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In the last decades, freshwater mussels belonging to the superfamily Unionoidea have increasingly been endangered, yet the concern regarding their conservation is equally growing (Bogan 1993, Strayer 2000, Strayer & Malcom 2007, Geist 2015, Lopes-Lima et al. 2016). Their particular life cycle, in which before being autonomous, the larvae (glochidia) needs to fix to a specific fish host to develop to the next stage, limits greatly their dispersion and propagation. There are many reasons for their population decline: (1) pollution, which is one of the major threats, (2) the introduction of new species that can act as competitors (mussels and fish), (3) a reduction in their hosts' population, and (4) multiple other factors of anthropogenic origin (Bogan 1993, Strayer 2000).

Freshwater mussels are key elements in freshwater systems because their suspension-feeding behavior is fundamental for cleaning the water and helping the recirculation of particles in rivers, lakes, and streams. In addition, their shells can be used as support structures for other organisms' proliferation (animals, plants, and algae) (Bogan 1993, Strayer et al. 2004). Aggressions affecting the environment are potentiated in these animals because of bioaccumulation processes and to their sessile lifestyle, making freshwater mussels ideal monitoring species,

very sensitive to environmental perturbations (Farris & Van Hassel 2006, Hartmann et al. 2015).

In a changing world where adaptation is a recurrent phenomenon, the increase of stressors in natural habitats raises the question whether environmental alterations may or may not have implications in the immune system of organisms. There have been reports suggesting that an immune modulation occurs (Mydlarz et al. 2006); however, in the long term, it is likely that the effects may cause deeper transformations. Freshwater mussels are not only being loaded with toxins because of the water pollution but are also being subjected to a microbial pool that is very labile.

Unionids, being suspension feeders, end up using large amounts of bacteria as a food source. Nevertheless, sometimes they are also able to establish symbiotic relationships either mutualistic or antagonistic with bacteria (Grizzle & Brunner 2009, Antunes et al. 2010). In the early stages of development (glochidia and juveniles), unionid mussels are prone to bacterial infectious diseases (Grizzle & Brunner 2009). Bacterial diseases such as vibriosis are much more studied in marine bivalves than in freshwater mussels because the economic implications are greater in the former case (Pruzzo et al. 2005, Mateo et al. 2009). When mussels reach the adult stage, they become much more resistant toward bacterial diseases, especially because they already have a mature innate immune system.

The immune system of unionid mussels, based on innate immunity (Danilova 2006, Mydlarz et al. 2006), comprises exterior barriers such as the shell and mucus on the surface of the tissues, and the internal cellular component and respective circulatory fluids. Particularly, blood cells (hemocytes) play a fundamental role in defense, especially through mechanisms associated with phagocytosis (Blaise et al. 2002, Canesi et al. 2002, Hong et al. 2006) and detoxification processes (Soares-daSilva et al. 2002), as well as the plasma, which is composed of humoral components.

The discovery of antimicrobial peptides (AMPs) and other molecules with antimicrobial properties from many organisms has been increasing (Cheng-Hua et al. 2009, Smith et al. 2010). Previous studies have already identified these peptides in invertebrates, such as crayfish (Pacifastacus leniusculus) (Jiravanichpaisal et al. 2007), but mostly in marine bivalves such as oysters (Crassostrea gigas) (Bachere et al. 2015) and mussels (Mytilus galloprovinciallis) (Mitta et al. 1999, 2000). It is very likely that freshwater mussels may also produce natural products possessing antibacterial activities.

Therefore, the aim of the present study was to evaluate the potentiality of several species of Portuguese freshwater mussels to produce antibacterial compounds, by testing their fluids, (plasma and extrapallial fluid) cells, and mucus against reference and multidrug-resistant bacterial strains of well-known human pathogens. Four species within the family Unionidae were selected from two freshwater systems: Anodonta cygnea from a shallow lagoon (Barrinha - Mira), and Anodonta anatina, Potomida littoralis, and Unio delphinus from a river (Tamega river).


Location and Collection of Freshwater Mussels

The species of freshwater bivalves used in this study were collected in northern Portugal in the Spring of 2014: Anodonta cygnea was collected from the Mira lagoon (40[degrees]27'22" N, 8[degrees]48'7" W) and Anodonta anatina, Unio delpinhus, and Potomida littoralis from the Tamega river (41[degrees]24'52" N, 7[degrees]57'51" W). They were kept in aerated tanks with dechlorinated water and acclimatized in these conditions for 2 wk before performing the assays, so the starting point of all tested species was the same, independent of their origin. These animals were fed daily with a microalgae diet (a mix of chlorophyte species). The organisms were considered healthy when the surface of the shell was smooth and shiny and when they were able to close the valves on disturbance.

Cells and Fluids Extraction

Hemolymph from three to six organisms of each species (Anodonta cygnea, Anodonta anatina, Potomida littoralis, and Unio delphinus) was carefully extracted using a 21G needle (Braun) attached to a 2-mL sterile syringe (Braun), by inserting it between the valves across the inner layer of the mantle, in the so-called intraepithelial space, avoiding contact with other surfaces. Each hemolymph sample was maintained on ice immediately after collection; no anti-aggregation solution was added to avoid introducing bias. The whole procedure was conducted using sterile material and in an aseptic environment.

The cells were isolated from the plasma by centrifugation at 200x g, 4[degrees]C for 10 min (Antunes et al. 2014), and resuspended in 50 mM phosphate buffer saline (PBS) for the susceptibility disc diffusion assay or in tryptic soy broth (TSB--Biokar Diagnostics, Allonne, France) for further use in the biofilm assays. The final concentration of cells was around [10.sup.6] cells/mL. The plasma was filtered through a 0.22-(im membrane filter to eliminated eventual contaminants.

The extrapallial fluid could only be collected from the Anodonta species because this species is big enough to ensure a safe extraction by inserting the needle between the shell and the mantle; the fluid was filtered and kept on ice until use.

Mucus Collection

Mucus was collected from the foot surface using a sterile blade and further suspended in 50 mM PBS, vortexed, and kept on ice (it was not filtered because it was too viscous).

Bacterial Water Quality Analysis at the Collection Points

Water (n = 19) and biofilm (n = 19) samples were previously collected from 2009 to 2011, in a total of 10 surveys from Mira lagoon and nine surveys from Tamega river to evaluate the natural habitat of the tested freshwater bivalves in terms of total mesophilic aerobic bacteria, Escherichia coli and Enterococcus spp., from the natural habitat. Biofilm was collected from the surface of rocks and macrophytes immersed in the water, by scraping it with a brush into a 50-mL falcon tube.

Water samples of 100 mL were filtered through 0.45-[mu]mpore-size membrane filters (Millipore Corporation, USA), which were then placed on tryptone bile X-glucuronide agar (TBX) (BioKar Diagnostics, Beauvais, France) and on Slanetz and Bartley agar (SB) (Oxoid, Basingstoke, UK) for Escherichia coli and Enterococcus spp. enumeration, respectively. The TBX plates were incubated at 37[degrees]C for 24 h and the SB plates at 37[degrees]C for 48 h. Enterococcus spp. colonies were confirmed in kanamycin aesculin azide agar (Liofilchem, Roseto degli Abruzzi, Italy), incubated at 44[degrees]C for 4 h.

For the determination of total aerobic mesophilic bacteria, dilutions of the water samples were made in tryptone salt and then 1 mL of each dilution was incorporated in plate count agar medium (Biokar Diagnostics) and incubated at 30[degrees]C for 72 h. The number of colony-forming units (CFUs) were counted and recorded.

The biofilm was diluted in tryptone salt and analyzed by incorporation in the abovementioned media (TBX, SB, and plate count agar) using the respective conditions of incubation.

Biochemical Quantifications on the Plasma from the Unionidae Species

Subsamples of the plasma from Anodonta cygnea, Unio delphinus, and Potomida littoralis were used for the quantification of total proteins (Bradford method), calcium (Quanti-Chrom Calcium Assay kit; Bioassays Systems), phosphates (PiBlue Phosphate assay kit; Bioassays Systems), and total lipids (Total lips Liquid kit; FAR Diagnostics, after extraction using methanol and chloroform).

Antibacterial Susceptibility Testing

Agar Disc Diffusion Method

For screening the potential antibacterial activity of cells, fluids, and mucus obtained from the four bivalve species, four reference strains of bacteria (Staphylococcus aureus ATCC 25923, Bacillus subtilis ATCC 6683, Pseudomonas aeruginosa ATCC 27853, and Escherichia coli ATCC 25922) were initially used. Fresh bacterial cultures were used to prepare an inoculum equivalent to 0.5 McFarland.

Mueller-Hinton agar (BioKar Diagnostics) plates were inoculated with the bacterial inocula. Then, small sterile paper discs (6 mm in diameter) were equidistantly attached to the agar (five per plate) and loaded with 15 [micro]L of the sample (cells, fluids, or mucus). The plates were kept for 30 min at room temperature before incubation at 37[degrees]C for 18-24 h (Pereira et al. 2015). A negative control was made using PBS.

The plates were observed under a stereoscope (SZ61 stereoscope; Olympus America Inc., Center Valley, PA) and if zones of inhibition occurred around the discs, the diameter of such zones was measured. Photographs of zones of inhibition were also taken (PD70; Olympus America Inc.). The samples that have caused any bacterial inhibition around the discs were further studied against other potential pathogenic bacterial strains: Salmonella enterica Typhimurium CECT 443, Klebsiella pneumoniae ATCC 13883, Acinetobacter baumannii ATCC 19606, Listeria monocytogenes ATCC 19111, Enterococcus faecalis ATCC 29212, and multidrug-resistant clinical strains of Pseudomonas putida, and Klebsiella spp., following the same procedure.

Antibiofilm Activity Assay

Biofilm Biomass Quantification

The fractions that had given positive results in the previous assay, which were derived from the hemolymph, the cellular fraction, and the plasma fraction (pure or diluted in the ratio of 1:2 and 1:4 in TSB), were also tested for antibiofilm activity. Briefly, biofilms of the following strains were formed in the wells of 96-well plates: Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, and Enterococcus faecalis ATCC 29212. The initial inocula were at a concentration of 1 X [10.sup.6] CFU/mL in TSB, to which the respective samples to be tested had been added. Six to eight replicate wells were used for each condition tested. The positive control consisted only the bacterial inoculum. The microplates were incubated at 37[degrees]C for 24 h. The biofilm biomass was quantified through the crystal violet staining, as described by Gomes et al. (2014).

Microscopic and Viability Analyses of Biofilms

The conditions that have hampered or decreased the biofilm formation were then also replicated for a qualitative assessment by microscopic visualization, after staining with the Live/Dead BacLight viability kit (Molecular Probes; Life Technologies, Carlsbad, CA). For this qualitative assay, three Anodonta cygnea were used.

Biofilms of Staphylococcus aureus ATCC 25923, Pseudomonas aeruginosa ATCC 27853, and Escherichia coli ATCC 25922 were formed in 35-mm-diameter polystyrene plates using TSB (control) and TSB supplemented with cellular fractions and plasma (pure or diluted in the ratio of 1:2 and 1:4 in TSB). The plates were incubated at 37[degrees]C for 24 h. After that, the supernatant phase was removed from each plate, the biofilms were washed with PBS, stained with 500 ([micro]L of the mixture of SYTO 9, and propidium iodide and incubated for 20 min at room temperature in the dark; then, the biofilms were rinsed and examined under a fluorescence microscope (BX41 Microscope; Olympus America Inc.) (Gomes et al. 2014).


The statistical significance of differences between biofilms of controls and biofilms in the presence of hemolymph, the cellular fraction, or the plasma fraction were evaluated using Student's t-test. Probability levels <0.05 were considered statistically significant.


Microbial Quality of the Water at the Collection Points of the Bivalves

The annual variations of Escherichia coli, Enterococcus spp., and total mesophilic aerobic bacteria in the water and in the biofilm collected from Tamega river and Mira lagoon are shown in Figure 1. It is possible to observe that counts in water and biofilm do not present the same profile of progression. Tamega river, in comparison with Mira lagoon, presented higher fluctuations in all counts, as well as the highest values of E. coli and Enterococcus spp. in the water and biofilm samples and of total aerobic mesophilic bacteria in the biofilm. Enterococcus spp., E. coli, and total aerobic mesophilic bacteria maximums occurred in the biofilm fraction.

Plasma Composition

Regarding the quantification of the components of the plasma (Fig. 2), it was possible to observe that the plasma of the three species tested presented similar values in terms of calcium and total lipids. The species Unio delphinus presented higher differences in the quantity of the components tested, compared with the other species and the highest values concerning phosphates, whereas Potomida littorallis presented a lower content in terms of total proteins.

Antibacterial Activity

Regarding the results from the agar disc diffusion assay (Fig. 3), it was observed that the cellular fraction of some bivalve species could cause bacterial inhibition, but not against all bacteria tested. The zones of inhibition were generally small. The mucus did not cause any bacterial inhibition; in fact, greater bacterial proliferation around the disc was recorded, which probably means that the mucus harbors the bacteria itself. A similar result was observed with the extrapallial fluid samples.

The most promissory species was Anodonta cygnea from Mira lagoon because its cellular fraction caused greater zones of inhibition in Bacillus subtilis ATCC 6683 and Pseudomonas aeruginosa ATCC 27853, as well as multidrug-resistant isolates, such as Pseudomonas putida and Acinetobacter baumannii (Fig. 3A).

Potomida littoralis cellular fraction also inhibited Bacillus subtilis ATCC 66683, Pseudomonas aeruginosa ATCC 27853, Listeria monocytogenes ATCC 19111, Acinetobacter baumannii ATCC 19606, and Salmonella enterica Typhimurium CECT 443 (Fig. 3B). The cellular fraction of Anodonta anatina showed similar results as the previous bivalve species, inhibiting the same strains; however, this inhibition was greater in the case of 5. enterica Typhimurium CECT 443 and lesser in the case of A. baumannii ATCC 19606 (Fig. 3C). Unio delphinius, although being collected from the same place, at the Tamega river, did not cause any bacterial inhibitions in this assay (results not shown).

Antibiofilm Activity

Regarding the biofilm formation assay, the results were more evident, with almost all tested conditions being significantly different from the control (P < 0.05, Fig. 4). In the presence of the plasma of the bivalves, even when diluted, less biofilm was formed. Interestingly, the cellular fraction hampered the biofilm formation solely by Escherichia coli ATCC 25922, which was the bacterial strain tested that apparently had more difficulty to form a biofilm both in the presence of the cellular fraction and higher dilution of plasma of Anodonta cygnea. Moreover, A. cygnea showed great ability to also inhibit the biofilm formation of Staphylococcus aureus ATCC 25923, with all its components tested able to cause a lower biofilm production than the control. The biofilm of Pseudomonas aeruginosa ATCC 27853 was mostly inhibited when in contact with plasma; the diluted plasma caused the highest inhibition. Regarding Enterococcus feacalis ATCC 29212, A. cygnea caused inhibition only through pure plasma and plasma diluted in the ratio of 1:2; the higher dilution of the plasma and the cellular fraction caused higher biofilm production than the control (Fig. 4A). Although belonging to the same genus, Anodonta anatina showed slightly different results, presenting a significant capacity to inhibit the biofilm formation by S. aureus ATCC 25923 and E. coli ATCC 25922, especially in the presence of undiluted and diluted 1:2 plasma, however, causing little inhibition in the biofilm formed by P. aeruginosa ATCC 27853. On the contrary, the cellular fraction of A. anatina was even able to significantly increase the biofilm biomass in respect to the control. The cellular fraction equally increased the biofilm biomass of S. aureus ATCC 25923 (Fig. 4B).

In terms of biofilm production, Unio delphinus and Potomida littoralis (Fig. 4C, D) caused a similar effect in the three different bacterial strains tested. Staphylococcus aureus ATCC 25923 suffered the highest inhibition in the biofilm formation (the highest value was recorded for the more diluted sample of plasma). Pseudomonas aeruginosa ATCC 27853 biofilm was inhibited in the presence of all fractions of these two species; nonetheless, the plasma diluted in the ratio of 1:2 caused the highest inhibition whereas the cellular fractions showed little effect on the biofilm formation. Regarding Escherichia coli ATCC 25922, the results were more inconsistent; although the diluted plasma produced a marked reduction on the biofilm formed, the undiluted plasma of U. delphinus led to a higher production of biofilm in comparison with the control. Hemocytes from P. littorallis and U. delphinus inhibited significantly the biofilm production by S. aureus ATCC 25923 (P < 0.05).

Viability Assay

The live/dead fluorescence assay (Fig. 5) allowed a qualitative analysis along with a viability assessment of the biofilms formed in the presence of different biological fractions of the freshwater bivalves. It was visible that no biofilm was formed in the presence of the plasma (nondiluted and at both dilutions) of Anodonta cygnea. The undiluted plasma and the plasma diluted in the ratio of 1:2 in TSB inhibited almost totally the growth of the three bacterial strains tested; even the more diluted fraction of the plasma could inhibit greatly the biofilm formation by Staphylococcus aureus ATCC 25923 and Escherichia coli ATCC 25922. Pseudomonas aeruginosa ATCC 27853, despite being able to form a biofilm in the presence of the plasma, the biomass produced was lower than the one formed in the control condition. The cellular fraction showed almost no inhibition, the big red cells (Fig. 5A5, B5, C5) correspond to dead hemocytes, which may have been used as a substrate, favoring the proliferation of the three bacterial strains tested.


Invertebrate animals have an innate immune system mainly composed by the circulating cells in their body fluids, such as hemocytes, which are responsible for mechanisms such as phagocytosis (Mitta et al. 1999, Antunes et al. 2010, 2014). In previous studies, it was demonstrated that Escherichia coli and Enterococcus faecalis were not tolerated by Anodonta spp., being eliminated by the bivalve's granulocytes, which proved the action of the cellular immunity (Antunes et al. 2010, 2014); however, it is quite evident that humoral substances must also be present in the plasma, allowing it to play a significant role in the immunity defense, as shown by the present study, which revealed an antibiofilm activity of the plasma of all unionid species tested.

Cellular fractions could cause bacterial inhibition, although only to a relatively small extent, in some cases. Nevertheless, there were cases in which the cellular fractions stimulated the bacterial growth, which was probably due to the fact that the conditions used did not allow maintaining the hemocytes viable long enough. There is a gap in the literature in terms of the optimal conditions to maintain these cells viable. In the article by Hinzmann et al. (2013), some procedures were recommended such as the use of appropriate anti-aggregation solutions and low temperature (keeping the cells on ice); still, the several attempts to put these cells into the culture failed (data not shown). Moreover, this work focused on the bacterial conditions, so nothing was added to avoid interference with the bacterial growth and the temperature selected was the optimal for the bacteria. Thus, the use of cellular fractions for assessing biological activities needs to be optimized. Nonetheless, the agar disc diffusion method showed that the cellular fraction from Anodonta spp. and Potomida littoralis inhibited Bacillus subtilis ATCC 6683 and Pseudomonas aeruginosa ATCC 27853, meaning that these fractions may comprise inhibitory components, which could have been originated from cells. Moreover, the agar disc diffusion assay showed that these species of freshwater mussels had also the potential to inhibit multidrugresistant strains, such as multidrug-resistant isolates of Pseudomonas putida and Acinetobacter baumannii. Obviously, the agar disc diffusion assay is a basic screening assay that has several limitations, e.g., only a small volume of sample can be tested, and some substances do not diffuse in the agar medium. Probably, this can be one of the reasons explaining why in some cases no inhibition was recorded using the plasma fraction in the disc whereas very high inhibition was recorded in the biofilm quantification assay.

Taking into account the results regarding the effect of bivalves' biological fractions toward the biofilm formation by bacteria, it is likely that these bivalves developed molecular mechanisms capable of defending them from bacteria that could develop a biofilm inside them, putting at stake their survival. As no antibacterial effect on the bacterial growth was observed, the antibiofilm activity of plasma components was probably due to their interference in the communication/quorum-sensing system of the bacterial cells or due to specific physicochemical characteristics of that body fluid which do not favor the adhesion of bacteria to a surface. Moreover, because the diluted plasma still had the same effects, it may indicate that the components interfering with the biofilm formation are able to do it even at a low concentration. The fact that the inhibition of biofilm formation by these plasma components was found in diverse bivalve species and toward bacterial strains that are typically not present in their living water (e.g., Staphylococcus aureus) is suggestive of an innate nonspecific mechanism; otherwise, the bivalve species inhabiting waters with higher microbial contamination would have expressed a more pronounced inhibitory capacity against bacteria. The cellular fraction obtained from Anodonta cygnea was able to inhibit the biofilm formation by Escherichia coli, but not by the other bacterial strains, which means that it may have particular compounds targeting specifically the biofilm formation by E. coli. This hypothesis need to be further explored. Little is known on the chemical identification of these substances as well as on their mechanisms of action. Several antimicrobial compounds, namely AMPs, have been identified in cultivated species, e.g., mytilins in mussels (Mytilus edulis and Mytilus galloprovicialis) (Charlet et al. 1996, Mitta et al. 2000) or defensins from oysters (Crassostrea virginica and Crassostrea gigas) (Anderson & Beaven 2001, Bachere et al. 2015) and big defensin from the scallop Argopecten irradians (Zhao et al. 2007). Thus, the studies on AMPs have been increasing because these peptides have the potential to replace actual antibiotics as new antimicrobial drugs with application on aquaculture (Cheng-Hua et al. 2009).

Similar studies on freshwater mussels were reported by Estari et al. (2011) regarding Lamellidens marginalis and by Santhiya and Sanjeevi (2014) using Parreysia corrugatea. Estari et al. (2011) tested fluids and tissue extracts from L. marginalis, which were diluted in solvents (water, chloroform, acetone, or methanol) and followed protein extraction protocols; among the different extracts tested, they obtained antimicrobial inhibition against Staphylococcus aureus, Streptococcus pyroenes, Morganella morganii, Bacillus subtilis, Escherichia coli, Proteus vulgaris, and Crassostrea albicans. The other study reported bacterial inhibition of tissue extracts through the diffusion agar method against pathogenic bacteria, such as S. aureus, B. subtilis, E. coli, Pseudomonas aeruginosa, and Klebsiella pneumonia (Santhiya & Sanjeevi 2014). Regarding freshwater bivalves, the molecules that may be involved in this antibacterial response are still unidentified; however, breakthroughs in this field may start to appear, because a big defensin gene was recently identified in the freshwater mussel Hyriopsis cumingii (Wang et al. 2014).

Herein, Anodonta cygnea gave more marked results in comparison with the other bivalve species, which, however, slightly followed the same pattern. The freshwater bivalve A. cygnea inhabits a more constant environment: the water from Mira lagoon, from where only A. cygnea was collected, presented lower fluctuations in the microbial counts in respect to the water samples analyzed from Tamega river. The lagoon provides the appropriate conditions for this species to persist, allowing it to be integrated in the "least concern" conservation category. The species Unio delphinus and Potomida littoralis came from a river with higher fluctuation in terms of bacteria load (Escherichia coli and Enterococcus spp.) and this may explain the less-pronounced antibacterial results and the more fragile conservation status of these species, "near threatened" and "endangered," respectively (IUCN red list). The species Anodonta anatina, equally with "least concern" conservation status, showed the lower antibacterial potential when compared with A. cygnea.

Although the plasma composition was not determined in the present study, it surely must be further explored to understand the immune system of this highly endangered group of species and to identify potential antimicrobial compounds. Because the bacterial burden is increasing in their natural habitat (Costa et al. 2013) and threatening even more their survival, it is expectable that freshwater bivalves may find strategies to overcome that hostile situation by producing antimicrobial substances; therefore, further studies in this field are fundamental to fully understand how humoral factors act in the innate immune response.


This work was supported by the Portuguese Foundation for Science and Technology (FCT) with a research grant to M. H. (SFRH/BD/76265/2011). This article was also supported by the project INNOVMAR-Innovation and Sustainability in the Management and Exploitation of Marine Resources (NORTE-01-0145-FEDER-000035, within Research Line NOVELMAR), by North Portugal Regional Operational Programme (NORTE 2020), under the PORTUGAL 2020 Partnership Agreement, and through the European Regional Development Fund (ERDF).


Anderson, R. S. & A. E. Beaven. 2001. Antibacterial activities of oyster (Crassostrea virginica) and mussel (Mytilus edulis and Geukensia demissa) plasma. Aquat. Living Resour. 14:343-349.

Antunes, F., M. Hinzmann, M. Lopes-Lima, J. Machado & P. Martins da Costa. 2010. Association between environmental microbiota and indigenous bacteria found in hemolymph, extrapallial fluid and mucus of Anodonta cygnea (Linnaeus, 1758). Microb. Ecol. 60:304-309.

Antunes, F., M. Hinzmann, M. Lopes-Lima, P. Vaz-Pires, S. Ferreira, B. Domingues & J. Machado. 2014. Antibacterial effects of Anodonta cygnea fluids on Escherichia coli and enterococci multi-drugresistant strains: environmental implications. Toxicol. Environ. Chem. 96:880-889.

Bachere, E., R. D. Rosa, P. Schmitt, A. C. Poirier, N. Merou, G. M. Charriere & D. Destoumieux-Garzon. 2015. The new insights into the oyster antimicrobial defense: cellular, molecular and genetic view. Fish Shellfish Immunol. 46:50-64.

Blaise, C, S. Trottier, F. Gagne, C. Lallement & P. D. Hansen. 2002. Immunocompetence of bivalve hemocytes as evaluated by a miniaturized phagocytosis assay. Environ. Toxicol. 17:160-169.

Bogan, A. E. 1993. Freshwater bivalve extinctions (Mollusca: Unionoida): a search for causes. Am. Zool. 33:599-609.

Canesi, L., G. Gallo, M. Gavioli & C. Pruzzo. 2002. Bacteria-hemocyte interactions and phagocytosis in marine bivalves. Microsc. Res. Tech. 57:469-476.

Charlet, M., S. Chernysh, H. Philippe, C. Hetru, J. A. Hoffmann & P. Bulet. 1996. Innate immunity. Isolation of several cysteine-rich antimicrobial peptides from the blood of a mollusc, Mytilus edulis. J. Biol. Chem. 271:21808-21813.

Cheng-Hua, L., Z. Jian-Min & S. Lin-Sheng. 2009. Review of advances in research on marine molluscan antimicrobial peptides and their potential application in aquaculture. Molluscan Res. 29:17-26.

Costa, P. M., L. Loureiro & A. J. Matos. 2013. Transfer of multidrugresistant bacteria between intermingled ecological niches: the interface between humans, animals and the environment. Int. J. Environ. Res. Public Health 10:278-294.

Danilova, N. 2006. The evolution of immune mechanisms. J. Exp. Zool. B Mol. Dev. Evol. 306:496-520.

Estari, M. S. J., B. S. Kumar, T. Bikshapathi, A. S. Reddy & L. Venkanna. 2011. In vitro study of antimicrobial activity in freshwater mussel (Lamellidens marginalis) extracts. Biol. Med. (Aligarh) 3:191-195.

Farris, J. L. & J. H. Van Hassel. 2006. Freshwater bivalve ecotoxicology. Taylor & Francis. 408 pp.

Geist, J. 2015. Seven steps towards improving freshwater conservation. Aquat. Conserv. Mar. Freshw. Ecosyst. 25:447-453.

Gomes, N., L. Bessa, S. Buttachon, P. Costa, J. Buaruang, T. Dethoup, A. Silva & A. Kijjoa. 2014. Antibacterial and antibiofilm activities of tryptoquivalines and meroditerpenes isolated from the marine-derived fungi Neosartorya paulistensis, N. laciniosa, N. tsunodae, and the soil fungi N.fischeri and N. siamensis. Mar. Drugs 12:822-839.

Grizzle, J. M. & C. J. Brunner. 2009. Infectious diseases of freshwater mussels and other freshwater bivalve mollusks. Rev. Fish. Sci. 17:425-467.

Hartmann, J. T., S. Beggel, K. Auerswald, B. C. Stoeckle & J. Geist. 2015. Establishing mussel behavior as a biomarker in ecotoxicology. Aquat. Toxicol. 170:279-288.

Hinzmann, M. F., M. Lopes-Lima, J. Goncalves & J. Machado. 2013. Antiaggregant and toxic properties of different solutions on hemocytes of three freshwater bivalves. Toxicol. Environ. Chem. 95:790-805.

Hong, X. T., L. X. Xiang & J. Z. Shao. 2006. The immunostimulating effect of bacterial genomic DNA on the innate immune responses of bivalve mussel, Hyriopsis cumingii Lea. Fish Shellfish Immunol. 21:357-364.

Jiravanichpaisal, P., S. Y. Lee, Y. A. Kim, T. Andren & I. Soderhall. 2007. Antibacterial peptides in hemocytes and hematopoietic tissue from freshwater crayfish Pacifastacus leniusculus: characterization and expression pattern. Dev. Comp. Immunol. 31:441-455.

Lopes-Lima, M., R. Sousa, J. Geist, D. C. Aldridge, R. Araujo, J. Bergengren, Y. Bespalaya, E. Bodis, L. Burlakova, D. Van Damme, K. Douda, E. Froufe, D. Georgiev, C. Gumpinger, A. Karatayev, U. Kebapci, I. Killeen, J. Lajtner, B. M. Larsen, R. Lauceri, A. Legakis, S. Lois, S. Lundberg, E. Moorkens, G. Motte, K. O. Nagel, P. Ondina, A. Outeiro, M. Paunovic, V. Prie, T. von Proschwitz, N. Riccardi, M. Rudzite, M. Rudzitis, C. Scheder, M. Seddon, H. serefiisan, V. Simic, S. Sokolova, K. Stoeckl, J. Taskinen, A. Teixeira, F. Thielen, T. Trichkova, S. Varandas, H. Vicentini, K. Zajac, T. Zajac & S. Zogaris. 2016. Conservation status of freshwater mussels in Europe: state of the art and future challenges. Biol. Rev. Camb. Philos. Soc. 92:572-607.

Mateo, D. R., A. Siah, M. T. Araya, F. C. Berthe, G. R. Johnson & S. J. Greenwood. 2009. Differential in vivo response of soft-shell clam hemocytes against two strains of Vibrio splendidus: changes in cell structure, numbers and adherence. J. Invertebr. Pathol. 102:50-56.

Mitta, G, F. Vandenbulcke, F. Hubert & P. Roch. 1999. Mussel defensins are synthesised and processed in granulocytes then released into the plasma after bacterial challenge. J. Cell Sci. 112:4233-4242.

Mitta, G, F. Vandenbulcke, F. Hubert, M. Salzet & P. Roch. 2000. Involvement of mytilins in mussel antimicrobial defense. J. Biol. Chem. 275:12954-12962.

Mydlarz, L. D., L. E. Jones & C. D. Harvell. 2006. Innate immunity, environmental drivers, and disease ecology of marine and freshwater invertebrates. Annu. Rev. Ecol. Evol. Syst. 37:251-288.

Pereira, A. L., L. J. Bessa, P. N. Leao, V. Vasconcelos & P. Martins da Costa. 2015. Bioactivity of Azolla aqueous and organic extracts against bacteria and fungi. Symbiosis 65:17-21.

Pruzzo, C, G. Gallo & L. Canesi. 2005. Persistence of vibrios in marine bivalves: the role of interactions with haemolymph components. Environ. Microbiol. 7:761-772.

Santhiya, N. & S. B. Sanjeevi. 2014. Antibacterial activity of freshwater mussel Parreysia corrugata (Muller 1774) from lower Anaicut Reservoir, India. Int. J. Pharm. Life Sci. 5:3899-3902.

Smith, V. J., A. P. Desbois & E. A. Dyrynda. 2010. Conventional and unconventional antimicrobials from fish, marine invertebrates and micro-algae. Mar. Drugs 8:1213-1262.

Soares-da-Silva, I. M., J. Ribeiro, C. Valongo, R. Pinto, M. Vilanova, R. Bleher & J. Machado. 2002. Cytometric, morphologic and enzymatic characterisation of haemocytes in Anodonta cygnea. Comp. Biochem. Physiol. A 132:541-553.

Strayer, D. L. 2000. The ecology of freshwater molluscs. Nature 406:126.

Strayer, D. L., K. A. Hattala & A. W. Kahnle. 2004. Effects of an invasive bivalve (Dreissena polymorpha) on fish in the Hudson River estuary. Can. J. Fish. Aquat. Sci. 61:924-941.

Strayer, D. L. & H. M. Malcom. 2007. Shell decay rates of native and alien freshwater bivalves and implications for habitat engineering. Freshw. Biol. 52:1611-1617.

Wang, G. L., X. L. Xia, X. L. Li, S. J. Dong & J. L. Li. 2014. Molecular characterization and expression patterns of the big defensin gene in freshwater mussel (Hyriopsis cumingii). Genet. Mol. Res. 13:704715.

Zhao, J., L. Song, C. Li, D. Ni, L. Wu, L. Zhu, H. Wang & W. Xu. 2007. Molecular cloning, expression of a big defensin gene from bay scallop Argopecten irradians and the antimicrobial activity of its recombinant protein. Mol. Immunol. 44:360-368.


(1) Department of Aquatic Production, Abel Salazar Institute for the Biomedical Sciences, University of Porto, Rua de Jorge Viterbo Ferreira No. 228, Porto 4050-313, Portugal; (2) Interdisciplinary Centre of Marine and Environmental Research, University of Porto, Terminal de Cruzeiros do Porto de Leixoes, Av. General Norton de Matos s/n, Matosinhos 4450-208, Portugal; (3) Department of Chemistry and Biochemestry, Faculty of Sciences, University of Porto, Rua do Campo Alegre, Porto 4169-007, Portugal; (4) Mountain Research Centre, School of Agriculture, Polytechnic Institute of Braganca, Campus de Santa Apolonia, Braganqa 5300-253, Portugal

(*) Corresponding author. E-mail:

DOI: 10.2983/035.037.0110
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Article Details
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Author:Hinzmann, Mariana; Bessa, Lucinda J.; Teixeira, Amilcar; Costa, Paulo Martins Da; Machado, Jorge
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:4EUPR
Date:Apr 1, 2018

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