Marine surface microlayer as a source of enteric viruses.

MARINE SURFACE MICROLAYER AS A SOURCE OF ENTERIC VIRUSES Ana Catarina Bispo Prata Tese de Doutoramento em Ciências do Mar e do Ambiente 2014 Ana Catarina Bispo Prata MARINE SURFACE MICROLAYER AS A SOURCE OF ENTERIC VIRUSES Tese de Candidatura ao grau de Doutor em Ciências do Mar e do Ambiente Especialidade em Oceanografia e Ecossistemas Marinhos submetida ao Instituto de Ciências Biomédicas de Abel Salazar da Universidade do Porto. Programa Doutoral da Universidade do Porto (Instituto de Ciências Biomédicas de Abel Salazar e Faculdade de Ciências) e da Universidade de Aveiro. Orientador – Doutor Adelaide Almeida Categoria – Professora Auxiliar Afiliação – Departamento de Biologia da Universidade de Aveiro Co-orientador – Doutor Newton Carlos Marcial Gomes Categoria – Investigador Principal do CESAM Afiliação – Departamento de Biologia da Universidade de Aveiro i LEGAL DETAILS In compliance with what is stated in the legislation in vigor, it is hereby declared that the author of this thesis participated in the creation and execution of the experimental work leading to the results here stated, as well as in their interpretation and writing of the respective manuscripts. This thesis includes one scientific paper published in an international journal and three articles in preparation originated from part of the results obtained in the experimental work referenced as: • Prata C, Ribeiro A, Cunha A , Gomes NCM, Almeida A, 2012, Ultracentrifugation as a direct method to concentrate viruses in environmental waters: virus-like particles enumeration as a new approach to determine the efficiency of recovery, Journal of Environmental Monitoring, 14 (1), 64-70. • Prata C, Cunha A, Gomes N, Almeida A, Surface Microlayer as a source of health relevant enteric viruses in Ria de Aveiro. • Prata C, Cunha A, Gomes N,Nunes, T, Almeida A, Marine aerosols as a source of health relevant virus. • Prata C, Coelho F, Ladeiro B, Cunha A, Gomes N, Almeida A, Influence of UV light exposure on viral abundance. iii Table of Contents A gradecimentos v Figures Index vii Tables Index ix Abreviations list xi Resumo xiii Abstract xv T h esis outline xvii Chapter 1 – Introduction 1 1.1. Enteric virus in the aquatic system 4 1.2. Factors affecting viral survival in the marine environment 9 1.3. Enteric virus transmission from the marine environment 11 1.3.1. Role of Surface Microlayer in viral transmission 12 1.3.2. Transmition of human virus by aerosols 14 1.4. Detection of enteric virus in the aquatic system 20 1.4.1. Viral concentration methods 20 1.4.2. Identification and quantification methods 24 Chapter 2 - Ultracentrifugation as a direct method to concentrate virus in environmental waters: virus-like particle enumeration as a new approach to determine the efficiency of recovery 51 Chapter 3 - Surface microlayer as a source of health relevant enteric viruses in Ria de Aveiro 71 Chapter 4 - Marine aerosols as a source of health relevant virus 91 Chapter 5 - Influence of UltraViolet light exposure on viral abundance 109 Chapt er 6 – Final Considerations 125 v Agradecimentos Esta viagem termina agora e surge o momento de agradecer aos que passaram pelo meu caminho, durante estes cinco anos, contribuindo de algum modo para a realização deste trabalho. A todos o meu mais sincero obrigada. Começo por agradecer à minha orientadora, Dra Adelaide Almeida e ao co-orientador, Newton Gomes, pela disponibilidade mostrada ao longo de todo o trabalho, pela ajuda na resolução de problemas e pela opinião crítica relativamente a todas as questões de trabalho. Aos colegas de laboratório, pela ajuda dada em campo, no laboratório, ou simplesmente por descontraírem o ambiente quando era necessário. Aos amigos promarianos que desde o primeiro dia se mostraram disponíveis para ajudar em tudo o que fosse necessário. Juntos inicámos esta viagem e juntos estamos a terminá-la (uns mais cedo do que outros). Muito obrigada por estarem sempre presentes: Allan Souza, Fátima Pinto, Helena Oliveira, Inês Rosa, Joana Rocha, Martina Ilarri, Rita Polónia, Rosa Reboreda, Rute Pinto e Sérgio Prats. Agradeço aos melhores pais do mundo, os meus, por acreditarem em mim, por me terem apoiado em mais esta jornada da minha vida e por estarem lá, sempre que preciso. A todos os meus amigos, mas em especial à Sónia, ao Francisco, ao João e à Inês, por me terem apoiado, ouvido os desabafos e dado força nos momentos difíceis, para seguir em frente. À Mariana, ao André e à família Antunes, por terem entrado na minha vida e marcado a diferença. Obrigada do fundo do coração. Ao Henrique, por teres reaparecido na minha vida, por me fazeres acreditar que há algo melhor. Obrigada por seres assim, perfeito para mim. xiii Resumo Os vírus entéricos são um importante grupo de agentes patogéncicos, responsáveis por um grande número de doenças no Homem. Podem existir naturalmente no ambiente aquático ou ser transportados até lá. Alguns destes vírus são considerados emergentes no ambiente aquático, representando um elevado risco para a saúde pública, economia e ecologia. Assim, é de extrema importância, conhecer os seus reservatórios naturais, bem como o modo como podem ser transferidos a partir do ambiente aquático para a atmosfera, a fim de minimizar os riscos por eles provocados. Apesar do elevado número de vírus presente no ambiente aquático, apenas uma pequena parte causa infeções no Homem, mesmo quando presentes em baixa concentração, uma vez que apresentam uma baixa dose infeciosa. A concentração das amostras de água representa um importante passo aquando da deteção dos vírus entéricos. No entanto, não existe um método ideal para concentrar vírus em amostras de água, dependendo sempre do objetivo do trabalho em questão. Com este trabalho, verificou-se que o método da ultracentrifugação é um bom método para a concentração de águas ambientais uma vez que foram obtidas taxas de recuperação entre 66 e 76%. Por outro lado, este método não adiciona compostos químicos, o que permite usá-lo para detetar posteriormente vírus usando métodos moleculares. A microcamada superficial (SML) constitui a interface entre a água e o ar, acumulando microrganismos e partículas. Sabe-se que constitui um reservatório natural para vários microrganismos, a partir do qual são libertados aerossóis para a atmosfera, levando assim os materiais acumulados na SML, incluindo vírus. Foram obtidos neste trabalho valores de enriquecimento de vírus entéricos na SML, relativamente à água subjacente (UW), superiores a 2, tanto na zona marinha como na zona salobra. Comparando esses valores para a SML, relativamente aos aerossóis, o enriquecimento é de cerca de 26 na zona marinha e cerca de 30 na zona salobra, mostrando assim que a SML funciona como reservatório para os vírus entéricos e que estes são transmitidos, do ambiente aquático para a atmosfera, através dos aerossóis. Relativamente à abundância dos vírus entéricos estudados, observou-se que rotavirus e enterovirus são mais abundante nos meses mais frios e que HAV apresenta abundância semelhante nos meses quentes e frios. As variações na abundância dos vírus entéricos foram explicadas pelas variações de temperatura da água e pela concentração de nitratos e de nitritos, o que sugere que estes vírus têm origem xiv terrestre, sendo transportados para o ambiente marinho através das águas de escorrência provocadas pelas chuvas, durante os meses mais frios. Atualmente as alterações climáticas e as suas consequências nos ecossistemas são consideradas um problema grave, pelo que foi testada neste trabalho a influência da exposição à luz UV, na abundância de vírus entéricos de RNA. Observámos que os rotavírus foram mais resistentes à UVR do que os enterovirus (p = 0.03 < 0.05 and p = 0.41 < 0.05, respectivamente), podendo as diferenças observadas entre os dois grupos estar relacionada não só com o tipo de RNA, dsRNA nos rotavirus e ssRNA nos enterovirus, mas também com a estrutura da cápside, constituída por três camadas nos rotavírus e apenas uma nos enterovírus. xv Abstract Enteric viruses are an important group of pathogenic agents responsible for a large number of diseases in humans. They can exist naturally in the aquatic environment or be transported from terrestrial environment to the water. Viruses present in this group are currently considered emerging in the aquatic environment, representing a hazard to public health, economy and ecology. It is therefore of outmost importance to know their natural reservoirs, as well as how they can be transferred from the aquatic environment to the atmosphere in order to minimize the risks posed by them. Despite the high number of viruses present in the aquatic environment, only a small part is responsible for human infections. However, even at low concentrations viruses are responsible for infections, since they have a low infectious dose. The concentration of water samples is an important step for the detection of enteric viruses. However, there is no ideal method to concentrate viruses in water samples, always depending of the goal of the work. In this work, we found that the ultracentrifugation method is a good method for the concentration of viruses in environmental waters, once it was obtained recovery rates between 66 and 76%. On the other hand, this method does not add any chemical compound, allowing posterior viral detection by molecular techniques. Surface microlayer (SML) corresponds to the interface between water and air, accumulating microorganisms and particles. It is known that constitutes a natural reservoir for enteric viruses from which aerosol are released, transporting materials accumulated in the SML. In this study the enrichment values for SML in respect to underwater (UW) were higher than 2 in the marine zone and in the brackish water zone. The enrichment of SML regarding to aerosols, was about 26 in the marine zone and about 30 in the brackish water zone for the SML, showing that the SML acts as a reservoir for enteric virus and that these are transmitted from the aquatic environment into the atmosphere through aerosols. Regarding the abundance of enteric viruses studied, rotavirus and enterovirus were more abundant during cold months and HAV presents similar abundance in cold and warm months. Variations in the abundance of enteric virus were explained by differences in water temperature and in nitrates and nitrites concentration, suggesting terrestrial origin of these viruses, wich are transported to maritime system by rain water runoff, during cold months. xvi Nowadays we face serious problems of climate change and the consequences of changes in the ecosystems are a major problem. The influence of UV light exposure in the abundance of enteric viruses was tested and it was found that only enterovirus showed a significantly reduction in their abundance (p = 0:03 <0.05 and p = 0:41 < 0.05, respectively). Differences observed betwenn the two groups may be related with the RNA type, dsRNA for rotavirus and ssRNA for enterovirus, but also with the capside structure, with three layes for rotavirus and only one for enterovirus. xvii Thesis Outline The viruses in the aquatic system are among the best fitted living beings to become emerging pathogens and are able to survive and remain infectious for long periods. Most of these microorganisms present low infectious dose, representing a major threat for human health and a problem for economy and environmental ecology. Regarding to this problems, it becomes urgent to determine the source of viral contaminations and the way of their transference and transport, in order to evaluate their association with health risk and implement the required measures. This work pretends to understand how surface microlayer (SML) acts as a reservoir for enteric viruses in aquatic system, the way they are transferred to the atmosphere and the influence of UV light on viral abundance. In order to achieve the knowledge pretended, several specific goals were established: • To determine the best concentration method to detect viruses by molecular methods. • To determine the role of SML as a reservoir of enteric virus in the aquatic system. • To determine the best method for aerosols formation and recovery. • To determine the role of aerosols in the transference of enteric viruses from the aquatic environment to the atmosphere. • To determine the influence of UV light on enteric viral abundance in the aquatic system. Chapter 1 presents a careful revision of the state of the art for enteric viruses in the aquatic system, the factors affecting their survival, the way they are transmitted to the atmosphere and the methods used to detect them. Since only a small number of viruses are epidemiologically relevant to human health, direct viral detection is not possible, being necessary to concentrate large volumes of water before proceeding to their detection. There are several concentration methods for viruses but none of these methods is considered perfect. Since the choice of the concentration method depends on the sample tested and the subsequent analysis to be made. xviii Chapter 2 includes a laboratorial work part, to compare two concentration methods (organic flocculation and ultracentrifugation) and decide which one would be used in the subsequent work. Chapter 3 includes a study on enteric viral abundance in both SML and underwater (UW) in order to determine the role of SML and aerosols in accumulation and transference of enteric virus. In virology, much is already known about the viruses present in the aquatic system that represent a health hazard for humans. However, little is known about their natural reservoirs, namely the SML. In this chapter it was also evaluated the physical and chemical parameters that may affect viral abundance. Chapter 4 includes a laboratorial work section to develop a protocol for aerosols collection and a field work section, where aerosols were collected and viral abundance determined. It is known that aerosols constitute the main vector for transport of viruses across the air-water interface, being their composition dependent on SML composition. Chapter 5 presents a laboratorial study about the UV light influence on enteric viruses abundance. Viral survival may vary, depending on viral type. However, there are several factors that may affect that survival. Chapter 6 presents all the conclusions of this thesis and some future work considerations. Chapter 1 - Introduction Chapter 1 3 1. Introduction At the present, poor water quality due to the presence of viruses still represents a major threat for human health (Symonds and Breitbart, 2014). It becomes urgent to trace and characterize the type and origin of viral contamination in order to evaluate their association with health risk and implement the required measures (Bosch 1998, Griffin et al, 2003, Bosch et al. 2005, Myrmel et al. 2006). Viruses are the simplest form of life, consisting of genetic material, which may be either DNA or RNA, surrounded by a protein coat and, in some cases, by a membranous envelope, being characterized by their simple organization and their unique mode of replication (White et al, 1994, Mahony et al. 2008). Unlike cellular organisms, viruses do not contain all the biochemical mechanisms for their own replication; they need to use the biochemical mechanisms of a host cell to synthesize and assemble their separate components. Viruses vary in their stability. Some are very stable and survive well outside the host body while others do not survive well and, therefore, usually require close contact for transmission being readily destroyed in the environment (White et al. 1994, Mahy et al. 2009). Among microorganisms, viruses are best fit to become emerging pathogens since they are able to adapt not only by mutation but also through recombination and reassortment and can thus become able to infect new hosts and to adapt to new environments (Desselberger 2000, Tabor 2007, LaRosa et al, 2012). Mutations contribute to changes in viral pathogenicity, since viruses have a simple structure allowing mutations to occur easily and viral explosive replication magnifies the mutation (Tabor, 2007). RNA viruses are particularly susceptible to these modifications since their host lack molecular “proofreading” mechanisms to correct these mutations and replication errors (Tabor 2007; Domingo, 2010). Some health relevant enteric viral groups are nowadays considered to be emerging waterborne pathogens (LaRosa et al, 2012; Woods, 2013), increasing the concern about the discharge of human enteric viruses in not only fresh water but also in estuarine and marine waters (Lee and Kim, 2002; Hamza et al, 2009; Rodríguez-Díaz et al, 2009). Chapter 1 10 pH Enteric viruses, in general, are very stable at the pH of most natural waters (pH 5 to 9) (Melnick et al. 1978, Duizer et al. 2004, Cannon et al. 2006, Bosch 2007), but most enteric viruses are more stable at low pH (3 to 5) (Melnick et al. 1978, Duizer et al. 2004, Thurston-Enriquez et al. 2005, Cannon et al. 2006, Bosch 2007), because the increase in pH has a direct toxic effect by fragmenting the nucleic acids (Bosch 2007, Abdel-Moety et al. 2008). It has been suggested that the sensitivity of enteric viruses to pH may be strain dependent (Bosch 2007). Enterovirus can survive at very high pH (11.0 to 11.5) and very low pH (1.0 to 2.0) for short periods of time. Adenovirus and rotavirus are sensitive to inactivation at a pH of 10.0 or greater (Bosch 2007). Calicivirus are inactivated between pH 3 and pH 5 (Abdel-Moety et al. 2008). However, Duizer et al. (2004) shown that 3 h at pH 2.7 was not enough to completely inactivate norovirus. UV light The ultraviolet light of sunlight can inactivate viruses by causing cross-linking among the nucleotides (Suttle et al. 1992, Noble et al. 1997, Bosch 2007). Because UV radiation from the sun is present in the environment, natural defense mechanisms have evolved in microorganisms that allow UV inactivated microorganisms to reverse UVinduced damage through such repair pathways as photoreactivation and dark repair (Koivunen and Heinonen-Tanski, 2005; Quek and Hu, 2008; Hu et al, 2011). A microorganism UV resistance depends on the UV dose applied and its ability to protect itself from UV light and to repair damages (Sommer et al., 2001; Hu et al, 2011). UV disinfection is highly effective against most viral pathogens at low doses (40 to 60 mJ/cm 2 ) commonly rendered in water treatment plants (Guo et al, 2010). However, viruses are more resistant to UV radiation than many other pathogens, because of their low molecular weight (Suttle et al. 1992, Bosch et al. 2006, Suttle, 2007). The effectiveness of UV radiation in the inactivation of Calicivirus is comparable to that of the Enterovirus and less effective than that of Adenovirus (Thurston-Enriquez et al, 2003; Duizer et al. 2004). Salinity Although salinity does not present a direct effect on viral survival (Lo et al, 1976; Fong et al, 2005; Bosch et al, 2006), many studies relate that virus almost always survive longer in freshwater than in seawater (Wetz et al, 2004; Bosch, 2007; Seo et al, 2012). This may be due to the fact that high salinity concentrations are virucidal (Lo et al, Chapter 1 11 1976; Fong et al, 2005; Bosch et al, 2006) and because salinity increase viral aggregation, which appears to contribute to the loss of viral titre (Wetz et al, 2004; Bosch, 2007; Seo et al, 2012). Adsorption to sediments The association with sediments does not act only as reservoirs of human enteric viruses but also as a source from which viruses can be released into the water column by storm action and dredging activities (Rao et al, 1984; Bosch et al, 2005; Fong et al, 2005; Bosch, 2007). While viruses associated with small-size particulate material (<3 µm) tend to float in the water column, viruses adsorbed onto large/medium particles (>6 µm) readily settle down in the bottom sediment (Grabow, 1996; Bosch et al, 2005; Fong et al, 2005). Adsorption of enteric viruses to sediments has been demonstrated to prolong the survival of Enterovirus (Bosch, 2007), which remain infectious for 19 days compared to 9 days for unassociated enteroviruses in the water column (Griffin et al,2003). The increased virus survival in the presence of sediment has important implications in the marine environment, because fecal contamination of coastal areas results in contamination of shellfish harvesting areas, accumulation of solid-associated viruses in sediments, with sediments acting as virus reservoirs, and accumulation of viruses in shellfish (Bosch et al, 2006). Additionally, viruses concentration by molluskan bivalves is enhanced by the presence of particulate material (Bosch et al. 2006). 1.3. Enteric virus transmission from the marine environment Viruses are a major cause of water-related disease (Bosch et al, 2008) and humans are exposed to marine enteric viruses through the consumption of shellfish grown in contaminated sea waters; to a lesser extend through sewage-polluted recreational waters, through the sea bathing and water sport activities, that are becoming increasingly popular and nowadays are expanded beyond the traditional summer seasons; and through marine aerosols (Bosch et al. 2005, Lipp et al. 2002, Bosch et al. 2006, Bosh et al. 2008). Although much is known about the aquatic specific viruses that put in danger human populations, little is known about their natural reservoirs, namely the surface microlayer (SML), and their transmission to the atmosphere through aerosols. Since many pathogenic viruses for humans circulate in the marine environment and some of them are considered emerging pathogens (Albinana-Gimenez et al., 2006; Chapter 1 12 Muscillo et al, 2008; LaRosa et al, 2012), it is pertinent to determine their density in SML, underlying water (UW) and atmosphere in order to assess the potential transmission via aerosols. 1.3.1. Role of SML in viral transmission Aquatic surface microlayers are just a few tens of micrometers deep at the air–water interface and are physicochemically distinct from the UW below (Cunliffe et al, 2011). The primary interest in the structure and function of the SML is based on its crucial role in exchange processes of gases and matter across the air-water interface (Agouguè et al, 2004). The location of SML makes it a highly dynamic system. Atmospheric inputs to surface films include wet/dry deposition, air–sea gas and aerosol transfer. Exchange with the atmosphere is strongly influenced by the physicochemical and microbiological nature of SML (Cunliffe et al, 2011) (Figure 1.1). Water column processes regulate the accumulation of material in the SML and small molecules and larger particles can accumulate at to form a film that extends into the UW (Cunliffe et al, 2011). In comparison with the UW, SML is physically more stable because of surface tension (Hardy, 1982) and the presence of a biogenic gelatinous film layer constitute a microbial habitat where compounds and particles can concentrate (Cunliffe et al, 2011). The SML is generally enriched in organic materials, which might stimulate biological growth. However, it has been shown that density, activity and diversity of microorganisms in the SML can be higher, similar or lower than those in UW. Unlike in the UW, microorganisms at SML receive maximal UV radiation, which has the potential to cause direct nucleic acid damage or indirect damage via the formation of destructive intermediates such as reactive oxygen species (Cunliffe et al, 2011). Organisms present in the SML may have developed strategies to survive in this habitat exposed to extreme conditions, and toxic heavy metals (Hardy 1982, Williams et al. 1986; Liss and Duce, 1997), but although these organisms may be exposed to the stress of SML, they can also be protected by the organic matrix present in this layer. SML habitats are unique as they interact with both the atmosphere and the hydrosphere simultaneously (Cunliffe et al, 2011). Chapter 1 13 Figure 1.1: Inputs, outputs and processes whithin aquatic surface microlayer (Cunliffe et al, 2011). Although viruses are the most abundant biological entity, representing the largest reservoir of genetic diversity, scarce information about their natural reservoirs, such as SML, is available. It is known that virus in SML are as abundant as in UW (Parada et al, 2005), but until now, there is no study on human pathogenic viruses in the SML. This can be, in part, due to difficulties in SML collection relatively to UW, which represents a greater problem for viral detection, since large volumes of water samples are needed. The most commonly used devices for SML collection are mesh screens, often referred to as Garret screens (Garrett, 1965), glass plates (Harvey, 1965; Harvey & Burzell, 1972) and membranes (Crow et al., 1975; Kjelleberg et al., 1979), all of which whith inherent advantages and disadvantages (Cunliffe et al, 2011) (Figure 2). Mesh screens rely on the collection of water in the interstitial spaces of a mesh. Metal or plastic mesh is stretched over and secured to a hand-held frame. The mesh screen is oriented horizontally and lowered through the SML into the UW, before being slowly withdrawn in the same way (Figure 2). Water retained within the mesh is then poured into a sampling vessel for processing as required (Cunliffe et al, 2011). Glass plates (typically 20–30 cm 2 ) exploit the adhesion of a SML sample to a clean glass surface. Unlike the mesh, the glass plate is oriented vertically, lowered through the SML into the UW and slowly raised back out. The adhered sample is then removed using a wiper blade (e.g. neoprene) (Cunliffe et al, 2011) (Figure 1. 2). Chapter 1 14 Figure 1.2: Mesh screen and membrane surface microlayer samplers in use (Cunliffe et al, 2011), at left. Glass plates surface microlayer sampler in use, at right. SML can also be sampled by adhesion to sterile membrane filters placed directly onto the water surface (Crow et al., 1975; Kjelleberg et al., 1979). Various types of membrane material are available; like polycarbonate and PTFE have been used (Crow et al, 1975; Kjelleberg et al, 1979; Cunliffe et al, 2011).Membranes present some distinct advantages over mesh screens and glass plates such the fact that they do not come into contact with the UW (Figure 2) (Cunliffe et al, 2011). The potential for contamination is virtually eliminated using forceps; individual membranes are lifted from the water surface with the SML attached and placed into storage vessels for processing. 1.3.2. Transmission of viruses by aerosols Marine erosols are generally defined as colloidal systems of liquid or solid particles suspended in a gas (Hinds, 1999; Baron and Willeke, 2001; Fuzzi et al, 2006; Després et al, 2012), which can contain microorganisms and are generated through natural processes in the marine environment, like wave action (Fannin et al, 1985) or bubbles bursting through the SML (Aller et al, 2005; Morris et al, 2011). The size of aerosols can range from several nanometers to a few hundred micrometres in aerodynamic diameter (Cox and Wathes, 1995; Hinds, 1999; Jaenicke, 2005; Pöschl, 2005; Després et al, 2012) and viruses are transported in the smallest aerosols, with physical diameter as low as 20 nm (Dongsheng, 2006; Després et al, 2012). However, viruses are not commonly airborne as individuals and are more likely attached to other suspended particles (Després et al, 2012). Marine aerosols can be classified as large respiratory droplets and droplet nuclei (Mubareka et al, 2009). Large respiratory droplets are > 5-15 µm in diameter and are involved in short-range transmission. Droplet nuclei are < 5 µm and are responsible for Chapter 1 15 transmission over great distances (long-range airborne transmission) (Mubareka et al, 2009). Marine aerosols are formed primarily by the eruption of rising bubbles through SML (Aller et al, 2005; Morris et al, 2011) and contain high concentration of salts, organic matter, proteinaceous material, gel particles, microorganisms and viruses (MathiasMaser & Jaenicke, 1994; Posfai, Li, Anderson & Buseck, 2003; St. Louis and Pelletier, 2004; Kuznetsova et al., 2005; Brooks et al, 2011; Morris et al, 2011; Danovaro et al, 2012). Bubbles eventually rise through the water column and upon reaching the sea surface, burst and eject aerosol droplets into the atmosphere, delivering the material carried by the bubbles to SML and to the atmosphere (Blanchard, 1975; Blanchard & Syzdek, 1982; Aller et al, 2005). Thus, the composition of marine aerosols formed from bursting bubbles at the sea surface changes in response to the SML composition (O’Dowd et al., 2004; Russell et al., 2010; Cunliffe et al, 2011). Bubble bursting and aerosol formation is an important transport mechanism for SML components (Russell et al., 2010). Bubbles generate the major portion of marine aerosols that are easily suspended and transported, through long distances, in the lower atmosphere (Gutafsson & Franzen, 2000; Grammatika & Zimmerman, 2001). Aerosols formation constitutes the main vector for transport of viruses across the airsea interface, playing an important role in long-distance dispersal and being responsible for the distribution of viruses (Mathias-Maser & Jaenicke, 1994; Posfai et al, 2003; St. Louis and Pelletier, 2004; Li et al, 2008; Brooks et al, 2011; Danovaro et al, 2012), since aerosols may remain suspended in the atmosphere for weeks (Dueñas, Fernandez, Carretero, Liger and Cñete, 2004). Any microorganism, and also viruses, can become airborne (Verrault et al, 2008). If the aerodynamic size of an infectious particle is appropriate, it can remain airborne, come into contact with humans or animals, and potentially cause an infection (Verrault et al, 2008). The probability of an airborne microorganism-laden particle causing an infection depends on its infectious potential and its ability to resist the stress of aerosolization (Verrault et al, 2008). Several studies have shown that marine aerosols large enough to contain organic particles, microbes and viruses can be transported for hundreds of kilometers from their source (Klassen and Roberge, 1999; Moorthy et al, 1998; Chow et al, 2000). Aerosols can be single virus, aggregates of one or several types of particles, or attached to non-biological particles (Sun and Ariya, 2006; Morris et al, 2011). While single virus particles exist in the air, they tend to aggregate rapidly (Verrault et al, 2008), with aggregation speed depending on the size distribution of the airborne particles, the concentration of the aerosol, and of the thermodynamic conditions (142; Chapter 1 16 (Verrault et al, 2008). Organic and inorganic materials in viral aerosols can affect the size of the aerosolized particles and their infectious potential. Many factors, such as relative humidity, temperature, radiation, aerosolization medium, exposure period, chemical composition of the air and also the sampling methods, can affect the infectivity of airborne viruses (Verrault et al, 2008). When a virus is contained in an aerosol, their infectivity is increased, compared to a virus free in the air, since the aerosol prevent desiccation and protects from temperature and sunlight (Li et al, 2008) and so, aerosols of pathogenic viruses pose an increased threat to health (Adams et al, 1982). For the past 200 years, the field of aerobiology has explored the abundance, diversity, survival and transport of microorganisms in the atmosphere (Morris et al, 2011). Yet, the atmosphere still presents a frontier for pioneering microbiologists (Verrault et al, 2008; Morris et al, 2011). This is due mainly to the difficulty in collecting and analyzing airborne biological contaminants, which is an even greater problem for viral detection (Verrault et al, 2008). The perceived lack of research might also be connected to the fact that before the development of molecular biological methods (e.g. PCR) to detect genetic material of microorganisms (Alvarez et al., 1995; Peccia and Hernandez, 2006), only viable viruses could be found in air samples (Morris et al, 2011). Another possible reason for the scarcity of publications containing information on viable viruses in atmospheric aerosol is inactivation of viruses in the atmosphere under the influence of different environmental factors (changes in temperature, relative humidity, solar radiation, etc.). Aerosols sampling techniques have been improved greatly over the years, giving more ability to tackle the important health issue of airborne viruses. However, the lack of standardization has to be addressed, as it limits the development of general recommendations for sampling of airborne viruses (Verrault et al, 2008). Studies to date have rarely included quantitative analyses of total viral load. As far as the knowledge of viruses in the aerosol is concerned, most of the studies have been focused on the analysis of specific pathogenic viruses and their infectivity (Ijaz et al, 1987; Sagripanti and Lytle, 2007). Only two studies on marine aerosol have reported information on the abundance of viruses and their enrichment factor in comparison with the SML (Aller et al, 2005; Kuznetsova et al, 2005). The probability of detecting airborne viral pathogens is dependent on three primary factors: the concentration of airborne viruses in the environment, the ability of the airsampling system to recover airborne particles (collection efficiency), and the analytical sensitivity of the diagnostic assay(s) used to detect the target pathogen in the sample (Herman et al, 2006; Verrault et al, 2008). Chapter 1 17 The sampling and collection of atmospheric aerosols requires a great understanding of the physical principles that govern interactions with suspended particles. As such, particle size is by far the most important characteristic for choosing a sampling procedure for airborne particles (Nicholson, 1995; Després et al, 2012). A representative sample should contain nanoparticles with larger particles (Verrault et al, 2008). However, the abiotic factors are also important to evaluate the possible formation of aerosol and the subsequent transmission of viruses from SML to the atmosphere. It is known that different environmental conditions, including relative humidity, temperature, and UV radiation, wind velocity and particulate matter, influence the characteristics of the viruses, including concentration and infectivity (Li et al, 2008). Collection of living, or viable, material requires care in order not only to sample microorganisms appropriately but also to keep them alive until the desired analysis can be performed (Després et al, 2012). Different methods have been developed based on the attachment properties to surfaces of airborne particles. The most used air samplers are based on impact on solid surfaces, impingement, and filtration (Figure 3). All of them have been successfully used for virus detection, but have advantages and disadvantages (Verrault et al, 2008; Bosch et al, 2011). While culture is often used to determine viral concentrations and most sampling methods affect viral infectivity, culture methods become inadequate for calculating the true concentrations of infectious airborne viruses. Technologies such as PCR can be used to detect viruses in air samples even when they are no longer infectious (Verrault et al, 2008). Figure 1.3: Different bioaresol samplers (Verrault et al, 2008). A: Impact samplers; B: Impinge samplers and C: Filter samplers. Chapter 1 18 Aerosols collection by impact samplers or solid impactors The impact samplers are usually more efficient at capturing large particles. Andersen and slit samplers accelerate the particles through narrow holes or slits. The streamline moves toward a solid surface and abruptly changes direction (Figure 1.3.A). The inertia of the particles deviate them from the airflow and impacts them on the surface, which usually holds a petri dish with a culture medium. The medium is either washed to collect the particles or used directly for plaque assays (Verrault et al, 2008). Slit samplers are used mostly to determine aerosol concentrations of microorganisms as a function of time. The accelerated particles are impacted onto a rotating petri dish containing a culture medium with a liquid layer, making it possible to determine the time when each particle was sampled (Verrault et al, 2008). Impact samplers are easy to use, but dehydratation or impact trauma can affect viral survival. Flow rate and sampling duration are crucial when using these samplers (Bosch et al, 2011). Aerosols collection by impinge samplers or liquid impactors The all-glass impingers (AGI) are the most often used samplers for the capture of airborne viruses (Verrault et al, 2008). The liquid impinge offer the possibility of extended sampling times (Després et al, 2012) and it works by accelerating airborne particles through a narrow orifice placed at a fixed distance from the bottom of a flask containing a liquid. A pressure drop is created in the flask and forces the air to enter through the inlet of the impinger. The air enters horizontally through a glass tube, which curves to a vertical position, forcing the air to change direction and flow downward. The diameter of the tubing abruptly narrows and acts as a critical flow orifice, accelerating the air passing through it to sonic velocity. The curve in the tube is intended to trap the larger particles by inertial impaction and mimics the airway of the human nose (Herman et al, 2006; Verrault et al, 2008) (Figure 1.3.B). The formation of small bubbles in the liquid of the impinger can also help to sample very small particles by diffusion. However, the reaerosolization of particles due to the scavenging properties of the air bubbles can be a problem, especially for hydrophobic particles (Verrault et al, 2008). After sampling, the medium can be concentrated or directly decontaminated, purified, and analyzed (Bosch et al, 2011). The recovery efficiency of this method is high because it avoids dehydration and the liquid facilitates the extraction of genetic material, for subsequent analysis (Verrault et al, 2008; Bosch et al, 2011). However, flow rate and the composition of the collection fluid are critical for virus recovery (Bosch et al, 2011).The gentler sampling process leading to better recovery of infective viruses Chapter 1 19 seems to be the main reason for the wide use of AGI samplers in aerovirology (Verrault et al, 2008). A number of variables are known to affect impinger collection efficiency. These include impinger design (Cage et al, 1996; Donaldson et al, 1982; Herman et al, 2006; Jensen et al, 1992), sampling time (Lin and Li, 1998), and the composition of collection medium (Lin et al, 2000). Sampling times reported for the recovery of airborne pathogens using air impingers are variable, ranging from minutes to hours (Terzieva et al, 1996; Donaldson et al, 1987; Lin and Li, 1998; Lin and Li, 1999; Herman et al, 2006). Likewise, flow rates described for the collection of airborne viruses are variable, ranging from 12.5 liters/min (Elazhary and Derbyshire, 1979) to 450 liters/min (Dee et al, 2005; Herman et al, 2006). Collection media described for the recovery of airborne pathogens in air impingers are diverse including deionized water (Lin et al, 1997), buffered solutions (Juozaitis et al, 1994) and mineral oil (Lin et al, 2000). Compounds added to collection medium to improve pathogen recovery include various proteins (Stolze et al, 1989) and antifoaming agents (Karim et al, 1985; Schoenbaum et al, 1990; Chinivasagam and Blackall, 2005; Herman et al, 2006). According to Harstad (1965), liquid impingers are the least destructive samplers, being 18% more efficient, although 30% to 48% of the sample is physically lost (Després et al, 2012; Herman et al, 2006; Verrault et al, 2008). Aerossols collection by filter samplers In filter samplers, the air passes through a filter and airborne particles are retained (Bosch et al, 2011) (Figure 1.3.C). Filter efficiency is based on the following five basic mechanisms: (i) interception, (ii) inertial impaction, (iii) diffusion, (iv) gravitational settling and (v) electrostatic attraction (Hinds, 1999). Filtration efficiency improves with increasing and decreasing particle size (Verrault et al, 2008). Many different types of filters have been used to sample airborne viruses. They differ mainly in composition, pore size, and thickness (tightly packed cotton, cellulose filters (0.45µm pore size), PTFE filters (2.0µm pore size) and gelatin filters) (Verrault et al, 2008; Després et al, 2012). Polycarbonate filters are much less efficient than gelatin or PTFE filters (Burton et al, 2007). The low filtration efficiency of polycarbonate filters may be due to the structure of the filter. The contact area of filters with uniform cylindrical pores, such as polycarbonate filters, is much smaller than that of filters with a complex structure, such as PTFE filters, where the probability of adherence is greater because airborne particles are exposed to a greater surface area (Verrault et al, 2008). Chapter 1 26 fluoresce when bound to complementary sequences in the amplified region (Fong and Lipp, 2005). The procedure is less time-consuming because a confirmation step such as agarose gel electrophoresis and additional hybridization are generally not required. The entire analysis can be done in a closed system, which may reduce the potential for contamination (Fong andLipp, 2005). However, the cost of a real-time PCR instrument is still substantially more than that of a conventional PCR instrument, and in some cases, real-time PCR has been shown to be less sensitive than conventional RT-PCR and nested PCR (Noble et al, 2003; Fong and Lipp, 2005). Pyrosequencing To determine the identity of the genomes obtained, the PCR products are often cloned and sequenced (Boreson et al, 2004; Maron et al, 2005; Després et al, 2007; Fierer et al., 2008; Bowers et al., 2009; Fröhlich-Nowoisky et al, 2009; Georgakopoulos et al., 2009; Després et al, 2012). Pyrosequencing uses luciferase to generate light for detection of the individual nucleotides added to the nascent DNA, and the combined data are used to generate sequence readouts (Després et al, 2012). Species can often be identified by comparing the obtained sequences with those that are already available in online databases like the National Center for Biotechnology Information (NCBI) (Després et al, 2012; Zao et al, 2013). The high demand for low-cost sequencing in recent years has led to the development of high-throughput sequencing technologies. In these technologies, the sequencing process is parallelized for several samples, and thus in a short time thousands or millions of sequences are produced (Després et al, 2012). The advent of wholecommunity genome sequencing is rapidly changing the way viral and microbial diversity are assayed. Using this approach, it is possible to rapidly characterize the metabolic diversity and community structure of any microbial ecosystem (Angly et al, 2006). Moreover, quick and accurate identification of microbial pathogens is essential for both diagnosis and response to emerging infectious diseases. High-throughput sequencing has recently emerged as a powerful approach to identify both known and novel viruses in clinical specimens (Zao et al, 2013). Integrated cell culture PCR (ICC-PCR) While PCR-based methods offer many advantages in sensitivity, specificity, and efficiency over cell culture, they still cannot provide information on the infectivity of viruses detected with the reliability of cell culture. (Fong and Lipp, 2005). Cell culture combined with PCR (ICC-PCR) is an approach that has been used to overcome most Chapter 1 27 of the disadvantages associated with both conventional cell culture and direct PCR assays (Reynolds, 2004; Rodríguez et al, 2009). Detection relies on an initial biological amplification of viral nucleic acid, followed by amplification via PCR (Reynolds et al, 1994; Rodríguez et al, 2009). Viruses are allowed to replicate in cell culture for short periods followed by PCR amplification, which dramatically reduces the time necessary for infectious viral detection (Reynolds, 2004; Rodríguez et al, 2009). ICC-PCR has also the advantage of detecting viable viruses that do not produce cytophatic effects (CPE). The sensitivity obtained with ICC-PCR is comparable to that obtained in cell culture after a second passage in cell culture (Blackmer et al, 2000; Rodríguez et al, 2009). In addition, fewer problems are encountered with inhibitory compounds that may be contained in environmental concentrates (Chapron et al, 2000; Rodríguez et al, 2009). The use of ICC-PCR has been described for the detection of enteroviruses (Reynolds et al, 1996; Rodríguez et al, 2009), hepatitis A virus (Reynolds et al, 2001;Jiang et al, 2004; Rodríguez et al, 2009), enteric adenoviruses (Lee and Kim, 2002; Rodríguez et al, 2009), and astroviruses (Reynolds et al, 2001; Rodríguez et al, 2009). 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Three independent assays were done for each situation in each sampling date. Nucleic acids were extracted from water samples using the geneMAG-RNA/DNA kit, a magnetic RNA/DNA purification kit (Chemicell™), according to the instructions of the manufacturer. For nucleic acid purification the Geneclean kit (MP Biomedicals, LLC) was used according to the instructions of the manufacturer. Detection of adenovirus and rotavirus was performed in a Labnet TC9600-G thermocycler and amplification products were separated by electrophoresis in a 2% agarose gel stained with ethidium bromide and detected with a UV transilluminator. Detection of Adenovirus Detection of adenovirus was made by a nested PCR technique using the primers described by Allard et al, 2001. Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2 mM Buffer taq, 1.5 mM MgCl2, 1.28 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.4 mM of each primer (StabVida) (hex1deg 50-GCC SCA RTG GKC WTA CAT GCA CAT C-30 and hex2deg 5’-CAG CAC SCC ICG RAT GTC AAA-3’). After the first PCR, 5 µL of PCR product were added to 20 µL of a new reaction mixture consisting of 1 mM Buffer taq, 1.5 mM MgCl2, 1.28 mM dNTPs, 0.4 mM of each primer (nehex3deg 5’ GCC CGY GCM ACI GAI ACS TAC TTC 3’ and nehex4deg 5’ CCY ACR GCC AGI GTR WAI CGM RCY TTG TA 3’) and 0.1 U µmL -1 of Taq polymerase. All primers sequences are found between base pair position 21 and position 322 in the coding region of the hexon gene. The first set of primers creates a 301-bp product and the second set of primers creates a 171-bp product. The amplification was carried out for 45 cycles at 94ºC for 10 s, 55ºC for 30 s and 72C for 20 s after initial denaturation at 94ºC for 4 min. A final extension step was performed at 72ºC for 5 min. The amplification products were separated by electrophoresis in a 2% agarose gel stained with ethidium bromide and detected with a UV transilluminator. Detection of Rotavirus Detection of rotavirus was made with a PCR technique using the primers described by Villena et al, 2003 and using the SuperScript™ II RT for the synthesis of cDNA, according to the manufacturer instructions. Five microliters of sample were added to 20 µl of the reaction mixture, which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U µmL -1 of Taq polymerase (Fermentas) and 0.48 mM of each primer Chapter 2 59 (StabVida) (VP6-3 5’-GCT TTA AAA CGA AGT CTT CAA C-3’ and VP6-4 5’-GGT AAA TTA CCA ATT CTT CCA G-3). Rotavirus primers positions is found between the position 187 and position 166 of human strain Wa [accession number K02086]), creating a product of 186-bp. The amplification was carried out for 40 cycles at 94ºC for 10 s, 50ºC for 30 s and 72ºC for 20 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. The amplification products were separated by electrophoresis in a 2% agarose gel stained with ethidium bromide and detected with a UV transilluminator. Quantification of Adenovirus in concentrated water samples by qPCR The quantification of adenovirus was performed in an iQ5 thermocycler and standards were obtained from a serial dilution of a suspension of Adenovirus, with a known initial number of copies (obtained by nucleic acid quantification with nanodrop). The qPCR was performed using the same protocol as that described for detection of Adenovirus using the PCR technique, but with a reaction mixture of 2x iQ ™ SYBR ™ Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6 mM MgCl2, SYBR Green I, fluorescein and stabilizers) and 0.4 mM of each primer (nehex3deg and nehex4deg), to which was added 5 µl of sample. Statistical analysis For the enumeration of virus-like particles (VLP) and for the detection and quantification of the two enteric viruses, three sub-samples were used for both concentration methods and for both water types of each sampling date. For the enumeration of VLP, for each subsample, 3 replicates were analysed and for the detection and quantification of the enteric virus only 2 replicates were done for each sub-sample. The results of the three sub samples were averaged and the standard deviation was calculated. The differences between the efficiency of recovery, determined by the enumeration of VLP, of the two concentration methods were analyzed by one-way ANOVA to check for significant differences between methods. The difference between Adenovirus quantification in recreational waters concentrated by the two methods was also evaluated using one-way ANOVA. Only the data with normal distribution (assessed by the Kolmogorov–Smirnov test) and with homogeneity of variances (assessed by Levene’s test) were used. A value of p < 0.05 was considered significant. Statistical analyses were performed by using SPSS (SPSS 15.0 for Windows, SPSS Inc., USA). Chapter 2 60 Results Efficiency of viral recovery for the two concentration methods The recovery efficiency for sewage samples (Fig. 2.1a) and recreational samples (Fig. 2.1b) was higher with the ultracentrifugation than with the flocculation method. Ultracentrifugation showed average recovery efficiency of 69% for wastewater and of 76% for recreational water samples. The corresponding recovery efficiencies with the flocculation method were 38% and 22% for wastewater and recreational water, respectively. The differences between the efficiency of recovery for the two sampling dates were not significant for wastewater (p = 0.86 and p = 0.16 respectively), but for recreational waters there were significant differences in the efficiency of recovery with the two concentration methods for both sampling dates (p = 0.01 and p = 0.00, respectively). Detection of enteric viruses in water samples The adenovirus and rotavirus A in wastewater and recreational water samples were found in all samples after concentration by ultracentrifugation and by flocculation. Quantification of Adenovirus in recreational water samples The presence of adenovirus in recreational water samples after concentration was tested by qPCR (Fig. 2.2). This group of virus was present in all samples after concentration. The differences between the quantification of adenovirus, for the two concentration methods, for the two sampling dates, were not significant (p = 0.057 and p = 0.868, respectively). Fig. 2.1: (a) Recovery rate of VLP, for both concentration methods, in sewage waters, for two sampling dates. (b): Recovery rate of VLP, for both concentration methods, in recreational waters, for two sampling dates. Values represent the mean of three independent experiments; error bars represent the standard deviation. Chapter 2 61 Fig. 2.2: Quantification of adenovirus in recreational water samples after concentration by flocculation and ultracentrifugation for the two sampling dates. Values represent the mean of three independent experiments; error bars indicate the standard deviation. Discussion Environmental samples must be sufficiently concentrated to allow efficient detection of very few viruses in a large volume of water (Schwab et al, 1995). However, there is no perfect viral concentration method for water samples (Bosch et al, 2008; AlbinanaGimenez et al, 2009), the method should be chosen so that, despite its limitations, it is adequate to achieve the aim of the work. In the present work, the ultracentrifugation and the flocculation methods were compared, considering further use of the samples for molecular analysis such as PCR or pyrosequencing. The results of this study show that the ultracentrifugation method, usually used as a secondary step for virus concentration (Steward, 2001; Fout et al, 2003; Kovac et al, 2009; Sylvain et al, 2009), is an adequate approach to concentrate virus directly from environmental waters and that the determination of the VLP by microscopy is a simple, fast and cost effective method to evaluate the recovery efficiency of the concentration method. The ultracentrifugation method recovered about 30% more viruses in residual waters and about 50% more in recreational waters than the organic flocculation method, one of the most currently used approaches to concentrate viruses from environmental waters (Calgua et al, 2008; Guttman-Bass and Armon, 1983; Guttman-Bass and Nassen, 1984; Shields and Farrah, 1986; virobathe, 2011). Moreover the ultracentrifugation method is simpler and faster than the flocculation approach. During flocculation, HCl is added to water samples to adjust the water pH to 3.5 and skim milk solution is added to allow virus adsorption. It is, however, well known that during the Chapter 2 62 concentration step, inhibitory substances are concentrated along with the virus and, consequently, the addition of HCl and skim milk solution may increase the inhibitory effect on subsequent PCR reactions. In ultracentrifugation, only the non-added inhibitory substances are concentrated. Although for recreational water this effect can be negligible, for wastewaters, which have large concentrations of inhibitory substances, purification of the concentrated samples is always required (Guo et al, 2009; Moussani et al, 2009). For more clean environmental waters this can be an inconvenience relative to the ultracentrifugation method. The incubation period of more than 16 hours used for flocculation may also affect the structure of the viral community. It is well known that bacteriophages represent a large fraction of the virioplankton (Bettarel et al, 2008; Miki and Jacquet, 2008; Suttle and Fuhrman, 2010) and their replication cycle is frequently less than 1 hour (Bettarel et al, 2008; Madigan and Martinko, 2006), allowing these virus to undergo several replications during the period of concentration by flocculation. In contrast, viruses that infect eukaryotic cells have replication times of around 40 hours and, consequently, it is unlikely that they replicate during the incubation period. These facts are not important if the concentration method is used to detect specific enteric virus by PCR because specific primers are used, but if the concentrated water samples are used to study the structure of the viral community, the results will not reflect the structure of the original community. In contrast, in the ultracentrifugation method, as water samples are filtered with 0.2 mm membranes in order to remove bacteria prior to centrifugation, all cellular hosts are removed and viral replication is avoided, even for bacteriophages that have shorter life cycles. However, the water filtration before ultracentrifugation can cause viral loss due to membrane clogging. The replacement and/or the employment of largesize membranes, as used in this study, can overcome this problem. Moreover, when ultracentrifugation is used to detect specific enteric viruses, water filtration is not necessary and, consequently the loss of virus by membrane clogging is avoided. Therefore, the concentration by ultracentrifugation provides a more realistic picture of the viral community structure than the flocculation method. Although it is not practical to ultracentrifuge large volumes of water, it is possible to reduce the final volume of concentrated samples to a few microlitres and, consequently, efficiently concentrate virus in environmental waters. In this study, water samples of 0.5–1.0 L were concentrated 1000–2000 times in a volume of 200 mL, while by flocculation water samples of 10 L were concentrated 100–1000 times to a final volume of 10 mL. The large amount of flocs formed during the precipitation with the skim milk prevents reduction of the concentrated water volume of the flocculated samples to less than 10 mL. Moreover, techniques of nucleic acid amplification are now the most common way for viral detection Chapter 2 63 in water since they are rapid and more sensitive than traditional cell culture methods (Hovi et al, 2007; Pinto et al, 2007; Rodriguez-Diaz et al, 2009). Consequently, a small volume of water is sufficient to detect a specific virus. In fact, in this study enteric viruses were detected by PCR in recreational waters at sites where, in previous studies, their presence had been undetectable by cell culture and immunological methods (Alcântara and Almeida, 1995). The results show that both flocculation and ultracentrifugation methods are adequate to concentrate water samples for detection and quantification of enteric viruses in environmental waters. rotavirus A and adenovirus were present in all samples concentrated by either of the two concentration methods and the number of Adenovirus in recreational waters, quantified by qPCR, was similar for both concentration methods in the three independent assays of each of the two sampling dates. The estimation of the recovery efficiency based on the enumeration of VLP by epifluorescence was similar to that achieved in other studies based on the addition of specific enteric viruses to water samples and further quantification by plaque assay (Guttman-Bass and Nassen, 1984; Shields and Farrah, 1986; Haramoto et al, 2005; Kovac et al, 2009) or qPCR (Calgua et al, 2008; Muscillo et al, 2008; Hamza et al, 2009; Kovac et al, 2009). The enumeration of VLP by epifluorescence is, however, a simpler, faster and cheaper approach relative to the traditional plaque assay and qPCR approaches. Moreover, using the VLP, it is possible to evaluate the concentration of the whole viral community and not of a specific virus, as it happens when the viruses are added to the samples and detected by plaque assay and PCR techniques. In contrast to the plaque assay enumeration, using VLP counts it is considered the infective and non-infective viruses. However, since the techniques of nucleic acid amplification are now the most common way for viral detection/quantification in environmental waters (Tsai et al, 1993; Villena et al, 2003; Hovi et al, 2007; Pinto et al, 2007) and these techniques amplify nucleic acids of both infective and non-infective viruses, the recovery efficiency based on the enumeration of VLP is not an inconvenience, reflecting even better the efficacy of the concentration methods to recover viruses in environmental waters. Consequently, the determination of VLP is a good alternative to determine the recovery efficiency of the viral concentration methods. Chapter 2 64 Acknowledgements The authors would like to thank the University of Aveiro and Centre for Environmental and Marine Studies (CESAM) for funding the study. Financial support to C. Prata was provided by the Portuguese Foundation for Science and Technology (FCT) in the form of a PhD grant (SFRH/BD/33390/2008), São José Nascimento of University of Porto and Albert Bosh of University of Barcelona, for providing viral stocks used in this work as positive controls. 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Chapter 3 74 Introduction Health relevant enteric viral groups are considered to be waterborne pathogens (Noble and Fuhrman, 1998; Sedmak et al, 2005; Albinana -Gimenez et al, 2006; Myrmel et al, 2006), increasing the concern over the discharge of human enteric viruses into not only freshwater but also into estuarine and marine environments (Lee and Kim, 2002; Hamza et al., 2009; Rodríguez-Díaz et al, 2009). This group of viruses is stable in water and can exist naturally in the aquatic environment or could be transported to this environment through sewage outfall and vessel wastewater discharge (Grabow 1996, Pianetti et al. 2000, Griffin et al. 2003, Suttle 2005, Bosh et al. 2006, Suttle 2007, Lugoli et al. 2009). Although viruses can survive in water, as they are obligate intracellular parasites they cannot multiply in the environment (Wyn-Jones and Sellwood 2001). However, they can maintain infectivity, even after long periods in the environment, surviving for more than 130 days in seawater at temperatures between 20 and 30ºC (Rzezutka & Cook, 2004), increasing the probability of human exposure by recreational contact (Fong and Lipp, 2005). Although it is well known that viruses are of particular concern to human health, scarce information about their natural reservoirs, such as surface microlayer (SML) is available. Despite some studies on infectivity, survival, abundance, diversity and interactions of viruses with their hosts in the SML (Weinbauer 2004), showing that these are as abundant as in the underwater (UW) (Parada et al, 2005), until now, there is no study on human pathogenic virus in the SML. The sea SML is generally defined as the uppermost millimeter of the ocean (Liss and Duce 1997) and corresponds to the interface between the surface of the water and the air, forming a physically stable environment, but climatically variable. Water column processes regulate the accumulation of material in the SML and so, small molecules and larger particles can accumulate to form a film that extends into the UW (Cunliffe et al, 2011). The location of the SML makes it a highly dynamic system (Cunliffe et al, 2011) and in comparison with the UW, SML is physically more stable due to surface tension (Hardy, 1982) and the presence of a biogenic gelatinous film layer, constituting a microbial habitat where compounds and particles can be concentrated (Cunliffe et al, 2011). Therefore, the SML is generally enriched in organic materials, which might stimulate biological growth. However, it has been shown that organism density, activity and diversity in the SML can be higher, similar or lower that in UW. As SML interact with both the atmosphere and the hydrosphere simultaneously (Cunliffe et al, 2011), this layer may play an important role in microbial concentration and transmission across the air-water interface, acting as a reservoir for microorganisms, such as enteric viruses, in the aquatic environment. Chapter 3 75 N1 I6 The aim of this study was to evaluate the role of SML as a source of three groups of enteric viruses (rotavirus, enterovirus and HAV) in the aquatic environment. Material and methods Study area and sample collection Ria de Aveiro is a branched estuarine ecosystem, located in the northwest coast of Portugal, connected to the Atlantic by a narrow opening and separated from the sea by a sand bar. The system covers an area of 66 to 83 km 2 at low and high tide, respectively (Silva, 1994; Dias et al, 2000). Water samples were collected in a marine zone (MZ) (N1), in Canal de Navegação and in a brackish water zone (BWZ) (I6), in Canal de Ilhavo, both in Ria de Aveiro (Figure 3.1) in four dates of 2010 (two in the cold season (January and February) and two in the warm season (June and July)). In each sampling date, three independent samples were collected at the SML and UW. SML samples were collected with plexiglass and glass plates (0.25 m wide, 0.35 m long, and 4 mm thick). Prior to sample collection, the plates were rinsed with ethanol and sterile distilled water. The plates were introduced vertically through the SML and withdrawn in the same position. The excess of water was allowed to drain for about 5 s. Approximately 5 mL of water were collected each time the plate was introduced into the SML. The water adhering to the plate was subsequently removed from both sides with a wiper blade system (Harvey and Burzell 1972). The estimated thickness of the collected SML, determined from the volume of collected sample and the area of both sides of the plate, was approximately 60 μm, which is in the range reported by Harvey and Burzell (1972) for glass plate samplers. For SML samples, 2.5 L were collected. Figure 3.1: The estuarine system Ria de Aveiro with an indication of the sampling stations. Station N1 in Canal de Navegação represents the marine zone, and statio n I6, in Canal de llhavo, represents the brackish water zone. Chapter 3 76 For UW samples, 5 L were collected by submerging a sterilized glass bottle 15 cm in the water column. Three samples of SML and of UW were collected in each date. Water samples were maintained at 4 °C until analysis. Physical and chemical variables Phisical parameters were only determined for UW, because the collection for the SML is a labourious process during which changes in water temperature and salinity may occur. Water temperature and salinity were determined in the field with a WTW (Wissenschaftlich TechnischeWerkst¨atten, Germany) Cond330i/SET. The concentration of suspended solids (seston) was performed after filtration of aliquots of 500 mL for UW and of 150 mL for SML through Whatman GF/F (47 mm diam.) preweighted, pre-combusted filters. The filters were dried at 60°C for 24 h and seston content was calculated as the increase in weight (Parsons et al. 1989). For each nutrient, three replicates of 5 mL were filtered through GF/F Whatman filters (47 mm diameter) and frozen for later analysis. Ammonium (NH 4+ ) was quantified following the indophenol blue method (Dias et al, 2000). Nitrates (including nitrites) were determined by the sulfanilic acid method after reduction of nitrates to nitrites in a cadmium column (Lorezen, 1967). Phosphates were determined by the molibdate blue method (Koroleff, 1970). Chlorophyll a was determined espectrophotometrically after extraction by 90% acetone (Yentsch & Menzel 1963). Three replicates of 500 mL and three replicates of 150 mL were filtered for UW and SML, respectively, with GF/F Whatman filters. The filters were stored at -20ºC, until analysis, with 10 mL acetone 90%. A standard curve was determined, at an emission wavelength of 660 nm and 440 nm for excitation wavelength. Water samples concentration Water samples were concentrated, using the ultracentrifugation method, as described by Prata et al (2012). Samples were centrifuged (Beckman Optima TM , LE-80K Ultracentrifuge, rotor 50.2 Ti) at 100000 g for 1 hour at 20°C and the pellet was ressuspended in 200 µl of 1x PBS. Nucleic acids extraction Nucleic acids were extracted from concentrated water samples from SML and UW, using the geneMAG-RNA/DNA kit, a magnetic RNA/DNA purification kit (Chemicell ™), according to the instructions of the manufacturer. cDNA synthesis was made using a NZY Chapter 3 77 First-Strand cDNA Synthesis Kit (Nzytech), using the forward primers described below in the PCR technique. The nucleic acids were stored at -80°C until analysis. Enteric virus detection and quantification All viral groups were detected using a TPersonal thermocycler (Biometra) for PCR technique. The amplification products were separated by electrophoresis in a 2% agarose gel stained with ethidium bromide and detected with UV transilumination. The quantification was made, using a stepone plus 46 weel thermocycler (Applied Biosystems). Standards were obtained from a serial dilution of a suspension of each viral group, with a known initial number of copies, obtained by nucleic acid quantification with nanodrop. Negative controls were made, by using sterilized miliQ-water, instead of nucleic acids. Detection of rotavirus Detection of rotavirus was made by a PCR technique using the primers described by Villena et al, 2003. Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (VP6-3 5’: GCT TTA AAA CGA AGT CTT CAA C: 3’ and VP6-4 5’: GGT AAA TTA CCA ATT CTT CCA G: 3’). Rotavirus primers position is found between the positions 187 and 166 of human strain Wa (accession number K02086), creating a fragment of 186-bp. The amplification was carried out for 40 cycles of 94ºC for 10 s, 50ºC for 30 s and 72ºC for 20 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. Detection of enterovirus Detection of enterovirus was made by a PCR technique using the primers described by by Beld et al, 2004. Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (Entero 1 5’- CCC TGA ATG CGG CTA AT -3’ and Entero 2 5’- ATT GTC ACC ATA AGC AGC CA - 3’). Enterovirus primers sequences used for amplification are located in the conserved 5’ noncoding region of the EV sequence. Nucleotide positions 452 to 468 for Entero-1 and 579 to 597 for Entero-2, creating a fragment of 172-bp. Chapter 3 78 The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Detection of HAV Detection of HAV was made by a PCR technique using the primers described by Tsai et al, 1993. Five microliters of sample were added to 20 µl of the reaction mixture used which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (HAVC-R 5’- CTC CAG AAT CAT CTC CAA C - 3’ and HAVC-L 5’- CAG CAC ATC AGA AAG GTG AG -3’). The primers for HAV capsid amplify a 192-bp. The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Quantification of rotavirus The quantification of rotavirus was made by Real time-PCR using the primers described by Villena et al (2003). Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2x iQ ® SYBR ® Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6mM MgCl 2 , SYBR Green I, fluorescein and stabilizers) and 0.48 µM of each primer (StabVida) (VP6-3 5’: GCT TTA AAA CGA AGT CTT CAA C: 3’ and VP6-4 5’: GGT AAA TTA CCA ATT CTT CCA G: 3’). Rotavirus primers positions is found between the position 187 and position 166 of human strain Wa [accession number K02086]). The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. Quantification of enterovirus The quantification of enterovirus was made by Real time-PCR using the primers described by Beld et al (2004). Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2x iQ ® SYBR ® Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6mM MgCl 2 , SYBR Green I, fluorescein and stabilizers) and 0.48 µM of each primer (StabVida) (Entero 1 5’- CCC TGA ATG CGG CTA AT -3’ and Entero 2 5’- ATT GTC ACC ATA AGC AGC CA - 3’). The sequences of the primers used for Chapter 3 79 amplification are located in the conserved 5’ noncoding region of the EV sequence at the nucleotide positions 452 to 468 for Entero-1 and 579 to 597 for Entero-2. The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Quantification of HAV The quantification of HAV was made by Real time-PCR using the primers described by Tsai et al (1993). Five microliters of sample were added to 20 µl of the reaction mixture consisting of 2x iQ ® SYBR ® Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6mM MgCl 2 , SYBR Green I, fluorescein and stabilizers) and 0.48 µM of each primer (StabVida) (HAVC-R 5’- CTC CAG AAT CAT CTC CAA C - 3’ and HAVC-L 5’- CAG CAC ATC AGA AAG GTG AG -3’). The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. Statistical analysis The statistical analysis of microbiological data was performed using the SPSS21 software. To explain the variation of viral abundance in water, stepwise multiple regression analysis was used. Water temperature, salinity, seston, nutrients and chlorophyll a were used as independent variables. Enrichment factors Enrichment factors were calculated dividing the value of viral abundance in SML by the value of viral abundance in UW. Results Physical and chemical parameters The physical and chemical data obtained both in field and in the laboratory, are presented in Table 3.1. Salinity ranged from 20.3 to 35, in the marine zone and from 0.82 to 33 in the brackish water zone, with the higher values in the warm months. The water temperature varied between 10.8 to 21.2°C in the marine zone and between 7.9 to 19.2°C in the brackish water zone, being the values higher in warm months (Table 3.1). Chapter 3 80 Table 3.1: Physical data in cold (January and February) and warm (June and July) months, in the marine and brackish water zone, in both SML and UW. Sampling date Salinity Water temperature (ºC) MZ BWZ MZ BWZ January 27.5 0.82 10.8 7.9 February 20.3 1.7 10.9 8.2 June 35 34 20.6 18.3 July 33 30 21.2 19.2 MZ – Marine zone; BWZ – Brackish water zone; SML – Surface microlayer; UW – Under water Table 3.2: Averaged values and standard deviation of the three samples and for seston and chlorophyll a data in cold (January and February) and warm (June and July) months, in the marine and brackish water zone, in both SML and UW. January February June July MZ SML 70.20± 0.33 49.20± 0.31 98.40± 0.80 66.70± 0.84 Seston (mg L -1 ) UW 44.50± 0.14 38.40± 0.31 54.44± 0.94 48.00± 0.60 BWZ SML 42.50± 0.33 48.89± 0.29 98.44±0. 16 71.00±0. 36 UW 37.00±0. 62 32.50±0. 40 67.00±0. 33 53.20±0. 84 Chlorophyl a (µg L -1 ) MZ SML 1.57±0.64 0.96±0.20 2.40±0.01 2.22±0.15 UW 1.30±0.10 0.50±0.24 1.70±0.11 1.60±0.18 BWZ SML 2.31±0.03 2.37±0.06 2.30±0.04 2.38±0.01 UW 1.00±0.08 1.85±0.09 1.99±0.11 2.16±0.06 MZ – Marine zone; BWZ – Brackish water zone; SML – Surface microlayer; UW – Under water In Table 3.2 are presented the data obtained for seston and chlorophyll a. Seston values showed a seasonal pattern, with higher concentration in warm months. Seston concentration in the marine zone presented values ranging from 38.40 to 98.40 mg L -1 and ranging from 32.50 to 98.44 mg L -1 in the brackish water zone. There is not a wellestablished pattern between the two areas studied, but in both SML has higher seston concentration than UW, with enrichment factors varying from 1.15 to 1.81. The Chlorophyll a results presented in Table 3.2 show a clear seasonal pattern, with higher values in warmer months, ranging from 0.50 µg L -1 to 2.37 µg L -1 in cold months Chapter 3 81 and from 1.60 µg L -1 to 2.40 µg L -1 in warmer months It was observed a higher concentration of chlorophyll a in the brackish water zone, than in the marine zone. Chlorophyll a values were higher in the SML, than in UW, with an enrichment factor of 1.40 in the marine zone and of 1.34 in the brackish water zone. Nutrients The nutrients concentration is showed in Table 3.3. The values were higher in cold months. Amonium concentration ranged from 1.52 to 7.12 µM, nitrates from 2.32 to 17.21 µM, nitrites from 0.15 to 4.90 µM and phosphates from 0.18 to 1.89 µM (Table 3.3). The nutrients concentration was higher in the SML compared with the UW, with a medium enrichment factor of 1.30 and 1.09 for amonium, of 1.64 and 1.34 for nitrates, of 1.33 and 1.92 for nitrites and of 1.66 and 1.42 for phosphates, in marine and brackish water zone, respectively. Table 3.3: Nutrients data in cold (January and February) and warm (June and July) months, in the marine and brackish water zone, in both SML and UW. Sampling Day Ammonium (µM) Nitrates/Nitrites (µM) Phosphate (µM) MZ BWZ MZ BWZ MZ BWZ SML UW SML UW SML UW SML UW SML UW SML UW January 2.44 2.22 3.23 3.03 6.92 4.9 17.79 12.05 1.49 1.32 0.23 0.18 February 2.28 2.09 4.83 4.83 14.01 7.08 16.71 10.96 0.76 0.68 1.89 0.46 June 2.62 1.99 7.12 6.32 3.35 2.46 4.88 4.46 0.40 0.37 1.32 0.78 July 2.83 1.52 6.83 6.15 2.86 2.76 4.38 4.06 0.82 0.55 0.61 0.57 MZ – Marine zone; BWZ – Brackish water zone; SML – Surface microlayer; UW – Under water Enteric viruses The three viral groups were found in all water samples and Figure 3.2 presents the results of viral abundance for the three groups of enteric virus tested. Rotavirus is the most abundant group (range between 10 8 – 10 10 copies L -1 in cold months and between 10 1 – 10 2 copies L -1 in warm months), followed by enterovirus (range between 10 7 copies L -1 in cold months and 10 1 copies L -1 in warm months). HAV was the less abundant group (range between 10 1 – 10 2 copies L -1 all year). Chapter 3 82 All viral groups presented higher concentration in cold months, except for HAV which was almost constant along the year. Viral concentration was higher in the SML, for the three viral groups tested, with enrichment factors varying from 1.01 to 6.37 in the marine zone and from 1.12 to 15.23 in the brackish water zone for rotavirus, from 1.01 to 2.80 in the marine zone and from 1.03 to 5.05 in the brackish water zone for enterovirus and from 1.02 to 2.15 in the marine zone and from 1.17 to 1.44 in the brackish water zone for HAV. There is not clear pattern for the enrichment factors (Table 3.4). Enrichment factors may be underestimated, because SML, as a reservoir for organic matter and other PCR inhibitors, may have greater abundance than it was observed. Figure 3.2: Enteric viruses abundance, for the four sampling dates (two in cold months: January and Februry and two in warm months: June and July), in the marine zone (at left) and brackish water zone (at right), for both SML and UW. Values represent the mean of three replicates and error bars represent standard deviation. Chapter 3 83 Table 3.4: Enrichment factors for rotavirus, enterovirus and HAV in four sampling dates, in the marine and brackish water zones of Ria de Aveiro. SML/UW Sampling Date r otavirus e nterovirus HAV MZ BWZ MZ BWZ MZ BWZ January 6.37 1.63 1.01 1.03 1.02 1.31 February 1.01 6.09 1.10 1.08 2.15 1.44 June 2.77 1.53 2.74 5.05 1.86 1.17 July 1.18 1.12 2.80 1.59 1.40 1.23 MZ – Marine zone; BWZ – Brackish water zone The results of the stepwise multiple regressions analysis are presented in Table 3.5. None of the independent variables explained the variation of viral abundance for HAV, in both marine and brackish water zones and for rotavirus in the brackish water zone. Water temperature explained almost 100% of viral abndance variation for enterovirus in the brackish water zone and for enterovirus in the marine zone. Nitrites concentration explained almost 100% of viral abundance variation for Rotavirus in the marine water zone. Table 3.5: Regression equations for viral abundance obtained from stepwise multiple-regression analysis, for marine and brackish water zones. (Dependent variables: enteric virus abundance; independent variables: physical and chemical parameters (salinity, water temperature, DOC, seston, chlorophyll a and nutrients)). Dependent variable Independent variables Regression equation Adj. R 2 rotavirus MZ (β= 0.998; p=0.002) Nitrites Rot= 1.34x10 7 x – 1.86x10 6 0.993 BWZ Not explained by any independent variable enterovirus MZ (β= -0.0998; p= 0.002) Water temperature Ent=3.68x10 6 x+7.71x10 7 0.993 BWZ (β= -0.998; p= 0.002) Water temperature Ent=-3.27x10 6 x+6.14x10 7 0.995 HAV MZ Not explained by any independent variable BWZ Not explained by any independent variable MZ – Marine zone; BWZ – Brackish water zone Chapter 3 90 Chapter 4 - Marine aerosols as a source of health risk enteric virus Chapter 4 93 Marine aerosols as a source of health risk enteric virus 1 Catarina Prata, 1 Ângela cunha, 1 Newton Gomes, 2 Teresa Nunes, 1 Adelaide Almeida* 1 Department of Biology and CESAM, University of Aveiro, Campus Universitário de Santiago, 3810-193 Aveiro, Portugal 2 Department of Environment and CESAM, University of Aveiro, Campus Universitário de Santiago, 3810-193 Aveiro, Portugal *Corresponding author: E-mail: [email protected], Tel.: +351234370350, Fax: +351234372587 ABSTRACT Viruses are nowadays considered to be emerging pathogens in the marine environment. Due to their small infectious dose, it is of extreme importance to evaluate if they can be transferred from water to air through aerosol formation. The aim of this study was to evaluate the role of aerosols in the transference of health relevant viruses from marine waters to air. It was developed a method to create and recover aerosols, using Virus-like particles (VLP) to access the efficiency of the method. It was used a polycarbonate membrane (0.2 µm pore) to remove virus from the air used do generate aerosols, with an efficiency of almost 80%. The abundance of three enteric virus groups (enterovirus, rotavirus and HAV) was quantified in underwater (UW), surface microlayer (SML) and in aerosols simulated by bubbling in marine and brackish waters zones of Ria de Aveiro (Portugal). The three enteric viruses abundances were higher in SML relatively to the UW and all of them were detected in aerosols, although at concentrations significantly lower than those found in the SML. The data indicate that SML is an important source of viruses entering the atmosphere trough aerosols formation. INTRODUTION Some health relevant enteric viral groups are nowadays considered to be emerging marine waterborne pathogens (La Rosa et al, 2012; Woods, 2013), since they are able to adapt not only by mutation but also through recombination and reassortment and can thus become able to infect new hosts and to adapt to new environments (La Rosa et al, 2012). Enteric viruses can be transmitted to humans through marine waters used for recreational purpose or from waters used to shellfish growing, but also through aerosols transmition from the sea surface microlayer (SML). The sea SML is generally defined as the uppermost millimeter of the ocean (Liss and Duce 1997). Consequently, the primary interest in the Chapter 4 94 structure and function of the SML is based on its crucial role in exchange processes of gases and matter across the air-water interface (Agougué et al, 2004; Cunliffe et al, 2011; Donaldson and George, 2011). Marine aerosols are formed primarily by the eruption of rising bubbles through the SML (Aller et al, 2005) and constitute the largest fraction of atmospheric aerosol loading over the remote open oceans, contributing to regional air quality (Brooks et al, 2011) and through sea-to-air transfer of bacteria and viruses (Mathias-Maser and Jaenicke, 1994; Posfai et al, 2003; Brooks et al, 2011), transporting microbes for long distances (Moorthy et al, 1998; Klassen and Roberge, 1999; Chow et al, 2000; Gutafsson and Franzen, 2000; Grammatika and Zimmerman, 2001). Aerosols may remain suspended in the atmosphere for weeks (Dueñas et al, 2004) and it is known that many microorganisms, as well as viruses, can remain infectious outside their hosts for prolonged periods of time (Verrault et al, 2008). Despite the lack of information for viral survival in aerosols, there are evidences that at least some marine bacteria remain viable after overland transport in aerosols (200 m inland, collected at 30 m above sea level) (Marks et al, 2001), despite exposure to ultraviolet radiation, changes in temperature and salinity and other immediate stresses. Since some health relevant viruses are waterborne emerging pathogens in the marine environment and their infectious doses are small, it is of extreme importance to understand if it is possible that these pathogens pass from water to air, and if they can be disperse inside the aerosols (Bosch et al, 2011). Studies on the aerobiology of infectious diseases have been rather limited (Roy and Milton, 2004), mainly due to the difficulty in collecting and analyzing airborne biological contaminants, which is an even greater problem for viruses (Verrault et al, 2008; Danovaro et al, 2011). The aim of this study was to evaluate the role of aerosols in the transference of enteric virus from marine waters to air. For this, the abundance of three enteric viruses (enterovirus, rotavirus and HAV) was quantified in UW, SML and in aerosols, simulated by bubbling water, in two different locations (marine zone and brackish water zone) of Ria de Aveiro (Portugal). The abundance of virus-like particles (VLP) was also determined in order to develop the aerosol collection protocol, to decide the collection medium, the sampling period and the membrane to filtrate the air to be used to form the aerosols. The VLP was also determined in the UW, SML and in the simulated aerosols. Chapter 4 95 Material and Methods Development of aerosols collection protocol in laboratory Aerosols collection medium Destilated miliQ water was tested as an aerosols collection medium (Yu et al, 2010), The air was collected in 10 mL of distelated water, by impingement, for 5 hours with a flow rate of 2 L min -1 . The collection medium was tested using the bundance of VLP. Three samples were prepared. Determination of membrane efficiency to remove viruses from the bubbling air A 0.2 µm pore polycarbonate membrane was tested to remove the viruses from the bubbling air used to produce the aerosols. Ten milliliters of collection medium were placed in a glass chamber and the filtered air was used to bubble the water, at a controlled flow rate of 2 L/min, for 5 hours. The same procedure was done in the absence of the membrane (control). Three samples were processed with polycarbonate membranes and three without membrane. The polycarbonate membrane efficiency was determined using the abundance of VLP for all samples with and without membrane. Sampling time for aerosols formation in laboratory The aerosols formation was tested using 10 mL of water sample collected in a brackish zone of Ria de Aveiro (Station I6, located in Canal de Ílhavo, see information below), in three diferrent dates in January 2012, by submerging a glass bottle 15 cm in the water column. Sampling times tested were 30 minutes and 1 hour of air filtration, in order to determine the minimal time needed to form aerosols containing an enough amount of VLP. The aerosols sampler used was an all glass impinge with the nozzle positioned 10 mm above the bottom and suction was applied to the small side arm. Ten milliliters of water sample were placed in a glass chamber of the impinge and 10 mL of miliQ water were used to recover the aerosols formed by bubbling filtered air into the water sample, at a controlled flow rate of 2 L min -1 . For each sampling time, three samples and a negative control of the air, consisting in filtered air bubbling directly into the miliQ water, were used. The VLP was determined in the samples and in the control by epifluorescence microscopy. Concentration of aerosols samples After aerosols formation and recovery, samples were concentrated, using the ultracentrifugation method, as described by Prata et al (2012). Briefly, samples were centrifuged (Beckman Optima TM , LE-80K Ultracentrifuge, rotor 50.2 Ti) at 100,000 x g for 1 Chapter 4 96 N1 I6 hour at 20°C and the pellet was ressuspended in 200 µl of 1x PBS, which was used to count the VLP by epifluorescence microscopy. Determination of VLP in aerosols samples VLP abundance was determined using a modified epifluorescence method described by Noble and Fhurman (1998). Water samples were filtered by a 0.2 µm polycarbonate membrane and then by a 0.02 µm Al2O3 Anodisc, which were than stained in the dark for 10 minutes with SYBR gold dye (0.25%). VLP were enumerated in an epifluorescence up-right microscope Imager.Z1 (Zeiss) with 38 HE GFP filter. For each sample two slides were made and a minimum of 200 particles per slide were counted. Three independent assays were done for each sampling date. Field work Study site Ria de Aveiro is a branched estuarine ecosystem, located in the northwest coast of Portugal, connected to the Atlantic by a narrow opening and separate from the sea by a sand bar. The Ria covers an area of 66 to 83 km 2 at low and high tide, respectively (Silva, 1994; Dias et al, 2000). Three samples of SML, UW and aerosols were collected in two locations: marine zone (Station N1) and brackish water zone (Station I6) in Canal de Ílhavo (Figure 4.1), both in Ria de Aveiro, three times, between July and August of 2012. Figure 4.1: The estuarine system Ria de Aveiro with an indication of the sampling stations. Station N1 in Canal de Navegação represents the marine zone, and station I6, in Canal de llhavo, represents the brackish water zone. Chapter 4 97 In situ generation and collection of aerosols Aerosols were generated in situ using a small floating device. According to the results of obatained in the laboratory experiments, a 0.2 µm pore polycarbonate membrane was used to remove the viruses from the bubbling air used to produce the aerosols and the bubbling was done at 2 L min -1 for 1 hour. The aerosols formed were collected in 10 mL of miliQ water, with two replicates per sample in each date. An air control sample was also included for each sample. Three independent samples were collected for each sampling date. SML and UW sampling Samples of 0.5 L of SML were collect with plexiglass and glass plates (0.25 m wide, 0.35 m long and 4 mm thick). Prior to sample collection, the plates were rinsed with ethanol and sterile distilled water. The plates were introduced vertically through the SML and withdrawn in the same position. Excess water was allowed to drain for about 5 s. Approximately 5 mL of water were collected each time the plates were introduced into the SML. The water adhering to the plate was subsequently removed from both sides of the plate with a wiper blade system (Harvey and Burzell 1972). The estimated thickness of the collected SML, determined from the volume of collected sample and the area of both sides of the plate, was approximately 60 μm, which is in the range reported by Harvey and Burzell (1972) for glass plate samplers. Samples of 0.5 L of subsurface water were collected by submerging a glass bottle 15 cm in the water column. Three independent samples were collected for each sampling date. Samples were maintained at 4 °C until analysis. Concentration of water and aerosols samples Aerosols samples were concentrated, using the ultracentrifugation method described above in Concentration of aerosols samples. Abundance of VLP in water and aerosols samples VLP abundance was determined using the modified epifluorescence method described above in the development of aerosols collection protocol in laboratory. Nucleic acids extraction Nucleic acids were extracted from concentrated water and aerosol samples, using the geneMAG-RNA/DNA kit, a magnetic RNA/DNA purification kit (Chemicell ™), according to the instructions of the manufacturer. cDNA synthesis was made using a NZY First-Strand cDNA Synthesis Kit (Nzytech), using the forward primers described below in the PCR technique. The nucleic acids were stored at -80°C until analysis. Chapter 4 98 Enteric virus detection and quantification All viral groups were detected using a TPersonal thermocycler (Biometra) for PCR technique. The amplification products were separated by electrophoresis in a 2% agarose gel stained with ethidium bromide and detected with UV transilumination. Quantification was made using a stepone plus 46 weel thermocycler (Applied Biosystems) and standards were obtained from a serial dilution of a suspension of each viral group, with a known initial number of copies. Negative controls were made, by using sterilized miliQ-water, instead of nucleic acids. Detection of rotavirus Detection of rotavirus was made by a PCR technique using the primers described by Villena et al, 2003. Five microliters of sample were added to 20 µl of the reaction mixture which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (VP6-3 5’: GCT TTA AAA CGA AGT CTT CAA C: 3’ and VP6-4 5’: GGT AAA TTA CCA ATT CTT CCA G: 3’). Rotavirus primers position is found between the position 187 and position 166 of human strain Wa [accession number K02086]), creating a fragment of 186-bp. The amplification was carried out for 40 cycles of 94ºC for 10 s, 50ºC for 30 s and 72ºC for 20 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. Detection of enterovirus Detection of enterovirus was made by a PCR technique using the primers described by by Beld et al, 2004. Five microliters of sample were added to the reaction mixture which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (Entero 1 5’- CCC TGA ATG CGG CTA AT -3’ and Entero 2 5’- ATT GTC ACC ATA AGC AGC CA - 3’). Enterovirus primers sequences used for amplification are located in the conserved 5’ noncoding region of the EV sequence. Nucleotide positions 452 to 468 for Entero-1 and 579 to 597 for Entero-2, creating a fragment of 172-bp. The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Chapter 4 99 Detection of HAV Detection of HAV was made by a PCR technique using the primers described by Tsai et al, 1993. Five microliters of sample were added to the reaction mixture which consisted of 2 mM Buffer taq, 3 mM MgCl2, 0.08 mM of each dNTP, 0.1 U/µL of Taq polymerase (Fermentas) and 0.48 mM of each primer (StabVida) (HAVC-R 5’- CTC CAG AAT CAT CTC CAA C - 3’ and HAVC-L 5’- CAG CAC ATC AGA AAG GTG AG -3’). The primers for HAV capsid amplify a 192-bp. The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Quantification of Rotavirus Quantification of Rotavirus was made by Real time-PCR using the primers described by Villena et al (2003). Five microliters of sample were added to the reaction mixture used which consisted of 2x iQ ® SYBR ® Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6mM MgCl 2 , SYBR Green I, fluorescein and stabilizers) and 0.48 µM of each primer (StabVida) (VP6-3 5’: GCT TTA AAA CGA AGT CTT CAA C: 3’ and VP6-4 5’: GGT AAA TTA CCA ATT CTT CCA G: 3’). Rotavirus primers positions is found between the position 187 and position 166 of human strain Wa [accession number K02086]), creating a product of 186-bp. The amplification was carried out for 40 cycles of 94ºC for 10 s, 50ºC for 30 s and 72ºC for 20 s after initial denaturation at 95ºC for 9 minutes. A final extension step was performed at 72ºC for 7 min. Quantification of Enterovirus Quantification of enterovirus was made by Real time-PCR using the primers described by Beld et al (2004). Five microliters of sample were added to the reaction mixture which consisted of 2x iQ ® SYBR ® Green Super mix (2x reaction buffer with dNTPs, iTaq, DNA polymerase, 6mM MgCl 2 , SYBR Green I, fluorescein and stabilizers) and 0.48 µM of each primer (StabVida) (Entero 1 5’- CCC TGA ATG CGG CTA AT -3’ and Entero 2 5’- ATT GTC ACC ATA AGC AGC CA - 3’). The amplification was carried out for 40 cycles of 95ºC for 10 s, 55ºC for 20 s and 72ºC for 30 s after initial denaturation at 95ºC for 10 minutes. A final extension step was performed at 72ºC for 5 min. Chapter 4 106 • Danovaro R, Corinaldesi C, Dell’Anno A, Fuhrman J. 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Chapter 5 - Influence of UV light exposure on viral abundance in aquatic system Chapter 5 111 Influence of UV light exposure on viral abundance in aquatic system Catarina Prata, Francisco Coelho, Bruno Ladeiro, Ângela Cunha, Newton Gomes, Adelaide Almeida * Department of Biology and CESAM, University of Aveiro, Campus Universitário de Santiago, 3810193 Aveiro, Portugal *Corresponding author: E-mail: [email protected], Tel.: +351234370350, Fax: +351234372587 ABSTRACT Nowadays, one of the major issues related with global climate changes is the increase to ultraviolet light (UV) exposure of aquatic organisms. Over the next decades it is expected an increase in the exposure to damaging UV wavelengths by all aquatic organisms with far-reaching ecological consequences. The aim of this work was to evaluate the influence of increased exposure to UV light in the abundance of two groups of RNA enteric viruses (rotavirus and enterovirus) present in the SML of marine systems.To reach this objective an experimental life support system (ELSS), composed by several sediment-water microcosms, was used. The ELSS mimics fundamental aspects of biological activity in marine ecosystems, namely photoperiod, light intensity and tidal cycles and allows the control of ultraviolet radiation (UVR), temperature, salinity and pH. After 57 days of incubation, surface microlayer (SML) samples were collected in microcosms exposed to UVR and in non-exposed microcosms. The abundance of two enteric viruses, a doublestand RNA (rotavirus) and a single-stand RNA (enteroviruses) was determined in UVR exposed and non-exposed samples. No significant reduction in rotavirus abundance (p =0.41 >0.05) was found in exposed samples, reduction of less than 1 log, but for enterovirus a significant reduction (p = 0.03 < 0.05) in abundance, around 2 log, was observed. The results indicate that RNA enteric viruses present different resistance to UVR and that this difference can be related with RNA type but other viral characteristics can be also involved in UVR resistance of these viruses. As in this study a damaging UV wave-length with fluence similar to that reaching the surface of the planet was used, an increase in the exposure to UV wave-lengths, associated to global climate changes, would influence the viral communities, namely those more exposed to UVR, such as SML viral communities of marine environments, with extensive ecological consequences. Chapter 5 112 Introdution Climate change is increasingly recognized as a major risk to human health, with consequences occuring through direct and indirect routes and as a result of interactions with other changed environmental characteristics (Thomas et al, 2012). Interactions between climate change and stratospheric ozone will cause changes in the levels of ambient UVR in the future (Thomas et al, 2012). Ultraviolet light (UV) is electromagnetic radiation with wavelengths shorter than visible light that can induce damage in a variety of organisms. UV light can be divided into different ranges including UVA (400 to 320 nm), which is weakly affected by stratospheric ozone, UVB (320 to 280 nm), which is strongly affected by ozone and UVC (280 to 100 nm) or short-range UV. UVC is considered to be germicidal, because at this range, UV light is mutagenic to microorganisms like bacteria and also for viruses (Kowalski et al, 2000; Hirneisen et al, 2010; Zepp et al, 2011). UV light predominately damage the viral nucleic acid (Nuanualsuwan and Cliver 2003; Hirneisen et al, 2010), but at high enough doses (>1000 mWs/cm2) can also affect the capsid proteins, making the genome susceptible to DNases and RNases present in the environment (DeSena and Jarvis 1981; Smirnov and others 1983; Hirneisen et al, 2010). Enteric viruses that contain genomic RNA undergo successive morphological changes in the viral capsid when exposed to UV light (Katagiri et al 1967; Miller and Plagemann 1973; De Sena and Jarvis 1981; Hirneisen et al, 2010). Many factors can enhance or decrease UV action, such as the type of nucleic acid of the viruses, viral proteins, type of host cell, viral strain, virus aggregation and experimental conditions (Fino and Kniel 2008; Hirneisen et al, 2010). The effect of UV light on microorganisms depends on the UV dose applied, of its ability to protect itself from UV light and to repair damages (Sommer et al., 2001; Hu et al, 2011). Because UV radiation from the sun is present in the environment, natural defense mechanisms, such as photoreactivation and dark repair, have evolved in bacteria and other microorganisms, allowing UV injured microorganisms to reverse UV-induced damages (Koivunen and Heinonen-Tanski, 2005; Quek and Hu, 2008; Hu et al, 2011). Furthermore, viruses with double-stranded DNA or RNA are more stable when exposed to UV, than single-stranded DNA or RNA viruses, because their undamaged DNA or RNA strand may serve as a template for repair by host enzymes (Fong and Lipp, 2005; Hirneisen et al, 2010). In fact, double stranded DNA (e.g. adenovirus) or RNA viruses (rotavirus) have been frequently responsible for recreational waterborne disease outbreaks (Gerba et al. 2002; Guo et al, 2010; Di Bartolo et al, 2011). Chapter 5 113 Other factors, such as water temperature, pH, association with sediments, predators, concentration of particular matter, salinity and raining (Le-Guyader et al. 1983, Chuan et al. 1983, Goyal et al. 1984, Yates et al. 1985, Griffin et al. 2003, Bosch et al. 2005; Bosch 2007; Fong et al. 2005, Bosch et al. 2006, Suttle 2007, Lugoli et al. 2009), also affect virus survival in the marine environment. These factors may act individually, or they may interact with each other, affecting viral survival in different ways (Cutler et al, 2012). Environmental parameters such as temperature, precipitation, pH, salinity, and UV light irradiation have been changed as consequence of global climate changes (Kim et al, 2010). These factors potentially have important consequences for aquatic microorganisms including waterborne human pathogens (Häder et al, 2011). It is expected that over the next decades, the exposure of aquatic microorganisms to damaging UV wavelengths, particularly UV-B (280 to 320 nm) will increase with far-reaching ecological consequences (Andrady et al, 2010; Santos et al, 2012). The effects of UV-B (280–320 nm) on aquatic organisms depend on the dose of harmful radiation to which individuals are exposed, which in turn depends on the optical characteristics (i.e. UV-B transmittance) of the water body and organism positioning in the water column (Sommaruga 2003). In this work, the influence of exposure to UV light in the abundance of twou groups of RNA enteric viruses (rotavirus and entetovirus) present in SML of an estuarine system was evaluated. Material and methods Microcosm general description An ELSS, composed by several sediment-water microcosm (Figure 5.1), conceived by Coelho et al (2013) was used to evaluate UV light influence on viral concentration. The ELSS mimics fundamental aspects of biological activity in marine ecosystems, namely photoperiod, light intensity (including photosynthetically active radiation, PAR) and tidal cycles. Control of temperature, ultraviolet radiation (UVR), salinity and pH is also possible. The ELSS is divided into two frames of 16 microcosms (32 in total) (glass tanks 25 cm high, 28 cm length and 12.4 cm width, each with a maximum functional water volume of approximately 7 L) (Figure 5.1). Sediment was collected at the east margin of Mira channel (40°37′N, 8°44′W), one of the main channels of the Ria de Aveiro lagoon (Portugal), in May 2011. The Ria de Aveiro is a shallow mesotidal coastal lagoon connected with the Atlantic Ocean through a single inlet, and characterized by four main channels with several secondary narrow channels, inner basins and extensive intertidal areas (Dias et al., 2001). Plexiglass cores of Chapter 5 114 undisturbed sediment samples (10 cm deep, 27 cm length and 10.6 cm width) were collected and each core transferred directly into individual microcosms of the ELSS. Microcosms containing the sediment were taken back to the laboratory and connected to the ELSS less than 2 h after sampling. The ELSS was operated continuously during 57 days (Coelho et al., 2013). The ELSS system was programmed to simulate the specific characteristics of the estuarine system Ria de Aveiro, at the sampling site where and when sediment was collected. Salinity was adjusted to simulate the conditions recorded at the sampling location and kept constant (32.6 ± 1.5) during the experiment. The microcosms were exposed to a uniform semi-diurnal tidal regime, experiencing two high tides and two low tides daily. The minimum and maximum water levels above the sediment surface were approximately 5 cm (low tide) and 10 cm (high tide). During each tidal cycle about 50% of the water volume of each microcosm was exchanged (~1.5 l), using synthetic saltwater to compensate the water removed, thus simulating the water renewal percentage recorded for the sampling site. Water pH was adjusted to 8.0 and water temperature to 19 °C (Coelho et al., 2013). No water recirculation was employed in order to avoid cross contamination between experimental treatments and also laboratory artifacts that may be promoted by re-using the same water over time. Figure 5.1: Representation of experimental life support system (ELSS) used in the UVR experiments (Coelho et al, 2013). A – independent microcosm; B – saltwater reservoir; C – acidified saltwater reservoir; D – normal pH saltwater reservoir, E – water bath; F – refrigerator; G – lightning system (a vinyl frame can be included to isolate the light from the luminaires), H - pH control system. Chapter 5 115 UV light control system The ELSS is equipped with 4 Reef-SET ® , (Rees, Germany) programmable luminaire system for diurnal light cycle and controlled UV simulation. Each luminaire holds four UV fluorescent tubes (SolarRaptor, T5/54W, Rees, Germany) and four full spectra fluorescent tubes (AquaLight, T5/54W/10000K, Bramsche, Germany) disposed alternately under a reflector. The PAR value measured for the full spectrum fluorescent lamps is 260.50 ± 56.30 µmol photons m -2 s -1 and the UV-A (320–400 nm) and irradiance emitted by the UV lamps was 2875.91 ± 264.62 mW m -2 . To mimic summer photoperiod and light conditions at Portuguese latitudes during the experiment, a 14 h diurnal light cycle was simulated, with light intensity varying from 50 to 100% of the total fluorescent tube intensity. Since UV-A radiation is practically unaffected by changes in ozone depletion and plays an important role in biological systems, including photo-repair mechanisms (Bargagli, 2005), a similar amount of UV-A integrated irradiance was maintained constant among microcosms. The UV lamps were switched on for 4 hours a day at maximum intensity and UV-B component was filtered using a glass panel in the luminaire system. The UVR fluence used in the ELSS system corresponds to UV ligh that reach the surface of the planet in a summer day at the Portuguese latitudes. This UVB fluence is classified as a moderate intensity and is able of DNA damaging. SML sampling After the 57 days of incubation SML was collected using polycarbonate membranes, according to the method described by Cunliffe et al (2008). Briefly, a sterile polycarbonate membrane was placed directly onto the water surface, recovered with a tong and placed into a sterilized petri dish. SML samples were also collected in microcosms not exposed to UV light but exposed to PAR light during the 14 h diurnal light cycle. Three samples of each of three microcosms were collected and treated. Nucleic acids extraction Microorganisms were removed from polycarbonate membranes by using a mix of Zicornia and glass beads (0.10 g of 0.1 mm Zicornia beads, 0.20 g of 0.25-0.5 mm glass beads, 0.20 g of 0.75-1.0 mm glass beads and 2 glass beads of 2.85-3.45 mm). Polycarbonate membrane was cut and the beads were added in a 1.5 ml tube. One milliliter of lises buffer from the kit geneMAG-RNA/DNA kit, a magnetic RNA/DNA purification kit (Chemicell ™) was added to the tube which was centrifuged in Fastprep, at maximum velocity (6) for 40 s. Supernatant was removed to a clean tube and centrifuged for 1 Chapter 5 122 • Guo H, Chu X, Hu J, 2010, Effect of Host Cells on Lowand Medium-Pressure UV Inactivation of Adenoviruses, Applied and Environmental Microbiology, 76 (21) 7068– 7075. • Häder D-P, Helbling E W, Williamson C E, Worrest R C, 2011, Effects of UV radiation on aquatic ecosystems and interactions with climate change, Photochemical and Photobiological Science, 10, 242-260 • Hirneisen K A, Black E P, Cascarino J L, Fino V R, Hoover D G, Kniel K E, 2010, Viral Inactivation in Foods: A Review of Traditional and Novel Food-Processing Technologies, Comprehensive reviews in food science and food safety, 9, 3-20. • Hu X, Geng S, Wang X, Hu C, 2011, Inactivation and Photorepair of Enteric Pathogenic Microorganisms with Ultraviolet Irradiation, Environmental Engineering Science, 29 (6) 549-553. • Katagiri S, Hinuma Y, Ishida N, 1967, Biophysical properties of poliovirus particles irradiated with ultraviolet light, Virology, 32, 337–43. • Kim J, Park J, Kim P-G, Lee C, Kyungho K C, 2010, Implication of global environmental changes on chemical toxicityeffect of water temperature, pH, and ultraviolet B irradiation on acute toxicity of several pharmaceuticals in Daphnia magna, Choi. Ecotoxicology, 19:662–669 • Koivunen J, Heinonen-Tanski H, 2005, Inactivation of enteric microorganisms with chemical disinfectants, UV irradiation and combined chemical/UV treatments, Water Research, 39, 1519. • Kowalski W J, Bahnfleth W P, Witham D L, Severin B F, Whittam T S, 2000, Mathematical modeling of ultraviolet germicidal irradiation for air disinfection, Quant Microbiology, 2, 249–70. • Lawton J A, Estes M K, Prasad B V V, 2000, Mechanism of genome transcription in segmented dsRNA viruses, Advances in Virus Research, 55: 185–214, IN1–IN8, 215–229. • Le-Guyader F L, Apaire-Marchais V, Brillet J, Billaudel S, 1983, Use of Genomic Probes To Detect Hepatitis A Virus and Enterovirus RNAs in Wild Shellfish and Relationship of Viral Contamination to Bacterial Contamination, Applied and Environmental Microbiology, 59(11) 3963-3968. • Leveque N, Laurent A, 2008, A Novel Mode of Transmission for Human Enterovirus Infection Is Swimming in Contaminated Seawater: Implications in Public Health and in Epidemiological Surveillance, Clinical Infection, 47, 624–626. Chapter 5 123 • Lugoli F, Leopizzi M I, De Donno A, 2009, Viral Indicators of Hygienic-Sanitary Quality: Detection of Somatic Coliphages in the Southern Adriatic Sea, Water Qual Expo Health, 1, 171-177. • Miller R, Plagemann P, 1973, Effect of ultraviolet light on mengovirus: formation of uracil dimers, instability and degradation of capsid, and covalent linkage of protein to viral RNA, Journal of Virology, 13, 729–39. • Nuanualsuwan S, Cliver D O, 2003, Capsid Functions of Inactivated Human Picornaviruses and Feline Calicivirus, Applied and Environmental Microbiology, 69 (1) 350-357. • Quek P H, Hu J Y, 2008, Indicators for photoreactivation and dark repair studies for following ultraviolet disinfection, Journal of Industrial Microbiology Biotechnology, 35, 533. • Santos A L, Oliveira V, Baptista I, Henriques I, Gomes N C M, Almeida A, Correia A, Cunha A, 2012, Effects of UV-B Radiation on the Structural and Physiological Diversity of Bacterioneuston and Bacterioplankton, Applied and Environmental Microbiology, 78 (6) 2066 • Smirnov Y A, Rodrigues-Motto MP, Famadas MT, 1983, Protein-RNA interaction in encephalomyocarditis virus as revealed by UV light-induced covalent linkages, Journal of Virology, 45, 1048–55. • Sommaruga R, 2003, UVR and its effects on species interactions, Organisms and Ecosystems, Edited by E.W. Helbling and H. Zagarese, pp. 485-508, The Royal Society of Chemistry, Cambridge, UK. • Sommer R, Pribil W, Appelt S, Gehringer P, Eschweiler H, Leth H, Cabaj A, Haider T, 2001, Inactivation of bacteriophages in water by means of non-ionizing (UV253.7nm) and ionizing (gamma) radiation: A comparative approach, Water Research, 35, 3109. • Suttle C A, 2007, Marine viruses — major players in the global ecosystem, Nature reviews – microbiology, 5: 801-811. • Thomas P, Swaminathan A, Lucas R M, 2012, Climate change and health with an emphasis on interactions with ultraviolet radiation: a review, Global Change Biology, 18 (8) 2392–2405. • Villena C, El-Senousy W M, Abad F X, Pinto R M, Bosch A, 2003, Group A Rotavirus in Sewage Samples from Barcelona and Cairo: Emergence of Unusual Genotypes, Applied and Environmental Microbiology, 69 (7) 3919-3923. Chapter 5 124 • Wegley L, Edwards R, Rodriguez-Brito B, Liu H, Rohwer F, 2007, Metagenomic analysis of the microbial community associated with the coral Porites astreoides, Environmental Microbiology, 9 (11) 2707–2719. • Yates M V, Gerba C P, Kelley L M, 1985, Virus persistence in groundwater, Applied and Environmental Microbiology, 49(4): 778-781. • Zepp R G, Erickson III D J, Paulc N D, Sulzbergerd B, 2011, Effects of solar UV radiation and climate change on biogeochemical cycling: interactions and feedbacks, Photochemical and Photobiology Science, 10, 261-279. Chapter 6 – Final Considerations Chapter 6 127 Although there is a great amount of viruses in the aquatic system, only a few number is important for human health. As pathogenic viruses are present in low numbers in the aquatic environment, samples must be concentrated to allow efficient detection of very few viruses in a large volume of water (Schwab et al, 1995). However, there is no perfect viral concentration method for water samples (Bosch et al, 2008; AlbinanaGimenez et al, 2009), and it should be chosen to achieve the aim of the work. In this work, it was compared the ultracentrifugation and the organic flocculation methods and it was detected that ultracentrifugation method can recover about 50% more viruses in recreational waters than the organic flocculation method, one of the most currently used approache to concentrate viruses from environmental waters (Calgua et al, 2008; Guttman-Bass and Armon, 1983; Guttman-Bass and Nassen, 1984; Shields and Farrah, 1986; virobathe, 2011). Besides the higher recovery of viruses obtained by the ultracentrifugation method, compared with the organic flocculation method, the first presents some other advantages that made it the best method to use in this work: is a simple and fast method, does not interfere with viral community structure, no chemicals are added to the sample avoiding PCR inhibition and it can reduce the final volume of sample to few microliters. In virology there is still scarce information in some areas. Nowadays, much is already known about the viruses present in the aquatic system that represent a health hazard for humans, but little is known about their natural reservoirs, namely the surface microlayer (SML), and their transmission to the atmosphere through aerosols, mainly because of the difficulty in collecting and analyzing airborne virus (Verrault et al, 2008). The sea SML corresponds to the interface between the surface of the water and the air and is known to accumulate particles and microorganisms and plays a crucial role in exchange processes of gases and matter across the air-water interface. The gases and matter transferred from the SML are transported across the air-water interface by aerosols. So, aerosols composition depends on SML composition. With this work it was found that SML acts as a major source of microorganisms to the atmosphere and aerosols constitute an important mechanism of viruses dispersion. In order to form and collect aerosols in field work, a protocol was developed. Polycarbonate membranes (0.45 µm) were tested to filtrate the air used to form aerosols, miliQ water and 1xPBS were compared as a collection medium and two times (30 minutes and 1 hour) of aerosols formation were tested. The polycarbonate Chapter 6 128 membranes were efficient in removing viruses from the air, with a reduction of about 80% and they do not interfere with viral community, since values of VLP between 10 9 and 10 10 VLP L -1 (data not shown), which is according to the literature (viral concentration range between 10 7 L -1 and 10 11 L -1 in coastal areas (Wommack and Colwell, 2000) were detected. The collection medium chosen was miliQ water, mainly because of the background fluorescence of 1x PBS in the epifluorescence microscopy. The time needed to form aerosols in laboratory was of 1 hour. So, the protocol established for aerosols formation and recovery in field work was formation of aerosols for 1 hour, with air filtered by a polycarbonate membrane bubbling in the water sample and collection of the aerosols formed in miliQ water. The five groups of enteric viruses tested were detected in water and aerosols in both the marine and the brackish water zones. Viral abundance obtained was according to which is indicated in the literature and the patterns of variation were also according to that described in the literature. Rotavirus and Enterovirus presented a peak in cold months, as in other studies (2000; Guix et al, 2002; Bosch et al, 2008), and HAV presented similar density in all sampling dates. Rotavirus and Enterovirus ranged between 10 2 – 10 7 copies L -1 and HAV range between 10 1 – 10 2 copies L -1 . The values obtained in other works described in the literature were of 10 1 to 10 8 copies L -1 for Rotavirus (Li et al, 2010; Chigor and Okoh, 2012; Vieira et al, 2012), 10 1 to 7 copies L -1 for Enterovirus (Zhang et al, 2010; Chigor and Okoh, 2012; Aslan et al, 2013) and 10 1 to 10 9 copies L -1 for HAV (Chigor and Okoh, 2012; Keuckelaere et al, 2013). The variation found in viral abundance is better explained by temperature and nitrites plus nitrates concentration, decreasing viral concentration with the increase in temperature and incrasing with the increase in nitrites plus nitrates concentration. This suggests that enteric viruses have mainly terrestrial origin reaching the marine environment through runoff namely during the winter months. Enteric viruses were more abundant in the SML and less abundant in the aerosols, with significant differences between the water layers and the aerosols (p < 0.05), both in the marine and in the brackish water zones.Enteric viruses were about 6 and 18 times more abundant in the SML than in UW, in marine and brackish water zones, respectively and 14.63 and about 46 times more abundant in the SML than in the aerosols, for marine and brackish water zones, respectively. These results are according the literature since virus concentration in the air is low (Després et al, 2011) and the SML is generally enriched in microrganisms (Cunliffe et al, 2011) and is agreement to the work of Aller et al (2005), that indicates that the SML acts as a major Chapter 6 129 source of microorganisms to the atmosphere and aerosols constitute an important mechanism of viral dispersal to long distances. Viral survival may vary, depending on viral type. However, there are several factors that may affect that survival, like exposure to UV light, water temperature, pH, salinity, adsorption to sediments, predators, presence of particular matter, salinity and raining (Le-Guyader et al. 1983, Chuan et al. 1983, Goyal et al. 1984, Yates et al. 1985, Griffin et al. 2003, Bosch et al. 2005; Bosch 2007; Fong et al. 2005, Bosch et al. 2006, Suttle 2007, Lugoli et al. 2009). In this work the influence of exposure to UV light on viral abundance was tested, but usually the studies about the effect of UV exposition on viruses test only viral infectivity and not changes in viral abundance. The results of this study showed that for Rotaviruses no significant reduction was found in viral abundance but a significant reduction was found for Enterovirus (p = 0.03 < 0.05). The difference observed between the two viruses can be related to RNA type, dsRNA in Rotavirus and ssRNA in Enterovirus, but also with the structure of the viral capsid, a three-layered capsid in Rotavirus and a single capside in Enterovirus. It is well known that dsDNA are more resistant to environmental factor such as UVR, and as a complex capsid difficult the RNA damage mechanism, Rotavirus is more resistant to UVR than Enteroviruses. This may explain the reduced number of Enterovirus waterborne outbreaks relatively to Rotavirus (Leveque et al. 2008). The development of scientific work for a PhD thesis must generate knowledge that contributes in some way to create value, whether social, economic or environmental. Since enteric viruses have low infectious dose, being responsible for numerous hospitalizations, due to gastroenteritis, it is extremely important to know how these viruses can be transmitted to humans, in order to minimize this scourge. The finds that SML is a reservoir of enteric viruses in the marine environment and that there is potential for their transference from water to air. This is an important knowledge in preventing waterborn viral infection. This work provides the basis for future studies. Future work will pass to: • Determine the percentage of the UV light exposed remaining viruses that are still capable of successfully infect host cells. • The time and UV necessary to reduce significantly the abundance of all enteric viruses tested. 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