Shorebird low spillover risk of mosquito-borne pathogens on Iberian wetlands
Sara Pardal1, José A. Alves2, Líbia Zé-Zé3, Hugo Osório3, Afonso Rocha1, Ricardo J. Lopes4, Pete Potts5,
Fátima Amaro3, Francisco Santiago-Quesada6, Juan M. Sanchez-Guzman6, José Masero6, Maria J. Alves3,
Javier Pérez-Tris7, Jaime A. Ramos1 and Luisa Mendes1
IMAR – Marine and Environmental Research Center (IMAR/CMA), Department of Life Sciences,
University of Coimbra, Apartado 3046, 3001-401 Coimbra, Portugal
School of Biological Sciences, University of East Anglia, Norwich Research Park, Norwich, NR4 7TJ,
United Kingdom
Centre for the Study of Vectors and Infectious Diseases Doutor Francisco Cambournac, National Health
Institute Doutor Ricardo Jorge, IP, Av. Liberdade n°5, Apartado 2965-575 Águas de Moura, Portugal
CIBIO, Centro de Investigação em Biodiversidade e Recursos Genéticos, InBIO Laboratório Associado,
Universidade do Porto, 4485-661 Vairão, Portugal.
Farlington Ringing Group, Solent Court Cottage, Chilling Lane, Warsash, Southampton, SO31 9HF,
United Kingdom
Department of Anatomy, Cell Biology and Zoology, Faculty of Sciences, University of Extremadura,
06006 Badajoz, Spain
Departamento de Zoología y Antropología Física, Facultad de Biología, Universidad Complutense,
28040 Madrid, Spain
Corresponding author: Sara Pardal, e-mail address: [email protected]; telephone number: (+351)
Shorebirds were sampled in four Iberian wetlands (Fig. 1) during different stages of their annual
cycle: autumn migration (September – October), wintering (November – February), spring migration
(March-May) and breeding (June – August). With the exception of the Black-winged Stilt that is present
throughout the year, all studied species breed in northern Europe and winter on Sub-Saharan areas, using
Iberian wetland as staging during spring and autumn movements (Catry et al. 2011; Delany et al. 2009).
However, a small proportion of Common Redshank Tringa totanus, Common Greenshank Tringa
nebularia and Common Snipe Gallinago gallinago may remain on Iberia for the winter (Delany et al.
2009). Captured birds were marked with a numbered metal ring, aged and measured. A small blood
sample (160 µl) was taken from the braquial vein with a heparinized microhematocrit capillary tube. For
flaviviruses antibodies detection, approximately 80 µl of blood was transferred to a serological disc
Schleicher & Schuell (12.7 mm) and a small drop to a NUNC tube filled with a RNA Latter solution to be
used on RT-PCR screening. The serological discs were left to dry at ambient temperature and then kept at
-4°C. NUNC tubes stayed overnight at -4°C and then stored at -80°C until analysis. The remaining 80 µl
of blood was transferred to a 1.5 ml eppendorf filed with 96% ethanol for laboratory detection of
Haemosporidians screening
Total genomic DNA from the avian blood samples was extracted by standard ammonium acetate
protocol (Pérez-Tris and Bensch 2005), quantified in NanoDrop and diluted to our working concentration
(25 ng/µL). To determine good quality DNA to proceed for parasite identification, birds were sexed by
products were run on 2% agarose gel for band visualization and sex determination of each sample.
Samples were screened for haemosporidians using a two step nested PCR method. On both steps
PCR reactions were set up in 25 µL total volume including 25 ng of template DNA (tDNA), 1.5 mM
MgCl2, 2.5 µl of 10x PCR buffer II, 400 mM of each deoxynucleoside triphosphates, 0.6 mM of each
primer, and 0.625 U of AmpliTaq DNA polymerase (Applied Biosystems, Foster City, CA, USA). The
first step of the nested PCR consists in 20 cycles of DNA preamplification using the primers HaemNF
and HaemNR2 that combine with the outside of the target fragment. On the final step, the DNA is
amplified during 35 cycles using the primer pairs HaemF and HaemR2, designed to amplify and sequence
a partial fragment of about 479 base pairs (bp) of mtDNA cit b gene of Plasmodium and Haemoproteus
parasites. Afterwards 3 µL of the PCR products were set to run on 2% agarose gel stained with ethidium
bromide and TAE buffer.
Contaminations were controlled by including a negative control during extraction and positive
and negative controls (water) during PCR. No negative controls amplified. Negative controls were run
again on a second nested PCR to confirm results. Afterwards 3 µL of the PCR products were runned on
agarose gel stained with ethidium bromide and TAE buffer. Samples with amplification were precipitated
and sequenced. The output sequences were visualized and edited on FinchTV software (Geospiza Inc.,
Seattle, WA; Windows MacOS X, UNIX), aligned using ClustalW in BioEdit (Hall 1999) and MEGA4
(Tamura 2007) and compared with sequences stored at GenBank International Nucleotide Sequence
Database (NCBI) and MalAvi database (Bensch et al. 2009) for lineage identification.
ELISA assays
For flaviviruses antibodies detection, 900 µL of PBS solution was added to serological disc to
obtain a working concentration of 1:10 blood serum. Samples were screened by an in-house epitopeblocking ELISAs (Eb-ELISA), with monoclonal antibody MAb 3.1112G, according to Blitvich et al.
(2003). The percent inhibition of MAb binding was calculated and an inhibition value  30% was
considered to indicate the presence of viral antibodies. Moreover all samples were run again with a
competitive ELISA assay according to manufacturer’s instructions (ID Screen® West Nile Competition,
ID VET, Montpellier, France) specific to antibodies directed against the Pr-E envelope protein of WNV.
The ID VET diagnostic kit is designed to detect antibodies directed against Pr-E containing an
epitope common to Japanese Encephalitis antigenic complex. Thus, is expected that this assay can also
recognize other closely related specific antibodies produced by other flaviviruses of the same antigenic
complex such as WNV. The Eb-ELISA used in this study is directed to immunodominant epitopes on the
NS1 protein of flaviviruses, which does not require the use of multiple reporter or capture antibodies, and
showed to have good levels of specificity when the most efficient monoclonal antibody MAb 3.1112G
was used Blitvich et al. (2003).
From the 340 screened samples, the only twenty one positive samples were obtained with the
Eb-ELISA (Blitvich et al. 2003). All the samples including the ones with detectable antibody levels from
Eb-ELISA came negative with ELISA ID VET assay. We were therefore unable to identify which
flavivirus was present on the screened birds.
Bensch S, Hellgren O, Pérez-Tris J (2009) MalAvi: a public database of malaria parasites and related
haemosporidians in avian hosts based on mitochondrial cytochrome b lineages. Molecular Ecology
Resources 9: 1353 – 1358
Catry T, Alves J A, Andrade J, Costa H, Dias M P, Fernandes P, Leal A, Lourenço P M, Martins R C,
Moniz F, Pardal S, Rocha A, Santos C D, Encarnação V, Granadeiro J P (2011) Long-term declines of
wader populations at the Tagus estuary, Portugal: a response to global or local factors? Bird Conservation
International 21 (04): 438-453. doi: http://dx.doi.org/10.1017/S0959270910000626
Delany S, Scott D, Dodman T, Stroud D (2009) An Atlas of Wader Populations in Africa and Western
Eurasia. Wetlands International, Wageningen, The Netherlands
Hall T A (1999) BioEdit a user friendly biological sequence alignment editor and analysis program for
Windows 95/98/NT. Nucleic Acid Symp. 41: 95-98
Pérez-Tris J, Bensch S (2005) Diagnosing genetically diverse avian malarial infections using mixedsequence analysis and TA-cloning. Parasitology 131: 15-23
Tamura K, Dudley J, Nei M, Kumar S (2007) Mega4: Molecular Evolutionary Genetics Analysis
(MEGA) software version 4.0. Molecular Biology and Evolution 24: 1596-1599
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