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Experimental evaluation of infection, dissemination, and transmission rates for two West Nile virus strains in European Aedes japonicus under a fluctuating temperature regime
research-article
Author(s):
Eva Veronesi
1
,
,
Anca Paslaru
1 ,
Cornelia Silaghi
1
,
2 ,
Kurt Tobler
3 ,
Uros Glavinic
1 ,
Paul Torgerson
4 ,
Alexander Mathis
1
Publication date (Electronic):
28 April 2018
Journal:
Parasitology Research
Publisher:
Springer Berlin Heidelberg
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Abstract
West Nile virus (WNV) is continuously spreading in Eastern and Southern Europe. However, the extent of vector competence of Aedes japonicus (Theobald, 1901) is controversial. In this work, we elucidated the dynamics of virus growth in this invasive mosquito species. Females of Ae. japonicus were reared from eggs collected in the field in Switzerland and fed on bovine blood spiked with two WNV lineage 1 strains (FIN, Italy; NY99, USA). Fully engorged females were incubated for 14 days under a fluctuating temperature regime of 24 ± 7 °C (average 24 °C), 45–90% relative humidity, which is realistic for a Central European mid-summer day. Infection, dissemination, and transmission rates were assessed from individual mosquitoes by analyzing the abdomen, legs and wings, and saliva for the presence of viral RNA. Saliva was also investigated for the presence of infectious virus particles. Overall, 302 females were exposed to WNV strain FIN and 293 to strain NY99. A higher infection rate was observed for NY99 (57.4%) compared to FIN (30.4%) ( p = 0.003). There was no statistical evidence that the dissemination rate (viral RNA in legs and wings) was different between females infected with FIN (57.1%) compared to NY99 (35.5%) ( p = 0.16). Viral RNA load of FIN compared to NY99 was significantly higher in the hemocoel ( p = 0.031) of exposed females but not at other sites (legs and wings, saliva). This is the first study describing the vector competence parameters for two WNV strains in a European population of Ae. japonicus. The high dissemination and transmission rates for WNV under a realistic temperature regime in Ae. japonicus together with recent findings on its opportunistic feeding behavior (mammals and birds) indicate its potential role in WNV transmission in Central Europe where it is highly abundant.
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Most cited references35
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West Nile fever--a reemerging mosquito-borne viral disease in Europe.
Z Hubalek, J Halouzka (1999)
West Nile virus causes sporadic cases and outbreaks of human and equine disease in Europe (western Mediterranean and southern Russia in 1962-64, Belarus and Ukraine in the 1970s and 1980s, Romania in 1996-97, Czechland in 1997, and Italy in 1998). Environmental factors, including human activities, that enhance population densities of vector mosquitoes (heavy rains followed by floods, irrigation, higher than usual temperature, or formation of ecologic niches that enable mass breeding of mosquitoes) could increase the incidence of West Nile fever.
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Lineage 1 and 2 Strains of Encephalitic West Nile Virus, Central Europe
Tamás Bakonyi, Éva Ivanics, Károly Erdélyi … (2006)
Geographically, West Nile virus (WNV) is the most widespread member of the Japanese encephalitis virus (JEV) complex within the genus Flavivirus and the family Flaviviridae. The first strain (B 956) was isolated from a human patient in the West Nile district of Uganda in 1937; later the virus was also detected in several mosquito species, horses, humans, and other hosts in Africa, Europe, Asia, and Australia (where it has been named Kunjin virus) ( 1 – 3 ). WNV was introduced into the United States in 1999, and it spread quickly over large parts of North America and reached Mexico ( 4 – 7 ). The clinical impact of WNV varies in different regions. In the Old World, WNV causes relatively mild infections with influenzalike symptoms or no apparent disease ( 2 ); encephalitis and fatalities in the human population, horses, or poultry are spasmodic ( 3 , 8 , 9 ). In the New World, WNV exhibits increased virulence among the local wild bird populations and causes more frequent severe central nervous system symptoms and deaths in humans and horses ( 6 , 10 ). Although exactly how WNV was introduced into New York is unclear, phylogenetic comparison of the viral nucleic acid sequences has shown a close relationship between the American WNV isolates and strains isolated from encephalitic geese and storks in Israel in 1998 ( 11 – 13 ). Experimental infections of rodents indicated that the neurovirulence of WNV correlates with its genotype, and the North American strains are highly neurovirulent for mice ( 14 ). WNV shows relatively high levels of sequence diversity. Comprehensive studies on the phylogenetic relatedness of WNV strains show that they form at least 2 main lineages ( 15 – 17 ). Lineage 1 is composed of WNV strains from different geographic regions, and it is subdivided into at least 3 clades. Clade A contains strains from Europe, Africa, the Middle East, and America; clade B represents the Australian (Kunjin) strains; and clade C contains Indian WNV isolates. Lineage 2 contains the B 956 prototype strain and other strains isolated so far exclusively in sub-Saharan Africa and Madagascar. In addition to the 2 major WNV lineages, we recently proposed 2 lineages for viruses that exhibited considerable genetic differences to the known WNV lineages: lineage 3 consists of a virus strain isolated from Culex pipiens mosquitoes at the Czech Republic/Austria border (named Rabensburg virus), and lineage 4 consists of a unique virus isolated in the Caucasus. These 2 viruses, however, may also be considered independent flaviviruses within the JEV complex ( 18 ). WNV has been known to be present in central Europe for a long time. Seroprevalence in humans was reported in several countries, including Hungary, and WNV strains were isolated from mosquitoes, humans, migrating birds, and rodents during the last 30 years ( 3 ). Until 2003, however, WNV infections in Hungary have never been associated with clinical symptoms, although a severe outbreak of West Nile encephalitis in humans was reported in 1996 and 1997 in neighboring Romania. In late summer 2003, an outbreak of encephalitis emerged in a Hungarian goose flock, resulting in a 14% death rate among 6-week-old geese (Anser anser domesticus). Based on histopathologic alterations, serologic investigations, and nucleic acid detection by reverse transcription–polymerase chain reaction (RT-PCR), WNV was diagnosed as the cause of the disease ( 19 ). Chronologically and geographically related to the outbreak in geese, a serologically confirmed WNV outbreak was also observed in humans, which involved 14 cases of mild encephalitis and meningitis ( 20 ). One year later, in August 2004, a goshawk (Accipiter gentilis) fledgling showed central nervous system symptoms and died in a national park in southeastern Hungary. When histopathologic methods and RT-PCR were used, WNV antigen and nucleic acid were detected in the organs of the bird. Furthermore, the virus was isolated after injection of suckling mice. Here we report the sequencing and phylogenetic results of these 2 encephalitic WNV strains that emerged recently in central Europe. Materials and Methods Brain specimens from one 6-week-old goose, which died during the encephalitis outbreak in a Hungarian goose flock, and brain samples from a goshawk, which also died from encephalitis, were used for WNV nucleic acid determination. The brain samples were homogenized in ceramic mortars by using sterile quartz sand, and the homogenates were suspended in RNase-free distilled water. Samples were stored at –80°C until nucleic acid extraction was performed. Viral RNA was extracted from 140 μL of brain homogenates by using the QIAamp viral RNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer's instructions. First, a universal JEV-group specific oligonucleotide primer pair designed on the nonstructural protein 5 (NS5) and 3´-untranslated regions (UTR) of WNV (forward primer: 5´-GARTGGATGACVACRGAAGACATGCT-3´ and reverse primer: 5´-GGGGTCTCCTCTAACCTCTAGTCCTT-3´ [21]; ) was applied on the RNA extracts in a continuous RT-PCR system employing the QIAGEN OneStep RT-PCR Kit (Qiagen). Each 25-μL reaction mixture contained 5 μL of 5× buffer (final MgCl2 concentration 2.5 mmol/L), 0.4 mmol/L of each deoxynucleoside triphosphate, 10 U RNasin RNase Inhibitor (Promega, Madison, WI, USA), 20 pmol of the genomic and reverse primers, 1 μL enzyme mix (containing Omniscript and Sensiscript Reverse Transcriptases and HotStarTaq DNA polymerase) and 2.5 μL template RNA. Reverse transcription was carried out at 50°C for 30 min, followed by a denaturation step at 95°C for 15 min. Thereafter, the cDNA was amplified in 40 cycles of heat denaturation at 94°C for 40 s, primer annealing at 57°C for 50 s, and DNA extension at 72°C for 1 min, and the reaction was completed by a final extension for 7 min at 72°C. Reactions were performed in a Perkin-Elmer GeneAmp PCR System 2400 thermocycler (Wellesley, MA, USA) and in a Hybaid PCR Sprint thermocycler (Thermo Electron Corporation, Waltham, MA, USA). After RT-PCR, 10 μL of the amplicons was subjected to electrophoresis in a 1.2% Tris acetate-EDTA-agarose gel at 5 V/cm for 80 min. The gel was stained with ethidium bromide; bands were visualized under UV light and photographed with a Kodak DS Electrophoresis Documentation and Analysis System using the Kodak Digital Science 1D software program (Eastman Kodak Company, Rochester, NY, USA). Product sizes were determined with reference to a 100-bp DNA ladder (Promega). Where clear PCR products of the previously calculated sizes were observed, the fragments were excised from the gel, and DNA was extracted by using the QIAquick Gel Extraction Kit (Qiagen). Fluorescence-based direct sequencing was performed in both directions on PCR products. Sequencing of PCR products was carried out with the ABI Prism Big Dye Terminator cycle sequencing ready reaction kit (Perkin-Elmer), according to the manufacturer's instructions, and an ABI Prism 310 genetic analyzer (Perkin-Elmer) automated sequencing system. Nucleotide sequences were identified by Basic Local Alignment Search Tool (BLAST, http://www.ncbi.nlm.nih.gov/blast) search against gene bank databases. Based on the sequence information obtained from the amplification products, complete WNV sequences that exhibited the highest nucleotide identities with the Hungarian genotypes were selected from the GenBank database to design primers that amplify overlapping RT-PCR products covering the entire genome of the strains. Oligonucleotide primers were designed with the help of the Primer Designer 4 for Windows 95 (Scientific and Educational Software, Version 4.10; Microsoft, Redmond, WA, USA) and were synthesized by GibcoBRL Life Technologies, Ltd. (Paisley, Scotland, UK). Detailed information on all primers is in the Tables A1 and A2. PCR amplification products were directly sequenced in both directions; the sequences were compiled and aligned to complete genome sequences of selected representatives of WNV lineages 1a, 1b, 2, and putative lineages 3 and 4 (listed in Table). Phylogenetic analysis was performed by using the modified neighbor-joining method (ClustalX [22]; ), and trees were constructed to demonstrate the relationship between the Hungarian WNVs and other WNV strains (Figure). Figure Phylogenetic tree based on the complete nucleotide sequences of selected West Nile virus strains demonstrating the genetic relatedness of these strains (abbreviations are listed in Table). Boxes indicate different lineages and clades. The Hungarian strains reported in this article are highlighted with gray background). RabV, Rabensburg virus; JEV, Japanese encephalitis virus. Scale bar depicts degree of relatedness. Table West Nile virus strains included in the phylogenetic analysis Name Code Accession no. Isolation Year Host Origin Lineage, clade WNV HNY1999 NY99a AF202541 1999 Human New York, USA 1a WNV NY99flamingo38299 NY99b AF196835 1999 Flamingo New York, USA 1a WNV IS98STD Is98 AF481864 1998 Stork Israel 1a WNV goose-Hungary/03 Hu03 DQ118127 2003 Goose Hungary 1a WNV Italy1998Equine It98 AF404757 1998 Horse Italy 1a WNV RO9750 Ro96 AF260969 1996 Culex pipiens Romania 1a WNV VLG4 Rus99a AF317203 1999 Human Volgograd, Russia 1a WNV LEIV-Vlg99-27889 Rus99b AY277252 1999 Human Volgograd, Russia 1a WNV PaH001 Tu97 AY268133 1997 Human Tunisia 1a WNV PaAn001 Fr00 AY268132 2000 Horse France 1a WNV Eg 101 Eg51 AF260968 1951 Human Egypt 1a WNV Chin-01 Chin01 AY490240 1950s ? Russia 1a WNV Kunjin MRM61C Kunjin D00246 1960 Cx. annulirostris Australia 1b WNV Sarafend Sarafend AY688948 Laboratory strain 2 WNV B956 (WNFCG) Ug37 NC_001563 1937 Human Uganda 2 WNV goshawk-Hungary/04 Hu04 DQ116961 2004 Goshawk Hungary 2 Rabensburg virus (97-103) RabV AY765264 1997 Cx. pipiens Czech R. 3? WNV LEIV-Krnd88-190 Rus98 AY277251 1998 Dermacentor marginatus Caucasus, Russia (Georgia?) 4? The nucleotide sequences of the Hungarian WNV strains goose-Hungary/03 (Hu03) and goshawk-Hungary/04 (Hu04) were submitted to the GenBank database. They are available under accession numbers DQ118127 and DQ116961, respectively. Results In this study, the complete genome sequences of WNV strains derived from a 6-week-old goose, which died in 2003 during an outbreak of encephalitis in a Hungarian goose flock (strain goose-Hungary/03), and from a goshawk, which also died from encephalitis in the same region 1 year later (strain goshawk-Hungary/04), were determined, aligned, and phylogenetically analyzed. The genome of the goose-Hungary/03 strain is composed of 10,969 nucleotides (nt) and contains 1 open reading frame between nucleotide positions 97 and 10,398, coding for a 3,433 amino acid (aa)–long putative polyprotein precursor. The complete genomic sequence of the virus was subjected to a BLAST search against gene bank databases. The highest identity rates (98% at the nucleotide and 99% at the amino acid level) were found with WNV strains isolated in 1998 in Israel and in 1999 in the United States. In addition, phylogenetic analysis was performed to indicate the relationships between the Hungarian goose–derived WNV strain and selected representatives of WNV clades and clusters. The resulting phylogenetic tree (Figure) confirmed the results of the BLAST search, i.e., the Hungarian goose–derived WNV strain is clustering close to the previously mentioned WNV strains isolated in the United States and Israel, which belong to lineage 1a of WNV. Other European WNV strains (isolated in Italy, France, and Romania) are more distant to the Hungarian strain; they form a separate cluster consisting of a Romanian/Russian and a French/Italian subcluster. The complete nucleotide sequence of the goshawk-Hungary/04 WNV strain is composed of 11,028 nt and contains 1 open reading frame between nucleotide positions 97 and 10,401, coding for a 3,434-aa putative polyprotein precursor. In BLAST search, the strain showed the highest (96% nt and 99% aa) identity to the WNV prototype strain B 956. Consequently, as the phylogram also indicates (Figure), this virus belongs to lineage 2 of WNV. Alignments of the available partial sequences from the E protein coding regions of other representatives of this cluster showed even higher identities (97%–98% nt and 100% aa) with WNV strains isolated in central Africa in 1972 (AnB3507, AF001563) and in 1983 (HB83P55, AF001557), respectively ( 15 ). More recently (in early August 2005), additional lethal cases of encephalitis occurred in birds of prey in the same place in which the goshawk died of West Nile encephalitis in 2004, involving up to a total of 3 goshawks and 2 sparrow hawks (A. nisus); 2 of the goshawks and 1 sparrow hawk died. Preliminary investigations detected WNV-specific nucleic acid in the brains of the birds. The partial nucleotide sequence of the 2005 virus (1,000 bp at the NS5´–3´-UTR regions) showed 99.9% identity with the goshawk-Hungary/04 strain (only 1 substitution at nucleotide position 9,376 [g→a] has been observed, which did not influence the putative amino acid sequence). Additional observation of the outbreak and investigations of the cases are in progress. Discussion The primary aim of our investigations was to show the genetic relatedness of the WNV strains detected in Hungary in the last 2 years and to estimate their clinical and epidemiologic impact. The phylogenetic analysis emphasizes the close genetic relationship of the goose-Hungary/03 strain with a WNV strain isolated in Israel in 1998 and the WNV strain introduced in New York in 1999, since the 3 WNVs form 1 single cluster within clade 1a of lineage 1. These strains caused outbreaks in birds, humans, and horses. Previous European WNV isolates exhibited lower identity values, e.g., the strain that was responsible for the Romanian outbreak(s) in 1996 and 1997 showed only 96% nt identity with the Hungarian goose-2003 strain, and in the phylogenetic tree the other European isolates form a separate cluster consisting of 2 subclusters (Figure). The earliest representatives of the Israel/USA/goose-Hungary/03 cluster were reported by Malkinson et al. ( 23 ) from ill and dead white storks (Ciconia ciconia) in Israel in 1998. These storks, however, had hatched in central Europe, and during their autumn migration southwards, strong winds had blown them off course, from their usual route to Africa, to southern Israel. Malkinson et al. suspected that these birds introduced the neurovirulent genotype of WNV to Israel from their hatching place. The wetlands of southeastern Hungary are foraging and nesting habitats for storks and many other wild bird species, and the goose farm, where the WNV outbreak occurred in 2003, is located in this region. These facts, together with the close phylogenetic relatedness of the Israeli/US/Hungarian WNV strains, strongly support the theory that storks carried the neurovirulent WNV strain from central Europe (that is, from Hungary) to Israel, which sheds new light on the introduction of WNV to New York. This virus could have originated in Israel (which is the generally accepted although not proven theory) or central Europe. In both cases, however, the virus seems to have its true origin in Europe. In a recent publication, Lvov et al. suggested that WNV could have been introduced into New York by ships traveling from Black Sea ports ( 24 ). When a WNV infection was detected in 2004 in a goshawk fledgling, which died from encephalitis in the same region of Hungary in which the outbreak in geese and humans occurred during the previous year, we anticipated a WNV strain more or less identical to the genotype detected there in 2003. The genomic sequence of this strain was not closely related to the sequence of the WNV strain detected in geese in the year before, however, but belonged to the group of central African lineage 2 WNV strains. A closely related strain from this cluster (ArB3573, AF001565, and AF458349) was identified as a neuroinvasive strain of WNV in a mouse model ( 14 ). To our knowledge, this report is the first on the emergence of a lineage 2 WNV strain outside Africa. Migratory birds that had overwintered in central Africa probably introduced this exotic strain to the wetlands of Hungary. On the other hand, as the goshawk is not a migratory species, and infection occurred in August, the African WNV strain must have already successfully adapted to local mosquito vectors. Consequently, this neurotropic, exotic WNV strain may become a resident pathogen in Europe with all the possible public health consequences. Our results indicate that the WNV strains that emerged in 2 consecutive years and caused avian deaths in Hungary are epidemiologically unrelated. Genetically distinct WNV strains are circulating simultaneously yet independently in local birds and thus most likely also in local mosquito populations within the same region. They cause sporadic cases of encephalitis and also raise the possibility of spreading to other European countries or even to other continents, as happened in 1999 with another WNV strain, which resulted in a public health catastrophe in America. In addition to the above 2 novel WNVs, we recently characterized another novel flavivirus of so far unknown human pathogenicity named Rabensburg virus, which has been isolated from Culex pipiens mosquitoes in 1997 and 1999 at the Czech Republic–Austria border, only a few hundred kilometers from the region where the Hungarian WNVs emerged. After the entire genome was sequenced, Rabensburg virus turned out to represent either a new (third) lineage of WNV or a novel flavivirus of the JEV group ( 18 ). Thus, several distinct WNV strains seem to circulate in central Europe. In 2001 another flavivirus of the JEV group, Usutu virus, which has never previously been observed outside Africa, emerged in Austria and resulted in deaths in several species of birds, especially Eurasian blackbirds (Turdus merula) ( 21 ). This virus became a resident pathogen in Austria and continues to disperse and cause deaths in blackbirds and other species of birds ( 25 , 26 ). The snowy winter and rainy spring of 2005 resulted in serious floods in the area in which the Hungarian WNV strains were identified. Since the floodplains and polders were under water, the conditions for mosquito development were ideal. The summer was also very rainy, which resulted in more floods in the region and continuous mosquito gradation. The most recent data imply that the lineage 2 WNV strain may have overwintered in Hungary, causing several clinical cases of encephalitis in Accipiter species in 2005 as well. The routine diagnostic techniques in most of the European public health and veterinary laboratories are designed to detect lineage 1 WNV strains. In a recent PCR external quality assurance multicenter test, <40% of the involved laboratories could detect lineage 2 WNV strains (Matthias Niedrig, pers. comm.). Therefore, a major goal of this article is to increase the scientific and public awareness of this potential public health threat for Europe and, perhaps, America. Furthermore, comprehensive investigations on the occurrence, ecology, and epidemiology of the different WNV strains circulating in central Europe, as well as the development of monitoring and surveillance programs, must be of highest priority. One may also speculate on environmental factors, such as climate change or global warming, that may have enhanced the recent emergence of viruses, which had previously been restricted to Africa, in new habitats and continents. Improved observation, reporting, and detection methods have also contributed to the apparent increasing emergence of these viruses.
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Temperature, Viral Genetics, and the Transmission of West Nile Virus by Culex pipiens Mosquitoes
A. Kilpatrick, Mark A Meola, Robin M. Moudy … (2008)
Introduction The interaction between pathogens, their vectors, and vertebrate hosts is a dynamic one, and evolution in any one of the three can significantly alter transmission dynamics. Theory suggests that pathogens will evolve to maximize their fitness, which is a function of transmissibility and virulence to the host [1],[2]. Pathogens that infect and replicate well in their vertebrate hosts and vectors may decrease the survival of both which may reduce their lifespan for transmission. At the same time, the distribution and intensity of transmission of vector-borne pathogens is strongly influenced by the interaction of temperature, vectors, hosts, and pathogen genetics. Temperature can determine both the latitudinal boundary and upper elevational limit of pathogen transmission if the extrinsic incubation period (EIP) is greater than the longevity of the vector [3]. Temperature also has been linked to changes in the intensity of transmission of pathogens [4],[5], which may be linked to temperature-induced changes in the EIP, the longevity, and the feeding rate of vectors [6],[7]. West Nile virus (WNV; Flaviviridae, Flavivirus), is a single-stranded positive-sense RNA virus that was introduced into the western hemisphere in 1999 and has subsequently spread throughout much of North, Central, and South America [8],[9]. It is primarily transmitted between birds (especially American robins, Turdus migratorius, in many areas [10]–[14]) and Culex mosquitoes [8],[15] and has caused at least 2,500 reported cases each year since 2002 for a total of 32,135 total reported cases, 11,243 cases of encephalitis, and 1,125 deaths, with an estimated 1.56 million infections and 310,000 illnesses from 1999–2007 [9], [16]–[18]. In addition, WNV has evolved over the past 7 years, and a genotype that was first isolated in 2001 (termed WN02) has displaced the introduced genotype (termed NY99) [19],[20]. WNV strains in the WN02 or North American dominant genotype have three consensus changes in the full length genome compared to NY99 [19],[20]. The rapid expansion of the WN02 genotype has been linked to a shorter extrinsic incubation period in Culex mosquitoes [21],[22], but the full mechanisms of displacement are not yet known. In particular, in previous studies WN02 genotypes were transmitted more efficiently than NY99 by Cx pipiens on 5 and 7 days post feeding, but not day 9 and by Cx. tarsalis from 5 to 14 days post feeding [21],[22]. The vector competence of mosquitoes characterizes their ability to transmit a pathogen after taking an infected blood meal. The fraction of vectors transmitting the pathogen is known to vary between populations of a species [23]–[25], and increase with time [26],[27] and temperature for WNV [28]–[30] and many other pathogens, including western equine encephalomyelitis virus and St. Louis encephalitis virus in Cx. tarsalis [28],[31], Rift Valley Fever virus in Aedes fowleri [32], Ockelbo virus in Culex spp [33] and Aedes spp [34], and African horse sickness virus, bluetongue virus, and epizootic hemorrhagic disease in Culicoides sonorensis [35]. However, the exact relationship between vector competence, temperature and the time since feeding on an infected host is not clear, and in other studies the influence of temperature on vector competence varies, sometimes depending on the mosquito species infected [33],[36],[37]. A degree day model developed for Cx. tarsalis has been used to model the effect of temperature on WNV transmission across North America [28],[38]. In this approach, a mosquito (or a fraction of a population of mosquitoes) feeding on an infected host becomes infectious after a time period at a certain temperature, termed the number of degree days. Degree days are often measured as the number of days since feeding multiplied by the temperature in degrees Celsius above a minimum temperature threshold (Tthr ) below which no transmission is assumed to occur. However, the exponential increase with temperature in chemical and molecular processes contributing to viral replication would suggest that the relationship between transmission and the number of degree days should be accelerating and would not be well described by a simple degree day model. Transmission would be expected to be higher at higher temperatures given the same number of degree days. For example, if Tthr = 14°C, transmission would be expected to be higher after seven days at 30°C (16° above the threshold temperature of zero transmission) than after 16 days at 21°C (7° above the Tthr ), even though the same number of degree days, 112, is the same in both cases. Here we explore the relationship between temperature and transmission of two genotypes of WNV (NY99 and WN02) and test the adequacy of a simple degree day model for WNV transmission by Culex pipiens, a key enzootic and bridge vector for WNV in the northern USA [15],[39]. Methods Viruses and mosquitoes Two strains of WNV were used, one belonging to genotype NY99 and one to WN02 (strain designations are NY99-3356 and WN02-1956, Genbank accession number, AF404756 and AY590210, respectively). Previous work suggested that there was little phenotypic variation between strains within genotype [21],[22]. NY99-3356 was passed twice in Vero (African Green Monkey kidney) cells, and WN02-1956 was passed once in Vero cells followed by one passage in C6/36 (Aedes albopictus) cells prior to use in these studies. Colonized Cx. pipiens were reared and maintained in the Wadsworth Center Arbovirus Laboratory BSL-2 insectary. The colony was established in 2002 from egg rafts collected in Pennsylvania (courtesy of Michael Hutchinson) and has been maintained continuously using defibrinated goose blood (Hema Resourse and Supply, OR) for egg production and 10% sucrose ad lib for maintenance at 27°C with 16∶8 L∶D light cycle and 85% humidity. All experiments with infectious virus were performed in the BSL-3 laboratories or insectaries at the Wadsworth Center Arbovirus Laboratories. Vector competence Seven day-old mosquitoes were deprived of sucrose and water for 48 h and then fed on a suspension of defibrinated goose blood (Hema Resourse and Supply, OR) plus a final concentration of 2.5% sucrose and either a NY99 or a WN02 virus, using a Hemotek feeding system (Discovery Workshops, UK). The WNV titer in the bloodmeals was 1.2–1.4×108 plaque-forming units (PFU)/ml. Mosquitoes were allowed to feed for up to 1.5 h at which time engorged mosquitoes were separated from unfed mosquitoes under CO2 anesthesia. Fully engorged mosquitoes were placed into 0.6L cardboard cartons, supplied with 10% sucrose ad lib, and held at the prescribed temperatures under 85% RH, photoperiod of 16∶8 (L∶D). Groups of 25 mosquitoes were removed at several different intervals post-feeding and anesthetized with triethylamine (Sigma, St. Louis, MO). The days sampled included days 4, 7, 10, 14, 18, 21, 24, 28, 31, 34, and 40 for 15°C, 18°C, and 22°C and additional early sampling at days 0.5, 1, 1.5, 2, 2.5, 3 for experiments at 32°C. Legs were removed and placed in 1.0 ml of mosquito diluent (MD; 20% heat-inactivated fetal bovine serum [FBS] in Dulbecco's phosphate-buffered saline plus 50 ug/ml penicillin/streptomycin, 50 ug/ml gentamicin, and 2.5 ug/ml Fungizone) and frozen at −80°C for subsequent assay. Salivary secretions were collected using a modified in vitro capillary transmission assay [40]. Briefly, mosquito mouthparts were inserted into a capillary tube containing approximately 10 µL of a mixture of 50% sucrose and FBS (1∶1) for 30 minutes, at which time the contents were placed into 0.3 ml MD in a microfuge tube. Bodies were placed in 1.0 ml MD and all samples were frozen at −80°C for subsequent assay. Bodies and legs were homogenized separately using a mixer mill (Qiagen, Valencia, CA) at 24 cycles/s for 30 s and then clarified by centrifugation. Samples were analyzed for the presence of infectious virus by plaque assay on Vero cells as previously described [41]. Statistical analyses We treated each group of 25 individual mosquitoes tested after a fixed time at a temperature as an experimental unit (data point) and the fraction of mosquitoes that were infected (# with virus in the body/# fed), had disseminated infections (# with virus in their legs/# fed), or transmitted (# expectorating virus/# fed) as dependent variables. We built regression models (using SPSS v 15.0) including degree days (DD = tT, where t = time or days since feeding, and T = temperature in degrees Celsius) and a genotype by DD interaction (to test for a temperature and time varying advantage of WN02) as independent variables. We note that this statistical model (and the one described below) assumes that infection and transmission are increasing functions of temperature and time since feeding, and statistical effects model differences between genotypes as differences in the rates of increase (slopes), rather than fixed differences (intercept or main effects). We believe intercept differences are less biologically realistic because infection, dissemination, and transmission all start at zero and increase with time and temperature. In essence, the statistical effect of viral genotype is assumed to influence the rate at which the probability of a group of mosquitoes transmitting and becoming infected increases with temperature and time. We also note that our degree model implicitly assumes the minimum temperature threshold is 0°C (since it uses the raw temperature), which is likely too low, as no transmission was observed at 10°C in Cx. tarsalis held for 110 days [28]. However, the fit of the data were much better using raw temperature than either (Temperature −10°C) or (Temperature −14.3°C) (the residual error from models in Table 2 with a threshold of 14.3°C and 10°C were 3.02 and 1.87, respectively, compared to 0.99 with a threshold of 0°C; all regressions had the same number of predictors). We arc-sin square-root transformed the three dependent variables to normalize the residuals. We omitted an intercept from the model because we assumed that, except for residual virus in the blood meal, infection and transmission would be 0 at degree-day 0. The qualitative conclusions presented below were identical using an intercept. We then tested the hypothesis that infection, disseminated infection, and transmission should accelerate with increasing temperature faster than the DD model, by regressing the residuals of the previous models against temperature. A priori, we hypothesized that the increase in transmission and infection with temperature would be a balance between chemical and kinetic processes that increase exponentially (i.e. as eT where T is temperature, and e is the base of the natural logarithm) and rate-limiting processes that would constrain viral replication. Since the residuals were significantly correlated with temperature for all three dependent variables (see Results) we attempted to determine if DD with a higher order temperature term would provide a better fit to the data. To facilitate model comparison we replaced the DD model term (tT) with a term that was the product of the days since feeding (t) and temperature (T), raised to the power n (tTn ), and compared models with increasing n. For the DD by genotype interaction we also used DD with temperature also raised to the nth power. Finally, the qualitative results of both our analyses were unaffected by using actual temperature to calculate degree days, or degrees above a previously reported [28] threshold for zero transmission of 14.3°C. Our own data, and those in ref. [28], show that low-level transmission occurs at 14–15°C (but after very long periods that may exceed mosquito lifespan in the field). Results We examined a total of 2075 Cx. pipiens mosquitoes in 83 groups of 25 individuals and examined midgut infection, disseminated infection, and transmission from 12 hours to 40 days post-feeding (Figure 1). At 32°C, we detected transmission at 12, 36, and 60 hours for the WN02 genotype, and on day 3 (72 hours) for the NY99 genotype of WNV (Figure 1C). Virus was also present in the legs (and abdomens) of these mosquitoes at these time points (Figure 1B) and was not present in the saliva of any of the mosquitoes that were not infected, so it is unlikely that mechanical transmission or regurgitation accounted for the virus detected in the transmission assays (see also Discussion, below). 10.1371/journal.ppat.1000092.g001 Figure 1 The relationship between genotype (NY99 and WN02), temperature, and days since feeding and the fraction of Culex pipiens mosquitoes infected (A), with disseminated infections (B), or transmitting WNV (C), after 0.5–40 days as the proportion of mosquitoes tested. The initial regressions indicated that the fraction of mosquitoes infected and the fraction with disseminated infections increased with degree days (DD = tT) since feeding (Figures 1,2; Table 1). However, neither was significantly different between genotypes (Figures 1,2; Table 1). In contrast, transmission of WNV by Cx. pipiens mosquitoes was significantly influenced by both DD and a genotype by DD interaction (Figures 1,2; Table 1). The coefficient of this last term indicated that the fraction of mosquitoes transmitting WNV increased faster for the WN02 genotype than the NY99 genotype (and the fitted function for WN02 was greater at all times and temperatures since both lines intersect the origin). In fact, the fraction of mosquitoes transmitting the WN02 genotype was greater than or equal to the fraction transmitting the NY99 genotype for all but two of the 42 time-temperature samplings (the two exceptions were at 22°C on days 7 and 10 where 1/25 mosquitoes transmitted the NY99 genotype but 0/25 transmitted the WN02 genotype). Thus, the WN02 genotype appeared to have an advantage at both high and low temperatures, and this advantage increased with time and temperature. 10.1371/journal.ppat.1000092.g002 Figure 2 Fitted relationships between the fraction of mosquitoes transmitting virus for two genotypes of WNV and time and temperature, based on the statistical model in Table 2(WN02: Tr = (sin(8.00tT4/108))2; NY99: Tr = (sin(5.32tT4/108))2. Each curve shows the fraction of mosquitoes transmitting at a fixed time period after feeding on WNV-infected blood (4, 7 or 14 days) with points showing increasing temperatures (12°C to 32°C, symbol every 2°C). 10.1371/journal.ppat.1000092.t001 Table 1 Regression analysis (no intercept) of midgut infection, disseminated infection, and transmission after arc-sin square root transformation with Degree Days (DD) and a genotype (GT) by Degree Day interaction as predictors. Term Transmission p-value Disseminated Infection p-value Infection p-value DD*103 0.62±0.084 <0.0005 1.20±0.16 <0.0005 2.0±0.28 <0.0005 GT-DD*103 −0.27±0.12 0.029 −0.32±0.23 0.17 −0.11±0.39 0.77 Residual Error 3.41 12.9 37.5 Total Error 6.38 24.7 82.9 Coefficient±1SD is given. However, the residuals of regressions for all three dependent variables was significantly correlated with temperature (all p<0.001), suggesting that a degree day predictor using a linear product of temperature and incubation period (tT) was not fully capturing the temperature-dependent acceleration in infection and transmission. In the second statistical analysis, we found that both transmission and disseminated infection was best predicted by a model including DD with temperature raised to the 4th power (tT4), and a DD by genotype interaction (Table 2). The results were the same if the fraction transmitting was expressed as the fraction of infected mosquitoes transmitting (arc-sin square root transformed fraction of infected transmitting, DD: 9.09×10−8 tT4; p <0.0005; DD-genotype interaction: −3.30×10−8; p<0.0005). Infection was also best predicted by a model including DD with temperature raised to the 4th power (tT4), but was not significantly influenced by the DD by genotype interaction (Table 2). The residual error in these second set of regressions was substantially lower compared to the first statistical analysis, with the same number of independent variables (Tables 1,2). The significant negative coefficient for DD by genotype interaction term for transmission and disseminated infection again indicates that the fraction of Cx. pipiens infected with and transmitting genotype WN02 increased faster than mosquitoes transmitting NY99, as illustrated by the raw data (Figure 1), and the fitted relationships (Figure 2). Thus, WN02 would have a significant advantage over NY99 under warmer conditions after the same incubation period. 10.1371/journal.ppat.1000092.t002 Table 2 Regression analysis (no intercept) of midgut infection, disseminated infection, and transmission after arc-sin square root transformation with Degree Days (DD) and a genotype (GT) by Degree Day interaction as predictors, as in Table 1, except DD term was tT4 (t = days since feeding on WNV-infected blood; T = temperature). Term Transmission (tT4 ) p-value Disseminated Infection (tT4 ) p-value Infection (tT4 ) p-value DD*108 8.00±0.46 <0.0005 14.7±0.80 <0.0005 22.2±2.3 <0.0005 GT-DD*108 −2.68±0.65 <0.0005 −2.2±1.1 0.05 0.43±3.2 0.89 Residual Error 0.99 3.0 24.2 Total Error 6.38 24.7 82.9 Coefficient±1SD is given. Discussion The relationship between temperature and the transmission of pathogens has gained substantial attention recently, because projected changes in global temperature may increase the health burden of some diseases [42]. We have shown that, in the laboratory, increases in temperature have a two-fold impact on WNV transmission. First, as has been shown previously, increasing temperatures significantly increased viral infection, dissemination, and transmission, most likely through increased viral replication. Our study used the plaque assay which measured the presence of infectious virus and not the presence of unpackaged viral RNA, to test for infection, dissemination, and transmission in the mosquito. As a result, since the replication cycle is completed more quickly at higher temperatures, this will lead to greater concentration of infectious virus above the limit of detection in each mosquito. This is the case for replication in all tissues, and as such, increased temperature would affect not only infection kinetics, but dissemination and transmission kinetics as well. Second, warmer temperatures increased the advantage of the WN02 genotype over the NY99 genotype virus, and this advantage accelerated with temperature. Thus, the WN02 genotype appears to be better adapted to warmer temperatures than NY99, and NY99 was better adapted to warm conditions than a South African strain of WNV in Cx. tarsalis [28]. This result highlights the importance of understanding vector-pathogen-environment interactions and the role of pathogen evolution in influencing transmission. We also have shown that the advantage of WN02 over the NY99 genotype extends beyond day 7 post infection in Cx. pipiens, as we had observed in Cx. tarsalis [22]. The disparate results between our study and previous research that indicated no difference on day 9 [21] is likely due to extending the experiments past day 9 (up to day 40 at some temperatures) and including additional experiments resulting in much larger sample sizes. Nonetheless, our results support the earlier assertion that the WN02 genotype has an advantage over the NY99 genotype in the laboratory. Our results refine the WNV temperature-transmission relationship and show that WNV transmission in mosquitoes accelerates nonlinearly with the extrinsic incubation temperature, suggesting that even a small increase in temperatures can have a significant impact. They show that traditional degree day models for WNV may not accurately describe the impact of temperature on transmission. Instead, transmission may be more accurately modeled using degree day functions that include a temperature term raised to a power greater than 1. For WNV, we found that a degree day term with temperature raised to the fourth power, tT4 , was most accurate in explaining variation in transmission in our data. The implications of this difference are that even relatively small changes in temperature (e.g. the 2°C projected change in global temperatures [43]) have the potential to substantially increase transmission, and traditional degree day models used to investigate the potential impact of global warming will thus underestimate the effects of warming on transmission of WNV by mosquitoes. For example, if we fit a linear degree day model, tT, to our data, an increase from 28°C to 30°C would be predicted to increase temperature only 0.9% (from 12.1% to 13.0%), whereas the fitted model with tT4 this increase from 28°C to 30°C would actually increase transmission 7.8% (from 11.4% to 19.2%). In our study, we occasionally detected infectious virus in both salivary secretions and the legs of mosquitoes only 12 hours after feeding on an infected blood meal. Although under normal conditions the WNV replication cycle requires 10–12 hours [44], it is known that the virus replicates more quickly at higher temperatures [45]. Thus it may be possible that sufficient levels of replication took place in some mosquitoes held at 32°C to result in dissemination and transmission very quickly after feeding. It is equally possible that at high temperatures the cell junctions of the epithelium of the midgut were disrupted or increasingly permeable creating a rapid mechanism for midgut escape, possibly via leakage of virus, as has been observed with other virus-mosquito pairs [46]–[48]. This would have facilitated early escape of the virus to the legs, and subsequent infection of the salivary glands. Since the only mosquitoes that had virus in their salivary secretions were those that had virus in their legs, this argues against either mechanical transmission due to residual virus on the proboscis or regurgitation of virus from the midgut during the capillary transmission assay. Nonetheless, we cannot entirely rule out these other explanations, and furthermore, it is unlikely that mosquitoes would feed again 12 or 24 hours after the initial blood meal unless they had only obtained a partial or interrupted blood meal. It should be noted that our study did not evaluate the impact of mosquito rearing temperature, as all immature stages were maintained at 27°C. Previous work showed that vector competence for several flaviviruses, including Murray Valley [49], Japanese encephalitis [50], and St. Louis [51] encephalitis viruses, and dengue [52] and yellow fever [53] viruses, was depressed by maintaining adults at temperatures lower than those they experienced during larval development. In contrast, transmission of two alphaviruses, eastern equine encephalitis [54] and western equine encephalomyelitis [55], were not observed to decrease when adults were maintained at temperatures lower than the rearing temperature, and early season populations were considerably more susceptible to infection that those collected during midsummer. These results contribute to our broader understanding of how factors can generate spatial and temporal variation in transmission of pathogens. The transmission of WNV by Cx. pipiens has been shown to be influenced by host availability [12], mosquito genetic ancestry [56], and now the interaction of temperature and viral genotype. A key goal of future research will be to link the temperature-transmission patterns observed in the laboratory to patterns of transmission in the field. This should enable more accurate predictions of the impact of climate and climate change on the transmission of WNV and other vector borne pathogens.
Author and article information
Contributors
Eva Veronesi:
ORCID: http://orcid.org/0000-0003-3411-7891
+ 41 44 6358532
,
eva.veronesi@uzh.ch
Journal
Journal ID (nlm-ta): Parasitol Res
Journal ID (iso-abbrev): Parasitol. Res
Title:
Parasitology Research
Publisher:
Springer Berlin Heidelberg
(Berlin/Heidelberg
)
ISSN
(Print):
0932-0113
ISSN
(Electronic):
1432-1955
Publication date
(Electronic):
28
April
2018
Publication date PMC-release: 28
April
2018
Publication date
(Print):
2018
Volume: 117
Issue: 6
Pages: 1925-1932
Affiliations
[1
]ISNI 0000 0004 1937 0650, GRID grid.7400.3, National Centre for Vector Entomology, Institute of Parasitology, Vetsuisse Faculty,
, University of Zürich, ; Winterthurerstr. 266a, 8057 Zürich, Switzerland
[2
]GRID grid.417834.d, Present Address: Institute of Infectology, , Friedrich-Loeffler-Institut, ; Greifswald Isle of Riems, Greifswald, Germany
Author notes
Author information
Eva Veronesi
http://orcid.org/0000-0003-3411-7891
Article
Publisher ID: 5886DOI: 10.1007/s00436-018-5886-7
PMC ID: 5949136
PubMed ID: 29705877
SO-VID: 39d44fc1-efee-49c5-9ae2-d05234ea1e05
Copyright © © The Author(s) 2018
License:
Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.
History
Date
received
: 9
October
2017
Date
accepted
: 18
April
2018
Categories
Subject:
Original Paper
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issue-copyright-statement © Springer-Verlag GmbH Germany, part of Springer Nature 2018
ScienceOpen disciplines: Parasitology
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ScienceOpen disciplines: Parasitology