species have yet been discovered. The coral ecosystem consists of sponges, coral, corallivorous, crustaceans, molluscs and other organisms that live beneath the reef ecosystem. Sponges are among the oldest metazoans which can be divided into four classes: Hexactinellida, Calcarea, Demospongiae and Homoscleromorpha [3]. Sponges feed on plankton that include bacteria, algae, protozoa and microscopic animals which transfer carbon flow to higher trophic levels [4]. These sponges are also known to be food sources for organisms, for example, fish, crustaceans, sea urchins, star fish and molluscs [4]. Sponge's morphology is very diverse with a colorful array ranging from amorphous types, to branching, with a great variety in length and size [5]. Moreover, sponges are plentiful and functionally essential for coral reef systems [6]. They play a crucial role in numerous ecosystems such as substrate accretion [7] and erosion [8,9]. As described by Aerts [10], out of 128 sponge species, 30 interact with corals in coral over-growth [9]. As reviewed by Bell [6], functional roles that are played by sponges in Caribbean coral include increasing coral survival by binding live corals to reef frame and preventing entry to their skeletons by excavating organisms, nutrient cycling, bioerosion reworking of solid carbonate, primary production via microbial symbionts, removing prokaryotic plankton at water column and providing food sources for organisms.

Corallivores are known as fishes that feed on live corals in reef ecosystems. These coral-feeding fishes have shown that almost 80% of their feeding is based on coral, which assumes that they are dependent on coral for their survival [11]. They can be classified as polyps-feeders, removing coral tissues. Persistent predation caused by coral fish aggravate

**Citation:** Ambalavanan, L.; Iehata, S.; Fletcher, R.; Stevens, E.H.; Zainathan, S.C. A Review of Marine Viruses in Coral Ecosystem. *J. Mar. Sci. Eng.* **2021**, *9*, 711. https://doi.org/ 10.3390/jmse9070711

Academic Editor: Snježana Zrnˇci´c

Received: 7 May 2021 Accepted: 25 June 2021 Published: 27 June 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

the effects of coral disturbance and slow down reef recovery [11]. As reviewed by Cole [11], the corallivorous species can vary among habitats and geographic areas. Coral colonies as a food source for these corallivores need to find a balance between feeding severity and coral regeneration [11].

Meanwhile, coral reefs are among the most biodiverse biological systems on the planet, the persistence of which relies on the reef-building capacity of scleractinian corals. The reefbuilding corals accrue almost 250 million years with hundreds of scleractinian coral species that exhibit in multiple colony sizes, shapes and life spans, providing a wide assemblage of territory for molluscs, fish and crustaceans [12]. Corals are made up of networks of polyps and gastrovascular system that consist of tissue layers that contain numerous cell types such as epidermis mesoglea, gastrodermis, coelentron, gastrodermis mesoglea and calicoblastic epithelium [12]. According to Tresguerres and Barott [13], the corals' oral ectoderm connects to the external environment and is involved in the production of the mucus layer which helps corals in capturing prey and guarding against nematocysts. Hence, corals filter the surrounding food particles and other particles in seawater including pathogens such as viruses.

Corals are either hermatypic (reef-building) or ahermatypic (non-reef-building). Hermatypic corals flourish in shallow waters which contain millions of zooxanthellae. *Symbiodinium* are colloquially known as zooxanthellae. Due to the complex beneficial interactions among coral hosts, unicellular algae and dinoflagellates *Symbiodinium* spp., and their different microbiomes, coral reefs flourish in oligotrophic tropical waters [14]. Coral beneficial interaction with *Symbiodinium* algae permits the coral to harness power from daylight (photosynthesis), as fixed natural carbon is moved to the host while the algae obtain inorganic supplements reprocessed from the host's metabolism, for example ammonium and carbon dioxide [15]. These resources supply the coral with vitality for growth, reproduction and respiration [14]. Zooxanthellae conducts photosynthesis from sunlight as it captures and transfers 95% of the energy generated to the coral polyps. This relationship between zooxanthellae and corals supports both the reef and the zooxanthellae, sustained itself by nutrients such as inorganic substances and acquiring refuge in exchange. Global climate change can also stress the coral, collapsing the symbioses, which leads to bleaching (paling) or loss of zooxanthellae [14].

Cziesielski et al. [16] discovered the first mass coral bleaching in 1988 and over the past 4 years alone prominent coral reef ecosystems such as the Great Barrier Reef have lost about 50% of shallow-water corals [17]. Coral reefs depend on a differing consortium of free-living and host-related microorganisms for the capture, maintenance, and reuse of nutrients and minor components that permit these environments to flourish in the marine environment, which can be compared to a desert [18,19]. Climate change can trigger disruption of the natural microbiome which could lead to a state of dysbiosis with the exposure of opportunistic and probably pathogenic taxa, resulting in an inclining incidence of disease, bleaching, and host mortality [20,21]. It has been proposed that the microbiome could play a central role in coral reef reclamation, including marine viruses [21]. These changes in the microbial community cause the aggravations that could act as early warning signals, since these movements may anticipate visual indications of bleaching and tissue necrosis [22–25]. Coral reefs have endured uncommon losses and are critically threatened by continuous elevation in ocean surface water temperature and ocean acidification due to climate change, despite viral infection in corals and their worldwide significance [26,27].

According to Thurber et al. [28], the coral holo-biont contains diverse viral-like particles (VLPs) which were first observed in stony corals (cnidarians) and the sea anemone [29]. It is reported that larger viruses could infect dinoflagellate endosymbionts of the corals due to environmental factors. Virus-associated corals infect the coral tissues and the coral surface microlayers (CSM) [30–33]. Cram et al. [34] predicted that high viral infection is the source of marine bacterial mortality that affects shape together with structure of the microbial host community in the marine ecosystem and transfers genetic material between microorganisms, known as horizontal gene transfer (HGT) [35]. Viruses also

encode auxiliary metabolic genes (*AMG*s) that increase microbial host growth and fitness [36–38]. Viruses exist as both pathogen and symbiont of metazoans, controlled by the environmental factors and mutual relationship between the host and virus itself [39,40]. The ecological roles of the virus communities in health and disease in corals are still inadequately recognized despite the availability of advanced molecular technology. According to Thurber et al. [28], the stress-effect of viral infections damages the coral tissues that drive carbon from the benthos into the water column in the reef ecosystem. Marine bacterioplankton blooms trigger viral production in reef waters that releases more carbon and nitrogen into the environment. These occurrences induce viral production and mortality in corals and lead to coral reef decline [28]. Generally, corals are involved in the biogeochemical cycles in the reef ecosystems due to the release of dissolved organic carbon (DOC) [28]. The presence of DOC stimulates bacterial growth and it has been reported to enhance the DOC flux processes, and nutrients released from CSM have negative feedback on the coral reefs [41,42]. A shift in the relative abundances of eukaryotic viruses and phages has been reported by Correa et al. [43] (lab-stressed corals, Hawaii and Florida) and Soffer et al. [44] (bleached, disease and healthy corals, Caribbean reefs). The shifts in the relative abundance of virus families may also play a role in coral mortality and disease. For instance, the Circoviridae family is one of the most abundant viruses in corals that causes a disease known as the white plague [28].

According to Sweet et al. [45], viral agents may act in various manners, including critical infection, reactivation of inert disease, immune suppression (declining of the immune system of the host or weakening of the immune system due to natural aging), where it is able to destroy algae symbionts under environmental pressure and may participate in coral bleaching reactions. Nonetheless, some of these viruses are possibly host-specific to the symbiotic algae and may perform as primary pathogens in coral disease. In conjunction with these known groups, various viral sequences have been distinguished that could nt be appointed to any known families of viruses, demonstrating that the coral microbiome is a rich environment for novel viral revelations [45].

The bacteriophage-adhering-to-mucus (BAM) model had been hypothesized and named due to the observation of enrichment of phage in mucus occurring through interaction of mucin glycoproteins and Ig-like proteins that realm on phage capsids [45]. These BAM models assimilate a mechanism in supporting the specific relationship of mutualistic bacteria with host, and dispenses an evolutionary structure for a process of specific co-advancement of a phage-bacterial-host related microbiome. Therefore, evidence has demonstrated the role of viruses in the control of coral-related bacterial communities, where viral lysis rates rise in the surface mucus layer and complex associations are presented in the bacteriophage-adhering-to-mucus model. Viruses were also indirectly involved in controlling pathogenic bacterial populations and disease pervasiveness [45]. Wood-Charlson et al. [46] showed that diverse viral families had been detected from coral and symbiotic microbes by metagenomics. They reported that these viruses are likely to play numerous, commensal roles and are parasitic, regarding the health of coral reefs. Regardless of these assorted varieties, a number of taxonomic classifications are generally found in corals, including bacteriophages belonging to an order of the Caudovirales, and eukaryotic nucleocytoplasmic large DNA viruses (NCLDVs) associated with families such as Phycodnaviridae, Mimiviridae, Poxviridae and Iridoviridae, as well as Polydnavridae and Retroviridae [43,46]. Other than the families that affect unicellular algae, there are few a number of coral-associated families known to infect plants and/or fungi and protists including Geminivirdae, Nanoviridae, Tymoviridae, Potyviridae, Tombusviridae, Caulimoviridae, Alphaflexiviridae, Endornaviridae, Partitiviridae and Reoviridae [46].

#### **2. Progress of Marine Virus Research**

Marine viruses are found to be abundant in the ocean in many organisms as they play vital roles in global geochemical cycles [47]. Viruses are widely distributed in various environments including extreme environments, such as hydrothermal vents, cold springs and hypoxic saline conditions. They exist widely in seawater [48] and sediment [49].

Spencer et al. [50] discovered the first bacteriophage from the marine environment. Viruses recycle nutrients together with organic matter via a process known as the viral shunt. The cellular materials released as particular or dissolved organic material are not directly available for utilization by organisms from higher trophic levels but are primarily utilized by predominantly heterotrophic bacteria, although some efforts have shown nutrients released in this manner are rapidly assimilated by eukaryotic plankton [51]. Spatio-temporal dynamics of these viruses accounted for 10% of phytoplankton mortality within the course of phytoplankton blooms and additionally stimulated recycling of nutrients, together with organic matter, via viral shunt [52–54]. Research advances in these environments have encountered marine viruses and their hosts and organisms including plankton and aquatic invertebrates.

The Kill the Winner (KtW) hypothesis proposed that viruses are sustained via host specific infection including plankton and lysis, the most abundant microorganisms in the environment [55]. Numerous efforts have been made to identify and isolate viruses from cultivated microorganisms and have increased understanding of genomic structure and the host range of marine viruses, especially from Cyanobacteria (such as *Prochlorococcus* and *Synechococcus*) [56–59]. The RNA-dependent RNA-polymerase (*RdRp*) gene is an essential protein encoded in the genomes of all RNA-containing viruses such as Picornalike viruses and DNA containing viruses such as T4-like myoviruses (gene marker g23 and g20) have limited the research on viral diversity to genome fingerprinting [60–63]. Thereby, metagenomics studies were introduced, to overcome the bottleneck of cultivation and the lack of universal markers, in the early twenty-first century. Breitbart et al. [64] reported that both single-gene-based and genome-fingerprinting-based methods have revealed the temporal and spatial dynamics of marine viral communities.

Metagenomics-based studies have enabled scaling of the information on viral genomics and sequencing of the fragmented nucleic acid from seawater and marine sediments [65]. However, several important issues, despite the progress in the study of of marine viruses, remain unexplored, for instance, the examination of specific-host interactions, expansion of spatio-temporal marine viral studies, linking the environmental viruses with their hosts and enhancement of knowledge particularly regarding deep-sea viral communities and viral auxiliary metabolic genes (AMGs) [38]. Advanced informatics and theoretical research have unveiled the biological basis of complex host range patterns and explicated largely unknown viral sequences in marine ecosystems [66,67]. Regarding this, the progress of marine viruses from marine sponges are recommended so that we know the abundance or presence of viral communities that have been detected, making it easier for other researchers to refer to.

#### **3. Marine Viruses and Their Host**

The major impact of viruses in the marine environment began when it was discovered that seawater contains around 101◦ viruses per liter [68]. In the mid-1970s, viruses or virus-like particles (VLPs) were reported in numerous taxa of algae. In the 1980s, a group of large double stranded(ds) DNA-containing viruses (Chloroviruses) was discovered. These viruses infect and replicate in unicellular, eukaryotic, symbiotic, chlorella-like green algae known as Micratinium (formerly known as Zoochlorellae) (Chlorophyta). As described by Breitbart et al. [69], viral abundances are known to be tightly linked to their host and are generally more abundant in phytoplankton and Cyanobacteria than in any other microorganisms.

In the past, large-scale spatial investigations of viral dispersion in the Pacific and South Atlantic Oceans have utilized flow cytometry to reveal that viral abundance in 200 m of the water surface column is high in tropical and subtropical districts yet lower in Antarctic waters [70,71]. A virus "hot-spot" was recognized in the mid-scope region of the North Pacific supporting the hypothesis that large-scale distribution patterns of viruses are influenced by host distributions and physical procedures [71]. Brum et al. [72] used cultivation-independent methods such as metagenomics and revealed that the Southern Ocean is dominated by lysogenic viruses, which are termed as "seasonal time bombs" since they can shift to a lytic cycle as their bacterial host production rises. Roux et al. [73] collected virome data that identified virally encoded AMGs (Auxiliary Metabolic Genes) in surface waters that appear to be involved in nitrogen and sulfur cycling.

The Pacific Ocean Virome (POV) [74] is another significantly important curated dsDNA virome dataset that comprises 32 quantitatively representative viromes collected from different depths and seasons in transects from coastal waters to open-ocean waters in the Pacific Ocean [74]. POV study revealed that the richness of viruses in the Pacific reduced from deeper to surface waters, from winter to summer, and, in surface layers, with distance from the shore. Other than that, data collected by the TARA ocean expedition were used to develop a global virome map of dsDNA viruses sampled in surface and deep-ocean waters [73], along with the genomic data from the Malaspina 2010–11 Circumnavigation Expedition, which has assessed pelagic processes along the Indian, Pacific, and Atlantic Oceans [75]. The authors found that 38 of 867 viral clusters were locally or globally abundant and represented almost 50% of the viral population in any given Global Ocean Virome (GOV). Such huge datasets are critical in developing our insights into marine viruses on a global scale and are important in divulging uncultivated novel viruses and initiating important marine ecosystem models.

Hence, the International Committee on Taxonomy Viruses (ICTV) have classified and named viruses according to their taxonomy and binomial species [76]. The key factors that are used for the identification and classification of viruses include the display of capsid protein and viral morphology via TEM [77]. Nineteen families of unassigned viruses were reported, whereas approximately 5000 marine viruses were assigned to 26 families according to ICTV [67].

Moreover, viruses can be grouped based on their hosts such as bacteria, animal, archaea, algae and plant viruses (Table 1). Authors have also discovered single stranded DNA (ssDNA) and double stranded RNA (dsRNA) viruses other than the double stranded DNA (dsDNA), abundant in the ocean with marine organisms as their hosts.


Marine viruses and their hosts in the marine environment adapted from He et al. [67] and King et al.

 [78].

**Table 1.**

The concentration of free virions decreases with depth in seawater [47] with a total abundance of more than 103◦ [47], whereas the concentration of marine virio-plankton in surface seawater is typically billions per milliliter [54,114]. The published data on viral abundances in seawater are mostly found in freshwater, coastal water, open ocean, deep ocean and estuarine waters, but the discovery of marine viruses in Asia is still at an early stage.

Zhong et al. [115] have discovered four targeted viruses (e.g., *Cyanomyovirus*, T4-like *myovirus*, Cyanophage, Phycodnavirus) in Lake Annecy and Lake Bourget. Molecular detection such as PCR-denaturing gel gradient electrophoresis (DGGE) had been used to discover viruses in the lake water. DGGE banding pattern analysis revealed the different similarities of the viruses in both lakes. The results demonstrated the presence of 70% of *mcp* phycodnaviruses (Lake Bourget), 45% of *polB* phycodnaviruses (Lake Bourget) and 60% (Lake Annecy), respectively, 45% of psb A cyanophages (Lake Bourget and Lake Annecy), 45% of cyanomyo-viruses in Lake Bourget and 75% of T4-like myo-viruses in Lake Bourget. The total bands in common that showed the presence of the viruses in both lakes were as follows: *mcp* (85%), *polB* (54%), *g20* cyano-myovirus (52%), *psb A* cyanophages (39%) and *g23* of T4-like myo-viruses (75%), respectively. Previously, this method had focused on marine ecosystems where only a single gene marker was used typically to identify the presence of the viruses in the marine environment, *g20* [116–118], *polB* [118,119] or *mcp* [120–122]. Parvathi et al. [123] demonstrated the dynamics of virus infection of autotrophic plankton in Lake Geneva over a 5-month period and the abundances were determined via flow cytometry and PCR-DGGE method identification. The viral signature genes used by Parvathi et al. [123] were similar to those used by Zhong et al. [115] to identify the viral abundances in surface water. Parvathi et al. [123] have discovered that the abundances of picocyanobacterial hosts was in concurrence with cyanophages, which were higher in late summer.

Luo et al. [124] discovered metagenome sequence data that yielded about 16787 virus populations, and 1352 of these were identified as putative temperate phages with temperate phage marker genes. Moreover, about 12 complete archaeal virus genomes and 25 genome fragments of eukaryotic viruses have been identified [124]. They discovered that the depth of ocean inferred the Virus:Cell ratio (VC) temporal variability where, as the depth of ocean decreased, there were more temperate phages present compared to other viruses. The temperate phage peaked at a photic zone in the ocean at about 150–250 m. This peak indicates increased temperate phage productivity relative to deeper waters, with slowly growing hosts. A high VC variability driven by temporal resource variability indicates phages with more episodic virus particle production [124]. Thus, this shows that the ecology and biogeochemistry of microbial communities were impacted by viruses across the ocean.

#### **4. Coral Viruses**

Reef-building corals lay the foundation for the structure and biodiversity of the coral reef ecosystem. These huge biological structures can be seen from space and are the culmination of complex developments. The interaction between coral micro-polyps and their unicellular symbiotic algae is closely related to microorganisms such as bacteria, archaea, fungi and viruses. Reef building corals have existed in various forms for more than 200 million years, and human-induced conditions have been reported to threaten their function and durability [12]. The coral reef ecosystem is at the forefront of people's attention in the Anthropocene; for instances corals, the main species, are sensitive to human interference ranging from local activities (such as overfishing, coastal development and pollution) to worldwide phenomena (e.g., climate change and ocean acidification) [12].

Bacteria, Achaea and eukaryotic viruses are cosmopolitan and exist all over the world's oceans [125]. According to Thurber and Correa, [126], a few published studies have directly evaluated VLPs associated with shallow-water scleractinian (stony) corals, the primary architects of the coral reef ecosystem. According to Paul et al. [127], VLPs

were detected in surface water, sediments and several invertebrate groups in tropical coral reefs and seagrass beds, and changes in terms of distance from coast, seasonality and salinity were recorded in the abundance of these particles. The first detection of a marine cnidarian-based VLP is based on the temperate plumose sea anemone, an old specimen of *Metridium senile* [127]. Since VLP is visualized in the tissues of the anemone, *Metridium senile* does not form symbioses with algal partners, and VLP may infect the anemone itself and/or organisms outside the anemone. Wilson et al. [128] studied the first characterisation of viruses associated with corals where VLPs were detected in both healthy and heat-stressed colonies, coral *Pavona danai, Acropora formosa* and *Stylophora pistillata*. Tail-less VLPs, hexagonal and about 40 to 50 nm in diameter, were present around the corals. A large number of VLPs were related to heat shock treatment, prompting a viral outbreak in these coral fragments. One of the most fascinating papers on coralassociated viruses in the study of VLPs was related to the coral surface microlayer of *Acropora muricata, Porites lobata* and *Porites australiensis* [129]. VLP were divided into five groups (tail phage, polyhedron/spherical, lemon-shaped, filamentous and unique VLP) and 17 subgroups based on the morphological similarity to the previous described viruses which have potential hosts existing in the coral surface microlayer, including algae, cyanobacteria, archaea, fungi and the coral animal.

A previous meta-analysis [46] found approximately 60 virus families in corals around the world as recognized by ICTV. A number of dsDNA ssDNA type viruses were identified from coral-holobiont sequence data sets and the coral transcriptomes generated from extracted holobiont RNA were dominated by 36–78% of RNA viruses that includes ssRNA viruses (1–13%) and dsRNA virus (2–31%). The analysis made by Wood-Charlson et al. [46] demonstrated that dsDNA viruses from the order Caudovirales were the only group of viruses that were dominant in all of the data sets (ICTV, 2012). Besides, most of the sequences from the coral-related viral metagenomes were dsDNA and ssDNA bacteriophages that show significant matches to the NCBI'S RefSeq Virus database.

Weynberg et al. [130] have identified that about 84% of the sequences belonged to dsDNA viruses and approximately 85.6% were ssRNA viruses from *Alternaria tenuis* (Fugi, Ascomycota). Most of the ssRNA viruses matched with the major capsid protein (MCP) gene from dinoflagellate-infecting ssRNA virus (HcRNAV) which supported the observation by Correa et al. [131] in *Montastrea cavernosa* (Animalia, Cnidaria). Studies which used approaches such as transmission electron microscopy and flow cytometry revealed the presence of VLPs associated with cultures of *Symbiodinium* (Miozoa, Dinophyceae) cells from corals. According to Thurber et al. [28], core coral viromes comprise 9–12 families in three viral lineages known as dsDNA group I, ssDNA group II and retrovirus group IV [43]. A typical virus related to the Herpesvirales order reported to be a member of the core coral virome showed 98% sequence similarities to the Herpesviridae virus [43]. Similar authors stated that only 10% of ssDNA (Circoviridae) and ssRNA (Caulimoviridae) were represented in the data set, probably due to the methodological differences among the studies. TEM-based studies identified VLP morphotypes that were comprised of enveloped, icosahedral capsids ranging between 120 to 150 nm in diameter from Kane'ohe ¯ Bay in Hawaii in the United States, as well as in the corals that reside within a cellular vacuole alongside VLPs, and indicated its atypical herpes-like viral particles.

The morphological identification of VLPs was performed in the tissues of *Acropora muricata* colonies infected with healthy and white syndrome (WS) via TEM. This was the first study of cnidarians, which included temporal and spatial components. The characteristics of these dominant VLPs and their existence at multiple sampling time points led Patten et al. [30] to assume that the colony of the *Acropora muricata* is suffering from persistent infection of Phycodnaviridae and/or Iridoviridae. Buerger et al. [132] described the degradation of *Symbiodinium* cells and linked this to the abundance of VLPs in the coral. Table 2 shows the coral-related viruses reported in the reef ecosystem. As suggested in the case of virus-induced coral bleaching and yellow blotch disease, the virus itself causes the disease and therefore directly interacts with the coral itself, whereas the indirect processes involve bacteriophages that interact with the prokaryotic community. Phages may go through the horizontal transfer of virulence genes, increasing the virulence of the infected bacterium, which then causes coral diseases. In addition, bacteriophages may infect and lyse pathogenic bacteria, reduce the impact of disease and become a part of the coral microbiome, or reduce the external influences from the overall organism of the coral, for instance, manual application in phage therapy [132].




**Table 2.** *Cont.*

Wood-Charlson et al. [46] have discovered Phycodnaviridae, Marnaviridae and Alvernaviridae, which were the best characterized group of algal viruses, via metagenomic studies [149,150]. Nanoviridae and Geminiviridae are common viruses that have been found in almost every coral-associated virus study. These viruses are usually linked with sewage, which may highlight the connection between certain types of virus and environmental degradation [44]. Therefore, the abundance of viruses in corals is proportional to the concentration of local inorganic nutrients and human population centers [135,151]. According to Futch et al. [148], certain "human-specific" viruses such as Adenovirus have also been shown to exist in coral surface mucus layer, possibly due to human pollution, including sources of fecal pollution such as boats and contaminated groundwater associated with septic systems and injection wells.

According to the metagenome studies conducted by Wang et al. [152], the top five viral families that have been found in *Siderastrea sidereal* coral were Myoviridae (25.98%), Siphoviridae (9.26%), Mimiviridae (7.89%), Baculoviridae (7.61%) and Poxviridae (5.81%) whereas the overall mean number of viral abundances was about 1.14%. Moreover, similar studies have discovered thermal anomalies related to microbiome shifts, and the order Caudovirales has been found to be in high proportion in certain samplings in warmer months (July and August). Caudovirales have been found consistently in coral virome [28,46]. The member of the family known as Poxviridae is often found in marine coral viromes [140]

that infect marine invertebrates yet has been found in coral, *Siderastrea sidereal,* where thermal stress increased and coral health decreased [152].

A viral outbreak in both coral species, *Acropora aspera* and *Acropora millepora,* was caused by the Herpesvirales order which has been commonly identified in these coral species. These herpesvirus-like VLPs are composed of an enveloped and circular capsid that hosts on coral epidermal and gastro-dermal cells. Moreover, the second most common abundant eukaryotic virus annotations were the NCLDVs (Nucleocytoplasmic Large DNA Viruses) including, Phycodnaviridae, Mimiviridae, Poxviridae, Iridoviridae, Marseillevirus and Ascoviridae. The phages that were found to dominate in the *Acropora aspera* virome were Sipho- and Myoviridae [43].

According to Cardenas et al. [141], the viral community composition in Red Sea corals detected via metagenomic study included 97 viral families, which were found across meta-transcriptomics. The most abundant viral families were Siphoviridae, Mimiviridae and Retroviridae (dsDNA viral families) found in eukaryotes and bacteria, whereas ss-RNA viral families such as Qinviridae, Nyamiviridae and Solinviviridae were present in meta-transcriptome studies. Parvoviridae was found to be highly abundant in stresstolerant coral virome with massive growth in *Acanthastrea echinata, Diploastrea heliopora*, *Fungia* sp., and *Plerogyra sinuosa*. Dicistroviridae was observed in the outgroup samples of *Millepora platyphylla*, *Xenia* sp., and *Stylophora pistillata*. Similar studies have revealed the relative abundances of *Picobirnaviridae* and *Siphoviridae* that were most accounted for in viromes of *Galaxea fascicularis, Mycedium elephantotus*, and *Pachyseris speciosa* when compared to *Acropora cutherea*, *Pocillopora verrucosa*, and *Stylophora pistillata* [141].

#### **5. Coral Fish Viruses**

Coral disease outbreaks are the main cause of coral death and subsequent coral reef degradation [153]. It is reported that corals are susceptible to viral infections and the organisms involved mainly contribute to the HGT (Horizontal Gene Transfer) of viral particles to coral, for instance, fish, invertebrates and macroalgae [126]. Coral feeding fish (e.g., parrotfish and butterfly fish) and invertebrates [154–156] are another reasonable mechanism for the introduction of viruses into individual coral communities. According to Rotjan and Lewis [154], coral tissue mortality can be classified into two categories, where mortality of coral tissue occurred due to parrotfish grazing scars over the colony surface or mortality from unknown causes. Evidence provided by Bettarel et al. [157] that the grazing activity of the specific corallivorous gastropod *Drupella rugosa* damaged coral polyps were the result of predation on coral microbial associates is still obscure. Similar studies have revealed the presence of dsDNA virus from the coral *Acropora Formosa* which mainly belongs to uncultured Mediterranean phage uvMed, while others were Myoviridae, Podoviridae, Siphoviridae and Microviridae. The coral predators were suspected to assist in disease transmission either by acting as vectors or stressors on coral microbiota, while the presence of *Drupella rugosa* has corresponded to disease such as white syndrome, skeletal eroding band disease and black band disease [157].

Corallivorous fish act as vectors for disease transmission [158]. Aeby and Santavy [158] have determined that the coral-feeding butterflyfish, *Chaetodon capistratus* was involved in the colony transfer of black band disease. In aquaria, the presence of *Chaetodon acapistratus* increased the rate of spread from infected *Montastraea faveolata* to uninfected fragments. Both protected corals that were exposed to fish predation suffered from black band disease. Therefore, oral transmission of pathogens directly and/or via indirect fecal transmission may spread from colony to colony. This kind of direct feeding may actually be beneficial because it can reduce the extent and progression of the disease [158].

Thurber et al. [142] proposed that coral related viruses known as herpes-like viral were found to shift in response to abiotic factors when exposed to decreasing pH, elevated nutrients and thermal stress. Similar studies suggest that coral-related viruses target hosts such as protists, fungi, plants and metazoans. It is also found that invertebrates infecting viruses were mostly Baculoviruses and Polydnaviruses that predominantly infect the arthropods [159,160].

According to the study by Cherif [161], the molecular detection of Lymphocystis Disease Viral partial genome (LCDV-Sa) in Tunisian gilthead sea bream (*Sparus aurata*) caused fatal, chronic, rare and slowly developing disease which affects more than 150 marine and freshwater fish [162–164]. The etiological agent of LCD (Lymphocystis Disease Virus) belongs to genus *Lymphocystivirus,* family Iridoviridae [165,166]. *Lymphocystivirus* have been known to affect marine species such as *Holacanthus* spp., wrasses *Halichoeres* spp., grunts *Haemulon* spp., pinfish *Lagodon rhomboides*, puffers *Canthigaster* and *Sphoeroides* spp., porcupine fish *Diodon hystrix*, and many more, as stated by Stoskopf., [167]. Another study revealed LCDV strains from clownfish, *Amphiprion percula* in Indonesia. Lymphocystis in Atlantic croaker Micropogonias undulates and sand seatrout *Cynoscion arenarius* have been reported in Mississippi estuaries from 1966–1969 [168]. According to Bowden et al. [169], lymphocystis was observed in red drum *Sciaenops ocellatus* and other species in the Gulf of Mexico. In addition, a rhabdoviral infection was first described in 1983 in gray angelfish (*Pomacanthus arcuatus*) and French angelfish (*Pomacanthus paru*) collected from the Florida Keys. Table 3 below shows the reef-associated fish that have been infected with the virus.

**Table 3.** Lymphocystis disease virus (LCDV), detection method and its host in a coral ecosystem.


#### **6. Marine Sponge Viruses**

Marine sponges (phylum *Porifera*) are metazoans and have been distributed all over the world in the aquatic environment since 600 million years ago [175,176]. Marine sponges are a rich source of biotechnologically potential compounds [177,178]. There is a lack of information regarding the role of sponge-related virus communities and the impact of the virus on sponge holo-biont is still unclear [179–183]. Sponge microbiome composition is shifted by environmental factors such as climate changes [184] and host sponge habitat [185] in nine sponge species [186,187]. Fan et al. [188] suggested that marine sponge symbionts lose their metabolic functional potential during the early stages of heat stress and hence destabilize the sponge holo-biont before visual signs of stress occur in the host animal. Global climate change has had a significant effect on the associated microbial communities. For example, the rise of temperature in the ocean controls the link between viruses and the cells they infect, where the growth rate of prokaryotes increases, the length of lytic cycle decreases and burst size increases, resulting in higher virus production rate [189–191]. Thus, changes in climate have direct and indirect effects on marine viruses, including impact of cascade on biogeochemical cycles, food webs and the metabolic balance of the ocean [191]. The main focus of increased studies in sponges is due to their abundance as a high source of biologically active secondary metabolites and a focal point for various aspects of research in organism origin and evolution [179,192,193].

Despite the fact that the diversity and importance of viruses in sponge-associated microorganisms are still largely unknown, a virus associated with the Lake Baikal sponge (*Lubomirskia baikalensis*) was part of the first study using cyanophage related marker gene (*g20* gene) [194] and *g23* gene for the detection of T4-like bacteriophage (Butina T.V., unpublished data). Table 4 represents marine sponge related viruses and types of method used to identify marine viruses.

Marine sponge related VLPs were investigated from the Great Barrier Reef (*Carteriospongia foliascens*, *Stylissa carteri*, *Xestospongia* sp., *Lamellodysidea herbacea*, *Cymbastela marshae*, *Cinachyrella schulzei*, *Pipestela candelabra* and *Echinochalina isaaci*) and Red Sea (*Xestospongia testudinaria*, *Amphimedon ochracea*, *Hyrtios erectus*, *Crella* (Grayela) *cyathophora*, and *Mycale* sp.) [195]. Pascelli et al. [195] revealed various types of marine sponge-associated viruses via transmission electron microscopy (TEM). Almost fifty VLP morphotypes were found in sponge tissues and mucus or surface biofilm where the viruses possessed an icosahedral symmetry morphology with diameter ranging between 60–205 nm. Moreover, the same author confirmed the presence of bacteriophage in sponge species assigned to three Caudovirales families (based on capsid symmetry and tail shape). Virus families found from marine sponge tissue and mucus in the Great Barrier Reef were Podoviridae and Siphoviridae, including Myoviridae and Inoviridae from marine sponge meso-hyl, sponge tissues and sponge mucus in the Red Sea [195].

Fan et al. [196] observed the presence of cyanophage in high abundances in sponge *Stylissa* sp. 445, suggesting that the cyanobacteria may have a lysogenic relationship with their host [197]. A few double-stranded DNA (dsDNA) viruses were abundant in a diseased branch of endemic sponge *Lubomirskia baikalensis*, for instance, Siphoviridae, Myoviridae, Phycodnaviridae, Poxviridae, and Mimiviridae; while Podoviridae, Iridoviridae and Herpesviridae can be found in healthier *Lubomirskia baikalensis* sponge [198]. The order Caudovirales with tailed bacteriophages influenced all datasets, yet the distribution of these taxa varied between holo-bionts. Phycodnaviridae, Poxiviridae, Mimiviridae, and Herpesviridae dominated all of the viromic datasets which comprised almost 98% of reads, with other unassigned viruses in Baikal sponge. Podoviridae viruses were more abundant in *Lubomirskia baikalensis* (Sv3h) healthy sponge and coral *Acropora millepora*. The presence of Herpesviridae comprised approximately 95–98% of metaviromic dataset in *Rhopaloeides odorabile* [198].

Batista et al. [199] recorded sponge Darwinella sp. and *Dysidea etheria,* with the highest abundances of Myoviridae virus detected from Arrarial do Cabo Bay site, South-Eastern Brazil, at low, upwelling and high anthropogenic influence. Laffy et al. [200] demonstrated that most dominant sponge-associated viruses detected were DNA-containing viruses, order Caudovirales. Butina et al. [201] presented virome datasets on *Baikalospongia bacillifera* species sponge in nine types of dsDNA virus families (Myoviridae, Phycodnaviridae, Siphoviridae, Poxviridae, Podoviridae, Mimiviridae, Herpesviridae, Baculoviridae and Inridoviridae) which comprises more than 70% of the identified virome sequences. Potapov et al. [202] reported that 37 nucleotide sequences of *g23* gene fragment (accession number MH576490-MH576574) were found in sponge from Lake Baikal. The data produced by Patapov et al. [202] revealed seven sequences included in Far T4 group that contains phages *(Escherichia* phage 121Q and RM378).

RNA viruses have been discovered in marine sponges, coastal water [203,204], benthic sediments [205], invertebrates [206] and vertebrates [207]. The dsRNA virus genome had been detected through advanced technology, for example, fragmented and primerligated dsRNA sequencing (FLDS), which obtained the full-length of the sequence of dsRNA [208–210]. Waldron et al. [211] revealed the presence of RNA virus families such as Narnaviridae, Dicistroviridae, Partitiviridae, Picobirnviridae, Picornaviridae, Tombusviridae, Nodaviridae and Herpesviridae via meta-transcriptome-FLDS analysis. Similarly, Urayama et al. [212] revealed the presence of RNA virus (Dicistroviridae) in the sponge, *Hymeniacidon* sp. Thus, they have acquired sequence encoding *RdRp* gene sequence, about 253 (2014) and 233 (2015), from Tokyo Bay which are related to nine dsRNA virus families. Of these sequences, about 78 were obtained from the sponge of *Hymeniacidon* sp. that were classified as an unassigned RNA family.


**Table 4.** Marine sponge associated viral communities and detection methods used to identify marine viruses.

#### **7. Detection Methods for Marine Viruses in Coral Ecosystem**

Diverse approaches have been used to identify and describe viruses in various organisms including transmission electron microscopy (TEM) as described in Tables 2–4 [129,213], PCR-based analyses [214], DNA in situ hybridization [215], immune-histochemistry [216], flow cytometry [217], next generation sequencing (NGS) [218] and metagenomic analyses [138,142].

Weynberg et al. [130] investigated coral associated viruses via metagenomic analysis where studies clarified that the method is a relatively promising tool for characterizing coral related viral communities. Molecular detection is frequently used by researchers in coral ecosystem associated marine viruses, which includes the viral metagenomic method and transmission electron microscopy. Sequencing of environmental DNA samples (metagenome) and sequence of other plankton viruses have revealed an unanticipated abundance of large DNA viruses associated with the marine environment [219]. Metagenomic analysis is an important tool for describing viruses, because many viral hosts are not suitable for cultivation [64] and this method does not require any of the gene markers for virus identification. Metagenomic studies have pointed out challenges and cataloged a group of worldwide viruses in corals and their symbionts [45] and this approach has widened the diversity of viral communities [130].

Although the field of coral virology is still at its early stage, some researchers have applied microscopy and genomic methods to examine the diversity and role of viruses in coral organisms. Evidence from microscopy studies has shown that virus-like particles (VLP) are present in all of the corals [29,220]. VLPs observed are likely to have been produced throughout the lytic replication phase of endogenous infections of coral animals or their microbiota [30,129]. Lawrence et al. [221] reported that the TEM approach revealed structures within corals which are marine viruses in the coral ecosystem.

Advanced molecular approaches have shown that viruses disperse in all types of environment. Sequencing of 16SRNA techniques is not recommended because they lack the gene to identify viruses, when viruses do not share common genes which significantly fit as phylogenetic markers [222]. Traditional techniques have been used to identify viruses, for example filtration, tissue culture, electron microscopy and serology. Traditional methods and advanced molecular techniques have contributed to the exploration of more viruses [222]. Electron microscopy is considered to be an expensive tool with a paucity of sensitivity. Polymerase chain reaction (PCR) only focuses on certain genes that use markers of related viruses, but PCR analyses are unable to identify complete novel viruses. In conjunction with this, metagenomic analyses are recommended to track viruses whether they are known or unknown, and they are easily detected via this method. Metagenomic

study was first used in marine environmental research, where the analysis was done in San Diego [64,222], and depended upon cloning of double stranded DNA genomes. Implementing this method will discover many unidentified viruses and uncultured novel viruses which will benefit researchers in this field. Due to the difficulty of identifying some of the marine environmental viruses, especially in the ocean, a metagenomic approach would be a major breakthrough in discovering new viruses.

#### **8. Conclusions**

Marine viral communities play a crucial role in biogeochemical cycles in the ocean, indirectly and directly. These viral communities induce mortality and disease in the reef ecosystem through abiotic and biotic factors which causes the reduction in the symbiotic relationship between the coral and their surrounding hosts. This review emphasizes marine viruses from the ocean, coral-associated viruses, marine sponge and coral fish viruses in reef ecosystems, examining previous research via traditional methods to modern advanced approaches. The findings of this review have enhanced the understanding of coral-virus interactions and enriched our understanding of reef-associated virus interaction and the diversity of viral communities in marine environments.

**Author Contributions:** Conceptualization, S.C.Z. and S.I.; resources, L.A.; writing—original draft preparation, L.A., R.F. and E.H.S.; writing—review and editing, S.C.Z., S.I.; supervision, S.C.Z. and S.I.; funding acquisition, S.I. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Fundamental Research Grant Scheme, Ministry of Higher Education, Malaysia (FRGS vot. no. 59535).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author. The data are not publicly available due to confidentiality.

**Acknowledgments:** The authors would like to thank Faculty of Fisheries and Food Sciences, Universiti Malaysia Terengganu for their immense support. The authors also would like to thank the anonymous reviewers and editors for their helpful and constructive comments.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Article* **Noble Pen Shell (***Pinna nobilis***) Mortalities along the Eastern Adriatic Coast with a Study of the Spreading Velocity**

**Željko Mihaljevi´c 1, Željko Pavlinec 1, Ivana Giovanna Zupiˇci´c 1, Dražen Orai´c 1, Aleksandar Popijaˇc 2, Osvin Pe´car 2, Ivan Sršen 2, Miroslav Beni´c 3, Boris Habrun <sup>3</sup> and Snježana Zrnˇci´c 1,\***


**Abstract:** Noble pen shells (*Pinna nobilis*) along the Eastern Adriatic coast were affected by mass mortalities similarly to the populations across the Mediterranean basin. Samples of live animals and organs originating from sites on Mljet Island on the south and the Istrian peninsula on the north of the Croatian Adriatic coast were analyzed using histology and molecular techniques to detect the presence of the previously described *Haplosporidium pinnae* and *Mycobacterium* spp. as possible causes of these mortalities. To obtain more information on the pattern of the spread of the mortalities, a study was undertaken in Mljet National Park, an area with a dense population of noble pen shells. The results of the diagnostic analysis and the velocity of the spread of the mortalities showed a significant correlation between increases in water temperature and the onset of mortality. Moderate to heavy lesions of the digestive glands were observed in specimens infected with *H. pinnae*. A phylogenetic analysis of the detected *Haplosporidium pinnae* showed an identity of 99.7 to 99.8% with isolates from other Mediterranean areas, while isolated *Mycobacterium* spp. showed a higher heterogeneity among isolates across the Mediterranean. The presence of *Mycobacterium* spp. in clinically healthy animals a few months before the onset of mortality imposes the need for further clarification of its role in mortality events.

**Keywords:** noble pen shell; mass mortality event; Croatia; *Haplosporidium pinnae*; *Mycobacterium* spp.; spreading velocity

#### **1. Introduction**

As in many parts of the Mediterranean area, the noble pen shell (*Pinna nobilis*) is the largest bivalve (60–120 cm) and an endemic inhabitant of shallow waters along the Croatian Adriatic coastline [1]. Over the last few decades, its numbers have drastically declined [2], and the species is now protected under Annex IV of the Habitats Directive, Annex II of the Barcelona Convention, and national legislation in Croatia and most Mediterranean countries. The decline has been attributed to uncontrolled trawling [3], illegal collection for food or souvenirs by amateur divers, and devastation of their natural habitats due to anthropogenic inputs [1,4]. Some coastal waters in the Mediterranean basin that are known as natural habitats of this protected species have been designated as marine parks, such as the Mljet Island National Park in Croatia or the Parque Natural de Cabo de Gata-Nijar [5] and the Parque Nacional Marítimo-Terrestre del Archipiélago de Cabrera in Spain [6].

Unfortunately, in early autumn of 2016, mass mortalities that affected the noble pen shell at several different points along the Spanish Mediterranean coast were reported [7], indicating the presence of a haplosporidian-like parasite within the digestive gland as a possible cause of the mortalities. Shortly thereafter, severe mortalities spread to most of the

**Citation:** Mihaljevi´c, Ž.; Pavlinec, Ž.; Zupiˇci´c, I.G.; Orai´c, D.; Popijaˇc, A.; Pe´car, O.; Sršen, I.; Beni´c, M.; Habrun, B.; Zrnˇci´c, S. Noble Pen Shell (*Pinna nobilis*) Mortalities along the Eastern Adriatic Coast with a Study of the Spreading Velocity. *J. Mar. Sci. Eng.* **2021**, *9*, 764. https://doi.org/ 10.3390/jmse9070764

Academic Editor: Valerio Matozzo

Received: 14 June 2021 Accepted: 9 July 2021 Published: 12 July 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Spanish coast, Corsica in France [8], several sites in Italy [9,10], and the Aegean Sea [11,12]. Recently, they were also reported in the Adriatic Sea, affecting the populations in Albania and Croatia [13–15], as well as in Turkey [16], Tunisia, and Morocco [17]. These mass mortality outbreaks are widespread and worrying. Consequently, *P. nobilis* was included in the red list of critically endangered species [18].

Studies on the etiology described the presence of *Haplosporidium* species that parasitized the digestive epithelium. As a result of the morphology and molecular phylogeny, the new species *Haplosporidium pinnae* was designated [8]. The same parasite (morphologically and phylogenetically) was reported to be associated with inflammation and heavy lesions in the digestive tubules that were a cause of a mass mortality event (MME) in the Gulf of Taranto (Ionian Sea) in southern Italy [9]. Nonetheless, Carella et al. [10] observed strong inflammatory lesions in the connective tissue surrounding the digestive system and gonads and linked these to the presence of *Mycobacterium* spp. In the same animals, *Haplosporidium pinnae* was observed. Hence, the MMEs were attributed to co-infection of the parasite and bacterium. This statement was supported by the findings of co-infections of the same pathogens in moribund pen shells collected from MME sites in Greece [11,12]. However, some other pathogens, such as bacteria from the different genera (e.g., *Vibrio*, *Rhodococcus* sp., and *Mycoplasma*) and parasites of the genus *Perkinsus* [16,19–24], were also detected in noble pen shells.

One of the most important habitats of *P. nobilis* in the southern Eastern Adriatic Sea is in Mljet National Park [25], which was two lake-like inlets: The Small Lake and Big Lake. Šileti´c and Peharda [26] found that the density of the individuals in this area appeared to be higher than in other parts of the Adriatic and Mediterranean areas, and that the population comprised primarily of adults and older individuals that were potentially aged 8–15 years [5,27]. To monitor the health status of the *P. nobilis* in the Adriatic Sea, we conducted an inspection of the *P. nobilis* populations, primarily in the Mljet National Park and in habitats along the Istrian peninsula, from April 2019 until May 2020.

The main objectives of this study are (1) to present the results of laboratory analyses of pen shells collected from habitats along the Istrian peninsula and the Mljet National Park; (2) to report the observations of a small-scale study carried out in the Mljet National Park to elucidate the rate of the spread of MMEs within dense populations; and (3) to implement and assess a new non-lethal method of mantle tissue sampling for diagnostic purposes.

#### **2. Materials and Methods**

#### *2.1. Sampling Sites and Sample Collection*

In this study, we collected noble pen shells from two habitats along the Eastern Adriatic coast: The Mljet National Park (MNP) and the Istrian peninsula. The sampling sites were determined according to the previously collected information on the natural habitats of *P. nobilis* (Figures 1 and 2; Table 1). Samples of live animals were collected in the MNP by scuba diving during April 2019, and they were delivered to the laboratory under cooling conditions with temperatures not exceeding 8 ◦C. The other samplings—both along the Istrian peninsula and in the MNP—were also carried out by scuba diving. All the samplings were carried out with the permission of the Croatian Ministry of Environmental Protection and Energy (CLASS UP/1-612-17/18-48/172; No. 517-05-1-1-18-4 of 21 December 2018 and CLASS UP/1-612-07/19-48/193; No. 517-05-1-1-19-3 of 11 September 2019).

**Figure 1.** Sampling sites along the Istrian peninsula in the northern part of the Eastern Adriatic coast.

**Figure 2.** Study site within the Mljet National Park with five sampling points and two transect areas.

During the sampling, the appearance of pen shells in their habitat was evaluated, and portions of digestive glands, gills, mantles, gonads, and muscles were collected separately and preserved in Davidson solution [28] for histological examination and in either EtOH or RNAlater (Thermo Fisher, Waltham, MA, USA) for molecular analysis. Exceptionally, 20 mantle tissue biopsies were taken during September 2019 with a non-lethal sampling method and were preserved in EtOH.

#### *2.2. Site and Design of the Small-Scale Study within the MNP*

The study was conducted from April to November of 2019 in the MNP, which is in the south-east Adriatic (Figure 2), a highly productive and biodiverse marine ecosystem within the Natura 2000 European Network of Protected Areas (code HR5000037). Five sampling points were selected according to the most abundant pen shell populations. Two sampling points (1 and 2) were situated in the Small Lake, a lake-like inlet connected by a shallow, narrow channel, while sampling point 3 was in the Big Lake, which was connected to the open sea by a slightly deeper, wider channel. Another two sampling points in addition to these lake-like inlets were Gonoturska Bay (sampling point 4) and Cape Lenga (sampling point 5). The distance between the most external sampling point

(5) and the most internal sampling points (1 and 2) was about four kilometers. Based on the discovery of *Haplosporidium pinnae* at the outer sites in April, a field experiment was set up to determine the pattern and rate of the spread of mortality within this dense population. As it was obvious that MMEs started to spread quickly through the inner part of the MNP, we set up the study with the aim of learning more about the pattern of the spread of the disease.

Two 100 m surveillance sentinel transects—transect 1 (T1) and transect 2 (T2)—were placed in the Small Lake (Figure 2) in September 2019, perpendicularly to the shoreline and extending towards a deeper part of the inlet (Figure 3). The deepest points along the transects were 3.8 m for T1 and 7.3 m for T2. All the clinically healthy pen shells were tagged, and their positions along the transects were recorded withina2m corridor on each side of the transect line, covering approximately 400 m2 along each transect. A caliper was used to determine the maximum unburied length (UL) and minimum dorsoventral height or minimum width (w), and the maximum height was determined using the formula Ht = 1.79 w + 0.5 + UL, as described by García-March and Vicente [29]. The data were recorded underwater on a plastic slate, while the depth was determined using a diving computer. Along transect T1, there were 75 individuals with the density of 20.8 per 100 m2, while along T2, there were 106 individuals with a density of 37.83 individuals per 100 m2. Every 2 weeks from September to December 2019, two divers clinically examined all the pen shells along both transects and noted the results of the clinical examinations and the numbers of live specimens. The temperature and the salinity of the sea were measured and noted daily.

**Figure 3.** Labeling healthy individuals along the transect.

#### *2.3. Non-Lethal Sampling*

As the mortality rate within the study area rose to 20% in September, biopsies of the mantle tissue from 20 randomly selected individuals along both transects were taken to assess the non-lethal sampling methodology and to obtain information on the prevalence of infections. The biopsies of the mantles were collected according to the method described by Sanna et al. [30]. Obtaining biopsies did not cause significant damage since the individuals' valves were held open with a wooden stick (diameter = 0.5 cm), which was placed in proximity (4–5 cm) to the hinge ligament, and about 50 mg of mantle tissue samples were removed using forceps. The stick was then removed, and the tissue sample was stored in a 5 mL tube. Once the diver returned to the surface, the tissue samples were preserved in EtOH and transferred to the laboratory in a refrigerated box.

#### *2.4. Statistical Analysis*

Differences in the morphometric data between transects were compared with a multivariant regression analysis using STATA 13.1 (Stata, College Station, TX, USA). Survival curves were calculated according to the Kaplan–Meier method [31].

#### *2.5. Laboratory Procedures*

The live mollusks that were sent to the laboratory were opened by cutting the adductor muscle according to the methods described by Morton and Puljas [32]. The in situ appearance of the organs was evaluated. Samples of the gills, mantle, digestive gland, gonads, and muscles were collected from each individual for both molecular and histological purposes and were immediately preserved in ethanol or Davidson fixative, respectively [28].

#### *2.6. Histological Analysis*

Samples of the organs fixed in Davidson solution were processed for histological examination. Tissues were dehydrated with an ethanol series, cleared in xylene, embedded in paraffin, sectioned at 3 μm, and mounted on Microme EC 350-2 slides (Thermo Scientific, Waltham, MA, USA). Mounted slides were heated to 60 ◦C, deparaffinized in xylene, and rehydrated in an ethanol series and distilled water, followed by staining with hematoxylin and eosin (H&E, Harris).

#### *2.7. DNA Extraction*

DNA was extracted from approximately 25 mg of the gill, mantle, digestive gland, gonads, and muscle tissues, respectively, which were preserved in either EtOH or RNAlater, using a MagMAX CORE Nucleic Acid Purification Kit (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's instructions on a KingFisher Duo Prime Purification System (Thermo Scientific, USA). The extracted DNA was stored at −20 ◦C until the analysis.

#### *2.8. PCR for Detection of Haplosporidium spp.*

For the detection of *Haplosporidium* spp., the *Haplosporidium*-specific primers described by Renault et al. [33] were used. We used 2 μL of extracted DNA and 0.5 μM of the primers HAP-F1 (5 -GTTCTTTCWTGATTCTATGMA-3 ) and HAP-R2 (5 -GATGAAYAATTGCAAT CAYCT-3 ). PCR reactions were performed using the GoTaq G2 Hot Start Colorless Master Mix (Promega, Madison, WI, USA) on a ProFlex PCR System (Applied Biosystems, USA) with a final volume of 20 μL. The temperature protocol involved enzyme activation at 95 ◦C for 2 min, followed by 40 cycles of denaturation at 94 ◦C for 30 s, primer annealing at 48 ◦C for 30 s, and elongation at 72 ◦C for 60 s, and the process was completed with a final elongation step at 72 ◦C for 5 min. The results of the PCRs were checked through electrophoresis on a QIAxcel system (Qiagen, Hilden, Germany).

#### *2.9. PCR for Detection of Mycobacterium spp.*

For the detection of *Mycobacterium* spp., we used two protocols. For the amplification of a segment of the *hsp65* gene, we used the primers Tb11 (5 -ACCAACGATGGTGTGTCCAT-3 ) and Tb12 (5 -CTTGTCGAACCGCATACCCT-3 ), which were first described by Telenti et al. [34]. The thermal protocol involved enzyme activation at 95 ◦C for 2 min, followed by 45 cycles of denaturation at 94 ◦C for 60 s, primer annealing at 60 ◦C for 60 s, and elongation at 72 ◦C for 60 s, and the process was completed with a final elongation step at 72 ◦C for 5 min. For the amplification of a segment of the mycobacterial 16S rRNA gene, we used the primers 246 (5 -AGAGTTTGATCCTGGCTCAG-3 ) and 264 (5 - TGCACACAGGCCACAAGGGA-3 ), which were first described by Böddinghaus et al. [35]. The thermal protocol was the same as that with the first set of primers, except for the annealing temperature, which was 55 ◦C. All the PCR reactions were performed using the GoTaq G2 Hot Start Colorless Master Mix (Promega) on a ProFlex PCR System (Applied Biosystems). For each reaction, we used 2 μL of extracted DNA and 0.5 μM primers with a final volume of 20 μL.

#### *2.10. Sequencing and Phylogeny*

To obtain longer sequences of the haplosporidian 18S rRNA gene for phylogenetic comparisons, for all the samples that tested positive for haplosporidian DNA, we per-

formed PCR with the primers HPN-F1, HPN-F3, HPN-R3, and 16SB, as described by Catanese et al. [7]. All the PCR products obtained were sequenced, and the resulting sequences were aligned and assembled in continuous reads using MAFFT version 7 [36]. For a phylogenetic comparison of *Mycobacterium* spp., the PCR products obtained using 246–264 primer pairs were sequenced. Direct Sanger sequencing was performed by Macrogen Europe (Amsterdam, The Netherlands). The sequences obtained [37] were identified using BLAST [38]. The sequences were aligned with MAFFT v7.388 [37] with the default parameters. The phylogenetic analysis was performed in MEGA 10.0.5 [39]. The best model was selected based on the lowest BIC score. Maximum likelihood trees were calculated with the following settings: Five discrete gamma categories, use of all sites, SPR level 5 heuristic method with a very strong branch-swap filter, the K2+G substitution model for the haplosporidian tree, and the HKY+G+I substitution model for the *Mycobacterium* spp. tree. Phylogeny was tested with the bootstrap method with 1000 replications. The obtained tree was visualized in iTOL [40].

#### **3. Results**

During the study, the health checks of noble pen shells (*Pina nobilis*) collected on 15 sampling sites were included, five in the MNP and 10 along the Istrian coast during 2019 and the first half of 2020 (Figures 1 and 2; Table 1).

**Table 1.** List of sampling points and samples collected during 2019 and 2020, which were analyzed using PCR for detection of *Haplosporidium* spp. (Haplo) and *Mycobacterium* spp. (Myco) and using histology.


Legend: MNP: Mljet National Park; I: Istrian peninsula; 2/2: Two positive animals out of two tested; n/d: Analysis not performed due to the absence of appropriate samples; \* EtOH: Ethanol; \*\* HCHO: Formaldehyde.

#### *3.1. Description of the Pen Shells Collected in Situ*

During the monitoring of the sampling sites included in this study, we observed the different appearances of the present noble pen shells. Healthy individuals, individuals with disease symptoms, and even empty shells were observed (Figure 4). The external symptoms were very scarce, and the affected animals displayed weakened reactions to stimuli, such as gaping (slower closing of shells) after touching or increased drifting and obvious shrinkage of the mantle, causing the illusion of a greater space between the shells than there was. At the sites with high mortalities of pen shells, many empty shells remained in an upward position, causing the illusion that the animals were still alive. In this situation, soft tissues were never found. When the sick animals were dissected, a certain amount of liquid was observed within the shells, and the mantle was darker than usual (Figure 5).

**Figure 4.** Clinical inspection of the live noble pen shells in situ, where both healthy and sick animals were in an upward position. Differentiation between them was made by checking their reactions to stimuli. The healthy specimens quickly closed their shells.

**Figure 5.** Darker mantle and shrunken soft tissues in a noble pen shell infected by *Haplosporidium pinnae*.

#### *3.2. Results of the Pilot Study (Morphometrics, Environmental Conditions, and Mortality Patterns)*

During the first health screening in the MNP, which was carried out in mid-April 2019, a total of six specimens were collected in the sampling points shown in Figure 2. Three of them were collected in the Small Lake (sampling points 1 and 2), and no changes in appearance were observed. However, two out of the three samples tested positive for *Mycobacterium* spp. The others were collected outside the lake-like inlets, one in Gonoturska (sampling point 4) and two around Cape Lenga (sampling point 5). In Gonoturska

(sampling point 4), there were no signs of unusual behavior, and a single sample tested negative for both pathogens. Around Cape Lenga (sampling point 5), a high mortality rate of up to 80% was noticed, which was based on the presence of unresponsive specimens and empty shells. In the other four sampling points, the mortality rate was about 9%, which is considered normal. Laboratory tests revealed the positive discovery of *Haplosporidium pinnae* in both individuals from sampling point 5—with both histology (Figure 7a–d) and PCR—while both specimens tested negative for *Mycobacterium* spp. (Table 1). At the beginning of September, a mortality rate of almost 100% was observed in the Big Lake, and a rate of about 20% was observed in the Small Lake. Following the spread of mortality in the Big Lake and when setting up the pilot study, we found that all the individuals were adults, and those along T1 had a mean height of 37.1 cm (±0,65), mean width of 15.28 cm (±0.35), and calculated mean maximum height of 64.56 cm (±1.11). The mean height of the individuals along T2 was 39.19 cm (±0.56), they had a width of 14.77 cm (±0.28), and the calculated maximum height was 65.64 cm (±0.76). The detailed morphometric properties of the individuals marked along both transects did not differ significantly (Supplement 1). At the beginning of the surveillance in April, the temperature of the sea was 15 ◦C, and it increased over the following months, reaching 27 ◦C in August, and slowly decreased over the study period from 25 ◦C at the beginning of September to 10 ◦C in December. During the whole study period, the salinity had an average value of 37.98%—ranging from 35.30 to 39.20%—on the surface or 38.04%—ranging from 35.30 to 39.00%—at a depth of 2 m.

The mortality rate during April 2019 at sampling point 5 was estimated to be 80%, and in the rest of the study area, it was estimated to be about 9% based on the number of empty shells (Figure 2). During the control diving on August 1, the mortality in the Big Lake was about 20%, and at the beginning of September, 100% mortality was observed in the Great Lake, also in addition to the increased mortality in the Small Lake (Table 2). At the time of setting up the transects, along T1, there were 62 dead individuals out of a total of 137 (Mt = 45.26%), and along T2, there were 28 dead individuals out of 134 (Mt = 20.90%). Over the study period, the mortality rate increased, resulting in the final survival of one individual from the T1 and three individuals from T2.


**Table 2.** Number of dead noble pen shells found along the transects from the setup of the pilot study (4 September 2019) until its completion (25 November 2019).

The Kaplan–Meier survival curves indicate two mass mortality events during the study period: The first was between 4 and 18 September, when 52 animals died, and the second was between 9 and 24 October 2019, when 57 animals died (Figure 6 and Table 2). By 25 November 2019, one of the initial 75 pen shells were alive along T1, corresponding to a survival rate of 1.33%, while three of the initial 106 pen shells were alive along T2, corresponding to a survival rate of 2.83%. The overall survival rate was 2.21%.

During the clinical checks, we observed that the mortalities affected certain individuals and spread diffusely to others in the close vicinity, affecting primarily those in the shallower area of the Small Lake and, later, those in the deeper parts.

#### *3.3. Necropsy and Histological Findings*

Six live animals were sacrificed for health evaluation, and organs from 37 additional animals were collected for molecular analysis (Table 1). Live animals were collected before the first notification of the MME, while the organs were collected prior, during, and after the events. On dissection, two of the animals collected in April showed a darker mantle and reduced soft tissues (Figure 5). The results of the histological analyses in the infected animals disclosed moderate to heavy infiltration of hemocytes into the connective tissues of the digestive gland and gonads, as well as light necrosis in the connective tissues of the gills, digestive gland, and muscle (Figure 7a). Uni-nucleated *Haplosporidium*-like parasite stages were observed in the connective tissues of the mantle (Figure 7b) and musculature (Figure 7c). Different *Haplosporidium*-like parasite stages, such as uni- or binucleate stages and plasmodia, were present in the connective tissues and epithelia of the digestive glands (Figure 7d,e). Intra-hemocytic stages of uninucleate cells were observed in the heavily infected specimens, in which sporogonia were observed in the epithelia of the digestive gland (Figure 7e,f). However, very few specimens were analyzed using histology, and sporulation was observed in the digestive gland in the sample collected in May 2020 in Vinkuranska vala in the Istrian peninsula.

#### *3.4. Molecular Analysis for the Presence of Haplosporidium spp. and Mycobacterium spp.*

Eleven out of the 43 animals tested positive for *Haplosporidium* spp. (Table 1). Since the DNA was isolated from the digestive glands, mantles, gonads, and muscles of the infected animals, mostly the digestive glands and gonads, followed by muscles and, in a few animals, mantles were positive for *Haplosporidium* spp. Here, it should be emphasized that four biopsies of mantles tested positive for *Haplosporidium* spp., as well. Furthermore, eight out of 43 samples tested positive for *Mycobacterium* spp. As in the case of *Haplosporidium* spp., two of the positive cases were found with biopsies.

**Figure 7.** Histological findings in the organs of noble pen shells: Moderate infiltration of hemocytes into the connective tissue of the digestive gland (**a**); uni- and binucleate cells of *H. pinnae* in the connective tissue of the mantle (**b**); uninucleate cells of *H. pinnae* in the muscle (**c**); uninucleate cells and plasmodia in the connective tissue of the digestive gland (**d**); intra-hemocytic stages of uninucleate cells in the connective tissue of the digestive gland (**e**); sporulation stages of *H. pinnae* in the epithelium of the digestive gland with a disruption of the digestive tubules (**f**).

#### 3.4.1. Sequencing of *Haplosporidium* spp. Isolates

All the *Haplosporidium*-positive PCR products (1451 bp) were successfully sequenced, and since the obtained sequences did not differ, only one haplotype was deposited in the GenBank (accession number MT367896). The sequences of the Croatian isolates (Figure 8) showed 99.8% identity with the Italian isolates deposited by Scarpa et al. [23] and Tiscar et al. (unpublished data) and 99.7% with the Spanish isolates [8], confirming the presence of *Haplosporidium pinnae* in all the positive noble shells.

**Figure 8.** Maximum likelihood phylogenetic tree for haplosporidians. The numbers on the branches represent bootstrap values.

3.4.2. Sequencing of *Mycobacterium* spp. Isolates

In addition, all the PCR products that tested positive for *Mycobacterium* spp. (985 bp) were sequenced, and they were identical. For this reason, only one haplotype was deposited in the GenBank (accession number MT367873). The obtained sequence was identical to the Italian one (Figure 9) obtained by Carella et al. [10].

**Figure 9.** Maximum likelihood phylogenetic tree for *Mycobacterium* spp. The numbers on the branches represent bootstrap values.

#### **4. Discussion**

The results of this study are in line with the results obtained by other Croatian researchers [14,15] and clearly show that MMEs have spread from the south to the middle parts of the Eastern Adriatic regions, as well as, ultimately, to the north. We first detected *Haplosporidium pinnae* in association with mortalities on the Croatian Adriatic coast in individuals collected in mid-April 2019 on the outer side of the Mljet Island, situated on the most southern point. At the same time, there were no mortality events among populations in the inner parts of the Mljet Island, and all the sampled animals tested negative for *H. pinnae,* but two individuals from the Small Lake tested positive for *Mycobacterium* spp., as shown in Table 1. Then, in May, mortalities and detection of samples from Elafiti Island (which is a bit more to the north) that were positive for *H. pinnae* and *Mycobacterium* spp. were reported [14]. During the summer period, mortalities were reported at different sites along the coast and close to the islands, mostly in the southern and middle parts of the Eastern Adriatic coast [15]. Interestingly, the data from the mentioned reports, as well as notifications from fishermen, divers, and other parties, indicated that the mortalities primarily affected habitats on the outer parts of the islands and were spreading towards sites that were closer to the mainland—from southern sites to northern sites. In 2019, there were no reports of mortalities in the northern part of the Eastern Adriatic coast, around the Istrian peninsula. The results of the laboratory analysis of the sampled animals supported the absence of both pathogens in the northern regions. Unfortunately, less than half a year later, during May 2020, the pathogens reached the northern sites along the Istrian peninsula and caused high mortalities of the natural populations (Table 1), as indicated by the diagnosis with *H. pinnae* and *Mycobacterium* spp.

Generally, it seems that the mortalities were triggered by an increase in the temperature in the spring, and they reached a high intensity when the sea had a high temperature during the summer. The results of the pilot study undertaken in the Small Lake of the Mljet National Park (Figure 2) support the hypothesis that the increase in mortalities was influenced by high temperatures. The MMEs did not affect populations in the inner parts of the lakes of the National Park from April until the beginning of August, when high mortalities were observed in the Big Lake. This was strange since the distance from the site of the first record of *H. pinnae* at Cape Lenga (Figure 2, sampling point 5) is about 4 km, and the water enters into the lakes from the open sea. At the same time, the MMEs spread to more distant sites in the Middle Adriatic. It seems that the inlet into the lakes does not allow the entry of large quantities of water, which also creates a barrier to the entry of pathogens and causes them to proliferate slowly. Most likely, the high temperature of the sea, which reached its maximum in August, strongly supported the extremely quick spread of the pathogen in the Big Lake and also affected the more isolated population in the Small Lake. The observations from the pilot study indicate that some regularity exists in the spread of the disease. The mortalities of individuals along T1, which was closer to the inlet of the water from the Big Lake, occurred earlier than those along T2. It was also observed that the spread of the disease was not linear, but rather diffuse, affecting primarily weaker individuals and spreading to others in close proximity in the dense population. A higher mortality rate was also observed in the shallower parts of the studied area. The difference related to the depth might be attributed to the higher water temperature in the shallower areas and the depletion of oxygen, which could additionally worsen the environmental conditions for the host. Additionally, it seems that higher water temperatures favor the propagation of both of the detected pathogens. *Mycobacterium* spp. require a higher water temperature for growth, as it is known that a representative of this genus, *Mycobacterium marinum*, a pathogen of marine fish, usually causes disease during warmer parts of the year with water temperatures above 25 ◦C [41]. Moreover, *Mycobacterium* spp. are opportunistic bacteria that cause chronic diseases, and it should be possible to detect their presence in healthy individuals long before they cause granulomatous lesions, as observed by Carella et al. [10] and Latos et al. [12]. Unfortunately, we analyzed very few samples using histology, and in our scarce histological slides, we did not observe the lesions described

by the aforementioned authors [10,12]. Interestingly, in the samples of mantle biopsies collected during the pilot study at the end of August, we detected *H. pinnae* in four out of the 20 tested animals and *Mycobacterium* spp. in two out the 20 animals, which could suggest that the parasite was more representative and more responsible for the MMEs compared to *Mycobacterium* spp. It should be emphasized that the pen shells in the Small Lake of the Mljet National Park did not show increased mortality for 6 months, although it was confirmed that they were infected with *Mycobacterium* spp. in April. Infection with *Mycobacterium* spp. does not appear to be sufficient to trigger a mass mortality event among pen shells, but its co-occurrence with *H. pinnae* contributes to high mortality. Similarly, Box et al. [42] concluded that high mortalities occur in noble pen shells that have been infected with *H. pinnae* and additionally aggravated by co-infection with *Mycobacterium* spp. or other Gram-negative bacteria.

From the results of morphometry, it was obvious that all the marked animals were adults, with the smallest reaching a maximum height of 37.9 cm. It seems that the older individuals were more susceptible to infection with *H. pinnae,* as previously reported by Vázquez-Luis et al. [7], as well as to cases of infection with other genera of *Haplosporidium,* where mortalities occurred in older individuals concurrently with the highest levels of infection [43].

The peak prevalence of the mortalities of Eastern oyster (*Crassostrea virginica*) caused by *Haplosporidium nelsoni* in Delaware Bay, USA [43] was reported in autumn. The mortalities observed in the pilot study with the pen shells peaked during the autumn and slowly decreased during the winter, similarly to those caused by *H. nelsoni*. From the Kaplan– Meier survival estimates (Figure 6), it was visible that the mortalities increased gradually and that the most numerous mortalities occurred at the end of September and again at the end of October. Unfortunately, only four animals survived until the end of November. Undoubtedly, the pathogens identified in our research were those detected as the cause of mortalities throughout the Mediterranean region. The phylogenetic analysis of our isolates of *H. pinnae* showed 99.8% similarity (Figure 8) with those described in the Ionian Sea [9] and 99.7% similarity with Spanish isolates [8]. The phylogenetic analysis showed that the 16S sequences of mycobacteria detected in *P*. *nobilis* were more distant from each other than some sequences of mycobacteria belonging to different species. For example, *M*. *sherrisii*, *M*. *stomatepiae*, *M*. *florentinum*, *M*. *triplex,* and *Mycobaterium* spp. from *P*. *nobilis* from Croatia (MT367873) and Italy (MH569646) are more similar to each other than the two strains of *Mycobaterium* spp. from *P*. *nobilis* from Italy (MH569647 and MH569649) (Figure 9). This suggests that the detected mycobacteria from *P*. *nobilis* belong to a heterogeneous group and are not members of the same species.

There are still doubts about what enabled the quick spread of the pathogen through the Mediterranean basin. It was undoubtable that the pathogen entered into the Adriatic Sea and spread from the south to the north by following the main flows of the currents [44]. It is hard to believe that only water currents carry the pathogen, and the pathogen could probably spread through pelagic larval stages of other intermediate hosts. Nevertheless, the histological finding of all the stages of *H. pinnae* in the same animal—uninucleate cells observed in the mantle, gills, gonads, digestive gland, and connective tissue, followed by plasmodia in the connective tissue of the digestive gland, and sporocysts in the epithelium of the digestive tubules (Figure 7a–f)—contributed to the direct transmission from one to another individual, as postulated by Catanese et al. [8]. The quick spread of mortalities within the dense population in the pilot study in the Mljet National Park suggests the direct transmission of *H. pinnae*. Since it is known that spores of *Haplosporidium* are persistent in the environment [45], the involvement of an intermediate host that is abundant in the marine environment and is capable of quick movement through water should be considered, and future studies should aim to prove this hypothesis.

Further joint efforts of marine biologists and invertebrate health experts should be engaged in aiming to understand why only adult specimens were present in particular marine areas, and the reasons were for the increased susceptibility of the noble pen shells to pathogens.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/jmse9070764/s1. Table S1: Morphometric characteristics of noble pen shells along the transect 1 (T1); Table S2: Morphometric characteristics of noble pen shells along the transect 2 (T2).

**Author Contributions:** Conceptualization, Ž.M.; data curation, M.B.; formal analysis, Ž.P. and I.G.Z.; funding acquisition, B.H.; investigation, A.P., O.P., and I.S.; methodology, D.O.; writing—original draft, S.Z. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** All the sampling was carried out with the permission of the Ministry of Environmental Protection and Energy (CLASS: UP/1-612-17/18-48/172, No. 517-05-1-1- 18-4 issued on 21 December 2018 and CLASS: UP/1-612-07/19-48/193, No. 517-05-1-1-19-3 issued on 11 September 2019).

**Data Availability Statement:** All the data are available upon reasonable request from the corresponding author.

**Acknowledgments:** The authors kindly acknowledge Angela Bradari´c for the technical support in the field and the laboratory work, Pavel Ankon for the instructions on non-lethal sampling, and Ana Car and Iris Dupli´c Radi´c for sharing the salinity data in the Small Lake during the pilot study. Special thanks go to Isabelle Arzul and the EURL for mollusk diseases (IFREMER, La Tremblade) for their confirmation of the detection of *Haplosporidium pinnae*.

**Conflicts of Interest:** The authors declare no conflict of interest.

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## *Article Bonamia exitiosa* **in European Flat Oyster (***Ostrea edulis***) on the Croatian Adriatic Coast from 2016 to 2020**

**Dražen Orai´c 1,\*, Relja Beck 1, Željko Pavlinec 1, Ivana Giovanna Zupiˇci´c 1, Ljupka Maltar 2, Tihana Miški´c 2, Žaklin Acinger-Rogi´c <sup>2</sup> and Snježana Zrnˇci´c <sup>1</sup>**


**Abstract:** The annual production of European flat oysters (*Ostrea edulis*) in Croatia is about 50 to 65 tons, and it has a long tradition. All Croatian oyster farms are subjected to the national surveillance program aiming to detect the presence of *Bonamia ostreae* and *Marteilia refringens* according to the Council Directive 2006/88/EC. Within the surveillance program, the first findings of the parasite *Bonamia* spp. occurred in 2016 in two production areas in the north and south of the Eastern Adriatic coast. The repeated findings of the parasite were noted up to 2020 but also on two additional sites in the north. The parasite was detected by cytological analysis of stained heart smears, histological examination, and PCR. PCR positive samples were sequenced for SSU rDNA gene, and BLAST analysis confirmed infection with *Bonamia exitiosa*. Attempts to prove the Pacific oyster as a putative vector of the parasite failed. The infection prevalence from 2016 until 2020 ranged from 3.3 to 20% in different sites. No mortalities were reported from the infected sites, and it seemed that infection of flat oysters with *B. exitiosa* did not affect their health. The study has not shown the source and way of infection spread, which imposes the need for more comprehensive molecular and epidemiological studies.

**Keywords:** *Bonamia exitiosa*; bonamiosis; Croatian Adriatic coast; *Ostrea edulis*; prevalence; surveillance

#### **1. Introduction**

The production of European flat oyster (*Ostrea edulis*) has a long tradition and economic significance in many countries of the Mediterranean basin and the East Atlantic coast. Along the Croatian coast, flat oysters have been cultivated for centuries, and in recent decades, the annual production has ranged from 50 to 65 t [1]. Production of *O. edulis* has stagnated since the 70s of the last century as a result of the introduction of pathogens into the susceptible population. Therefore, diseases have caused large losses and dramatically affected production [2–4]. One of these pathogens is the protozoan parasite *Bonamia ostreae*, belonging to the clade "microcell" within Haplosporidia [5]. Various methods have been developed to identify and confirm the presence of parasites of the genus *Bonamia*, but each of them has certain limitations. Microscopy of gill and heart-stained tissue impressions and histological slides are methods to identify and confirm *Bonamia* species. Identifying parasite species due to morphological similarity is time-consuming and requires experience and expertise. However, both these methods have low sensitivity and specificity [6–9], similar to TEM, which seems to be insufficient in the identification of species [10]. Diagnostic sensitivity has significantly improved with the implementation of molecular methods, such as PCR [11,12] or in situ hybridization.

Based on molecular studies of small ribosomal subunit rRNA genes, the genera *Bonamia* is phylogenetically positioned into a clade microcell within the genus *Haplosporidia*, and within it, includes four species: *B. ostreae*, *B. exitiosa*, *B. roughleyi*, and *B. perspora* [13–16].

**Citation:** Orai´c, D.; Beck, R.; Pavlinec, Ž.; Zupiˇci´c, I.G.; Maltar, L.; Miški´c, T.; Acinger-Rogi´c, Ž.; Zrnˇci´c, S. *Bonamia exitiosa* in European Flat Oyster (*Ostrea edulis*) on the Croatian Adriatic Coast from 2016 to 2020. *J. Mar. Sci. Eng.* **2021**, *9*, 929. https:// doi.org/10.3390/jmse9090929

Academic Editor: Patrizia Pagliara

Received: 23 June 2021 Accepted: 21 August 2021 Published: 27 August 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

All species of *Bonamia* cause infection of flat oyster hemocytes, leading to death [12]. In some cases, parasites are present in low prevalence and with little impact on the flat oyster population [17]. It has been reported that flat oysters exposed to parasites over a long period develop a certain degree of resistance [18]. *Bonamia exitiosa* was for the first time reported as the cause of the Chilean oyster's (*Ostrea chilensis*) high mortality in New Zealand when only 9% of affected flat oysters survived [19]. Catastrophic and rapid mortality was caused by the same parasite on triploids of the Suminoe oyster (*Crassostrea ariakensis*), introduced for experimental cultivation in 2003 in Bogue Sound, North Carolina [20], in an area where the crested oyster (*Ostreola equestris*) was an indigenous species [15].

This paper presents the findings, distribution, and prevalence of the *Bonamia exitiosa* in the samples of flat oyster analyzed within the national surveillance program for the infection with *Bonamia* parasites.

#### **2. Materials and Methods**

#### *2.1. Sampling*

The sampling points within production areas were determined through an intensive monitoring program in the period from 1998 to 2000. Each year, according to the national surveillance program, authorized veterinarians sampled 30 adult flat oysters on each of nine sampling points along the Croatian Adriatic coast (Figure 1, Table 1). The samples were packed in a transport refrigerator and delivered to the laboratory within 24–36 h.

**Figure 1.** Map showing European flat oyster (*O. edulis*) production areas with sampling points included in the national surveillance plan. A production area in the south with five sampling points and three production areas in the north Croatian Adriatic Sea tested positive (red numbers) for *B. exitiosa* in 2016–2020. The numbers refer to production areas/sampling sites listed in Table 2.


**Table 1.** Positive findings of *Bonamia exitiosa* within sampling area showing sampling dates, sampling points, and prevalence using different diagnostic techniques in Croatia during the period 2016–2020.

\* Samples with positive finding of *B. exitiosa.*

**Table 2.** Total number of oysters analyzed for the presence of *Bonamia* spp. in the period 2016–2020.


\* labels the number of the sampling site corresponding to the map of sampling sites.

All samples submitted within the surveillance program in the period 2016 to 2020, from the *B. exitiosa* positive sites are listed in Table 2 and sites are shown in Figure 1. Namely, from the site of the Medulin Bay during the mentioned period (2016–2020), 180 flat oysters were submitted; from the Mali Ston Bay with five sampling sites, there were 900 individuals, while 180 individuals were submitted from the sites from Lim Bay, Savudrija Bay, and Marina Bay, respectively.

After the first finding of *B. exitiosa* positive flat oysters in the northern part of the Eastern Adriatic coast, epidemiological surveillance was carried out. It was found that there were some populations of Pacific oyster (*Magallana* = *Crassostrea gigas*) present in the Medulin Bay production area, and during 2017, a sample of 18 animals was collected to examine them as a possible source of infection. In 2018, after the detection of *B. exitiosa* in Lim Bay, another batch of 30 Pacific oyster was tested for the presence of this parasite.

#### *2.2. Tissue Processing, Cytological and Histological Examination*

After dissection of each oyster submitted for diagnostics, including Pacific oysters, tissues were collected for cytology, histology, and molecular analysis.

For cytological diagnosis of *Bonamia* spp., a piece of the heart of each oyster was taken and briefly dried on absorbent paper. A series of heart imprints were made in two rows on a glass slide. Imprints of five specimens were made on one slide and stained with Hemacolor® staining kit according to manufacturer instructions (Merck, Darmstadt, Germany).

For histological examination, transversal sections of soft tissue, including gills and the digestive gland of each oyster, were placed in histo-cassettes and immersed in Davidson's formalin, alcohol, acetic acid (AFA) fixative. Davidson-fixed tissues were dehydrated through a graded series of ethanols, succeeded by xylene, and embedded in paraffin, sectioned at 3 μm, and mounted on Microme EC 350-2 slides (Thermo Scientific, Waltham, MA, USA). Mounted slides were heated to 60 ◦C, deparaffinized, and rehydrated in xylene, a graded series of alcohol, and finally, water, followed by staining with hematoxylin and eosin (H&E).

For detection of *Bonamia* spp. by PCR, gill samples were taken and preserved in ethanol 96% until DNA extraction.

Stained tissue impressions and histological sections were examined on a Zeiss Axiskop-2 binocular microscope (Carl Zeiss, Jena, Germany) at 400× and 1000× magnifications with immersion oil.

#### *2.3. Molecular Detection and Characterization*

#### 2.3.1. DNA Extraction

DNA used in all molecular analyses was extracted from gill tissues from each animal. From the samples collected from 2016–2019, DNA was extracted using MagMAX CORE Nucleic Acid Purification Kit (Thermo Fisher Scientific, USA) on KingFisher Flex System (Thermo Fisher Scientific, Waltham, MA, USA). From the samples collected in 2020, DNA was extracted using innuPREP AniPath DNA/RNA Kit—IPC16 (Analytik Jena, Germany) on InnuPure C16 touch (Analytik Jena, Jena, Germany). Both methods of extraction were done according to the manufacturer's instructions.

#### 2.3.2. PCR and Sequencing

In the period from 2016 to 2018, a modified method developed by Cochennec et al. [11] for the detection of *Bonamia* sp was used. The reaction mix contained 10 μL of GoTaq G2 Hot Start Master Mix (Promega, Madison, WI, USA), 60 ng of DNA as measured on a DS-11 Series Spectrophotometer (DeNovix, Wilmington, NC, USA), 0.5 μM of each primer (BO 5 3 , BOAS 5 3 ), and nuclease-free water to the final volume of 20 μL. The amplification was performed using ProFlex PCR System (Applied Biosystems, Waltham, MA, USA) with an initial denaturation at 95 ◦C for 2 min, followed by 35 cycles of denaturation at 94 ◦C for 1 min, primer annealing at 55 ◦C for 1 min, elongation at 72 ◦C for 1 min, and ending with the final elongation step at 72 ◦C for 7 min. The presence of PCR products was examined by electrophoresis on a QIAxcel system (Qiagen, Hilden, Germany) using the QIAxcel DNA Screening Kit. For species identification, direct Sanger sequencing of the PCR products was performed by Macrogen Europe. Sequence identity was confirmed by BLAST [21].

#### 2.3.3. Real-Time PCR

For the samples from 2019 and 2020, for the simultaneous detection of *B*. *ostreae* and *B*. *exitiosa* DNA, a probe-based real-time qPCR assay developed by the European Union Reference Laboratory (EURL) for Mollusc Diseases (Ifremer, La Tremblade, France) was

performed [22]. The reaction mix contained 10 μL of GoTaq® Probe qPCR Master Mix, 2X (Promega, Madison, WI, USA), 0.3 μM of each primer (BO2\_F 5 AAATGGCCTCTTCC-CAATCT 3 , BO2\_R 5 CCGATCAAACTAGGCTGGAA 3 , BEa\_F 5 GACTTTGACCATCG-GAAACG 3 , BEa\_R 5 ATCGAGTCGTACGCGAGTCT 3 ), 0.2 μM of each double-labelled probe (BO2\_probe 5 HEX—TGACGATCGGGAATGAACGC—BHQ-1 3 , BEa\_probe 5 FAM—GGCAGCGAATCGATGGGAAT—BHQ-1 3 ), 25 ng of template DNA as measured on a DS-11 Series Spectrophotometer (DeNovix, Wilmington, USA), and nuclease-free water to the final volume of 20 μL. The amplification was performed on qTower<sup>3</sup> thermal cycler (Analytik Jena, Jena, Germany) with the following program: 95 ◦C for 2 min for polymerase activation, followed by 40 cycles of denaturation at 95 ◦C for 15 s and annealing at 60 ◦C for 30 s. The fluorescence was recorded at the end of each cycle with HEX and FAM filters. To detect possible differences, all positive samples were amplified and sequenced using the same method described above.

#### 2.3.4. Sequencing and Phylogeny

Obtained sequences were analyzed using Geneious Prime 2020.0.1 software. For the phylogenetic analysis of the obtained sequences, similar sequences available in Gen-Bank [23] were used and identified using BLAST [21]. Sequences were aligned with MAFFT v7.388 [24] using default parameters. Phylogenetic analysis was performed in MEGA 10.0.5 [25]. The best model was selected based on the lowest BIC score. The maximum likelihood tree was calculated with the following settings: K2+G substitution model, five discrete gamma categories, used all sites, and SPR level 5 heuristic method with very strong branch swap filter. Phylogeny was tested by the bootstrap method with 1000 replications. The obtained tree was visualized in iTOL [26]. Four sequences belonging to *Minchinia* spp. were used as an outgroup.

#### **3. Results**

Until 2016, the results of cytological examination of heart imprints, histological slides of oysters' organs, and PCR tested negative for the presence of parasite *Bonamia* spp. There were no reported mortalities, and there were no symptoms of the disease on the oysters from the production areas included in the surveillance program. In the May of 2016, for the first time, there were positive findings of the parasite *Bonamia exitiosa* from two production areas (Figure 1, Table 2).

The first one was in the north of the Eastern Adriatic Coast in Medulin Bay where 3 out of 30 animals tested positive for *Bonamia* sp. using conventional PCR and heart imprints, and the second one was in the south of the Croatian Adriatic Sea in the Mali Ston Bay. In all five sampling points, there were *Bonamia* sp. positive samples; (in Mali Ston and Brijesta, 3 out of 30, respectively, by heart imprints and five and six using conventional PCR; in Bistrina and Bjejevica, 2 out of 30, respectively, in imprints and two and three by PCR; and Sutvid, 1 out of 30 by each technique. Following the first finding in Medulin Bay in May, 1 out of 30 animals sampled in November of the same year tested positive. In 2017, positive samples were detected again in Medulin Bay in May and November and also in October of 2020. In 2019, a conventional PCR was replaced with a new molecular method, real-time PCR developed by EURL for mollusc diseases, and validated through interlaboratory proficiency testing as a screening method in the surveillance program. All samples analyzed in 2019 and 2020 were tested using real-time PCR. Therefore, samples collected in Medulin Bay in October 2020 were screened using real-time PCR, and positive samples were tested using conventional PCR and submitted for sequencing. In Mali Ston Bay, there have been no positive findings since 2016. In June 2018, there were positive findings of *B. exitiosa* in 2 out of 30 flat oysters sampled in the Lim Bay detected by imprints and three by PCR and in the same number by each method (two by imprints and three by PCR out of 30 flat oysters) sampled in the Savudrija Bay, two sampling points in the Northeastern Adriatic coast. In Savudrija Bay, in 2020, 4 out of 30 flat oysters were found positive by real-time PCR, confirmed by conventional PCR and sequencing. However, all samples from Marina Bay tested negative for the presence of *Bonamia* sp. during the whole studied period using heart imprints, conventional and real-time PCR.

All pacific oysters collected in the Medulin Bay (*n* = 18) and from the Limski Bay (*n* = 30) tested negative for the presence of *Bonamia* sp by both used techniques.

Microscopic examination of heart imprints revealed mononuclear cells (Figure 2a) and, in fewer numbers, binuclear cells of light blue-stained cytoplasm and red-stained nuclei that were located centrally or near the center in mononuclear stages (Figure 2b).

**Figure 2.** (**a**) Heart imprints of *Ostrea edulis* infected by cells identified as *Bonamia exitiosa* within hemocyte; (**b**) Heart imprints of *Ostrea edulis* infected by mononuclear and binucleated cells identified as *Bonamia exitiosa.* Hemacolor staining, 1000× magnification.

Microscopic examination of histological slides showed weak to moderate infiltration of hemocytes in the connective tissue of gills, digestive glands, and gonads. Weak to medium necrosis with low to medium presence of *Bonamia*-like parasites (Mean size: 1.95 μm) were found in the same tissues. In a small number of infected animals, the connective tissue of the gonads had marked necrotic changes and *Bonamia*-like cells were found (Figure 3a,b).

**Figure 3.** (**a**) Histological section of *Ostrea edulis*; *Bonamia*-like parasite in necrotic tissue of digestive gland (**b**) Gonadal connective tissue cells infected with *Bonamia*-like parasite. H&E staining, 1000× magnification.

**Figure 4.** *Bonamia* spp. maximum likelihood phylogenetic tree. Bootstrap values higher than 75 are displayed.

#### **4. Discussion**

Until 2010, the diagnostic procedure for detection of *Bonamia* spp. was based on the results of heart imprints and histopathological evaluation of each oyster. Subsequently, since 2010, examinations of stained imprints and PCR screening methods for the same sample have been combined according to a dual detection strategy described by Carnegie and EU legislative [27,28]. In the period from 2010 to the first finding of *Bonamia* spp. positive oysters, 1870 individuals were tested in total.

In the spring of 2016, *Bonamia* parasite was detected for the first time by conventional PCR in the samples collected from the production area in the northern part of the Eastern Adriatic Sea, sampling point 7 (Figure 1). The sample consisted of 30 adult individuals (more than 2-y old), which were reported to be more susceptible to infection with *Bonamia* spp. compared to younger ages [29,30]. The microcellular finding was confirmed in stained heart imprints. The suspicion for the presence of *Bonamia ostreae* was set up, but sequencing revealed the presence of *Bonamia exitiosa*. Shortly following, the samples collected from the production area in the southern part of the Croatian Adriatic coast, the Mali Ston Bay, were submitted for diagnostics. In each of five samples (*n* = 150) from different sampling points, the same diagnostic procedure revealed that 17 flat oysters were PCR positive for *Bonamia* spp. Again, an affiliation of *B. exitiosa* was determined by sequencing of the PCR positive product. It was found that the prevalence of *B. exitiosa* in the different sampling points within Mali Ston Bay notably varied from 3.3% to 20.0% per sampling point (Table 2) when tested by PCR compared to prevalence of 3.3% to 13.3% after evaluation of stained heart imprints. In stained heart imprints, single cells with a central nucleus were observed by microscopic examination. However, outside hemocytes, there was a lower abundance of cells with two nuclei. This corresponds to the previously described imprints findings of *Bonamia exitiosa* in flat oysters [31–34]. Histopathological examination of positive PCR samples of *B. exitiosa* confirmed *Bonamia*-like cells in one-third of samples. In this study, the histological examination was proven as a less sensitive method in line with the finding of Lynch et al., who previously related the low prevalence of parasite to the low sensitivity [35]. It can also mean that environmental conditions were not favorable for the spread and development of infection or that our PCR positive findings were related to the detection of non-infective parasite stages [36].

Interestingly, *B. exitiosa* was detected in all five sampling points in the Mali Ston Bay only in spring of 2016 and never again until 2020, although surveillance was carried out regularly each year, and in total, 750 oysters were examined during this period and tested negative for the presence of *B*. *exitiosa*. At the same time, in the north of the Adriatic coast, *B. exitiosa* was confirmed for the first time by sequencing and heart imprints in spring 2016, and in following years, it was detected in another two production areas. In Medulin Bay, the findings of *B. exitiosa* continuously occurred from the spring of 2016 to autumn of 2020, except in 2019. During the seasons with positive findings of the parasite, the prevalence of the PCR positive samples varied from 3.3% to 16.7%. Unexpectedly, in 2019, there were no *B. exitiosa* positive/infected oysters, which could be explained with a low number of analyzed samples. There were only 90 individuals submitted for diagnostics of *Bonamia* spp.

It has been experimentally proven that *B. ostreae* favors lower sea temperature for survival [37] and higher prevalence of bonamiosis in oysters, and increased mortality in the colder period with lower sea temperatures has been described. On the contrary, in the case of *Bonamia* spp. in *C. ariakensis*, it has already been observed that the disease occurred in the warmer period when temperatures exceeded 20 ◦C [38]. Epidemiological circumstances of our *B. exitiosa* positive cases are unlike each of the reported cases, as no increase in mortalities or disease prevalence has been observed over the last five years.

For the reason mentioned above, the real impact of the infection is hard to evaluate. The prevalence of the *B. exitiosa* along the Croatian Adriatic Coast varied from 3.35% to a relatively high prevalence of 20% in the sampling point Brijesta in the Mali Ston Bay, and the recent recorded prevalence was 16.7% in the Medulin Bay, the site of the first finding (Table 2). These results show a higher prevalence of *B. exitiosa* compared to the findings in the study from the Manfredonia Bay in the southern part of the Italian Adriatic coast [32] in 2007, in which only two oysters out of a total of 750 were positive for *B. exitiosa*. There were no constancy in the prevalence on the sampling site over the studied period. It was shown that sequences of *B. exitiosa* detected in *O. edulis* from the Mediterranean area, including the one from Manfredonia Bay, Spain and also *O. stentina* in Tunisia [31,32,39], are 99.3% similar to those in Croatia, which supports the fact that the same strain is circulating throughout the Mediterranean basin.

Another concern is the answer to the question of how the flat oysters on Croatian production sites became infected with *B. exitiosa* It is reported that Pacific oysters are a vector or reservoirs of *Bonamia* spp. [40,41]. Although there is no farming of this species in the Croatian part of the Adriatic Sea, it has been present in the Lim Bay since 1963 [42]. This invasive species was also confirmed in the central Adriatic, but it was not found in the locality of Mali Ston, a southern production area with positive findings of *B. exitiosa* in the spring of 2016 [43]. The first positive finding of *B. exitiosa* in the Medulin Bay followed by a positive finding in Limski Bay in 2018 aroused suspicion of Pacific oysters as a source of infection for flat oysters. Unfortunately, the suspicion had to be discarded as both samples, one consisting of 18 Pacific oysters from the Medulin Bay in 2017 and another consisting of 30 Pacific oysters from the Lim Bay, tested negative for the presence of *Bonamia* spp. Furthermore, until now, there have been no records of mortality or introductions of new batches of either Pacific or flat oysters or any other mollusc species into the infected production areas. The possibility of some vector that might transfer the parasite across the Adriatic Sea should not be fully discarded, but it is hard to prove it.

Since the source of infection with *B. exitiosa* in flat oysters in all recorded production areas in Croatia remains unknown, another susceptible species may be considered a source of infection for the flat oyster. It is known that another oyster species, dwarf oysters (*O. stentina*), also exist in the Adriatic Sea [44,45], and phylogeographic research has confirmed that *B. exitiosa* has always been found in *O. stentina* [39]. The hypothesis could be made that *O. stentina*, an inhabitant in the Adriatic Sea, could be a source of *B. exitiosa*, as *O. stentina* in Tunisia were found to be positive for this parasite [39,46].

Moreover, it was not possible to get any additional information on the source of *B. exitiosa*'s origin from phylogenetic analysis. As it is visible from the phylogenetic tree, the similarity of the Croatian isolate of *B. exitiosa* to different previously sequenced isolates from Chile or Australia was 100%. The similarity of different *B. exitiosa* isolates found in different oyster species around the world emphasize the fact that the SSU rDNA gene is highly conserved. Therefore, the sequencing of additional genes or the whole genome should be carried out to provide us with more details on the phylogeny of the Croatian isolates.

More comprehensive molecular studies of the *B. exitiosa*, together with investigation of the natural population of *O. stentina* from production areas and natural beds along the Eastern Adriatic coast to confirm the natural-historical origin of the parasite *B. exitiosa*, will enable a better understanding of the pathogen's presence in the Croatian flat oyster production area.

**Author Contributions:** Conceptualization, D.O. and S.Z.; formal analysis, R.B., I.G.Z. and Ž.P.; investigation, R.B., Ž.P., I.G.Z., D.O. and S.Z.; resources, S.Z. and D.O.; data curation, L.M., T.M. and Ž.A.-R.; writing—original draft preparation, D.O.; writing—review and editing, S.Z., Ž.P. and R.B.; visualization, I.G.Z. and D.O.; supervision, S.Z. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Croatian Ministry of Agriculture—Veterinary and Food Safety Directorate General.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available upon a reasonable request from corresponding author.

**Acknowledgments:** We would like to thank Isabelle Arzul and Bruno Chollet from EURL for Molluscs Diseases for confirming the results of our findings.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Article* **Histopathologic Lesions in Bivalve Mollusks Found in Portugal: Etiology and Risk Factors**

**Daniel Pires 1, Ana Grade 2, Francisco Ruano <sup>2</sup> and Fernando Afonso 1,\***


**Abstract:** Bivalve mollusks are an important resource due to their socioeconomic value and to the historical and genetic value of some species. Two nationally important oyster species-Portuguese oyster (*Crassostrea angulata*) and Japanese oyster (*Crassostrea gigas*) from distinctive areas in Portugal were studied to evaluate their sanitary status. Oysters were sampled from four different sites in Portugal. Oysters collected from Japanese oyster populations were cultivated in a strong oceaninfluenced environment and Portuguese oyster populations were cultivated in wild-beds. The histopathological examination of both oyster species revealed the presence of parasites in gills, mantle epithelium, digestive gland tubules and connective tissue, with a moderate prevalence. In both populations was observed hemocytosis in the connective tissue, edema and metaplasia in the digestive gland and tissues necrosis. In wild populations from Sado and Mira estuaries the prevalence of mud blisters and gill lesions were higher than from populations produced on 0.50 m tables from mudflats. Biosecurity measures and diagnostic techniques are fundamental to control pathogenic agents, including the identification of pathogens at an early stage in their life cycles. This will prevent diseases and improve pathogen reduction on transport of animals from different countries and regions to new production areas to avoid the transmission of diseases.

**Keywords:** bivalve mollusks; oysters; histopathology; parasites

#### **1. Introduction**

In Portugal, in the late 20th century, the two most important commercial oysters were the flat Oyster (*Ostrea edulis*) and the Portuguese Oyster *(Crassostrea angulata)* [1]. Initially, flat oyster commerce was more important, but in the 20th century it was replaced by the Portuguese oyster. The culture of bivalve mollusks is an activity with high expression in Portuguese aquaculture. It represents 55% of the whole production, being the main species, clams, oysters and mussels [2].

There was a significant change in bivalve mollusks production. Initially, it was exclusively chosen for semi-intensive production in large areas. Nevertheless, at this moment there is an intensification of production in less area, increasing bivalve mollusks density. This option results in lower growth rates, lower product quality, lower fertility rates and increasing diseases [3].

Microbiological contamination of water where aquaculture production is present, in most cases, is the result of industries, urban activities and leisure activities, causing fecal contamination. [4–6]. The microbiological organisms present in bivalve mollusks are diverse, with different populations of bacteria (namely, *Escherichia coli*, *Clostridium perfringens*, *Vibrio parahaemolyticus*, *Salmonella* and *Listeria*) [7–9], enteric viruses (namely, norovirus, enteric calicvirus, hepatitis A virus and other enteroviruses) [10,11] and protozoa (namely, *Crypto*sp*oridia* and *Giardia*) [12,13]. Generally, these microorganisms are not harmful to bivalve mollusks and do not cause lesions or disease.

**Citation:** Pires, D.; Grade, A.; Ruano, F.; Afonso, F. Histopathologic Lesions in Bivalve Mollusks Found in Portugal: Etiology and Risk Factors. *J. Mar. Sci. Eng.* **2022**, *10*, 133. https:// doi.org/10.3390/jmse10020133

Academic Editor: Snježana Zrnˇci´c

Received: 1 October 2021 Accepted: 30 December 2021 Published: 20 January 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Several diseases can affect different bivalve mollusks species and their production. Epizootics caused by fungi, viruses and protozoa can affect the entire bivalve production, as in Portugal and France in the 1970s where massive mortality caused by an iridovirus affected the Portuguese oyster production. Since 2009 a herpes virus (OsHV-1) has been the cause of massive mortalities in Japanese oyster (*Crassostrea gigas*), first in France and all over the most important European production countries of this species, including Portugal [3]. Japanese oyster aquaculture is a major primary production sector in many countries, but the industry has been threatened by mortality events for the last five decades [14]. In Portuguese and Japanese oyster, the main pathogens are virus, namely iridovirus, responsible for gill disease that affected Portuguese oyster production in the 1970's, and herpes viruses, namely Ostreid-herpesvirus 1 μvar (OsHV-1) that is related with summer oyster disease and affects the Japanese and Portuguese oyster populations [15]. From the first detection in the Eastern oysters (*Crassostrea virginica*) [14] in 1972, herpes virus has been identified in at least 20 bivalve species and has caused massive mortalities [14]. Herpes virus infecting bivalve mollusks are virulent pathogens of both larval and seed oysters. Mortalities of juvenile oysters associated with detection of herpes viruses have been reported from France, New Zealand, Spain and the USA [16–19].

In flat oyster, there are important diseases, including bonamiosis that is caused by *Bonamia ostrea* and *Marteilia refringens* that causes marteliosis and viruses such as *Ostreidherpesvirus* 1 μvar [20,21].

Regarding mussels *Mytilus* sp., the main pathogens are protozoa, such as *Marteilia refringens* that is the causative agent of marteliosis and copepods, such as *Mytilicola intestinalis* which causes red worm disease [22,23].

Japanese clam, European clam and cockles have been mainly affected by protozoa, such as *Perkinsus olseni*, the causative agent of Perkinsiosis [24–27] and *Haplo*sp*oridium tapetis* [26,28]. *Perkinsus olseni* is responsible for the high mortality of cockleshell populations. Perkinsiosis is caused by the protozoan *P. olseni*. This disease is a very common disease in clams and has caused massive mortalities in clam populations, contributing to the decrease in their production and consequently affecting their socioeconomic value [29]. Typical lesions that occur include cell disorganization, autolysis and necrosis of cells, both within and near the lesion [24].

The high prevalence of this pathogen in clams can be attributed to several factors, such as degraded sediments, high animal densities per unit area, lack of physical barriers between different culture beds, and transfer of animals carrying the disease [1].

Biotoxins can affect the nervous, the gastrointestinal and the respiratory systems, being fatal in some severe cases. They originate in various microalgae and dinoflagellates that produce these toxic elements in their metabolism. Mollusk bivalves are filter-feeder animals, and they can retain and accumulate those microalgae and the biotoxins, and as such, they can be a source of contamination for humans [30,31].

The accumulation of biotoxins in aquatic organisms depends on their feeding activity, on their metabolism, and on their elimination rate, modifying the transfer of toxins in the trophic chain [32].

Contaminant metals can cause several problems in humans, and the contamination of bivalve mollusks with different concentrations of heavy metals can be dangerous, namely lead, cadmium and mercury. They can accumulate high concentrations from water, and from the sediments and they have a wide bioaccumulation range of heavy metals depending on the species [33–35].

The aim of the present work was to study the sanitary condition in oyster production in four different populations of oysters. The objective was to describe the general health status, studying anatomo and histopathological lesions and parasites, in the Portuguese oyster (*Crassostrea angulata*) and Japanese oyster (*Crassostrea gigas*) from different production areas in Portugal, in order to evaluate their health condition.

#### **2. Materials and Methods**

The map (Figure 1) shows the four different areas where oysters were collected.

**Figure 1.** The map shows the areas were oysters were collected in mainland Portugal: Aveiro lagoon (1), Sado estuary (2), Mira estuary (3) and Alvor lagoon (4).

Oysters (9–11 cm, aged 24 months old) were sampled from four different sites of the Portuguese coast. *Crassostrea gigas* is an exotic species that was introduced in Portugal, and so, its presence is restricted to some production areas (1 and 4). In areas 2 and 4, *Crassostrea angulata* were collected from wild beds.

Portuguese oysters from Sado estuary (*n* = 30) and Mira estuary (*n* = 30) and Japanese oyster from Alvor lagoon (*n* = 30) and Aveiro lagoon (*n* = 30) were collected in May 2019. In May, in the four sites, oysters have already spawned and are more susceptible to environment stressors and increasing chances of histopathological lesions.

Individuals were randomized collected in production areas present in four mesotidal estuarine areas indicated in the map (Figure 1). The areas are located on the west Portuguese coast (1, 2 and 3) and in the south Portuguese coast (4). All the four production areas are under a strong oceanic influence because of the proximity to the coastal water and they have an influence of fresh water. For production, oysters are placed inside plastic bags that are put on tables with a distance of 0.50 m from the bottom in areas 1 and 4. In Sado and Mira estuaries, oysters were collected in intertidal mudflats where oyster beds are located. In Aveiro lagoon there is a lagoon with several channels, stretching for 45 Km. Sado stuary includes a section of river, marshlands and channels and the dynamic tide extends for 65 Km. Mira estuary is a narrow estuary of the "Ria" pattern that extends for 40 km. Alvor lagoon has a water body and a mesotidal shallow lagoon that extends for 15 Km.

Oyster were collected during low tide at a distance from the mouth of 1 Km (1), 17 Km (2), 15 Km (3) and 1 Km (4). Temperature and salinity of the water were, respectively: (1) 18.6 and 31.5; (2) 19.1 and 33.1; (3) 19.7 and 33.6; (4) 20.1 and 31.9.

After being collected, oysters were immediately transported on ice to the laboratory. To survey the presence of lesions, parasites and diseases anatomic and histopathological examinations were used as main diagnostic methods. Oysters were opened with a knife and the samples, including all organs and tissues, were collected with a tweezer and a scalpel, Tissue samples were prepared for histopathology processing, following the protocol used in Pathology laboratory of IPMA. Samples were fixed in Davidson's fixative for 48 h, dehydrated and embedded in paraffin. Sections with 5 μm thick were stained with Hematoxylin-Eosin and mounted on a microscopic glass slide.

Histological preparations were carefully examined under light microscopy (Motic BA-410), looking for the presence of lesions and parasites in oysters.

#### **3. Results**

In most of the samples no lesions were observed in organs (Figure 2A–C) The macroscopic lesions (Figure 3A–C), microscopic lesions (Figure 4A–F,I) and parasites (Figure 4G,H) were recorded.

**Figure 2.** Histological sections stained with hematoxylin and eosin, where different tissues and organs are shown. (**A**) Connective tissue normal is present (Mag = 40×; bar = 200 μm) (*C. angulata* tissue). (**B**) The normal gill with filaments where a ciliary system is observed (*C. gigas* tissue) (Mag = 200×; bar = 20 μm). (**C**) The normal digestive gland presents a regular tubule (t) thickness, characterized by their dense areas and star shape (*C. gigas* tissue). (Mag = 40×; bar = 200 μm).

**Figure 3.** Macroscopical lesions observed in oysters. (**A**) Gills lesions (arrow) in *C. angulata* (bar = 3 cm). (**B**) Shell disease (arrow) in *C. angulata* (bar = 3 cm). (**C**) Mud blisters (arrows) in *C. gigas* (bar = 3 cm).

**Figure 4.** *Cont*.

**Figure 4.** Microscopical lesions observed in oysters (tissue sections are stained with hematoxylin and eosin). (**A**,**D**) An inflammatory response is observed with the presence of hemocyte in the connective tissue (arrows) (*C. gigas* tissue). (Mag = 40×; bar = 200 μm). (**B**) Necrosis of gill tissue (arrow) (*C. angulata* tissue) (Mag = 200×; bar = 20 μm). (**C**) Metaplasia of digestive gland (arrows) (*C. angulata* tissue). (Mag = 200×; bar = 20 μm). (**E**) Necrosis of gill tissue (arrows) (*C. angulata* tissue). (Mag = 100×; bar = 100 μm). (**F**) Metaplasia of digestive gland (arrows) (*C. angulata* tissue). (Mag = 100×; bar = 100 μm). (**G**) *Trichodina* sp. (arrows) in gill tissue (*C. angulata* tissue). (Mag = 100×; bar = 100 μm). (**H**) *Mytilicola* sp. (arrow) in intestine (*C. gigas* tissue). (Mag = 40×; bar = 200 μm). (**I**) Loss of epithelium of the digestive gland (arrows) (*C. gigas* tissue) (Mag = 200×; bar = 20 μm).

In samples of *C. gigas* from Aveiro lagoon, it was observed only 3% of oysters with edema, 3% with shell disease and 10% with *M. ostrea*.

Finally, in samples from Aveiro lagoon, a prevalence of 13% *Trichodina* sp. infections, 13% *Ancistrocoma* sp. infections and 10% *Mytilicola* sp. infections. Concerning tissue lesions and morphological changes, 26% of wild oysters showed necrosis, 45% showed hemocytosis, indicating an inflammatory process, mainly in connective tissue. It was also observed a prevalence of 3% of ceroidosis in the connective tissue, edema in 15 % and 18% showed metaplasia in the digestive gland. Regarding farmed oysters, the following lesions and prevalence were found: necrosis (6%) in different tissues, hemocytosis (27%), ceroidosis (5%), edema (18%) and metaplasia (5%).

Samples of wild *C. angulata* from Sado estuary show the following results (Tables 1 and 2): 20% presented mud blisters, mainly caused by *Polydora* sp.; 33% with gills lesions; 10% had shell disease associated with the presence of the fungus *Ostracoblabe implexa*; and 17% showed *Myicola ostrea* copepods.

**Table 1.** Prevalence of macroscopic lesions and parasites in different populations.



**Table 2.** Prevalence of microscopic lesions in different populations.

Portuguese oysters sampled in Sado estuary, it was found a prevalence of 30% infections with *Trichodina* sp., characterized by a shape similar to a disc, a circlet of eosinophilic denticles, a ciliary fringe and a horseshoe shaped nucleous, in gills and mantle epithelium. A prevalence of 20% infections with *Ancistrocoma* sp., characterized by a Spindle-shaped ciliates with large, granular and polymorphic nuclei, in the digestive gland tubules and connective tissue. Copepods (*Mytilicola* sp.), characterized by an elongated cylindrical shape and a redish colour were observed within the intestine of oysters with the prevalence of 17% [36].

Samples of wild *C. angulata* Mira estuary showed mud blisters (96%), 50% had gills lesions, 10% presented edema and 10% shell disease.

In samples from Mira estuary, it was found a prevalence of 13% infections with *Trichodina* sp., 30% infections with *Ancistrocoma* sp. and 3% infections with *Mytilicola* sp.

Finally, in samples of *C. gigas* from Alvor lagoon 10% presented mud blisters.

In farmed oysters sampled from Alvor lagoon, it was observed a prevalence of 23% *Trichodina* sp. infections and a prevalence of 27% *Ancistrocoma* sp. Copepod (*Mytilicola* sp.) were observed within the intestine of oysters at a prevalence of 3%.

#### **4. Discussion**

Cultivation and harvesting of bivalve mollusks are two important activities, and it is fundamental to preserve the natural ecosystems that support those activities and lead to increasing natural production and improving its quality. Bivalve mollusks production depends on the external factors [37], including water parameters, such as temperature variations, food change, biotoxins and pathogens (including parasites, bacteria, viruses) or anthropogenic, such as chemical contaminants (heavy metals, pesticides), organic contaminants (fecal contamination) and overexploitation of natural banks [1,38]. It is wildly known that the marine environment provides stress to oysters during their life cycle, namely changes in environmental parameters, changes in the availability of food and the display of various toxic pollutants [39–41]. Oysters are very resistant to changes of temperature and salinity that are present in the ecosystems where they live, namely estuaries and lagoons. As other euryhaline organisms, oysters can live in water where large variations of salinity occur. For short periods of time salinity values could range between 2% and 38% [1,42]. Similarly, oysters are resistant to temperature changes between 8 ◦C and 30 ◦C [1]. There are several factors that influence the growth of mollusk bivalves such as temperature, oxygen and the presence of different species of phytoplankton. Once they are filter-feeders, the quality and the quantity of filtered food define the outcome of their growth [43].

The regularly consumption of bivalve mollusks by different populations and the growth of shellfish aquaculture production show the importance of these animals and their monitoring must be undertaken to avoid diseases and mortality in oyster populations. Hereby we are presenting the results of the sanitary monitoring of oyster production in four different production areas. In general, it was observed that low numbers of parasites were present in oysters being *Trichodina* sp. and *Ancistrocoma* sp. infections the most common ones present in the areas that were studied. The presence of *Trichodina* sp., *Ancistrocoma* sp., *Mytilicola* sp. and *Myicola ostrea* are commonly observed in oyster populations and in massive infestations could lead to host weakness [36,44–46]. Lesions of the internal organs, such as metaplasia, may reflect physiological stress, contaminants or the presence of large parasitic loads [1,44]. In our study, hemocytosis, cerodosis and edema were mainly present

in the interstitial connective tissue. These lesions and those observed in the epithelium of the diverticula of the digestive gland, including the hemocytic infiltration and necrosis were usually associated and could be related with the presence of the parasites. In wild populations from Sado and Mira estuaries the prevalence of mud blisters and gill lesions were higher than from populations produced on tables 0.50 m from mudflats.

The identification, characterization and registration of pathologic processes in oysters constitute a set of important measures for sanitary control. Based on the results of the sanitary control appropriate biosecurity tools could be developed and implemented leading to the control of diseases spread and transmission. Finally, knowledge and ability to manage risk of different pathogen will contribute to the sustainability of oyster production and protection of animal and human health.

Aquatic ecosystems can be modified due to human activities such as overharvesting and destruction of substrates [47]. In addition, transfers of oysters, between countries and regions can spread diseases and invasive species [47–49]. Concurrently with increase of the socio-economic importance of mollusk farming in Portugal all associated risk factors are affecting this production. It is essential to have stricter control at all stages of production of the different bivalve mollusk species. This is the only way to improve economic performance without jeopardizing animal welfare and at the same time to prevent diseases in oyster populations and their dissemination to different areas. In Portugal, IPMA (https://www.ipma.pt/en/) (accessed on 14 September 2021) undertakes regular analysis to the water, including in the areas where animal species that are produced to be consumed by populations. In the areas where animals were collected no contaminants were detected. Biotoxins are not harmful to bivalve mollusks but, when they are consumed by humans, they can cause several serious problems. As such, the control and monitoring of levels of contaminants are fundamental to prevent any risk to public health.

Diseases are important risk factors, specific or not, of different species of bivalve mollusks, which are caused, in many cases, by non-compliance with basic management rules, namely the level of animal load in production areas, the length of time the bivalve mollusks remain in these areas, and the introduction of seeds of unknown origin. Effective biosecurity measures and correct diagnostic techniques are essential to control pathogenic agents. The identification of pathogens should include those at an early stage in their life cycles, to control and to prevent their multiplication and proliferation. In the production areas from Aveiro and Alvor lagoons these measures are implemented by producers and by the authorities. Producers understand that it is essential to prevent mortalities in bivalve mollusk populations, namely avoiding overcrowding and preventing diseases. In Sado and Mira estuaries, wild populations show that oysters are more susceptible to lesions than oysters produced on tables under the supervision of producers.

Research on genetic resistant bivalve populations to different pathogenic agents and the use of probiotic bacteria are important strategies to prevent diseases caused, namely by different *Vibrio* species [50,51] that can cause mortality outbreaks, that did not occur in the areas that were studied. Furthermore, restoration programs can be implemented in areas where ecosystems were negatively affected [49]. The success of bivalve mollusk production can only be achieved, if appropriate measures are applied, namely biosecurity, correct and efficient diagnostic methods, effective compliance with legislation and the dissemination of knowledge among populations.

**Author Contributions:** Methodology (F.A.; F.R.); Investigation (A.G.; D.P.); Writing—original draft preparation (D.P.; F.A.); Writing—review and editing (A.G.; F.R.). All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by MAR2020: MAR-02.05.01-FEAMP-0010.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

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