<table><thead><tr><td>Country</td><td>Institute</td><td>Address</td></tr></thead><tbody><tr><td>Netherlands</td><td>National Institute for Public Health and the Environment (RIVM)</td><td>Antonie van Leeuwenhoeklaan 9, PO Box 1, 3720BA Bilthoven</td></tr><tr><td>Norway</td><td>Norwegian Institute of Public Health</td><td>PO Box 4404 Nydalen, (0)403 Oslo</td></tr><tr><td>Poland</td><td>National Institute of Public Health/National Institute of Hygiene</td><td>24 Chocimska Street, (00)791 Warsaw</td></tr><tr><td>Portugal</td><td>Directorate General of Health</td><td>Alameda D. Afonso Henriques, 45- 2º, 1049-005 Lisbon</td></tr><tr><td>Romania</td><td>National Institute of Public Health</td><td>Dr. Leonte Anastasievici 1-3, (0)50463 Bucharest</td></tr><tr><td>Slovakia</td><td>Public Health Authority of the Slovak Republic</td><td>Trnavská cesta 52, SK-826 45 Bratislava</td></tr><tr><td>Slovenia</td><td>National Institute of Public Health (NIJZ)</td><td>Trubarjeva 2, SL - 1000 Ljubljana</td></tr><tr><td>Spain</td><td>National Centre of Epidemiology, Health Institute Carlos III/ Instituto de Salud Carlos III: National Centre for Microbiology</td><td>Monforte de Lemos, 5, ES-28029 Madrid/ Ctra. Majadahonda a Pozuelo, Km. 2, 28220 Madrid</td></tr><tr><td>Sweden</td><td>Public Health Agency of Sweden</td><td>Nobels väg 18, Solna, 17182 Stockholm</td></tr></tbody></table>

## Estimated influence of environmental change on the disease future trends

TBEV circulation in nature require the co-occurrence of several biotic components (competent tick vectors, competent reservoirs, and feeding hosts for ticks) that interplay in a complex way and whose abundance, temporal and spatial dynamics are affected or modulated by abiotic factors (e.g. climatic condition, habitat structure, food availability, etc.) other than by biotic and socio-economic factors (trophic interaction, wildlife management, etc.). Climate change is expected to affect all these components, starting from the spatial and temporal distribution of *I. ricinus* complex (154-157), and consequently with an expected increase of the probability of TBE occurrence at higher altitude and northern latitude, beyond the current edges of its range (158, 159). The northern shift and increased tick activity are related to milder winters and longer spring and autumn seasons combined with increased vegetative cover and rise of ungulates abundance, as already observed in Sweden, Norway and in the Baltic countries (160). Similarly, in the Alps and in the Carpathian Mountains of Central Europe, an altitudinal shift of *I. ricinus* from about 700 to 1200 m asl has also been already reported (161, 162). Similarly, forest expansion and altered synchronized seed crops that affect density and distribution of rodents and wild ungulates and that in turn drive up the density of infected ticks has been reported (94, 163-165). In some cases, enhanced human exposure to infected ticks driven by changes in socio-economical condition have been considered the most important driver of upsurge in TBE cases (71). Estimating and predicting the transmission risk of TBEV under a global change scenario is essential for planning public health interventions including vaccination programs. Most of the proposed models infer TBE morbidity against some indicative environmental variables (e.g., mean annual temperature, precipitation, NDVI, etc.) or population estimates of key hosts (e.g., rodents or ticks), or most commonly combining both (166-169), in order to get an estimate of future trends.
TBE interannual fluctuations in the number of cases are not yet well parametrized, due to the
difficulties to obtain information such as the vaccination coverage and human behaviour. The
actual trend is toward an increasing, facilitated by the fact that the human population is ageing
and growing in numbers and with a higher probability of exposure. Moreover, the expected
extension of tick questing season, the periodic variations in food availability for key hosts and the
consequent increase of infected ticks through non-systemic transmission (co-feeding) (168),
support this trend. At the same time, it has also been reported that climate warming will to some
extent reduce TBE incidence in Southern and lowland areas (170) as well as the impact of extreme
meteorological events or prolonged drought conditions on ticks' occurrence (171).

On a large European scale, the affected countries might exhibit different and contrasting
oscillations of TBE human cases according to the matching of several cycles (e.g., 2 to 6 years
for the tick *I. ricinus*, 2 to 5 years for *muridae* rodents, 4-6 years for beech masting production,
quasi-decadal for large and medium-sized herbivores, some insectivores and predators, quasi-
biennial for some climatic oscillations, (172).

The complexity of the disease system, at this large-scale, makes a long-term prediction too
uncertain to be realistic therefore the use of validated covariates that can forecast the disease trend
1-2 years in advance would be enough for the health system to be prepared.

Finally, to assess the actual effective occurrence of TBE and therefore make more robust
predictions of the potential future trend under a climate change scenario, quantitative data related
to the aforementioned biotic factors (host community composition, host density etc.) and other
abiotic factors (such as land use) should be retrieved at a high-resolution spatial scale.

References

1. Pulkkinen LIA, Butcher SJ, Anastasina M. Tick-borne encephalitis virus: A structural view. *Viruses*. 2018;10(7):1-20.

2. Heinz FX, Mandl CW. The molecular biology of tick-borne encephalitis virus. Review article. *APMIS Acta Pathol Microbiol Immunol Scand*. 1993;101(10):735-45.

3. Gritsun TS, Holmes EC, Gould EA. Analysis of flavivirus envelope proteins reveals variable domains that reflect their antigenicity and may determine their pathogenesis. *Virus Res*. 1995;35(3):307-21.

4. McMinn PC. The molecular basis of virulence of the encephalitogenic flaviviruses. *J Gen Virol*. 1997;78(Pt 11):2711-22.

5. Kofler RM, Heinz FX, Mandl CW. Capsid protein C of tick-borne encephalitis virus tolerates large internal deletions and is a favorable target for attenuation of virulence. *J Virol*. 2002;76(7):3534-43.

6. Yoshii K, Yokozawa K, Kariwa H, et al. A conserved region in the prM protein is a critical determinant in the assembly of flavivirus particles. *J Gen Virol*. 2012;93(1):27-38.

7. Khasnatinov MA, Tuplin A, Gritsun DJ, et al. Tick-borne encephalitis virus structural proteins are the primary viral determinants of non-viraemic transmission between ticks whereas non-structural proteins affect cytotoxicity. *PLoS ONE*. 2016;11(6).

8. King AMQ, Adams MJ, Carstens EB, et al. Virus taxonomy, classification and nomenclature of viruses. *Ninth report of the International Committee of Taxonomic of Viruses*. Amsterdam: Elsevier, 2011.

9. Gritsun TS, Lashkevich VA, Gould EA. Tick-borne encephalitis. *Antiviral Res*. 2003;57(1-2):129-46.

10. Amato-Gauci A, Zeller H. Tick-borne encephalitis joins the diseases under surveillance in the European Union. *Euro Surveill*. 2012;17(42):20299.

11. Fares W, Dachraoui K, Cherni S, et al. Tick-borne encephalitis virus in Ixodes ricinus (Acari: Ixodidae) ticks, Tunisia. *Ticks Tick-Borne Dis*. 2020; 12:101606.
12. Růžek D, Dobler G, Mantke OD. Tick-borne encephalitis: Pathogenesis and clinical implications. *Travel Med Infect Dis.* 2010;8(4):223-32.

13. Leonova GN, Belikov SI. Phylogenetic analysis and distribution of far eastern tick-borne encephalitis virus subtype (Flaviridae, Flavirus, TBEV-FE) from Asia. *Vopr Virusol.* 2019;64(5):250-6.

14. Kovalev SY, Kokorev VS, Belyaeva I V. Distribution of Far-Eastern tick-borne encephalitis virus subtype strains in the former Soviet Union. *J Gen Virol.* 2010;91(12):2941-6.

15. Beauté J, Spiteri G, Warns-Petit E, et al. Tick-borne encephalitis in europe, 2012 to 2016. *Euro Surveill.* 2018;23(45).

16. Tkachev SE, Babkin I V, Belokopytova PS, et al. Genetic diversity and geographical distribution of the Siberian subtype of the tick-borne encephalitis virus. *Ticks Tick Borne Dis.* 2020;11(2):1877-9603.

17. Adelshin R V., Sidorova EA, Bondaryuk AN, et al. "886-84-like" tick-borne encephalitis virus strains: Intraspecific status elucidated by comparative genomics. *Ticks Tick-Borne Dis.* 2019;10(5):1168-72.

18. Kovalev SY, Mukhacheva TA. Reconsidering the classification of tick-borne encephalitis virus within the Siberian subtype gives new insights into its evolutionary history. *Infect Genet Evol J Mol Epidemiol Evol Genet Infect Dis.* 2017;20171100(55):159-65.

19. Kozlova V, Demina V, Tkachev E, et al. Characteristics of the Baikal subtype of tick-borne encephalitis virus circulating in eastern Siberia. *Acta Biomed Sci.* 2018;3(4):53-60.

20. Dai XY, Shang GB, Lu S, et al. A new subtype of eastern tick-borne encephalitis virus discovered in Qinghai-Tibet Plateau, China. *Emerg Microbes Infect.* 2018;25(7):1751-2222.

21. Weststrate AC, Knapen D, Laverman GD, et al. Increasing evidence of tick-borne encephalitis (TBE) virus transmission, the Netherlands, June 2016. *Euro Surveill.* 2017;22(11):1560-7917.

22. Demina T V., Dzhioev YP, Verkhozina MM, et al. Genotyping and characterization of the geographical distribution of tick-borne encephalitis virus variants with a set of molecular probes. *J Med Virol.* 2010;82(6):965-76.

23. Ruzek D, Avšič Županc T, Borde J, et al. Tick-borne encephalitis in Europe and Russia: Review of pathogenesis, clinical features, therapy, and vaccines. *Antiviral Research.* 2019; 164, p. 23-51.

24. Stoefs A, Heyndrickx L, De Winter J, et al. Autochthonous cases of tick-borne encephalitis, Belgium, 2020. *Emerg Infect Dis.* 2021 Aug;27(8):2179-82.

25. Mohareb E, Christova I, Soliman A, et al. Tick-borne encephalitis in Bulgaria, 2009 to 2012. *Euro Surveill.* 2013;14(18):1560-7917.

26. Mansbridge CT, Osborne J, Holding M, et al. Autochthonous tick-borne encephalitis in the United Kingdom: A second probable human case and local eco-epidemiological findings. *Ticks Tick-Borne Dis.* 2022;13(1):101853.

27. Labuda M, Kozuch O, Zuffová E, et al. Tick-borne encephalitis virus transmission between ticks cofeeding on specific immune natural rodent hosts. *Virology.* 1997;235(1):138-43.

28. Randolph SE, Miklisova D, Lysy J, et al. Incidence from coincidence: patterns of tick infestations on rodents facilitate transmission of tick-borne encephalitis virus. *Parasitology.* 1999;118(Pt 2):177-86.

29. Randolph SE, Green RM, Peacey MF, et al. Seasonal synchrony: the key to tick-borne encephalitis foci identified by satellite data. *Parasitology.* 2000;121(01):15-23.

30. Tonteri E, Jaaskelainen Anu E, Tikkakoski T, et al. Tick-borne encephalitis virus in wild rodents in winter, Finland, 2008-2009. *Emerg Infect Dis.* 2011;17(1):72-5.

31. Pukhovskaya NM, Morozova O V, Vysochina NP, et al. Tick-borne encephalitis virus in arthropod vectors in the Far East of Russia. *Ticks Tick-Borne Dis.* 2018;9(4):824-33.

32. Benda R. The common tick Ixodes ricinus L. as a reservoir and vector of tick-borne encephalitis virus, I, II. *J Hyg Epidemiol Microbiol Immunol.* 1958;2:314-44.
33. Süss J. Tick-borne encephalitis 2010: epidemiology, risk areas, and virus strains in Europe and Asia-an overview. *Ticks Tick-Borne Dis.* 2011;2(1):2-15.

34. Lichard M, Kozuch O. Persistence of tick-borne encephalitis virus in nymphs and adults ofIxodes arboricola and its transmission to white mice. *Acta Virol.* 1967;11(5):480.

35. Krivanec K, Kopecký J, Tomková E, et al. Isolation of TBE virus from the tick Ixodes hexagonus. *Folia Parasitol* (Praha). 1988;35(3):273-6.

36. Jemeršić L, Deždek D, Brnić D, et al. Detection and genetic characterization of tick-borne encephalitis virus (TBEV) derived from ticks removed from red foxes (Vulpes vulpes) and isolated from spleen samples of red deer (Cervus elaphus) in Croatia. *Ticks Tick-Borne Dis.* 2014;5(1):7-13.

37. Valarcher JF, Hagglund S, Juremalm M, et al. Tick-borne encephalitis. *Rev Sci Tech.* 2015;34(2):453-66.

38. Takeda T, Ito T, Chiba M, et al. Isolation of tick-borne encephalitis virus from Ixodes ovatus (Acari: Ixodidae) in Japan. *J Med Entomol.* 1998;35(3):227-31.

39. Kozuch O, Nosek J. Transmission of tick-borne encephalitis (TBE) virus by Dermacentor marginatus and D. reticulatus ticks. *Acta Virol.* 1971;15(4):334.

40. Nosek J. The ecology and public health importance of Dermacentor marginatus and D. reticulatus ticks in Central Europe. *Folia Parasitol* (Praha). 1972;19(1):93-102.

41. Abdiyeva K, Turebekov, N Yegemberdiyeva, et al. Vectors, molecular epidemiology and phylogeny of TBEV in Kazakhstan and central Asia. *Parasit Vectors*. 2020;1(13):504.

42. Lickova M, Fumacova Havlikova S, Slavikova M, et al. Dermacentor reticulatus is a vector of tick-borne encephalitis virus. *Ticks Tick Borne Dis.* 2020;11(4):1877-9603.

43. Chitimia-Dobler L, Lemhofer G, Dobler G, et al. Repeated isolation of tick-borne encephalitis virus from adult Dermacentor reticulatus ticks in an endemic area in Germany. *Parasit Vectors*. 2019;12(12):1756-3305.

44. Kozuch O, Nosek J. Experimental transmission of tick-borne encephalitis (TBE) virus by Haemaphysalis concinna ticks. Vol. 24, *Acta virologica*. Slovakia; 1980. p. 377.

45. Yun SM, Song BG, Choi W, et al. Prevalence of tick-borne encephalitis virus in ixodid ticks collected from the republic of Korea during 2011-2012. *Osong Public Health Res Perspect.* 2012;3(4):213-21.

46. Michelitsch A, Tews BA, Klaus C, et al. In vivo characterization of tick-borne encephalitis virus in bank voles (Myodes glareolus). *Viruses*. 2019;11(11):1069.

47. Lindquist L. Tick-borne encephalitis. *Handb Clin Neurol.* 2014;123:531-59.

48. Lindquist L, Vapalahti O. Tick-borne encephalitis. *The Lancet*. 2008;371(9627):1861-71.

49. Gerlinskaya LA, Bakhvalova VN, Morozova O V, et al. Sexual transmission of tick-borne encephalitis virus in laboratory mice. *Bull Exp Biol Med.* 1997;123(3):283-4.

50. Knap N, Avšič-Županc T. Factors affecting the ecology of tick-borne encephalitis in Slovenia. *Epidemiology and Infection*. 2015;143:2059-67.

51. Waldenström J, Lundkvist A, Falk KI, et al. Migrating birds and tickborne encephalitis virus. *Emerg Infect Dis.* 2007;13(8):1215-8.

52. Hubálek Z, Rudolf I. Tick-borne viruses in Europe. *Parasitol Res.* 2012;111(1):9-36.

53. Jaenson TGT, Petersson EH, Jaenson DGE, et al. The importance of wildlife in the ecology and epidemiology of the TBE virus in Sweden: incidence of human TBE correlates with abundance of deer and hares. *Parasit Vectors*. 2018;29(11):1756-3305.

54. Rieger MA, Nübling M, Müller W, et al. Foxes as indicators for TBE endemicity - A comparative serological investigation. *Zentralblatt Bakteriol.* 1999;289(5-7):610-8.
55. Haemig PD, Lithner S, de Luna SS, et al. Red fox and tick-borne encephalitis (TBE) in humans: Can predators influence public health? *Scand J Infect Dis.* 2008;40(6-7):527-32.

56. Radda A. Die Zeckenenzephalitis in Europa. *Z FürAngewandte Zool.* 1973;60:409-61.

57. Böhm B, Schade B, Bauer B, et al. Tick-borne encephalitis in a naturally infected sheep. BMC Vet Res. 2017; 22:13(1):1-6.

58. Pfeffer M, Dobler G. Tick-borne encephalitis virus in dogs - is this an issue? *Parasit Vectors.* 2011;59:1756-3305.

59. Fouché N, Oesch S, Ziegler U, et al. Clinical Presentation and laboratory diagnostic work-up of a horse with tick-borne encephalitis in Switzerland. *Viruses*. 2021;13(8).

60. Conze TM, Bagó Z, Revilla-Fernández S, et al. Tick-Borne Encephalitis Virus (TBEV) infection in two horses. *Viruses*. 2021;13(9).

61. Da Rold G, Obber F, Monne I, et al. Clinical tick-borne encephalitis in a roe deer (Capreolus capreolus L.). *Viruses*. 2022;14(2):300.

62. Bagó Z, Bauder B, Kolodziejek J, et al. Tickborne encephalitis in a mouflon (Ovis ammon musimon). *Vet Rec.* 2002;150(7):218-20.

63. Schönbächler K, Hatt JM, Silaghi C, et al. Confirmation of tick-borne encephalitis virus in an european hedgehog (Erinaceus europaeus). *Schweiz Arch Tierheilkd.* 2018;161(1):23-31.

64. Süss J, Gelpi E, Klaus C, et al. Tickborne encephalitis in naturally exposed monkey (Macaca sylvanus). *Emerg Infect Dis J.* 2007;13(6):905.

65. Dagostin F, Tagliapietra V, Marini G, et al. Ecological and environmental factors affecting the risk of tick-borne encephalitis in Europe, 2017 to 2021. *Euro Surveill.* 2023;28(42).

66. Nuttall PA. Climate change impacts on ticks and tick-borne infections. *Biologia* (Bratisl). 2022;77:1503-12.

67. Labuda M, Randolph SE. Survival strategy of tick-borne encephalitis virus: cellular basis and environmental determinants. *Zentralblatt Bakteriol Int J Med Microbiol.* 1999;289:5-7.

68. Perkins SE, Cattadori IM, Tagliapietra V, et al. Empirical evidence for key hosts in persistence of a tick-borne disease. *Int J Parasitol.* 2003;33(9).

69. Carpi G, Cagnacci F, Neteler M, et al. Tick infestation on roe deer in relation to geographic and remotely sensed climatic variables in a tick-borne encephalitis endemic area. *Epidemiol Infect.* 2008;136(10):1416-24.

70. Rosà R, Tagliapietra V, Manica M, et al. Changes in host densities and co-feeding pattern efficiently predict tick-borne encephalitis hazard in an endemic focus in northern Italy. *Int J Parasitol.* 2019;20190900(49):779-87.

71. Sumilo D, Asokliene L, Bormane A, et al. Climate Change Cannot Explain the Upsurge of Tick-borne Encephalitis in the Baltics. *PLoS One.* 2007;6(2):1932-6203.

72. Andreassen A, Jore S, Cuber P, et al. Prevalence of tick borne encephalitis virus in tick nymphs in relation to climatic factors on the southern coast of Norway. *Parasit Vectors.* 2012;22(5):1756-3305.

73. Randolph SE. Tick-borne encephalitis incidence in Central and Eastern Europe: consequences of political transition. *Microbes Infect.* 2008;10(3):209-16.

74. Jaenson TGT, Eisen L, Comstedt P, et al. Risk indicators for the tick Ixodes ricinus and Borrelia burgdorferi sensu lato in Sweden. *Med Vet Entomol.* 2009;23(3):226-37.

75. Barandika Jesus F, Hurtado A, Juste Ramon A, et al. Seasonal dynamics of Ixodes ricinus in a 3-year period in northern Spain: first survey on the presence of tick-borne encephalitis virus. *Vector Borne Zoonotic Dis.* 2010;10(10):1027-35.
76. Daniel M, Malý M, Danielová V, et al. Abiotic predictors and annual seasonal dynamics of Ixodes ricinus, the major disease vector of Central Europe. *Parasit Vectors*. 2015; 8(478):1-12.

77. Cuber P, Andreassen Å, Vainio K, et al. Risk of exposure to ticks (Ixodidae) and the prevalence of tick-borne encephalitis virus (TBEV) in ticks in Southern Poland. *Ticks Tick-Borne Dis*. 2015;6(3):356-63.

78. Randolph SE. Dynamics of tick-borne disease systems: minor role of recent climate change. *Rev Sci Tech Int Off Epizoot*. 2008;27(2):367-81.

79. Burri C, Bastic V, Maeder G, et al. Microclimate and the zoonotic cycle of tick-borne encephalitis virus in Switzerland. *J Med Entomol*. 2011;48(3):615-27.

80. Tagliapietra V, Rosà R, Arnoldi D, et al. Saturation deficit and deer density affect questing activity and local abundance of Ixodes ricinus (Acari, Ixodidae) in Italy. *Vet Parasitol*. 2011;183(1-2).

81. Brugger K, Schiffner-Rohe J, Walter M, et al. Modeling the Distribution of Tbev-Infected Ticks in Germany To Estimate Tbe-Infections in Humans. *Value Health*. 2018;21(1):S379.

82. Daniel M, Kriz B, Danielova V, et al. Correlation between meteorological factors and tick-borne encephalitis incidence in the Czech Republic. *Parasitol Res*. 2008;103(1):113-932.

83. Rosà R, Tagliapietra V, Manica M, et al. Changes in host densities and co-feeding pattern efficiently predict tick-borne encephalitis hazard in an endemic focus in northern Italy. *Int J Parasitol*. 2019;49(10):779-87.

84. Tkadlec E, Václavík T, Široký P. Rodent host abundance and climate variability as predictors of tickborne disease risk 1 year in advance. *Emerg Infect Dis*. 2019;25(9):1738-41.

85. Cagnacci F, Bolzoni L, Rosà R, et al. Effects of deer density on tick infestation of rodents and the hazard of tick-borne encephalitis. I: Empirical assessment. *Int J Parasitol*. 2012;42(4).

86. Bolzoni L, Rosa R, Cagnacci F, et al. Effect of deer density on tick infestation of rodents and the hazard of tick-borne encephalitis. II: population and infection models. *Int J Parasitol*. 2012;42(4):373-81.

87. Palo RT. Tick-borne encephalitis transmission risk: Its dependence on host population dynamics and climate effects. *Vector-Borne Zoonotic Dis*. 2014 May 1;14(5):346-52.

88. Vanwambeke SO, Sumilo DS, Bormane A, et al. Landscape predictors of tick-borne encephalitis in Latvia: land cover, land use, and land ownership. *Vector-Borne Zoonotic Dis*. 2010;10(5):497-506.

89. Kiffner C, Vor T, Hagedorn P, et al. Factors affecting patterns of tick parasitism on forest rodents in tick-borne encephalitis risk areas, Germany. *Parasitol Res*. 2011;108(2):323-35.

90. Knap N, Avsic-Zupanc T. Factors affecting the ecology of tick-borne encephalitis in Slovenia. *Epidemiol Infect*. 2015;143(10):2059-67.

91. Kriz B, Daniel M, Benes C, et al. The role of game (wild boar and roe deer) in the spread of tick-borne encephalitis in the Czech Republic. *Vector-Borne Zoonotic Dis*. 2014;14(11):801-7.

92. Kolář J, Potůčková M, Štefanová E. Tick-born encephalitis risk assessment based on satellite data. *Acta Univ Carol Geogr*. 2016;51(2):155-67.

93. Uusitalo R, Siljander M, Pellikka P, et al. Modelling habitat suitability for occurrence of human tick-borne encephalitis (TBE) cases in Finland. *Ticks Tick Borne Dis*. 2020;11(5):1877-9603.

94. Rizzoli A, Hauffe HC, Tagliapietra V, et al. Forest structure and roe deer abundance predict Tick-borne encephalitis risk in Italy. *PLoS ONE*. 2009;4(2).

95. Zeimes CB, Olsson GE, Hjertqvist M, et al. Shaping zoonosis risk: Landscape ecology vs. landscape attractiveness for people, the case of tick-borne encephalitis in Sweden. *Parasit Vectors*. 2014;7(1).

96. Stefanoff P, Rubikowska B, Bratkowski J, et al. A predictive model has identified tick-borne encephalitis high-risk areas in regions where no cases were reported previously, Poland, 1999-2012. *Int J Environ Res Public Health*. 2018 Apr 4;15(4).
97. Kjaer Lene J, Klitgaard K, Bodker R, et al. Predicting and mapping human risk of exposure to ixodes ricinus nymphs using climatic and environmental data, Denmark, Norway and Sweden, 2016. *Euro Surveill.* 2019;24(9):1560-7917.

98. Keesing F, Ostfeld RS. Impacts of biodiversity and biodiversity loss on zoonotic diseases. *Proc Natl Acad Sci U S A.* 2021;118(17):1-8.

99. McCallum H, Dobson A. Disease, habitat fragmentation and conservation. *Proc Biol Sci.* 2002;269(1504):2041-9.

100. Allan BF, Keesing F, Ostfeld RS. Effect of forest fragmentation on Lyme Disease risk. *Conserv Biol.* 2003;17(1):267-72.

101. Krauss J, Bommarco R, Guardiola M, et al. Habitat fragmentation causes immediate and time-delayed biodiversity loss at different trophic levels. *Ecol Lett.* 2010;13(5):597-605.

102. Wilkinson DA, Marshall JC, French NP, et al. Habitat fragmentation, biodiversity loss and the risk of novel infectious disease emergence. *J R Soc Interface.* 2018;15(149).

103. Sumilo D, Bormane A, Asokliene L, et al. Socio-economic factors in the differential upsurge of tick-borne encephalitis in Central and Eastern Europe. *Rev Med Virol.* 2008;18(2):81-95.

104. Zlobin VI, Pogodina V V., Kahl O. A brief history of the discovery of tick-borne encephalitis virus in the late 1930s (based on reminiscences of members of the expeditions, their colleagues, and relatives). *Ticks and Tick-borne Dis.* 2017; 8(6):813-20.

105. Yoshii K, Song JY, Park SB, et al. Tick-borne encephalitis in Japan, Republic of Korea and China. *Emerg Microbes Infect.* 2017;20(6):1751-2222.

106. Daniel M, Rudenko N, Golovchenko M, et al. The occurrence of Ixodes ricinus ticks and important tick-borne pathogens in areas with high tick-borne encephalitis prevalence in different altitudinal levels of the Czech Republic Part II. Ixodes ricinus ticks and genospecies of Borrelia burgdorferi sensu lato complex. *Epidemiol Mikrobiol Imunol.* 2016;65(3):182-192.

107. Labuda M, Danielova V, Jones LD, et al. Amplification of tick-borne encephalitis virus infection during co-feeding of ticks. *Med Vet Entomol.* 1993;7(4):339-42.

108. Kunz C. Vaccination against TBE in Austria: the success story continues. *Int J Med Microbiol.* 2002;291 Suppl:56-7.

109. Kohl O, Kožuch O, Elečková E, et al. Family outbreak of alimentary tick-borne encephalitis in Slovakia associated with a natural focus of infection. *Eur J Epidemiol.* 1996;12(4):373-5.

110. Gresíková M, Sekeyová M, Stúpalová S, et al. Sheep milk-borne epidemic of tick-borne encephalitis in Slovakia. *Intervirology.* 1975;5(1-2):57-61.

111. Holzmann H, Aberle SW, Stiasny K, et al. Tick-borne encephalitis from eating goat cheese in a mountain region of Austria. *Emerg Infect Dis.* 2009;15(10):1671-3.

112. Caini S, Szomor K, Ferenczi E, et al. Tick-borne encephalitis transmitted by unpasteurised cow milk in western Hungary, September to October 2011. *Euro Surveill.* 2012;22(12):17-21.

113. Hudopisk N, Korva M, Janet E, et al. Tick-borne encephalitis associated with consumption of raw goat milk, Slovenia, 2012. *Emerg Infect Dis.* 2013;19(5):806-8.

114. Lipowski D, Popiel M, Perlejewski K, et al. A cluster of fatal Tick-borne encephalitis virus infection in organ transplant setting. *J Infect Dis.* 2017;215(6):896-901.

115. Ličková M, Havlíková SF, Sláviková M, et al. Alimentary infections by tick-borne encephalitis virus. *Viruses.* 2022;14(1).

116. Martello E, Gillingham EL, Phalkey R, et al. Systematic review on the non-vectorial transmission of Tick-borne encephalitis virus (TBEv). *Ticks Tick-Borne Dis.* 2022;13(6).

117. Gritsun TS, Frolova TV, Zhankov AI, et al. Characterization of a siberian virus isolated from a patient with progressive chronic tick-borne encephalitis. *J Virol.* 2003;77(1):25-36.
118. Ternovoi VA, Kurzhukov GP, Sokolov YV, et al. Tick-borne encephalitis with hemorrhagic syndrome, Novosibirsk region, Russia, 1999. *Emerg Infect Dis.* 2003;9(6):743-6.

119. Mickiene A, Laiskonis A, Günther G, et al. Tickborne encephalitis in an area of high endemicity in Lithuania: disease severity and long-term prognosis. *Clin Infect Dis.* 2002;35(6):650-8.

120. Mickiene A, Pakalniene J, Nordgren J, et al. Polymorphisms in chemokine receptor 5 and Toll-like receptor 3 genes are risk factors for clinical tick-borne encephalitis in the Lithuanian population. *PloS One.* 2014;9(9):1932-6203.

121. Kaiser R. Tick-borne encephalitis: Clinical findings and prognosis in adults. *Wien Med Wochenschr.* 2012;162(11-12):239-43.

122. Zajkowska J, Czupryna P, Pancewicz S, et al. Fatal outcome of tick-borne encephalitis - a case series. *Neurol Neurochir Pol. 2011;45(4):402-6.*

123. Barkhash AV, Voevoda MI, Romaschenko AG. Association of single nucleotide polymorphism rs3775291 in the coding region of the TLR3 gene with predisposition to tick-borne encephalitis in a Russian population. *Antiviral Res.* 2013;99(2):136-8.

124. Barkhash AV, Babenko VN, Voevoda MI, et al. Association of IL28B and IL10 gene polymorphism with predisposition to tick-borne encephalitis in a Russian population. *Ticks Tick Borne Dis.* 2016;7(5):808-12.

125. Barkhash AV, Perelygin AA, Babenko VN, et al. Variability in the 2'-5'-oligoadenylate synthetase gene cluster is associated with human predisposition to tick-borne encephalitis virus -induced disease. *J Infect Dis.* 2010;15(202):1813-8.

126. Barkhash AV, Perelygin AA, Babenko VN, et al. Single nucleotide polymorphism in the promoter region of the CD209 gene is associated with human predisposition to severe forms of tick-borne encephalitis. *Antiviral Res.* 2012;93(1):64-8.

127. Barkhash AV, Yurchenko AA, Romaschenko AG, et al. A matrix metalloproteinase 9 (MMP9) gene single nucleotide polymorphism is associated with predisposition to tick-borne encephalitis virus -induced severe central nervous system disease. *Ticks Tick Borne Dis.* 2018;9(9):763-7.

128. Rushton JO, Lecollinet S, Hubalek Z, et al. Tick-borne encephalitis virus in horses, Austria, 2011. *Emerg Infect Dis.* 2013;19(4):635-7.

129. Müller K, König M, Thiel HJ. (Tick-borne encephalitis (TBE) with special emphasis on infection in horses). *DTW Dtsch Tierarztl Wochenschr.* 2006;113(4):147-51.

130. Barrett PN, Schober-Bendixen S, Ehrlich HJ. History of TBE vaccines. *Vaccine.* 2003;21(SUPPL. 1):S41-9.

131. Harabacz I, Bock H, Jüngst C, et al. A randomized phase II study of a new tick-borne encephalitis vaccine using three different doses and two immunization regimens. *Vaccine.* 1992;10(3):145-50.

132. Girgsdies OE, Rosenkranz G. Tick-borne encephalitis: development of a paediatric vaccine. A controlled, randomized, double-blind and multicentre study. *Vaccine.* 1996;14(15):1421-8.

133. Zavadska D, Anca I, Andre F, et al. Recommendations for tick-borne encephalitis vaccination from the Central European Vaccination Awareness Group (CEVAG). *Hum Vaccines Immunother.* 2013;9(2):362-74.

134. Zavadska D, Odzelevica Z, Liepina L, et al. Tick-borne encephalitis: A 43-year summary of epidemiological and clinical data from Latvia (1973 to 2016). *PloS One.* 2018;13(11):1932-6203.

135. Demicheli V, Debalini MG, Rivetti A. Vaccines for preventing tick-borne encephalitis. *Cochrane Database Syst Rev.* 2009;1(59):1469-493X.

136. Kollaritsch H, Paulke-Korinek M, Holzmann H, et al. Vaccines and vaccination against tick-borne encephalitis. *Expert Rev Vaccines.* 2012;11(9):1103-19.
137. Xing Y, Schmitt HJ, Arguedas A, et al. Tick-borne encephalitis in China: A review of epidemiology and vaccines. *Vaccine*. 2017;1(35):1227-37.

138. Divé I, Veje M, Dobler G, et al. Tick-borne encephalitis virus (TBEV) infection in pregnancy: Absence of virus transmission to the fetuses despite severe maternal disease - A case study. *Ticks Tick-Borne Dis*. 2020;11(5).

139. Bjonholm E, Soderholm S, Stephansson O, et al. Tick-borne encephalitis in pregnant women: A mini narrative review. *New Microbes New Infect*. 2022;48:101017.

140. Kerlik J, Avdičová M, Musilová M, et al. Breast milk as route of tick-borne encephalitis virus transmission from mother to infant. *Emerg Infect Dis*. 2022;28(5):1060-1.

141. European Centre for Disease Prevention and Control. Tick-borne encephalitis. In: ECDC. Annual epidemiological report for 2020. European Centre for Disease Prevention and Control. Stockholm; 2022.

142. Randolph SE, Asokliene L, Avsic-Zupanc T, et al. Variable spikes in tick-borne encephalitis incidence in 2006 independent of variable tick abundance but related to weather. *Parasit Vectors*. 2008;1(1):1-18.

143. Gonzalez G, Bournez L, Moraes RA, et al. A One-Health Approach to Investigating an Outbreak of Alimentary Tick-Borne Encephalitis in a Non-endemic Area in France (Ain, Eastern France): A Longitudinal Serological Study in Livestock, Detection in Ticks, and the First Tick-Borne Encephalitis Virus. *Front Microbiol*. 2022;13.

144. Šumilo D, Bormane A, Vasilenko V, et al. Upsurge of tick-borne encephalitis in the Baltic States at the time of political transition, independent of changes in public health practices. *Clin Microbiol Infect*. 2009;15(1):75-80.

145. Godfrey ER, Randolph SE. Economic downturn results in tick-borne disease upsurge. *Parasit Vectors*. 2011;15(4):1756-3305.

146. Lalani T, Yun H, Tribble D, et al. A comparison of compliance rates with anti-vectorial protective measures during travel to regions with dengue or chikungunya activity, and regions endemic for Plasmodium falciparum malaria. *J Travel Med*. 2016;23(5).

147. Hasler T, Fehr J, Held U, et al. Use of repellents by travellers: A randomised, quantitative analysis of applied dosage and an evaluation of Knowledge, Attitudes and Practices (KAP). *Travel Med Infect Dis*. 2019;28:27-33.

148. Borşan SD, Trif SR, Mihalca AD. Recreational behaviour, risk perceptions, and protective practices against ticks: a cross-sectional comparative study before and during the lockdown enforced by the COVID-19 pandemic in Romania. *Parasit Vectors*. 2021;14(1):1-15.

149. Barp N, Di Nuzzo M, Contini C, et al. Clinical and laboratory findings in tick-borne encephalitis virus infection. *Parasite Epidemiol Control*. 2020;10(19):2405-6731.

150. Šmit R, Postma MJ. The burden of tick-borne encephalitis in disability-adjusted life years (DALYs) for Slovenia. *PLoS ONE*. 2015;10(12).

151. Fafangel M, Cassini A, Colzani E, et al. Estimating the annual burden of tick-borne encephalitis to inform vaccination policy, Slovenia, 2009 to 2013. *Euro Surveill*. 2017;22(16).

152. Europe. Commission Implementing Decision (EU) 2018/945 of 22 June 2018 on the communicable diseases and related special health issues to be covered by epidemiological surveillance as well as relevant case definitions. *Official Journal of the European Union* L 170, 6/7/2018.

153. Bogovic P, Strle F. Tick-borne encephalitis: A review of epidemiology, clinical characteristics, and management. *World J Clin Cases*. 2015;16(3):430-41.

154. Jaenson TGT, Jaenson DGE, Eisen L, et al. Changes in the geographical distribution and abundance of the tickIxodes ricinus during the past 30 years in Sweden. *Parasit Vectors*. 2012;10(8):1756-3305.
155. Medlock JM, Hansford KM, Bormane A, et al. Driving forces for changes in geographical distribution ofIxodes ricinus ticks in Europe. *Parasit Vectors*. 2013;6(1)1-11.

156. Alkishe AA, Peterson AT, Samy AM. Climate change influences on the potential geographic distribution of the disease vector tick Ixodes ricinus. *PLoS ONE*. 2017;12(12):1932-6203.

157. Rizzoli A. Changes expected in Ixodes ricinus temporal and spatial distribution in Europe. In: Nuttall P (Ed.). *Climate, ticks and diseases*. Wallingford, UK: CABI; 2021. p. 171-5.

158. Danielova V, Kliegrova S, Daniel M, et al. Influence of climate warming on tickborne encephalitis expansion to higher altitudes over the last decade (1997-2006) in the Highland Region (Czech Republic). *Cent Eur J Public Health*. 2008;16(1):4-11.

159. Suss J. Tick-borne encephalitis in Europe and beyond--the epidemiological situation as of 2007. *Euro Surveill*. 2008;26(13):1560-7917.

160. Lindgren E, Talleklint L, Polfeldt T. Impact of climatic change on the northern latitude limit and population density of the disease-transmitting European tick Ixodes ricinus. *Environ Health Perspect*. 2000;108(2):119-23.

161. Garcia-Vozmediano A, Krawczyk AI, Sprong H, et al. Ticks climb the mountains: ixodid tick infestation and infection by tick-borne pathogens in the Western Alps. *Ticks Tick-Borne Dis*. 2020;1(5):101489.

162. Daniel M, Danielova V, Kriz B, et al. Shift of the tick Ixodes ricinus and tick-borne encephalitis to higher altitudes in Central Europe. *Eur J Clin Microbiol Infect Dis*. 2003;22(5):327-8.

163. Krawczyk AI, Duijvendijk GLA van, Swart A, et al. Effect of rodent density on tick and tick-borne pathogen populations: consequences for infectious disease risk. *Parasit Vectors*. 2020;13(1):34.

164. Bregnard C, Rais O, Voordouw MJ. Climate and tree seed production predict the abundance of the European Lyme disease vector over a 15-year period. *Parasit Vectors*. 2020;13(1):1-12.

165. Bregnard C, Rais O, Voordouw MJ. Masting by beech trees predicts the risk of Lyme disease. *Parasit Vectors*. 2021;14(1):1-22.

166. Haemig PD, Sjöstedt De Luna S, Grafström A, et al. Forecasting risk of tick-borne encephalitis (TBE): Using data from wildlife and climate to predict next year's number of human victims. *Scand J Infect Dis*. 2011;43(5):366-72.

167. Palo RT. Tick-borne encephalitis transmission risk: Its dependence on host population dynamics and climate effects. *Vector-Borne Zoonotic Dis*. 2014;14(5):346-52.

168. Nah K, Bede-Fazekas Á, Trájer AJ, et al. The potential impact of climate change on the transmission risk of tick-borne encephalitis in Hungary. *BMC Infect Dis*. 2020;20(1).

169. Hönig V, Švec P, Marek L, et al. Model of risk of exposure to lyme borreliosis and tick-borne encephalitis virus-infected ticks in the border area of the Czech republic (South Bohemia) and Germany (lower bavaria and upper palatinate). *Int J Environ Res Public Health*. 2019;16(7).

170. Brabec M, Daniel M, Malý M, et al. Analysis of meteorological effects on the incidence of tick-borne encephalitis in the Czech Republic over a thirty-year period. *Virol Res Rev*. 2017;1(1):1-8.

171. Danielova V, Daniel M. Climate, ticks and tick-borne encephalitis in Central Europe. In: Nuttall P (Ed.). *Climate, ticks and diseases*. Wallingford, UK: CABI; 2021. p. 331-40.

172. Zeman P. Predictability of tick-borne encephalitis fluctuations. *Epidemiol Infect*. 2017;145:2781-6.
# TULARAEMIA

Maria Bellenghi (a), Claudia Cataldo (a), Alessandra Ciervo (b), Francesca Dagostin (c), Giovanni Marini (c), Annapaola Rizzoli (c), Valentina Tagliapietra (c), Luca Busani (a)

(a) Centro di Riferimento per la Medicina di Genere, Istituto Superiore di Sanità, Rome

(b) Dipartimento Malattie Infettive, Istituto Superiore di Sanità, Rome

(c) Centro Ricerca e Innovazione, Fondazione Edmund Mach, San Michele all'Adige, Trento

## Biological, ecological and molecular features
of the causative agent

### Disease name

Tularaemia.

### Disease agent

#### Common, scientific and Latin name

*Francisella tularensis* is the causative microbial agent of tularaemia, a rare zoonotic disease that affects many animal species and humans. *F. tularensis subsp. tularensis* is extremely virulent, whereas *F. philomiragia* and *F. tularensis subsp. novicida* are opportunistic pathogens, causing disease in immunocompromised patients.

### Taxonomy

The taxonomic tree of *F. tularensis* is:

- Domain: *Bacteria*

- Phylum: *Proteobacteria*

- Class: *Alphaproteobacteria*

- Order: *Thiotrichales*

- Family: *Francisellaceae*

- Genus: *Francisella*

*F. tularensis* includes three subspecies of significant clinical relevance for humans i.e., subspecies *tularensis* (type A), subspecies *holarctica* (type B), and subspecies *mediasiatica*. The genus *Francisella* also involves opportunistic pathogens that can cause disease in immunocompromised patients (i.e., *F. novicida*, *F. hispaniensis*, and *F. philomiragia*). In addition, the genus *Francisella* includes *Francisella*-like endosymbionts (FLE) found in ticks and ciliates (i.e., *F. persica*, *F. endociliophora*, and *F. adeliensis*).

### Disease agent characteristics

*F. tularensis* is a small (0.7-1.7 µm), Gram-negative coccobacillus, strict aerobes, non-motile, non-spore-forming, heterotrophic, oxidase negative and requires cysteine for growing. This microorganism is easy to spread by aerosol and due to its low infectious dose and its virulence, is included in the Centers for Disease Control and prevention (CDC) category A of the bioterrorism agent (https://emergency.cdc.gov/agent/agentlist-category.asp).
## Physiochemical properties

Francisella is associated with different clinical and environmental sources, in particular water environment. Experimental studies suggest a long-term survival from 1 to 70 days in various water environments. F. tularensis survival seems to be influenced by both water temperature and salinity. In water milieu F. tularensis subsp. holarctica remains cultivable up to 28 days at 8°C. Both type A and type B strains remain cultivable up to 8-10 days in fresh-water, or 30-42 days in brackish-water at 21°C. F. tularensis can be metabolically active in water but not cultivable in vitro on agar plates. This microbial ability was observed for up to 140 days in water. This condition is defined as ‘Viable But Non-Culturable’ (VBNC) and could be responsible for long-term survival of bacteria in the water environment. The VBNC state has been defined as a state from which bacterial cells cannot be cultured but maintain a metabolic activity and cellular integrity. In addition, the VBNC state may be reversible, as bacteria may become cultivable under favorable conditions. This reversion in the ability to grow on agar plates is called ‘resuscitation’ of VBNC bacteria. F. tularensis subsp. holarctica possesses a mechanosensitive channel that protects this bacterium from hypoosmotic shock when it is released from an infected animal to water. In vitro studies have demonstrated that both F. tularensis subsp. holarctica and F. tularensis subsp. tularensis can form biofilms which is compatible with its natural aquatic reservoir (1-7). Multiple experimental studies have focused on the interaction between Francisella species and several amoebae particular in Acanthamoeba castellanii. Francisella was localized within vacuoles in amoeba trophozoites but were also able to survive in amoebal cysts for several weeks. This latter finding suggested that amoeba could be a long-term reservoir of Francisella spp. in water environments. Mosquito larvae may also represent a long-term F. tularensis reservoir in the aquatic environment. It has been shown that these larvae present in water can ingest Francisellae and ingested bacteria can survive throughout the different maturation stages of these arthropods up to adult mosquitoes (8-9).

## Priority level for EU

Tularaemia is a communicable disease subjected to the Commission Implementing Decision (EU) 2018/945. Since 2008, data on human cases have been collected by the European Surveillance System (TESSy) and maintained by European Centre for Disease Control and Prevention (ECDC). In Europe, the surveillance of tularaemia is mandatory and surveillance systems are comprehensive with full national coverage in all EU Member States. In 2021, a total of 876 confirmed cases of tularaemia were reported among 26 Member States (10). In general Sweden is the country that reports the highest number of cases, followed by Finland. Many European countries have reported sporadic cases, imported cases or outbreaks in the past 30 years. The disease has been identified in new areas, with an increasing tendency to emerge and/or re-emerge locally in Europe. The range of known hosts has also expanded to include animal not previously linked to Francisella, such as red fox (Vulpes vulpes), wild boar (Sus scrofa), and raccoon dog (Nyctereutes procyonoides). However, surveillance systems in the European countries are not the same, consequently data should be interpreted with caution.

## Distribution of the pathogen

Tularaemia is widely distributed in humans, wildlife and arthropod vectors, it is endemic throughout most of European countries, Northern and Central Asia, and North America. The maps
in Figures 1 and 2 represent the distribution of reported cases of *F. tularensis* from human, animal or arthropod vectors. Compared to previously known maps, the pathogen has been found in some new countries in Africa and Asia.

<figure><img src="image_59.png" /><figcaption>**Figure 1. Global distribution of *F. tularensis* detected in human, animal and the environment (from 2000 to 2020)**</figcaption></figure>

<figure><img src="image_60.png" /><figcaption>**Figure 2. European distribution of *F. tularensis* detected in human, animal and the environment (from 2000 to 2020)**</figcaption></figure>
# Ecology and transmission routes

The distribution and maintenance of *F. tularensis* may include a system of small natural foci associated to a physical (climate, geology) and biological (flora, fauna) element and to a different mode of transmission.

In this scenario, the pathogen is maintained and can be diffused for long time. The pathogen distribution and its ability to persist in many different ecological niches is related to the high adaptability to different animal reservoirs, vectors and environmental conditions.

In nature, *F. tularensis* is characterized in a terrestrial and aquatic lifecycle. The terrestrial stage involves different animal reservoir, while the aquatic cycle includes mainly mosquitoes, mosquito larvae, and aquatic rodents. In particular, environmental conditions, the aquatic lifecycle may be prevalent over the terrestrial lifecycle, amplifying the persistence, the spreading and the maintenance of the microbial agent (11).

*F. tularensis* has been detected in a high number of wild species including lagomorphs, rodents, insectivores, carnivores, ungulates, marsupials, birds, amphibians, fish, and invertebrates.

## Vectors

*F. tularensis* natural infections have been documented in different arthropods, although only a subset of these have been identified as important in *F. tularensis* transmission to humans. Arthropods found infected in nature include ticks of the genera *Amblyomma*, *Dermacentor*, *Ixodes* and *Ornithodoros*, mosquitoes of the genera *Aedes*, *Culex*, *Anopheles* and *Ochlerotatus excrucians*, and flies from the *Tabanidae* family (*Tabanus* spp., *Chrisozona* spp. and *Chrisops* spp.). Ticks appear to play an important key role in the ecology of tularaemia among arthropods, as they may carry pathogens over several years and life stages, thus maintaining enzootic tularaemia foci between epizootic periods. The genus *Dermacentor reticulatus* seems to be the most frequent carrier of *F. tularensis* in Central Europe. In Sweden, mosquito-borne infections, which are related to the aquatic life cycle, are a common route for transmission of tularaemia in humans and can lead to major outbreaks. Studies have also provided evidence that mosquito larvae may be infected with *F. tularensis* via water, possibly by ingesting predatory protozoa. In Table 1 is reported the list of arthropod vectors for this pathogen. Specific bibliography for vectors (8, 9, 12-27, 29-37).

## Reservoirs

The ecology of tularaemia is not sufficiently clarified because of the different potential host species involved, and may depend on geographical factors, as well as on the susceptibility and sensitivity of respective organisms to the pathogen. Recent observations provide evidence that most of the animals do not serve as amplifying hosts, but are “incidental” dead-end hosts, and therefore may not play a role in the persistence and spread of the pathogen.

The European hare (*Lepus europaeus*), also known as the brown or field hare, is considered an important host of *F. tularensis* and a common vector for the pathogen’s transmission to humans in Europe. In case of chronic infection, European hares may act as long-term reservoir for *F. tularensis*, persistent risk of transmission to humans, either directly or via vectors. Experimental studies showed that infected voles may also show a protracted course of disease representing a prolonged source of environmental contamination. Specific bibliography for reservoirs (1, 14, 16, 21, 22, 34, 35, 38-45).
<table><caption>Table 1. List of arthropod vectors (%) for *F. tularensis* data analysis from literature</caption><thead><tr><td>Vector</td><td>% positive</td><td>Method</td><td>Tested</td></tr></thead><tbody><tr><td>Aedes cinereus</td><td>4.2-42.9</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Aedes intrudens</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Aedes punctor</td><td>20.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Aedes spp.</td><td>20.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Aedes sticticus</td><td>50.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Aedes vexans</td><td>50.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Amblyomma americanum</td><td>0.0</td><td>Pools</td><td>&gt;300</td></tr><tr><td>Amblyomma maculatum</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Amblyomma spp.</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Culex pipiens/torrentium</td><td>0-100.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Culiseta alaskaensis</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Culiseta annulata</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Dermacentor marginatus</td><td>0-33.3</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Dermacentor reticulatus</td><td>0-0.3</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Dermacentor reticulatus</td><td>0.7-0.9</td><td>EXP INF</td><td>&gt;300</td></tr><tr><td>Dermacentor variabilis</td><td>0.0</td><td>Pools</td><td>50-300</td></tr><tr><td>Haemaphysalis concinna</td><td>0.3</td><td>Pools</td><td>&gt;300</td></tr><tr><td>Haemaphysalis inermis</td><td>0.0</td><td>Pools</td><td>50-300</td></tr><tr><td>Haemaphysalis parva</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Haemaphysalis punctata</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Haemaphysalis sulcata</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Hyalomma aegyptium</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Hyalomma excavatum</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Hyalomma marginatum</td><td>0.0</td><td>Pools</td><td>50-300</td></tr><tr><td>Hyalomma marginatum marginatum</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Hyalomma marginatum rufipes</td><td>0.0</td><td>Pools</td><td>50-300</td></tr><tr><td>Hyalomma spp.</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Ixodes ricinus</td><td>0-2.2</td><td>Pools</td><td>50-300</td></tr><tr><td>Ixodes scapularis</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Ixodes spp.</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Rhipicephalus bursa</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr><tr><td>Rhipicephalus turanicus</td><td>0.0</td><td>Pools</td><td>&lt;50</td></tr></tbody></table>

Pools: results from the testing of pools of arthropods; EXP INF: experimental infection

# Drivers of disease emergence and spread

## Ecological drivers

### Water

Long-term survival of *F. tularensis* in various water environments has been confirmed by several studies. Consumption or contact with surface water from natural springs, ponds or rivulets, or from wells have been identified as significant risk factors for tularaemia infection. Water salinity and levels of macro- and micronutrients are considered important factors that can contribute to the survival of *F. tularensis* in water environments. Experimental studies demonstrated that survival was enhanced in brackish water compared to freshwater, probably because higher concentration of sulphur residues and salinity, that are usually required for the cultivation of *F. tularensis*. Because brackish-water and normal saline support the culturability of
*F. tularensis* compared to what observed in freshwater, salinity may be an important factor for stability; however, it is likely that the presence of sulphur would enhance the effects of salinity.

## Presence of animals

Indirect evidence of the role of animals was the association between mouse and hare faeces and risk of infection during two outbreaks in Kosovo. During the outbreaks, faeces of small rodents were regularly found by the investigation teams in products stored in food stores linked to affected households. It is of interest that presence or proximity of domestic animals is protective towards the risk of *F. tularensis* detection (5, 23).

In Table 2, 3 and 4 the main *F. tularensis* vector environmental and animal drivers are reported.

<table><caption>Table 2. List of *F. tularensis* vector covariates identified from the selected studies in decreasing order according to the number references</caption><thead><tr><td>Vector drivers</td><td>N. of papers# (n. 45)</td><td>% of impact* (n. 51)</td></tr></thead><tbody><tr><td>Tick</td><td>40</td><td>78.4%</td></tr><tr><td>Mosquito</td><td>8</td><td>15.7%</td></tr><tr><td>Other</td><td>3</td><td>5.9%</td></tr></tbody></table>

# number of papers extracted, the number of reference per covariate is higher because from one paper more covariates could have been extracted * % calculated on the total number of references.

<table><caption>Table 3. List of *F. tularensis* environmental covariates identified from the selected studies in decreasing order according to the number references</caption><thead><tr><td>Environmental drivers</td><td>N. of papers# (n. 31)</td><td>% of impact* (n. 36)</td></tr></thead><tbody><tr><td>Water (brackish/saline or fresh water)</td><td>16</td><td>44.44%</td></tr><tr><td>Land use (distance from animal market or main road; percentage grassland; sediment)</td><td>7</td><td>19.44%</td></tr><tr><td>Location (city or rural areas)</td><td>4</td><td>11.11%</td></tr><tr><td>Time</td><td>2</td><td>5.5%</td></tr><tr><td>Altitude</td><td>2</td><td>5.5%</td></tr><tr><td>Air</td><td>1</td><td>2.77%</td></tr><tr><td>Chemical characteristic</td><td>1</td><td>2.77%</td></tr><tr><td>Humidity</td><td>1</td><td>2.77%</td></tr><tr><td>Precipitation (total rainfall)</td><td>1</td><td>2.77%</td></tr><tr><td>Soil</td><td>1</td><td>2.77%</td></tr></tbody></table>

# number of papers extracted, the number of reference per covariate is higher because from one paper more covariates could have been extracted

* % calculated on the total number of references.

<table><caption>Table 4. List of *F. tularensis* animal covariates identified from the selected studies in decreasing order according to the number references</caption><thead><tr><td>Animal drivers</td><td>N. of papers# (n. 69)</td><td>% of impact* (n. 115)</td></tr></thead><tbody><tr><td>Rodent</td><td>28</td><td>24.3%</td></tr><tr><td>Hare</td><td>19</td><td>16.3%</td></tr><tr><td>Canidae</td><td>10</td><td>8.7%</td></tr><tr><td>Mustelidae</td><td>6</td><td>5.2%</td></tr><tr><td>Swine</td><td>6</td><td>5.2%</td></tr><tr><td>Deer</td><td>5</td><td>4.35%</td></tr><tr><td>Host</td><td>5</td><td>4.35%</td></tr><tr><td>Ursidae</td><td>5</td><td>4.35%</td></tr><tr><td>Bird</td><td>4</td><td>3.48%</td></tr></tbody></table>
<table><thead><tr><td>Animal drivers</td><td>N. of papers# (n. 69)</td><td>% of impact* (n. 115)</td></tr></thead><tbody><tr><td>Bovidae</td><td>4</td><td>3.48%</td></tr><tr><td>Rabbit</td><td>4</td><td>3.48%</td></tr><tr><td>Felid</td><td>3</td><td>2.6%</td></tr><tr><td>Sheep</td><td>3</td><td>2.6%</td></tr><tr><td>Bat</td><td>2</td><td>1.74%</td></tr><tr><td>Mammals</td><td>2</td><td>1.74%</td></tr><tr><td>Monkeys</td><td>2</td><td>1.74%</td></tr><tr><td>Anatidae</td><td>1</td><td>0.87%</td></tr><tr><td>Badger</td><td>1</td><td>0.87%</td></tr><tr><td>Camelidae</td><td>1</td><td>0.87%</td></tr><tr><td>Lynx</td><td>1</td><td>0.87%</td></tr><tr><td>Opossum</td><td>1</td><td>0.87%</td></tr><tr><td>Skunk</td><td>1</td><td>0.87%</td></tr><tr><td>Viverrid</td><td>1</td><td>0.87%</td></tr></tbody></table>

# number of papers extracted, the number of reference per covariates is higher because from one paper more covariates could have been extracted *% calculated on the total number of references.

## Land cover

Land cover is an important risk factor for arthropod vectors of *F. tularensis* as far as it can ensure the suitability is to live in newly urbanized/suburban areas, residences surrounded by areas dominated by grassland vegetation. A serosurvey in Rural Azerbaijan identified possible associations between village-level tularaemia seroprevalence and suitable tick habitats, annual rainfall, precipitation in the driest quarter, and altitude (25, 46, 47).

## Economical and socio-demographic drivers

Any difference was observed in the risk for females and males, but there is a gender issue related to the risk of professional exposure. Some activities, like hunting, wood cutting or other outdoor activities are associated to higher risk of infection. On the contrary, contact with farming animals or agriculture practices are considered at lower risk. The infection is more frequent in adults and young adults over 14 years of age, but in case of waterborne outbreaks, age ranged from 2 to 81 years can be included. The disease is more common in people residing in rural area compared to those who live in cities, and among risk factor associated to the infection, arthropod bites, drinking water from springs, handling dead animals, or having rodents in the house and surroundings are the most frequently reported. Specific bibliography for economical and socio-demographic drivers (5, 32, 47-56).

## Natural history of disease in humans and animals, including symptoms, morbidity and mortality

### Brief history of the pathogen and disease

*F. tularensis* has been recognized as a human pathogen since the beginning of the 20th century. Reports of diseases strongly resembling to tularaemia disease preceded to first confirmed report in 1911. For example, there were cases reported from Utah in 1908, from Norway of a disease called lemming fever during the 1890s, and of Yato-Byo (hare disease) from Japan as early as 1818. There is also a description from Norway from 1653 of a tularaemia-like disease of
lemmings. The first human cases were reported in 1914 in Ohio. As suggested by Edward Francis, the etiological agent became known as *Bacterium tularensen*, named after Tulare County where the disease was endemic among rodents. A similar disease was reported under different names from various regions of the United States during the next decade. Francis published a report in 1919 entitled "Deer-fly fever – a disease of man of hitherto unknown etiology." The same author wrote a comprehensive review in 1921, in which he suggested that the disease was transmitted to man from rodents by bites of blood-sucking insects. Moreover, he proposed the disease be named tularaemia. In the commemoration of the pioneering work of Francis, the bacterium eventually received its present-day name, *F. tularensis*. For many years, *F. tularensis* has been considered as a potential biological weapon and, in the 1940s and 1950s, it was one of the agents that was given the highest priority in the offensive programs of the United States and the Soviet Union, and it was stockpiled by the U.S. military even in the late 1960s. The high costs and high morbidity justify the inclusion of *F. tularensis* as an agent of the highest priority together with, for example, anthrax, smallpox, and pandemic influenza. Accordingly, it is one of six agents designated as a category A Select Agent by CDC. These traits are also reflected by the fact that *F. tularensis* is highly contagious for laboratory workers (57-59).

## Disease in humans

*F. tularensis* can be transmitted to humans via different routes, such as direct contact with infected animals (e.g., during animal processing, through ingestion of uncooked meat, and animal bites), arthropods (ticks, horseflies, and mosquitos) and through the consumption of contaminated water or inhalation of contaminated soil, occurring in farming works. The human to human contagious was not reported until now, and the tularaemia disease is transmitted to humans by direct contact with infected animals, tissues or fluids from infectious animals, by ingestion of contaminated water and/or food, or by inhalation of infective aerosols. Early diagnosis and treatment may be difficult because of the aspecific clinical manifestations and its similarity to a wide variety of acute infectious diseases. The acute phase of the disease is characterized by rapidly bacteria proliferation; the severity of the disease depends on the patient's immunological response. This response depends on cell-mediated immunity rather than antibodies, which are good indicators of exposure to the bacterium but do not play a crucial role in protection. The mean incubation period of tularaemia is 3-5 days, but may range from 1-21 days. Onset of tularaemia is sudden, occurring 1 to 10 (usually 2 to 4) days after exposure, with headache, chills, nausea, vomiting, fever of 39.5 to 40° C, and severe prostration. Extreme weakness, recurring chills, and drenching sweats develop. Illness ranges from mild to life-threatening.

Main forms of this disease are listed below:

- Ulceroglandular

This is the most common form of tularaemia and usually occurs following a tick or deer fly bite or after handing of an infected animal. A skin ulcer, accompanied by swelling of regional lymph nodes, appears at the site where the bacteria entered the body.

- Glandular

It is similar to ulceroglandular tularaemia but without the presence of ulcer. Similarly, is acquired through the bite of an infected tick or deer fly or from handling sick or dead animals.

- Oculoglandular

This form occurs when the bacteria enter through the eye. This can occur when a person is butchering an infected animal and touches his or her eyes. Symptoms include irritation and inflammation of the eye and swelling of lymph glands.
- Oropharyngeal

This form results from eating or drinking contaminated food or water. Patients with oropharyngeal tularaemia may have sore throat, mouth ulcers, tonsillitis, and swelling of lymph glands in the neck.

- Typhoidal

This form is characterized by general symptoms without the localizing ones of other syndromes.

- Pulmonary

This is the most severe or aerosolization of contaminated milieu form of tularaemia, occurring after inhalation. Symptoms include cough, chest pain, and difficulty breathing and occasional rales may be the only symptom in pneumonia. A dry, nonproductive cough is associated with a retrosternal burning sensation. In untreated cases, temperature remains elevated for 3 to 4 weeks and resolves gradually. Mediastinitis, lung abscess, and meningitis are rare complications.

Mortality is generally related to untreated cases and about 6% in untreated cases of ulceroglandular tularaemia; it is higher for type A infection and for typhoidal, septicemic, and pulmonary form. Death usually results from overwhelming infection, pneumonia, meningitis, or peritonitis. Relapses can occur in inadequately treated cases. One infection confers immunity.

## Disease in animals

Signs and symptoms of tularaemia in wild animals are not well documented and are mostly based on post-mortem examinations. The clinical presentation of tularaemia depends on the host species, subspecies of the bacteria, and route of infection. The incubation period is 1-10 days. Type A tularaemia is particularly pathogenic for lagomorphs, with fatal infections also reported in cats and nonhuman primates.

The most common finding upon necropsy is an enlarged spleen and pinpoint white necrotic lesions in the spleen and liver. The best-documented clinical cases are in domestic cats and dogs, captive monkeys, prairie dogs and laboratory animals. Sheep and cats may be subclinically infected or develop bacteraemia, fever, and respiratory infection. Cats may also develop ulceroglandular or oropharyngeal disease, presumably through exposure to infected prey items. Outbreaks in untreated lambs may have up to 15% mortality.

Clinical signs include:

- increased pulse and respiratory rates;

- coughing;

- diarrhea;

- oral ulceration;

- pollakiuria with lymphadenopathy and hepatosplenomegaly.

In particular, the European hare (Lepus europaeus), also known as the brown or field hare, is characterized by variable clinical courses ranging from acute deadly septicaemia to protracted courses with only subacute lesions in various organs. In case of chronic infection, histopathological examinations demonstrated differences in the pathogenicity of clade B.FTNF002-00 (subgroup of B.6 and specific for Western Europe) and clade B.13 (subgroup of B.12 and specific for Central and Eastern Europe): while infections with strains of clade B.13 were reported to be associated with polyserositis, affecting the kidneys, pleura, and pericardium, histopathological findings in hares infected with B.FTNF002-00 have been almost invariably characterized by splenitis and hepatitis. These results are in accordance with further observations
in experimentally infected rats, showing significant differences in weight loss, mortality rate, and time to recovery between the two genotypes.

Rodents are very susceptible to *F. tularensis*, and commonly present with severe infection, leading to early death. Nevertheless, experimental studies showed that infected voles may also show a protracted course of disease with chronic nephritis and bacteriuria and could therefore also serve as a prolonged source of environmental contamination.

# Availability of preventive, therapeutic and control measures, including licensed or pipeline vaccines

## Therapy in humans

Three situations are to be considered:

1. Accidental exposure: antibiotic treatment should be initiated within 24 h and a treatment period of 14 days is recommended with either ciprofloxacin and doxycycline;

2. Exposure most likely did not occur: an increased vigilance may be sufficient, including daily measurement of body temperature for 14 days and a readiness to treat if symptoms appear;

3. Incidental spread of *F. tularensis* by aerosol: potentially exposed persons should be instructed to be alert to the development of fever within 14 days of exposure, and treatment initiated if necessary.

The preferred drugs are Gentamicin (for moderate to severe disease), Doxycycline (for mild disease), Chloramphenicol (used only for meningitis because there are more effective and safer alternatives), and Ciprofloxacin (for mild disease). Continuous wet saline dressings are beneficial for primary skin lesions and may diminish the severity of lymphangitis and lymphadenitis. Surgical drainage of large abscesses is rarely necessary unless therapy is delayed. Intense headache usually responds to oral analgesics. No natural resistance in *F. tularensis* to antibiotics used for clinical therapy has been demonstrated. This is true for aminoglycosides, tetracyclines, chloramphenicol, and quinolones. Due to the potential use of *F. tularensis* for biological weapon, antibiotic resistance remains of concern. Consequently, methods for rapid determination of the susceptibility to various antibiotics, including aminoglycosides, tetracyclines, chloramphenicol, quinolones, and rifampicin are needed.

## Therapy in animals

Early treatment is important to minimize risk of fatality. Streptomycin, gentamicin, and tetracyclines are effective at recommended dosage levels. Gentamicin should be continued for 10 days. Because tetracycline and chloramphenicol are bacteriostatic, they should be continued for 14 days to minimize the risk of relapse. Because of the substantial sylvatic (wildlife and tick) component of the *Francisella* life cycle, control involves reducing arthropod infestations and limiting exposure to wildlife, for example by keeping cats and dogs indoors. In some jurisdictions, tularaemia in animals is reportable to public health authorities.

## Licensed or pipeline vaccines

There is currently no effective and safe vaccine available against *F. tularensis*.
## Other prevention measures

Prevention measures consist in avoiding ingestion, breathing and inoculation of the bacteria. This includes: avoiding drinking untreated surface water; using insect repellent and clothes covering legs and arms to avoid tick and mosquito bites; avoiding contact with dead animals, using gloves when handling wild animals especially skinning of diseased hares, wild rabbits and rodents; not mowing over sick or dead animals, cooking thoroughly game meat before eating; handling biological samples potentially contaminated with *F. tularensis* in biosafety level-3 (BSL-3) laboratories.

## Disease specific recommendations

No specific recommendations are reported for tularaemia.

## Epidemiological situation at different spatial scales: past and current trends

Tularaemia usually causes epizootics (local amplification with clusters of human cases), but in some areas there is an endemic trend, with regular occurrence of human infections. Sweden has consistent annual numbers of mosquito-borne infections, and Sweden, Finland and Norway reported about 66 percent of the entire number of cases notified in Europe between 2019 and 2020 (Figure 3).

<figure><img src="image_61.png" /><figcaption>Figure 3. Number of reported cases of tularaemia reported in 29 countries of the European Union and European Economic Area, 2019-2021 (data from http://atlas.ecdc.europa.eu/public/index.aspx)</figcaption></figure>
On the other hand, in Europe there are countries that did not notified cases in the last time period (UK, Ireland, and Iceland). Other European countries that consistently notified human cases were France, Hungary, the Czech Republic, and Germany, almost all with an increasing trend, probably due in part to better surveillance and increased public awareness. Outbreaks in humans are often sporadic and likely to be spatially and temporally variable, in addition to specific sites with potential for clusters of infection. Based on the reporting of human cases the major hotspots for tularaemia at the European level are located in Scandinavia and Central Europe. Data reporting in the period 2012-2020 (Figure 4) has been quite consistent through the years, and considering all the EU/EEA (European Union and European Economic Area) countries together, a slight increasing trend can be observed. Trends were more apparent in individual countries: Norway, France, Czech Republic and Germany showed a trend towards an increasing number of cases, while a trend of decreasing case numbers was apparent in Finland, Hungary and Slovakia, and no trends appeared in data from Sweden.

<figure><img src="image_62.png" /><figcaption>Figure 4. Reported cases of tularaemia reported in 29 countries of the European Union and European Economic Area, 2012-2021 (data from http://atlas.ecdc.europa.eu/public/index.aspx)</figcaption></figure>

Animal cases of tularaemia are voluntarily notified to the World Organization for Animal Health (WOAH), and in the period 2012-2020, 13 EU/EEA countries notified 1475 wild and domestic animal cases to the World Animal Health Information System (WAHIS). Germany notified more than 50 percent of cases and, together with Spain and Finland, accounted for 84 percent of total cases during the period (Figure 5).
<figure><img src="image_63.png" /><figcaption>Figure 5. Number of cases of tularaemia reported in animals in 13 countries of the European Union and European Economic Area, 2012–2020 (data from WOAH-WAHIS)</figcaption></figure>

Among the infected animal species, brown hare accounted for 64 percent and bank vole 10 percent. The other 26 species contributed less than 5% each. Most cases occurred in wildlife, but domestic animals, particularly dogs (4%), sheep (3%) and cattle (1%) also tested positive.

## Sociological and demographic dimension affecting susceptibility and exposure, including gender

Although the age-related incidence rate of tularaemia is unknown, the disease is known to occur at all ages. Males have a higher incidence in all age categories. In Sweden, an over representation among males has been attributed to their more frequent outdoor professional and leisure activities (60). In most countries where tularaemia is endemic, the disease is seasonal; its incidence seems to be highest during late spring, the summer months and early autumn (61-63). Often, the number of cases shows wide variations from one year to another and this is probably related to climatic factors such as temperature and precipitation.

However, there are virtually no data linking specific climatic conditions and outbreaks of tularaemia. This is an important area for future research that may yield important tools for predicting and possibly preventing outbreaks. Table 5 reports the most *F. tularensis* human covariates which impact on disease spreading, while in Figure 6 and 7 the gender and age distribution is shown.
<table><caption>**Table 5.** List of *F. tularensis* human covariates identified from the selected studies in decreasing order according to the number of references.</caption><thead><tr><td>Human drivers</td><td>n. of papers# (n. 23)</td><td>% of impact*(n. 77)</td></tr></thead><tbody><tr><td>Infection (rate of infections)</td><td>40</td><td>51.9%</td></tr><tr><td>Age (median age)</td><td>15</td><td>19.5%</td></tr><tr><td>Sex (male or female)</td><td>11</td><td>11%</td></tr><tr><td>Profession (healthcare workers, hunters, butchers, rangers, woodcutters)</td><td>7</td><td>9.1%</td></tr><tr><td>Outcomes (hospitalization rate or duration; fatality rate)</td><td>2</td><td>2.6%</td></tr><tr><td>Activity (schooling)</td><td>1</td><td>1.3%</td></tr><tr><td>Human density (n. of households)</td><td>1</td><td>1.3%</td></tr></tbody></table>

<figure><img src="image_64.png" /><figcaption>**Figure 6.** Distribution of *F. tularensis* % of cases by gender (data from TESSy-ECDC)</figcaption></figure>

<figure><img src="image_65.png" /><figcaption>**Figure 7.** Distribution of *F. tularensis* % of cases by age (data from TESSy-ECDC)</figcaption></figure>

<footnote># number of papers extracted, the number of reference per covariate is higher because from one paper more covariates could have been extracted *% calculated on the total number of references</footnote>
# Diagnostic procedures and notification systems used at local, national and European scale

## Diagnosis

Human: diagnosis of suspected tularaemia is based on a history of contact with rabbits, hares, rodents or exposure to arthropod vectors, the sudden onset of symptoms, and the characteristic primary lesion. Because this organism is highly infectious, samples and culture media from patients suspected of having tularaemia should be handled with extreme caution and, if possible, processed by a high-level biosafety containment-equipped laboratory with a level 3 rating.

Tularaemia can be difficult to diagnose. It is a rare disease, and the symptoms are aspecific and common to other illnesses. For this reason, it is important to share an epidemiology report and any likely exposures, such as tick and deer fly bites, or contact with sick or dead animals. The case definitions that describe the criteria for diagnosis of tularaemia are: suspect, presumptive and confirmed. The criteria for each are:

- Suspect: an exposure history consistent with risks known to be associated with tularaemia together with clinical symptoms compatible with tularaemia.

- Presumptive: suggestive clinical symptoms and a clinical sample that tests positive for tularaemia by antigen or DNA detection. A single positive serum is also considered presumptive.

- Confirmed: isolation and identification of *F. tularensis* from biological specimen, specific antigen or DNA detection.

The choice of specimen for diagnostic testing is dependent on the form of clinical illness; ulceroglandular, glandular, oculoglandular, oropharyngeal, respiratory, or typhoidal. The following specimens are acceptable for the various forms of illness as specified:

- Blood: Whole blood (for all clinical forms of illness);

- Serum: A first specimen should be collected in early course of infection, followed by a second specimen taken in the convalescent period (at least 14 days later and preferably 3-4 weeks after onset of symptoms). Serum is acceptable for all clinical forms of illness;

- Respiratory secretions: Pharyngeal swabs, bronchial/tracheal washes or aspirates, sputum, transthoracic lung aspirates, or pleural fluid collection (for respiratory, typhoidal, oropharyngeal forms of illness);

- Swabs: Swabs of visible lesions or affected areas should be collected (for ulceroglandular and oculoglandular forms of illness);

- Aspirates: Aspirates from lymph nodes or lesions (for ulceroglandular, glandular, and oropharyngeal forms of illness);

- Tissue biopsies: Tissue samples from lymph nodes (for ulceroglandular, glandular, and oropharyngeal forms of illness). Invasive sampling, such as incision of an affected lymph node, should be avoided during the acute stage of disease;

- Autopsy materials: Samples from visible abscesses and from lymph node, lung, liver, spleen, cerebrospinal fluid, and bone marrow.

Serology is commonly used for confirmation of tularaemia. Antibody responses against *F. tularensis* are generally detectable in patients 10–20 days post-infection. Agglutination, either microagglutination or tube agglutination, is the standard serological test used for determining the
presence of antibody to *F. tularensis*. More recently an Enzyme-Linked Immunosorbent Assay (ELISA) (directed against LipoPolySaccharide-LPS) combined with Western blot (against antigen extracted from whole killed cells) showed very good sensitivity and specificity for diagnosis of infection. Culture provides a conclusive diagnosis of infection and an invaluable resource for molecular epidemiology, subtyping and discovery of novel species and subspecies. Whenever possible, culture should be attempted. *F. tularensis* grows well on several types of cysteine/cystine-supplemented agar including enriched Chocolate Agar (CA), Cystine Heart Agar with 9% chocolatized Blood (CHAB), Buffered Charcoal Yeast Extract (BCYE), Thioglycollate-GlucoseBlood Agar (TGBA), and GC Agar II with 1% haemoglobin and 1% IsoVitaleX. A variety of molecular methods have been described for the molecular detection of *F. tularensis*. PCR can be a valuable diagnostic tool when organisms are non-cultivable or when culture is not recommended due to biosafety concerns. Consist conventional PCR assays targeted at the genes *fopA* or *tul4* encoding the outer membrane proteins. A multiplex real-time TaqMan PCR assay for *F. tularensis* has also been developed.

Animal: confirmation is by culture, serology, or PCR. When individual animals present with consistent clinical signs of septicemic disease, generalized or acute lymphadenopathy, or pneumonia, tularaemia must be considered a possible cause. Tularaemia should also be ruled out when large numbers of sheep show typical signs during periods of heavy tick infestation or when large numbers of rodents or lagomorphs are found dead. Affected sheep should be evaluated for tularaemia and tick paralysis, whereas the etiologic agents to consider in cats and small mammals should also include agents of plague and pseudotuberculosis. When tularaemia is suspected, laboratory personnel should be alerted as a precaution to reduce the risk of laboratory-acquired infection. The infective dose required to transmit this pathogen is extremely low; thus, risk of infection during necropsy or to laboratory personnel is significant, and special procedures and facilities are essential. Diagnosis of acute infection of tularaemia is confirmed by bacterial culture and identification of the bacterium, serology, or PCR. Organisms can be readily isolated from necropsy specimens by use of special media using stringent personal protection protocols. A direct or indirect fluorescent antibody test, or tube agglutination test with a single titer of ≥1:80, is presumptive evidence of exposure, whereas a 4-fold increase in antibody titer between acute and convalescent serum specimens confirms acute infection. PCR can be used to confirm infection rapidly.

The Commission Implementing Decision (EU) 2018/945 of 22 June 2018 on the communicable diseases and related special health issues to be covered by epidemiological surveillance as well as relevant case definitions.

## Infrastructure capacity to identify pathogens for each Member State

Tularaemia is among the communicable diseases that according to the Commission Implementing Decision (EU) 2018/945 are covered by epidemiological surveillance. It means that EU Member States are required to establish national capacity of detection and reporting of human cases. The decision provides a case definition and laboratory criteria for case confirmation that are at least one of the following three cases:

1) Isolation of *Francisella tularensis* from a clinical specimen.

2) Detection of *F. tularensis* nucleic acid in a clinical specimen

3) Detection of *F. tularensis* specific antibody response. Diagnosis is routinely made by clinical microbiology laboratories, and there is no specific European-wide reference
laboratory network or national laboratories in most EU countries. Due to the potential biological weapon role, in many countries there is the involvement of Defence Research Agencies or laboratories. In animals, the disease is not reportable at the EU level (Regulation (EU) 2016/429 and Commission Implementing Regulation (EU) 2020/2002) but is reportable at the international level to the World Organization for Animal Health (WOAH). Human surveillance is reported by each EU Member State to ECDC through the surveillance TESSy System.

## Estimated influence of environmental change on the disease future trends

Tularaemia is widely distributed in Europe and has local emergence and re-emergence trends in humans and wildlife. The disease has a clear seasonality in humans, but this pattern has not been conclusively demonstrated in wildlife. Assessment of eco-epidemiological trends and estimation of infection risk in relation to projected environmental and climate change are difficult. Different geographical areas have different ecosystems, which influence the epidemiology and disease presentation. Such factors include temperature and humidity, the presence of different types of arthropod vectors, and the variety of small rodent and other wildlife species present. The important role of vectors in tularaemia transmission in the northern Hemisphere and the seasonality of the disease mean that climate change may have an effect on tularaemia transmission patterns in highly endemic areas. Statistical models used to study climate change include mosquito abundance as the main variable; however, the role of animal reservoirs is not often considered in the estimates. Overall, in northern Europe, along a latitudinal gradient, climate change scenarios respectively indicate a future decrease in disease in some Southern areas and an increase in disease in more northern areas. For the rest of Europe, estimating the impact of climate on disease is more difficult. Among the aspects that need better understanding, knowledge of the specific variables that influence the activity of natural tularaemia foci in Europe is one that needs to be improved for effective monitoring of this disease.

## References

1. Keim P, Johansson A, Wagner DM. Molecular epidemiology, evolution, and ecology of Francisella. *Ann N Y Acad Sci.* 2007;1105:30-66.

2. Colquhoun DJ, Duodu S. Francisella infections in farmed and wild aquatic organisms. *Vet Res.* 2011;42:47.

3. Birkbeck TH, Feist SW, Verner-Jeffreys DW. Francisella infections in fish and shellfish. *J Fish Dis.* 2011;34:173-187.

4. Berrada ZL, Telford Iii SR. Survival of Francisella tularensis Type A in brackish-water. *Arch Microbiol.* 2011;193:223-226.

5. Broman T, Thelaus J, Andersson A-C, et al. Molecular detection of persistent Francisella tularensis subspecies holarctica in natural waters. *Int J Microbiol.* 2011;2011:851946.

6. Gilbert SE, Rose LJ. Survival and persistence of nonspore-forming biothreat agents in water. *Lett Appl Microbiol.* 2012;55:189-194.

7. Hennebique A, Boisset S, Maurin M. Tularemia as a waterborne disease: a review. *Emerg Microbes Infect.* 2019;8:1027-1042.
8. Bäckman S, Näslund J, Forsman M, et al. Transmission of tularemia from a water source by transstadial maintenance in a mosquito vector. *Sci Rep*. 2015;5.

9. Brugueras S, Fernández-Martínez B, Martínez-de la Puente J, et al. Environmental drivers, climate change and emergent diseases transmitted by mosquitoes and their vectors in southern Europe: A systematic review. *Environ Res*. 2020; 191.

10. European Food Safety Authority, European Centre for Disease Prevention and Control. The European Union One Health 2021 Zoonoses Report. *EFSA Journal*. 2022;20(12):7666:273 pp. https://doi.org/10.2903/j.efsa.2022.7666.

11. Hennebique A, Boisset S, Maurin M. Tularemia as a waterborne disease: a review. *Emerg Microbes Infect*. 2019;8:1027-1042.

12. Bonnet S, De La Fuente J, Nicollet P, et al. Prevalence of tick-borne pathogens in adult Dermacentor spp. Ticks from nine collection sites in France. *Vector Borne Zoonotic Dis*. 2013;13:226-236.

13. Duzlu O, Yildirim A, Inci A, et al. Molecular investigation of Francisella-like endosymbiont in ticks and Francisella tularensis in ixodid ticks and mosquitoes in Turkey. *Vector Borne Zoonotic Dis*. 2016;16:26-32.

14. Ebani VV, Rocchigiani G, Nardoni S, et al. Molecular detection of tick-borne pathogens in wild red foxes (Vulpes vulpes) from Central Italy. *Acta Trop*. 2017;172:197-200.

15. Bielawska-Drózd A, Cieślik P, Żakowska D, et al. Detection of Coxiella burnetii and Francisella tularensis in tissues of wild-living animals and in ticks of North-west Poland. *Pol J Microbiol*. 2018;67:529-534.

16. Tomaso H, Otto P, Peters M, et al. Francisella tularensis and other bacteria in hares and ticks in North Rhine-Westphalia (Germany). *Ticks Tick-Borne Dis*. 2018;9:325-329.

17. Gehringer H, Schacht E, Maylaender N, et al. Presence of an emerging subclone of Francisella tularensis holarctica in Ixodes ricinus ticks from South-Western Germany. *Ticks Tick-Borne Dis*. 2013;4:93-100.

18. Pistone D, Pajoro M, Novakova E, et al. Ticks and bacterial tick-borne pathogens in Piemonte region, Northwest Italy. *Exp Appl Acarol*. 2017;73:477-491.

19. Richard S, Oppliger A. Zoonotic occupational diseases in forestry workers - Lyme borreliosis, tularemia and leptospirosis in Europe. *Ann Agric Environ Med*. 2015;22:43-50.

20. Guardone L, Ebani VV, Verin R, et al. Molecular detection of arthropod-borne pathogens in Eurasian badgers (Meles meles) from the United Kingdom. *Animals*. 2020;10.

21. Gyuranecz M, Rigó K, Dán A, et al. Investigation of the ecology of francisella tularensis during an inter-epizootic period. *Vector Borne Zoonotic Dis*. 2011;11:1031-1035.

22. Lopes de Carvalho I, Toledo A, Carvalho CL, et al. Francisella species in ticks and animals, Iberian Peninsula. *Ticks Tick-Borne Dis*. 2016;7:159-165.

23. Janse I, van der Plaats RQJ, Husman AM de R, et al. Environmental Surveillance of Zoonotic Francisella tularensis in the Netherlands. *Front Cell Infect Microbiol*. 2018;8:2235-2988.

24. Toma L, Mancini F, Di Luca M, et al. Detection of microbial agents in ticks collected from migratory birds in central Italy. *Vector Borne Zoonotic Dis* 2014;14:199-205.

25. Egyed L, Elő P, Sréter-Lancz Z, et al. Seasonal activity and tick-borne pathogen infection rates of Ixodes ricinus ticks in Hungary. *Ticks Tick-Borne Dis*. 2012;3:90-94.

26. Wójcik-Fatla A, Zając V, Sawczyn A, et al. Occurrence of Francisella spp. in Dermacentor reticulatus and Ixodes ricinus ticks collected in eastern Poland. *Ticks Tick Borne Dis* 2015;6:253-257.

27. Lundström JO, Andersson A-C, Bäckman S, et al. Transstadial transmission of *Francisella tularensis holarctica* in mosquitoes, Sweden. *Emerg Infect Dis*. 2011;17:794-799.
28. Hubalek Z, Treml F, Juricova Z, et al. Serological survey of the wild boar (Sus scrofa) for tularaemia and brucellosis in South Moravia, Czech Republic. *Vet Med (Praha)*. 2002;47:2-3.

29. Hubálek Z, Rudolf I. *Francisella tularensis* prevalence and load in *Dermacentor reticulatus* ticks in an endemic area in Central Europe: *Francisella tularensis* in *D. reticulatus*. *Med Vet Entomol*. 2017;31:234-239.

30. Hodžić A, Fuehrer H-P, Duscher GG. First molecular evidence of zoonotic bacteria in ticks in Bosnia and Herzegovina. *Transbound Emerg Dis*. 2017;64:1313-1316.

31. Paul REL, Cote M, Le Naour E, et al. Environmental factors influencing tick densities over seven years in a French suburban forest. *Parasit Vectors*. 2016;9.

32. Dryselius R, Hjertqvist M, Mäkitalo S, et al. Large outbreak of tularaemia, central Sweden, July to September 2019. *Euro Surveill*. 2019;24(42):1900603. doi:10.2807/1560-7917.ES.2019.24.42.1900603.

33. Thelaus J, Andersson A, Broman T, et al. Francisella tularensis Subspecies holarctica occurs in Swedish mosquitoes, persists through the developmental stages of laboratory-infected mosquitoes and is transmissible during blood feeding. *Microb Ecol*. 2014;67:96-107.

34. Regier Y, Komma K, Weigel M, et al. Combination of microbiome analysis and serodiagnostics to assess the risk of pathogen transmission by ticks to humans and animals in central Germany. *Parasit Vectors*. 2019;12(1):11.

35. Pajoro M, Pistone D, Boccazzi IV, et al. Molecular screening for bacterial pathogens in ticks (Ixodes ricinus) collected on migratory birds captured in northern Italy. *Folia Parasitol (Praha)*. 2018;65.

36. Franke J, Fritzsch J, Tomaso H, Straube E, Dorn W, Hildebrandt A. Coexistence of pathogens in host-seeking and feeding ticks within a single natural habitat in Central Germany. *Appl Environ Microbiol*. 2010;76(20):6829-6836. doi:10.1128/AEM.01630-10.

37. Pascucci I, Di Domenico M, Dall'Acqua F, et al. Detection of lyme disease and Q fever agents in wild rodents in Central Italy. *Vector Borne Zoonotic Dis* 2015;15:404-411.

38. Hestvik G, Uhlhorn H, Westergren E, et al. Tularaemia in European brown hares (Lepus europaeus) and mountain hares (Lepus timidus) characterized by histopathology and immunohistochemistry: organ lesions and suggestions of routes of infection and shedding. *J Comp Pathol*. 2017;157:2-3.

39. Desvars A, Furberg M, Hjertqvist M, et al. Epidemiology and ecology of Tularemia in Sweden, 1984-2012. *Emerg Infect Dis*. 2015;21:32-39.

40. Otto P, Chaignat V, Klimpel D, et al. Serological Investigation of Wild Boars (Sus scrofa) and Red Foxes (Vulpes vulpes) As Indicator Animals for Circulation of Francisella tularensis in Germany. *Vector Borne Zoonotic Dis*. 2014;1:46-51.

41. Bártová E, Kučerová HL, Žákovská A, et al. Coxiella burnetii and Francisella tularensis in wild small mammals from the Czech Republic. *Ticks Tick Borne Dis*. 2020;11:101350.

42. Rossow H, Sissonen S, Koskela KA, et al. Detection of Francisella tularensis in voles in Finland. *Vector Borne Zoonotic Dis*. 2014;14:193-198.

43. Moinet M, Decors A, Mendy C, et al. Spatio-temporal dynamics of tularemia in French wildlife: 2002-2013. *Prev Vet Med* 2016;130:33-40.

44. Tadin A, Tokarz R, Markotic A, et al. Molecular survey of zoonotic agents in rodents and other small mammals in Croatia. *Am J Trop Med Hyg*. 2016;94:466-473.

45. Mihelčić M, Habuš J, Vucelja M, et al. Prevalence of *Francisella tularensis* in the population of small mammals species in continental forests of Croatia. *Šumar List*. 2018;142:481-485.

46. Richter WR, Sunderman MM, Wendling MQS, et al. Evaluation of altered environmental conditions as a decontamination approach for nonspore-forming biological agents. *J Appl Microbiol*. 2020;128:1050-1059.
47. Grunow R, Kalaveshi A, Kühn A, Mulliqi-Osmani G, Ramadani N. Surveillance of tularaemia in Kosovo, 2001 to 2010. *Euro Surveill.* 2012;17(28):20217. doi:10.2807/ese.17.28.20217-en.

48. Desvars A, Furberg M, Hjertqvist M, et al. Epidemiology and ecology of tularemia in Sweden, 1984-2012. *Emerg Infect Dis.* 2015;21(1):32-39. doi:10.3201/eid2101.140916.

49. Rossow H, Ollgren J, Hytonen J, et al. Incidence and seroprevalence of tularaemia in Finland, 1995 to 2013: regional epidemics with cyclic pattern. *Euro Surveill.* 2015;20(33):21209.

50. Jurke A, Bannert N, Brehm K, et al. Serological survey of Bartonella spp., Borrelia burgdorferi, Brucella spp., Coxiella burnetii, Francisella tularensis, Leptospira spp., Echinococcus, Hanta-, TBE- and XMR-virus infection in employees of two forestry enterprises in North Rhine-Westphalia, Germany, 2011-2013. *Int J Med Microbiol.* 2015;305(7):652-662. doi:10.1016/j.ijmm.2015.08.015.

51. Larssen KW, Afset JE, Heier BT, et al. Outbreak of tularaemia in central Norway, January to March 2011. *Euro Surveill.* 2011;16:19828.

52. De Keukeleire M, Vanwambeke SO, Cochez C, et al. Seroprevalence of borrelia burgdorferi, anaplasma phagocytophilum, and francisella tularensis infections in Belgium: Results of three population-based samples. *Vector Borne Zoonotic Dis.* 2017;17:108-115.

53. Jenzora A, Jansen A, Ranisch H, et al. Seroprevalence study of Francisella tularensis among hunters in Germany. *FEMS Immunol Med Microbiol. 2008;*53:183-9.

54. Zukiewicz-Sobczak W, Zwolinski J, Chmielewska-Badora J, et al. Prevalence of antibodies against selected zoonotic agents in forestry workers from eastern and southern Poland. *Ann Agric Environ Med.* 2014;21:767-770.

55. Tokarska-Rodak M, Weiner M, Szymanska-Czerwinska M, et al. Seroprevalence of selected zoonotic agents among hunters from Eastern Poland. *Pol J Microbiol.* 2018;67:233-236.

56. Rigaud E, Jaulhac B, Garcia-Bonnet N, et al. Seroprevalence of seven pathogens transmitted by the Ixodes ricinus tick in forestry workers in France. *Clin Microbiol Infect.* 2016;22:735.e1-735.e9.

57. Hirschmann JV. From squirrels to biological weapons: the early history of Tularemia. *Am J Med Sci.* 2018;356:319-328.

58. Maurin M, Gyuranecz M. Tularaemia: clinical aspects in Europe. *Lancet Infect Dis.* 2016;16:113-124.

59. Sjöstedt A. Tularemia: history, epidemiology, pathogen physiology, and clinical manifestations. *Ann N Y Acad Sci.* 2007;1105:1-29.

60. Eliasson H, Lindbäck J, Nuorti JP, et al. The 2000 Tularemia outbreak: a case-control study of risk factors in disease-endemic and emergent areas, Sweden. *Emerg Infect Dis.* 2002;8:956-960.

61. Olsufév NG. (Results and perspectives of the study of natural foci of tularemia in USSR). *Med Parazitol (Mosk).* 1977;46:273-282.

62. Centers for Disease Control and Prevention (CDC). Tularemia--United States, 1990-2000. *MMWR Morb Mortal Wkly Rep.* 2002;51(9):181-184.

63. Tärnvik A, Priebe H, Grunow R. Tularaemia in Europe: an epidemiological overview. *Scand J Infect Dis.* 2004;36:350-355.
# USUTU VIRUS INFECTION

Soushieta Jagadesh (a), Claudia Cataldo (b), Francesca Dagostin (c), Di Luca Marco (d), Annapaola Rizzoli (c), William Wint (e), Luca Busani (b)

(a) *International Society of Infectious Diseases, Boston (MA)*

(b) *Centro di Riferimento per la Medicina di Genere, Istituto Superiore di Sanità, Rome*

(c) *Centro Ricerca e Innovazione, Fondazione Edmund Mach, San Michele all'Adige, Trento*

(d) *Dipartimento Malattie Infettive, Istituto Superiore di Sanità, Rome*

(e) *Department of Biology, Environmental Research Group Oxford Ltd, Oxford*

## Biological, ecological, and molecular features of the causative agent

### Disease name

Usutu virus infection/ USUV infection.

### Disease agent

#### Common, scientific and Latin name

Usutu virus (USUV).

### Taxonomy

USUV, an RNA arbovirus, is member of the Japanese encephalitis serocomplex of the *Flaviviridae* family and of the *Flavivirus* genus (1-3), and is phylogenetically close to Japanese encephalitis virus (JEV) and West Nile virus (WNV) (4, 5).

Phylogenetic studies based on the nucleic acid sequence of the NS5 gene divided USUV strains isolated into eight lineages: three African and five European (6), and that the level of genetic relatedness was found dependent on their geographical origin and the host from which they have been isolated.

### Disease agent characteristics

USUV is an enveloped virus of approximately 40-60 nm in diameter, with a single-stranded positive sense RNA comprised of 11064 base pairs with a 5' N7-methylguanosine-triphosphate cap but lacking a polyA tail at the 3' end (7, 8).

The genome of USUV comprises a single open reading frame coding for a polyprotein of 3434 amino acids that, which following cleavage, generates three structural proteins (capsid C, pre membrane prM and envelope E) and eight non-structural (NS) proteins (NS1/NS1', NS2a, NS2b, NS3, NS4a, 2K, NS4b and NS5) (4). The capsid protein (C) forms the central body of the virion and is associated with the viral RNA. The prM protein is required for virion assembly and maturation of virions through the folding of the envelope glycoprotein (E) (9). The NS5 of the virus ensures viral RNA replication by its RNA-dependent RNA polymerase domain and a methyltransferase (MTase) domain, which catalyses the capping of new viral RNA molecules (7, 10).
## Priority level for EU

Usutu virus infection in humans is not a mandatory notifiable disease at the EU/EEA (European Union and European Economic Area) level.

The sequences of USUV strains obtained in Italy in 2009 from mosquitoes, birds and humans, demonstrated that the sequences obtained from human hosts clustered with the sequences obtained from birds, indicating an endemic distribution of USUV in Europe (11). In Europe, USUV was isolated for the first time in 2001 from dead blackbirds (*Turdus merula*) during an epizootic near Vienna, Austria (12-14). A retrospective analysis of the high mortality of blackbirds in Tuscany, Italy in 1996 was attributed to USUV (15). The virus has since been detected in epizootics in Western, Southern, and Central European countries in birds and mosquitoes (3, 16). Although USUV infections in humans remain asymptomatic, sporadic neuroinvasive disease cases have been reported in Europe, predominantly among immunocompromised and elderly patients (17, 18).

## Distribution of the pathogen

In 2015, USUV was reported from mosquitoes, birds or horses in 12 European countries (Germany, Austria, Belgium, Croatia, Spain, France, Greece, Hungary, Italy, the Czech Republic, Serbia and Switzerland) (19-23). USUV infection has also been serologically identified in Slovakia and in Poland in equine and avian populations (24, 25). In the summer of 2016, Belgium, France, Germany and for the first time, the Netherlands reported widespread USUV activity based on live and dead bird surveillance (6, 26, 27). In 2018, USUV spread rapidly across Western Europe suggesting a continuous geographical spread of the virus, along with the colonization of new ecological niches. As of September 2023, 15 out of 30 EU/EEA countries reported USUV circulation among avian and equine animals – Austria, Belgium (28, 29), Croatia, Czechia, France, Germany, Greece (30), Hungary, Italy, Luxembourg, the Netherlands, Poland, Slovakia, Slovenia and Spain (31, 32) (Figure 1) (33). USUV has been predominantly detected in ornithophilic mosquito species of the *Culex* genus – like *Cx. modestus*, *Cx. neavei*, *Cx. perexiguus*, *Cx. perfuscus*, *Cx. pipiens*, *Cx. quinquefasciatus*, *Cx. univittatus* – but also to other genera such as *Aedes albopictus*, *Ae. japonicus*, *Ae. minutus*, *Anopheles maculipennis*, *Culiseta annulata*, *Mansonia africana*, *Coquilletidia aurites*, *Ochlerotatus caspius* and *Oc. detritus*. *Cx. pipiens*, an ornithophilic species, which can also feed on mammals, including humans, is considered to be the main vector in Europe (34). Europe encompasses highly suitable areas for *Cx. pipiens*, ranging from Mediterranean regions to Northern countries (Figure 2). Previous studies have detected USUV in nine mosquito species across Europe (Table 1).

<table><caption>Table 1. Mosquito species found infected by USUV across Europe</caption><thead><tr><td>Species</td><td>Country (ref.)</td></tr></thead><tbody><tr><td>Aedes albopictus</td><td>Italy (35-37), Croatia (38)</td></tr><tr><td>Aedes japonicus</td><td>Austria (39)</td></tr><tr><td>Anopheles maculipennis</td><td>Italy (37)</td></tr><tr><td>Culex modestus</td><td>Czech Republic (40), Italy (41), Belgium (42)</td></tr><tr><td>Culex perexiguus</td><td>Spain (43)</td></tr><tr><td>Culex pipiens</td><td>Austria (39), France (22), Germany (21, 44, 45), Italy (35-37, 46-51), Serbia (20), Spain (52), Switzerland (53)</td></tr><tr><td>Culiseta annulata</td><td>Italy (37)</td></tr><tr><td>Ochlerotatus caspius</td><td>Italy (35, 37)</td></tr><tr><td>Ochlerotatus detritus</td><td>Italy (37)</td></tr></tbody></table>
<figure><img src="image_66.png" /><figcaption>Figure 1. Geographical distribution of Usutu virus infections in humans, animals (birds-equids) and mosquitoes in the European Union and European Economic Area</figcaption></figure>

<figure><img src="image_67.png" /><figcaption>Figure 2. Current 1-km probability of presence of Cx. pipiens across Europe, produced using random forest and boosted regression trees analyses (source ERGO group)</figcaption></figure>
## Ecology and transmission routes

As with a majority of arboviruses, the transmission dynamics of USUV is influenced by biological and environmental factors, such as distribution and population density of vector and reservoir species, the extrinsic incubation period, humidity, temperature, host immunity, etc. (8). USUV and WNV are genetically, antigenically, and epidemiologically closely related. There is a significant interest on the possibility of overlapping in transmission cycles influencing the spatiotemporal trends of circulation of the two viruses in Europe (1) as both viruses are mainly transmitted by *Culex* mosquitoes, with migratory birds acting as the major amplifying host. Viral RNA was detected simultaneously for both viruses (54, 55), confirming that co-infection does occur. However, the co-circulation and co-infection remains uncertain at a population level.

The transmission cycle of USUV involves the vectors, ornithophilic mosquitos and the amplifying hosts and reservoirs, birds (Figure 3).

<figure><img src="image_68.png" /><figcaption>Figure 3. Biological cycle of USUV infection</figcaption></figure>

Infected ornithophilic mosquitos feed on birds, often species belonging to *Passeriformes* and *Strigiformes* (56). The infected birds develop sufficient viremia to allow transmission of the virus to a new mosquito during a subsequent bite. Some mosquitos of the *Culex* species may also take their blood meal from a mammalian host including humans, equids dogs, etc.

## Reservoirs

USUV is maintained through an enzootic cycle, similar to WNV, between passerine birds mainly Eurasian blackbirds (*Turdus merula*) or magpies (*Pica pica*) and Strigiformes, such as the Great Gray Owl (*Strix nebulosa*) as amplifying hosts and ornithophilic mosquitoes as vectors. Thus, the main USUV natural hosts are birds, with infection being reported in 93 different species
belonging to 35 families (57). The most affected species is the Eurasian blackbirds (*Turdus merula*) followed by gray owls (*Strix nebulosa*), and house sparrows (*Passer domesticus*) (12, 58). Occasionally, USUV was detected in birds in captivity (59) in Germany (60) (i.e. marabou stock, ruddy shellducks, red-breasted geese, Humboldt penguins, laughing kookaburras, steamer ducks, greater flamingos, snowy owls, Ural owls, white storks, Egyptian vultures, and Eurasian eagle owls), France (61) (i.e. Abyssinian ground hornbills, common peafowls, emus, scarlet ibis, and greater rheas) and Slovenia (62) (i.e. pelicans, Eurasian eagle-owls, barn and snowy owls). The list of birds detected with USUV infections in Europe is summarized in Table 2 (1).

<table><caption>Table 2. Updated list of birds with USUV clinical infections – RT-PCR (+)</caption><thead><tr><th>Order</th><th>Common name</th><th>Scientific name</th><th>Countries with USUV</th></tr></thead><tbody><tr><td rowspan="15">Passeriformes</td><td>Blackbird</td><td><i>Turdus merula</i></td><td>Austria (13, 63, 64), Czech Republic (65), France (23), Germany (34, 66), Hungary (63), Italy (11, 41,67), Netherlands (26), Belgium (68), Switzerland (69)</td></tr><tr><td>Common starling</td><td><i>Sturnus vulgaris</i></td><td>Germany (34,66), Italy (11,41,67)</td></tr><tr><td>Song trush</td><td><i>Turdus philomelos</i></td><td>Austria (64), Germany (70) Spain (31)</td></tr><tr><td>Canary</td><td><i>Serinus canaria domestica</i></td><td>Germany (34)</td></tr><tr><td>House sparrow</td><td><i>Passer domesticus</i></td><td>Austria (13,64), Germany (66), Switzerland (69)</td></tr><tr><td>Blue (great) tit</td><td><i>Parus caeruleus (major)</i></td><td>Austria (13), Switzerland (69)</td></tr><tr><td>European greenfinch</td><td><i>Chloris chloris</i></td><td>Switzerland (69)</td></tr><tr><td>European robin</td><td><i>Erithacus rubecula</i></td><td>Austria (13), Switzerland (69)</td></tr><tr><td>Bullfinch</td><td><i>Pyrhula pyrrhula</i></td><td>Belgium (71)</td></tr><tr><td>Nuthatch</td><td><i>Sitta europaea</i></td><td>Austria (13)</td></tr><tr><td>Eurasian jay</td><td><i>Garrulus glandarius</i></td><td>Hungary (63), Italy (67)</td></tr><tr><td>Magpie</td><td><i>Pica pica</i></td><td>Italy (11,72)</td></tr><tr><td>Barn swallows</td><td><i>Hirundo rustica</i></td><td>Austria (14)</td></tr><tr><td rowspan="10">Strigiformes</td><td>Great grey owl</td><td><i>Strix nebulosa</i></td><td>Austria (13), Germany (60,70,73), Switzerland (69), France (74), Italy (75), Netherlands (26)</td></tr><tr><td>Long-eared owl</td><td><i>Asio otus</i></td><td>Germany (60), Italy (76)</td></tr><tr><td>Snowy owl</td><td><i>Bubo scandiacus</i></td><td>Germany (60), Switzerland (69)</td></tr><tr><td>Tengmaml's owl</td><td><i>Aegolius funereus</i></td><td>Switzerland (69), IT</td></tr><tr><td>Hawk owl</td><td><i>Surnia ulala</i></td><td>Switzerland (69), DE</td></tr><tr><td>Pygmy owl</td><td><i>Glaucidium passerinum</i></td><td>Switzerland (69)</td></tr><tr><td>Tawny owl</td><td><i>Strix aluco</i></td><td>Germany (60), Italy (76)</td></tr><tr><td>Eurasian scops owl</td><td><i>Otus scops</i></td><td>Italy (76)</td></tr><tr><td>Little owl</td><td><i>Athene noctua</i></td><td>Italy (76)</td></tr><tr><td>Coraciiformes</td><td>Common kingfisher</td><td><i>Alcedo atthis</i></td><td>Germany (77)</td></tr><tr><td></td><td>European bee-eater</td><td><i>Merops apiaster</i></td><td>Italy (78)</td></tr><tr><td rowspan="2">Piciformes</td><td>Great spotted woodpecker</td><td><i>Dendroconopus major</i></td><td>Belgium (71)</td></tr><tr><td>European green woodpecker</td><td><i>Picus viridis</i></td><td>Germany (66,79)</td></tr><tr><td rowspan="2">Charadriiformes</td><td>Inca tern</td><td><i>Larosterna inca</i></td><td>Germany (66,79)</td></tr><tr><td>Yellow-legged gull</td><td><i>Larus michahellis</i></td><td>Italy (41)</td></tr><tr><td>Accipitriformes</td><td>Greater spotted eagle</td><td><i>Aquila clanga</i></td><td>Italy (41)</td></tr><tr><td>Caprimulgiformes</td><td>Nightjar</td><td><i>Caprimulgus europaeus</i></td><td>Italy (41,80)</td></tr><tr><td>Pelecaniformes</td><td>Grey heron</td><td><i>Ardea cinerea</i></td><td>Italy (41)</td></tr><tr><td>Columbiformes</td><td>Collared dove</td><td><i>Streptopelia decaocto</i></td><td>Italy (41,80)</td></tr><tr><td>Galliformes</td><td>Red-legged partridge</td><td><i>Alectoris rufa</i></td><td>Italy (76)</td></tr></tbody></table>
## Vectors

Ornithophilic mosquito species of the *Culex* genus are the predominant vectors of USUV with *Cx. pipiens* being the main vector in Europe (see Table 1). Mosquito vectors are responsible for virus transmission between birds and to susceptible mammals, such as humans and horses, which are incidental dead-end hosts, with short-lasting and low-level viremia. The transmission rate of USUV not only varies between different mosquito species and virus strains but also their geographical location (40) (Table 3).

<table><caption>Table 3. Infection, dissemination and transmission rates for mosquitoes following oral exposure to USUV</caption><thead><tr><th>Species</th><th>Country</th><th>Infection rate<sup>a</sup></th><th>Dissemination rate<sup>b</sup></th><th>Transmission rate<sup>c</sup></th></tr></thead><tbody><tr><td>Cx. pipiens</td><td>Netherlands (81)</td><td>80%</td><td></td><td>69%</td></tr><tr><td>Cx. pipiens</td><td>Belgium (40)</td><td>16.2%</td><td>0</td><td>0</td></tr><tr><td>Aedes albopictus</td><td>Italy (46)</td><td>0</td><td></td><td></td></tr><tr><td>Cx. modestus</td><td>Belgium (40)</td><td>60%</td><td>66.7%</td><td>50%</td></tr><tr><td>Cx. pipiens</td><td>France (82)</td><td>4.3%</td><td>100%</td><td>100%</td></tr><tr><td>Ae. rusticus</td><td>France (82)</td><td>0</td><td>0</td><td>0</td></tr><tr><td>An. plumbeus</td><td>France (82)</td><td>0</td><td>0</td><td>0</td></tr><tr><td rowspan="5">Ae. albopictus</td><td rowspan="5">France (82)</td><td rowspan="5">-</td><td rowspan="5">-</td><td>4.2% at 10 dpi</td></tr><tr><td>29.2% at 14 dpi</td></tr><tr><td>12.5% at 17 dpi</td></tr><tr><td>16.7% at 21 dpi</td></tr><tr><td>16.7% at 28 dpi</td></tr><tr><td rowspan="3">Cx. pipiens biotype molestus</td><td rowspan="3">Germany (83)</td><td>80%</td><td>37.5%</td><td>100% at 10 dpi</td></tr><tr><td>66.7%</td><td>100%</td><td>75% at 21 dpi</td></tr><tr><td>81.3%</td><td>100%</td><td>15.4% at 16 dpi</td></tr><tr><td rowspan="2">Cx. pipiens biotype molestus</td><td rowspan="2">Serbia (83)</td><td>80%</td><td>100%</td><td>50% at 21 dpi</td></tr><tr><td>12.5%</td><td>100%</td><td>100%</td></tr><tr><td>Cx. torrentium</td><td>Germany (83)</td><td>-</td><td>-</td><td>18%</td></tr><tr><td>Cx. pipiens biotype pipiens</td><td>Netherlands (84)</td><td>-</td><td>-</td><td>30%</td></tr><tr><td>Cx. pipiens biotype molestus</td><td>Netherlands (84)</td><td>-</td><td>-</td><td>30%</td></tr></tbody></table>

dpi: day-post-infection

*Cx. pipiens* that were exposed to USUV and WNV simultaneously via infectious blood meal displayed significantly reduced USUV transmission compared to mosquitoes that were only exposed to USUV (from 15% to 3%), while the infection and transmission of WNV was unaffected (85). In contrast, when mosquitoes were pre-infected with USUV via infectious blood meal, WNV transmission was significantly reduced (from 44% to 17%).

## Incidental hosts

Humans and non-human mammals are incidental dead-end hosts.

These hosts are not involved in the transmission cycle as they correspond to epidemiological dead ends for viral propagation because the viremia in the infected mammals is not high enough to ensure transmission through mosquito bites.

Apart from humans, USUV (or antibodies against it) has been detected in several mammals, including:

<footnote><sup>a</sup> No. infected mosquito bodies/no. mosquitoes tested</footnote>

<footnote><sup>b</sup> No. infected mosquitoes with virus in organs/no. mosquitoes tested.</footnote>

<footnote><sup>c</sup> No. infected mosquitoes with virus in saliva/no. mosquitoes tested.</footnote>
- horses: Poland (25), Italy, Croatia, Serbia and Spain (86);

- dogs: Italy (87) and Slovenia (88);

- squirrels: Italy (89);

- bats: Belgium (28), Germany (90);

- wild boars: Serbia (91);

- roe deers: Serbia (91);

- various deer species: Spain (92);

- zoo mammals (i.e. Asian lions, maned wolves, Iberian wolves, grey and northwestern wolves, African wild dogs, chimpanzees, common elands, giant pandas, Malayan tapirs, white rhinoceros, guinea pigs, rabbits, and red foxes): France (61, 93), Spain (92, 94), Slovenia (62) and other Central European country (59).

USUV was detected for the first time in reptiles, green lizards, in Slovakia (95).

# Drivers of the disease emergence and spread

## Ecological drivers

Studies report an increase in mosquito-borne disease outbreaks including USUV in Europe (6, 96) in the last decade due to changes in the temperature and precipitation attributable to climate change (97, 98) and anthropogenic landscape modifications (99-101).

There is a lack of evidence on the impact of climate change on the epidemiology of USUV infection, particularly considering that the weather can influence bird movements, vector abundance and virus replication in the vectors.

*Cx pipens*, the main vector of USUV in humans, was observed to have a strong relation with temperature (102). The study demonstrated that higher temperatures lead to higher mosquito abundances regardless of soil type. Larval stages of *Cx. pipiens* and related sibling species can inhabit a wide range of habitats, including artificial habitats (103). Land cover in urban areas consists mostly of artificial surfaces, which offer plenty of breeding sites filled with a small volume of water without the risk of predation (104-106). Studies also observed that artificial and clay surfaces, frequent in urban landscapes, increased mosquito abundance due to water stagnation in comparison to sandy soils (102).

Climate, land use, and biodiversity loss impact the distribution of bird populations which has an indirect effect on USUV emergence and spread. European breeding birds have shifted their range by 2.4 km per year, on average in the recent three decades (107). Studies also demonstrate that *Strigiformes* (owl, amplifying hosts of USUV) will shift to higher altitudes due to anthropogenic climate change (108).

Further ecological studies are required to get an understanding of the relationship between climate variables and human case incidence and to refine models or predictions of outbreak occurrences. Modelling the epidemiology of WNV throughout Europe under different climate change scenarios (IPCC scenarios) could help mapping the potential distribution of USUV, as co-circulation of both viruses is frequent.
# Natural history of disease in humans and animals, including symptoms, morbidity and mortality

## Brief history of the pathogen and disease

Following its first identification in South Africa in 1959, USUV has been detected in avian, equid and mosquito species in several African countries: Central African Republic, Senegal, Ivory Coast, Nigeria, Uganda, Burkina Faso, Tunisia, and Morocco (109-112). The first case of human infection by USUV was reported in the Central African Republic in the 1980s and a second case was diagnosed in Burkina Faso in 2004 (109). For these two cases, mild clinical signs were reported: fever and skin rash.

Phylogenetic studies suggest that at least three USUV introductions have occurred in Europe along the migratory routes from Africa. The virus has been introduced in Spain on two occasions in the 1950s and then in the 1990s along an eastern Atlantic migratory route (113). A unique introduction in Italy and Austria has been reported in the 1980s along a Black Sea/Mediterranean migratory route (15, 113). USUV has been classified into eight lineages: African (Africa 1/2/3) or European (Europe 1/2/3/4/5). Most strains currently circulating in Europe belong to the European lineages of USUV. However, African 2 lineage of USUV has also been identified in France in a human patient (74) and in owls in Berlin Zoo in 2015 (114). Furthermore, African lineages continue to be introduced into the continent, like the African 2 and 3 lineages reported in 2018 in *Cx. pipiens* in the South of France (22). The Europe 1 lineage is thought to derive from a Senegalese strain that reached Spain and was implicated in the first avian epizootic in blackbirds Austria in 2001(14). Europe 2 lineage is an Austrian strain from 1993, reappearing during the autochthonous Italian cases of 2009-2010 and the Austrian and Hungarian cases in 2016 (20). European 3 lineage is from an Italian strain that circulated in 2007 that was responsible for the massive bird die-off observed in Germany in 2011/2013 (114), in France in 2015 (23), and in Belgium in 2016 (29). European 4 lineage comprises strains circulating in Italy in 2010 and 2015 (16). Finally, the Europe 5 lineage was isolated from infected birds in Germany in 2016 (16).

## Disease in humans

Humans are infected with USUV following a bite from an infected mosquito. No transfusion-associated USUV infection has been reported (115). Due to limited clinical data available on USUV infection, the incubation period of 3 to 12 days was estimated based on WNV infection as reference (116). Following incubation, a brief viraemic phase is triggered and the patient may develop symptoms (117). Most USUV infections in humans remain largely asymptomatic with sporadic cases with neurological manifestations due to neuroinvasive disease. Severe disease is often associated with compromised immune systems and advanced age (17, 18, 118). Impaired neurological functions were observed in neuroinvasive diseases such as meningitis, encephalitis, and polyneuritis (17, 18, 74, 118-120). Symptoms of neuroinvasive disease are nuchal rigidity, hand tremor, and hyperreflexia (54).

Complications of USUV fulminant hepatitis have been reported following USUV neuroinvasive disease (18) in immunosuppressed patients. One of the samples during the retrospective screening of CSF in Montpellier, France tested positive for USUV (74). The patient was diagnosed with idiopathic facial paralysis which was putatively associated with an acute USUV infection. Rarely, in benign cases, the symptoms of an acute USUV infection resemble that of a febrile illness (109, 118) with fever, arthralgia, myalgia, headache, and rash. WNV and
other arboviral diseases such as Japanese encephalitis (with history of travel to South-East Asia) to be considered as differential diseases.

## Disease in animals

Although USUV has been detected in equids and other mammals, clinical features and pathological lesions of USUV infection have been described only in birds (133). Infected birds present as non-specific clinical features like immobility, apathy, ruffled feathers and neurological signs such as ataxia, paresis, tremors, torticollis, inability to fly and seizures (134).

# Availability of preventive, therapeutic and control measures, including licensed or pipeline vaccines

## Therapy in humans

Treatment in humans is symptomatic. A recent study demonstrated interferon therapy was a potential therapeutic intervention in human and veterinary medicine (135). Favipiravir, a broad-spectrum viral RNA polymerase inhibitor, was shown to reduce USUV load in a mice model (136).

## Therapy in animals

Ivermectin has been suggested as a candidate for treatment of USUV infection in captive birds (137).

## Licensed or pipeline vaccines

To date, there are no authorized vaccines against USUV infections. A study described the protective effect of a recombinant DNA vaccine against lethal challenge with USUV in alpha/beta interferon receptor deficient mouse model (138).

Another study reported the protective effect of an attenuated WNV-dengue virus 2 chimeric vaccine against USUV in the same mouse model.

However, the efficacy and safety of these vaccine candidates in birds has not been evaluated. Considering the rarity of USUV associated disease in mammals, the use of immunoprophylaxis in humans is not considered to be justified.

## Other prevention measures

### Blood safety

In EU/EEA, 45 USUV positive blood donations have been reported in Austria (29 with a peak of 20 in 2018), Italy (6 in 2018), and Netherlands (10, with 8 in 2018). None of the EU/EEA countries implemented specific Substances of Human origin (SoHo) safety measures for USUV infection, except the deferral of proven flavivirus positive donations.
However, measures applied to secure the safety of blood and other SoHO donations in relation to the risk of WNV infection including WNV NAT testing could mitigate the likelihood of donor-derived USUV transmission, the areas of co-circulation of the two viruses (33).

## Mosquito control and other prevention strategies

Control measures against mosquitoes can be applied preventively (e.g., larval and adult control) before and during the transmission season as the principal vectors of USUV are *Culex* spp. mosquitoes. However, there is little evidence on the direct effect of preventive mosquito control on the intensity of USUV outbreaks (33).

Reactive mosquito control measures (e.g., insecticide treatments in affected areas during outbreaks) is a widely applied method, although the implementation (geographic coverage, timing, repetitions) greatly vary in depending on the legislation of the use of biocides, available resources, and the public and animal health impacts of the outbreaks in different countries.

The use of individual protective measures to prevent mosquito bites (e.g., repellents, nets) and the reduction of mosquito breeding sites (e.g., stagnant water around households) is facilitated by public awareness campaigns at the start of and during the transmission season.

Eleven EU/EEA countries performed mosquito control actions and/or citizen education programmes on mosquito control (33). Among these, Croatia, Czechia, France, Germany, Greece, Italy, Slovakia, and Spain performed both. Malta focused only on citizen education programmes, and Cyprus, Hungary, and Romania solely on control actions.

Citizen education programmes provided information on mosquitoes, bite prevention, and breeding. In certain countries such as Germany, Italy, and Spain, tools or apps have been provided to citizens for reporting mosquito presence and biting incidents.

In some countries the control measures are intensified during detected WNV circulation in avian and mammalian hosts. This also benefits the control of USUV in circulation.

## Epidemiological situation at different spatial scales: past and current trends

USUV is endemic in several EU/EEA countries. As of September 2023, 15 out of 30 EU/EEA countries reported USUV circulation among avian and equine animals.

However, human cases in Europe remains sporadic.

The first autochthonous human cases of USUV infection in Europe were reported in 2009 in Italy.

Since then, eight countries including Austria, Croatia, Czechia (131), France, Germany (127, 139), Hungary (132, 140), Italy, and the Netherlands have reported autochthonous cases of USUV infections in humans (33). These eight countries reported a total of 109 cases of acute USUV infection, most of which were reported by Italy (n=56, 54%), Austria (n=26, 25%) and the Netherlands (n=11, 11%) (Table 4).
<table><caption>Table 4. Description of human Usutu cases in EU/EEA countries</caption><thead><tr><th>Country</th><th>Year (ref.)</th><th>Cases</th><th>Age</th><th>Sex</th><th>c/o</th><th>Symptomatology</th><th>Population</th><th>Lineage</th></tr></thead><tbody><tr><td rowspan="10">Italy</td><td>2009 (17)</td><td>1</td><td>60+</td><td>F</td><td>+</td><td>Meningoencephalitis</td><td>Clinical case</td><td>EU1</td></tr><tr><td>2009 (18)</td><td>1</td><td>40+</td><td>F</td><td>+</td><td>Encephalitis</td><td>Clinical case</td><td>EU2</td></tr><tr><td rowspan="3">2008-9 (121)</td><td rowspan="3">3</td><td>40</td><td>M</td><td>+</td><td rowspan="3">Meningoencephalitis</td><td rowspan="3">Meningoencephalitis patients</td><td rowspan="3">-</td></tr><tr><td>73</td><td>M</td><td>+</td></tr><tr><td>54</td><td>F</td><td>+</td></tr><tr><td>2008-11 (120)</td><td>10</td><td>60+</td><td>M</td><td>+</td><td>Meningoencephalitis /asymptomatic</td><td>Meningoencephalitis patients + various healthy and sick subjects</td><td>-</td></tr><tr><td>2016-18 (115)</td><td>25</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>-</td></tr><tr><td>2017-18 (122)</td><td>9</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU2/3/4</td></tr><tr><td>2017-18 (118)</td><td>8</td><td>67</td><td>F</td><td>+</td><td>1 encephalitis, 6 fever, 1 viremia</td><td>Suspected infection and +ve donors</td><td>EU2</td></tr><tr><td>2018 (123)</td><td>2</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU2</td></tr><tr><td>2018 (124)</td><td>1</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU1</td></tr><tr><td>2022 (125)</td><td>6</td><td></td><td></td><td></td><td>2 fever, 1 asymptomatic</td><td>Clinical case</td><td>-</td></tr><tr><td rowspan="6">Croatia</td><td rowspan="3">2013 (119)</td><td rowspan="3">3</td><td>29</td><td>F</td><td>none</td><td rowspan="3">Meningoencephalitis</td><td rowspan="3">Meningoencephalitis patients</td><td rowspan="3">-</td></tr><tr><td>61</td><td>M</td><td>none</td></tr><tr><td>56</td><td>M</td><td>+</td></tr><tr><td rowspan="3">2018 (126)</td><td rowspan="3">3</td><td>25</td><td>-</td><td>none</td><td rowspan="3">Neuroinvasive disease</td><td rowspan="3">Neuroinvasive cohort</td><td rowspan="3">EU2</td></tr><tr><td>84</td><td>-</td><td>none</td></tr><tr><td>60</td><td>-</td><td>+</td></tr><tr><td>Germany</td><td>2016 (127)</td><td>1</td><td></td><td></td><td></td><td>Healthy</td><td>Blood donors</td><td>EU3</td></tr><tr><td rowspan="2">France</td><td>2016 (74)</td><td>1</td><td>39</td><td>M</td><td>none</td><td>Idiopathic facial paralysis</td><td>Patients with infectious and/or neurological signs</td><td>AF2</td></tr><tr><td>2022 (128)</td><td>1</td><td></td><td></td><td></td><td>Fever</td><td>Clinical case</td><td>-</td></tr><tr><td rowspan="4">Austria</td><td>2017 (129)</td><td>6</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU2</td></tr><tr><td>2018 (55)</td><td>18</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU2/AF3</td></tr><tr><td>2021 (130)</td><td>1</td><td>81</td><td>M</td><td>+</td><td>Meningitis</td><td>Clinical case</td><td>EU2</td></tr><tr><td>2018 (131)</td><td>1</td><td>46</td><td>W</td><td>none</td><td>Meningoencephalitis</td><td>Clinical case</td><td>-</td></tr><tr><td>Czech Republic</td><td>2018 (132)</td><td>1</td><td>40+</td><td>M</td><td>none</td><td>Aseptic meningitis</td><td>Clinical case</td><td>EU2</td></tr><tr><td>Netherlands</td><td>2018</td><td>7</td><td></td><td></td><td></td><td>Asymptomatic</td><td>Blood donors</td><td>EU3</td></tr></tbody></table>
**Sociological and demographical dimension affecting
susceptibility and exposure, including gender**

Severe neuroinvasive disease following USUV infection is associated with compromised immune systems and advanced age (17, 18, 118). However, no gender predilection has been reported.

**Diagnostic procedures and notification systems
used at local, national and European scale**

**Diagnosis**

Diagnosing USUV infection in humans relies on the following diagnostic techniques (1): i) the detection of viral RNA in blood and in cerebrospinal fluid (CSF); ii) the isolation of the virus in cell culture and/or; iii) indirect assay detecting anti-USUV antibodies (IgM and G) in the serum and the CSF of patients.

The diagnosis of USUV in humans is challenged by the limited availability of validated commercial tests. USUV serological assays are based on Enzyme-Linked Immunosorbent Assay (ELISA) tests or immunofluorescence tests developed by national reference laboratories, performed with viral antigens or virus isolates (141). However, these tests lack specificity due to the risk of serological cross-reactions with infections by closely related flaviviruses, such as WNV (123, 142). These tests need further confirmation by sero neutralization assays (123, 143). However, WNV NATs validated for donor screening can detect USUV with high sensitivity (144). Direct diagnosis of USUV by isolating the virus in cell cultures such as Vero, BHK-21, and C6/36 cultures and visualizing cytopathic effects. The presence of USUV RNA has been demonstrated in blood, urine, and CSF of patients with acute infection (17, 18, 74, 118-120). Molecular methods include USUV-specific real-time RT-PCR assays, either as a single-target or multiplex test, and broad-range flavivirus RT-PCR followed by amplicon sequencing (121, 144, 145).

**Infrastructure capacity to identify pathogens
for each Member State**

Twenty EU/EEA countries including Austria, Belgium, Croatia, Czechia, Estonia, France, Germany, Greece, Hungary, Ireland, Italy, Latvia, Liechtenstein, Lithuania, Malta, the Netherlands, Romania, Slovakia, Slovenia, and Spain have the laboratory capability to detect USUV infection, mostly by the national reference laboratories (146). Diagnosis of USUV infection include USUV specific PCR or broad-range pan-flavivirus PCR and sequencing from blood, tissue and/or CSF. Serological diagnosis detects USUV specific antibody response or the flavivirus antibody response using other cross-reactive serological methods, followed by a neutralization assay.

Four countries (Greece, Germany, Italy, and Norway) developed case definitions for USUV infection in humans based on clinical (encephalitis, meningitis with clear CSF,
polyradiculoneuritis (similar to Guillain-Barre'), acute flaccid paralysis) and laboratory criteria (detection of specific antibodies in serum of CSF, isolation or detection of virus's nucleic acid).

There is need for a standardized case definition for USUV infection in humans in EU/EEA countries.

Currently, USUV is not a major public health threat in the EU/EEA but monitoring virus circulation and its pathogenicity is important to early detect any change in the epidemiology of the disease (146). Thus, not a notifiable disease. Improvements in monitoring might be achieved with a standard case definition of USUV infection in the EU/EEA, and with an integrated approach, including humans, animals, and vectors in USUV surveillance for early detection. Such integrated surveillance systems exist for WNV surveillance in Austria, Croatia, Czechia, France, Germany, Greece, Italy, the Netherlands, and Spain (33). Amongst these nine countries, Croatia, Germany, and Italy have integrated USUV in their WNV surveillance.

## Estimated influence of environmental change on the disease future trends

It was unknown how climate change, land use modifications, and biodiversity loss impacts the epidemiology of USUV infection in humans, particularly considering that the weather can influence bird movements, vector abundance and virus replication in the vectors.

## References

1. Clé M, Beck C, Salinas S, Lecollinet S, Gutierrez S, Van de Perre P, et al. Usutu virus: A new threat? *Epidemiol Infect*. 2019;147:e232.

2. McIntosh B. Usutu (SA Ar 1776), nouvel arbovirus du groupe B. *Int Cat Arboviruses*. 1985;3:1059-60.

3. Bakran-Lebl K, Camp JV, Kolodziejek J, Weidinger P, Hufnagl P, Cabal Rosel A, et al. Diversity of West Nile and Usutu virus strains in mosquitoes at an international airport in Austria. *Transbound Emerg Dis*. 2022 Jul;69(4):2096-109.

4. Calisher CH, Gould EA. Taxonomy of the virus family Flaviviridae. *Adv Virus Res*. 2003;59:1-19.

5. Beck C, Jimenez-Clavero MA, Leblond A, Durand B, Nowotny N, Leparc-Goffart I, et al. Flaviviruses in Europe: complex circulation patterns and their consequences for the diagnosis and control of West Nile disease. *Int J Environ Res Public Health*. 2013 Nov 12;10(11):6049-83.

6. Cadar D, Lühken R, van der Jeugd H, Garigliany M, Ziegler U, Keller M, et al. Widespread activity of multiple lineages of Usutu virus, western Europe, 2016. *Euro Surveill*. 2017 Jan 26;22(4):30452.

7. Bakonyi T, Gould EA, Kolodziejek J, Weissenböck H, Nowotny N. Complete genome analysis and molecular characterization of Usutu virus that emerged in Austria in 2001: comparison with the South African strain SAAR-1776 and other flaviviruses. *Virology*. 2004 Oct 25;328(2):301-10.

8. Roesch F, Fajardo A, Moratorio G, Vignuzzi M. Usutu virus: an arbovirus on the rise. *Viruses*. 2019 Jul 12;11(7).

9. Smit JM, Moesker B, Rodenhuis-Zybert I, Wilschut J. Flavivirus cell entry and membrane fusion. *Viruses*. 2011 Feb 22;3(2):160-71.

10. Albentosa-González L, Clemente-Casares P, Sabariegos R, Mas A. Polymerase activity, protein-protein interaction, and cellular localization of the Usutu virus NS5 Protein. *Antimicrob Agents Chemother*. 2019 Dec 20;64(1).
11. Savini G, Monaco F, Terregino C, Di Gennaro A, Bano L, Pinoni C, et al. Usutu virus in Italy: an emergence or a silent infection? *Vet Microbiol*. 2011 Aug 5;151(3-4):264-74.

12. Weissenböck H, Kolodziejek J, Fragner K, Kuhn R, Pfeffer M, Nowotny N. Usutu virus activity in Austria, 2001-2002. *Microbes Infect*. 2003 Oct;5(12):1132-6.

13. Chvala S, Bakonyi T, Bukovsky C, Meister T, Brugger K, Rubel F, et al. Monitoring of Usutu virus activity and spread by using dead bird surveillance in Austria, 2003-2005. *Vet Microbiol*. 2007 Jun 21;122(3-4):237-45.

14. Weissenböck H, Kolodziejek J, Url A, Lussy H, Rebel-Bauder B, Nowotny N. Emergence of Usutu virus, an African mosquito-borne flavivirus of the Japanese encephalitis virus group, central Europe. *Emerg Infect Dis*. 2002 Jul;8(7):652-6.

15. Weissenböck H, Bakonyi T, Rossi G, Mani P, Nowotny N. Usutu virus, Italy, 1996. *Emerg Infect Dis*. 2013 Feb;19(2):274-7.

16. Vilibic-Cavlek T, Petrovic T, Savic V, Barbic L, Tabain I, Stevanovic V, et al. Epidemiology of Usutu Virus: The European Scenario. *Pathog Basel Switz*. 2020 Aug 26;9(9).

17. Pecorari M, Longo G, Gennari W, et al. First human case of Usutu virus neuroinvasive infection, Italy, August-September 2009. *Euro Surveill*. 2009;14(50):19446.

18. Cavrini F, Gaibani P, Longo G, Pierro AM, Rossini G, Bonilauri P, et al. Usutu virus infection in a patient who underwent orthotropic liver transplantation, Italy, August-September 2009. *Euro Surveill*. 2009 Dec 17;14(50):19448.

19. Ashraf U, Ye J, Ruan X, Wan S, Zhu B, Cao S. Usutu virus: an emerging flavivirus in Europe. *Viruses*. 2015 Jan 19;7(1):219-38.

20. Kemenesi G, Buzás D, Zana B, et al. First genetic characterization of Usutu virus from Culex pipiens mosquitoes Serbia, 2014. *Infect Genet Evol*. 2018;63:58-61. doi:10.1016/j.meegid.2018.05.012.

21. Scheuch DE, Schäfer M, Eiden M, et al. Detection of Usutu, Sindbis, and Batai viruses in mosquitoes (Diptera: Culicidae) collected in Germany, 2011-2016. *Viruses*. 2018;10(7):389.

22. Eiden M, Gil P, Ziegler U, et al. Emergence of two Usutu virus lineages in Culex pipiens mosquitoes in the Camargue, France, 2015. *Infect Genet Evol*. 2018;61:151-154.

23. Lecollinet S, Blanchard Y, Manson C, Lowenski S, Laloy E, Quenault H, et al. Dual emergence of Usutu virus in common blackbirds, Eastern France, 2015. *Emerg Infect Dis*. 2016 Dec;22(12):2225.

24. Csank T, Drzewnioková P, Korytár L, et al. A serosurvey of Flavivirus Infection in horses and birds in Slovakia. *Vector Borne Zoonotic Dis*. 2018;18(4):206-213. doi:10.1089/vbz.2017.2216

25. Bazanow B, van Vuren P, Szymanski P, Stygar D, Fracka A, Twardon J, et al. A survey on West Nile and Usutu viruses in horses and birds in Poland. *Viruses*. 2018 Feb;10(2).

26. Rijks JM, Kik ML, Slaterus R, Foppen R, Stroo A, IJzer J, et al. Widespread Usutu virus outbreak in birds in the Netherlands, 2016. *Euro Surveill*. 2016 Nov 10;21(45):30391.

27. Michel F, Fischer D, Eiden M, Fast C, Reuschel M, Müller K, et al. West Nile Virus and Usutu virus monitoring of wild birds in Germany. *Int J Environ Res Public Health*. 2018 Jan 22;15(1).

28. Benzarti E, Sarlet M, Franssen M, Cadar D, Schmidt-Chanasit J, Rivas JF, et al. Usutu virus epizootic in Belgium in 2017 and 2018: evidence of virus endemization and ongoing introduction events. *Vector Borne Zoonotic Dis*. 2020 Jan;20(1):43-50.

29. Garigliany M, Linden A, Gilliau G, Levy E, Sarlet M, Franssen M, et al. Usutu virus, Belgium, 2016. *Infect Genet Evol*. 2017 Mar;48:116-9.

30. Chaintoutis SC, Dovas CI, Papanastassopoulou M, Gewehr S, Danis K, Beck C, et al. Evaluation of a West Nile virus surveillance and early warning system in Greece, based on domestic pigeons. *Comp Immunol Microbiol Infect Dis*. 2014 Mar;37(2):131-41.
31. Höfle U, Gamino V, de Mera IGF, Mangold AJ, Ortíz JA, de la Fuente J. Usutu virus in migratory song thrushes, Spain. *Emerg Infect Dis.* 2013 Jul;19(7):1173-5.

32. Llorente F, Pérez-Ramírez E, Fernández-Pinero J, Soriguer R, Figuerola J, Jiménez-Clavero MA. Flaviviruses in game birds, southern Spain, 2011-2012. *Emerg Infect Dis.* 2013 Jun;19(6):1023-5.

33. European Centre for Disease Prevention and Control, European Food Safety Authority, Surveillance prevention and control of West Nile virus and Usutu virus infections in the EU/EEA. Stockholm: 2023.

34. Becker N, Jöst H, Ziegler U, Eiden M, Höper D, Emmerich P, et al. Epizootic emergence of Usutu virus in wild and captive birds in Germany. *PloS One*. 2012;7(2):e32604.

35. Calzolari M, Bonilauri P, Bellini R, Albieri A, Defilippo F, Maioli G, et al. Evidence of simultaneous circulation of West Nile and Usutu viruses in mosquitoes sampled in Emilia-Romagna region (Italy) in 2009. *PloS One*. 2010 Dec 15;5(12):e14324.

36. Tamba M, Bonilauri P, Bellini R, et al. Detection of Usutu virus within a West Nile virus surveillance program in Northern Italy. *Vector Borne Zoonotic Dis.* 2011;11(5):551-557. doi:10.1089/vbz.2010.0055.

37. Mancini G, Montarsi F, Calzolari M, Capelli G, Dottori M, Ravagnan S, et al. Mosquito species involved in the circulation of West Nile and Usutu viruses in Italy. *Vet Ital*. 2017 Jun 30;53(2):97-110.

38. Klobucar A, Savic V, Curman Posavec M, Petrinic S, Kuhar U, Toplak I, et al. Screening of mosquitoes for West Nile virus and Usutu virus in Croatia, 2015-2020. *Trop Med Infect Dis.* 2021 Apr 2;6(2).

39. Camp JV, Kolodziejek J, Nowotny N. Targeted surveillance reveals native and invasive mosquito species infected with Usutu virus. *Parasit Vectors*. 2019 Jan 21;12(1):46.

40. Soto A, De Coninck L, Devlies AS, Van De Wiele C, Rosales Rosas AL, Wang L, et al. Belgian Culex pipiens pipiens are competent vectors for West Nile virus while Culex modestus are competent vectors for Usutu virus. *PLoS Negl Trop Dis.* 2023 Sep 20;17(9):e0011649.

41. Calzolari MB Paolo; Bellini, Romeo; Albieri, Alessandro; Defilippo, Francesco; Tamba, Marco; Tassinari, Massimo; Gelati, Antonio; Cordioli, Paolo; Angelini, Paola; Dottori, Michele. Usutu Virus Persistence and West Nile Virus Inactivity in the Emilia-Romagna Region (Italy) in 2011. *PloS One*. 2013;8(5):e63978-NA.

42. Wang L, Rosales Rosas AL, De Coninck L, Shi C, Bouckaert J, Matthijnssens J, et al. Establishment of Culex modestus in Belgium and a glance into the virome of Belgian mosquito species. *mSphere*. 2021 Apr 21;6(2).

43. Vazquez A, Jimenez-Clavero M, Franco L, Donoso-Mantke O, Sambri V, Niedrig M, et al. Usutu virus: potential risk of human disease in Europe. *Euro Surveill*. 2011 Aug 4;16(31):19935.

44. Sieg M, Schmidt V, Ziegler U, Keller M, Höper D, Heenemann K, et al. Outbreak and cocirculation of three different Usutu Virus strains in Eastern Germany. *Vector Borne Zoonotic Dis.* 2017 Sep;17(9):662-4.

45. Jöst H, Bialonski A, Maus D, Sambri V, Eiden M, Groschup MH, et al. Isolation of Usutu virus in Germany. *Am J Trop Med Hyg*. 2011 Sep;85(3):551-3.

46. Puggioli A, Bonilauri P, Calzolari M, Lelli D, Carrieri M, Urbanelli S, et al. Does Aedes albopictus (Diptera: Culicidae) play any role in Usutu virus transmission in Northern Italy? Experimental oral infection and field evidences. *Acta Trop*. 2017 Aug;172:192-6.

47. Carletti F, Colavita F, Rovida F, Percivalle E, Baldanti F, Ricci I, et al. Expanding Usutu virus circulation in Italy: detection in the Lazio region, central Italy, 2017 to 2018. *Euro Surveill*. 2019 Jan;24(3).
48. Pautasso A, Radaelli MC, Ballardini M, Francese DR, Verna F, Modesto P, et al. Detection of West Nile and Usutu Viruses in Italian Free Areas: Entomological Surveillance in Piemonte and Liguria Regions, 2014. *Vector Borne Zoonotic Dis.* 2016 Apr;16(4):292-4.

49. Cerutti F, Giacobini M, Mosca A, Grasso I, Rambozzi L, Rossi L, et al. Evidence of mosquito-transmitted flavivirus circulation in Piedmont, north-western Italy. *Parasit Vectors*. 2012 May 22;5:99.

50. Busani L, Capelli G, Cecchinato M, Lorenzetto M, Savini G, Terregino C, et al. West Nile virus circulation in Veneto region in 2008-2009. *Epidemiol Infect*. 2011 Jun;139(6):818-25.

51. Grisenti M, Vázquez A, Herrero L, Cuevas L, Perez-Pastrana E, Arnoldi D, et al. Wide detection of Aedes flavivirus in north-eastern Italy--a European hotspot of emerging mosquito-borne diseases. *J Gen Virol*. 2015 Feb;96(Pt 2):420-30.

52. Busquets N, Alba A, Allepuz A, Aranda C, Ignacio Nuñez J. Usutu virus sequences in Culex pipiens (Diptera: Culicidae), Spain. *Emerg Infect Dis*. 2008 May;14(5):861-3.

53. Engler O, Savini G, Papa A, Figuerola J, Groschup M, Kampen H, et al. European surveillance for West Nile virus in mosquito populations. *Int J Environ Res Public Health*. 2013 Oct;10(10):4869-95.

54. Vilibic-Cavlek T, Kaic B, Barbic L, Pem-Novosel I, Slavic-Vrzic V, Lesnikar V, et al. First evidence of simultaneous occurrence of West Nile virus and Usutu virus neuroinvasive disease in humans in Croatia during the 2013 outbreak. *Infection*. 2014 Aug;42(4):689-95.

55. Aberle SW, Kolodziejek J, Jungbauer C, Stiasny K, Aberle JH, Zoufaly A, et al. Increase in human West Nile and Usutu virus infections, Austria, 2018. *Euro Surveill*. 2018 Oct;23(43).

56. Farajollahi A, Fonseca DM, Kramer LD, Marm Kilpatrick A. "Bird biting" mosquitoes and human disease: a review of the role of Culex pipiens complex mosquitoes in epidemiology. *Infect Genet Evol*. 2011;11(7):1577-1585. doi:10.1016/j.meegid.2011.08.013.

57. Benzarti E, Linden A, Desmecht D, Garigliany M. Mosquito-borne epornitic flaviviruses: an update and review. *J Gen Virol*. 2019 Feb;100(2):119-32.

58. Bakonyi T, Erdélyi K, Ursu K, Ferenczi E, Csörgo T, Lussy H, et al. Emergence of Usutu virus in Hungary. *J Clin Microbiol*. 2007 Dec;45(12):3870-4.

59. Buchebner N, Zenker W, Wenker C, Steinmetz HW, Sós E, Lussy H, et al. Low Usutu virus seroprevalence in four zoological gardens in central Europe. *BMC Vet Res*. 2013 Aug 6;9:153.

60. Michel F, Sieg M, Fischer D, Keller M, Eiden M, Reuschel M, et al. Evidence for West Nile virus and Usutu virus infections in wild and resident birds in Germany, 2017 and 2018. *Viruses*. 2019 Jul 23;11(7).

61. Constant O, Bollore K, Clé M, et al. Evidence of exposure to USUV and WNV in zoo animals in France. *Pathogens*. 2020;9(12):1005.

62. Kvapil P, Račnik J, Kastelic M, Bártová E, Korva M, Jelovšek M, et al. A Sentinel serological study in selected zoo animals to assess early detection of West Nile and Usutu virus circulation in Slovenia. *Viruses*. 2021 Apr 6;13(4).

63. Bakonyi T, Erdélyi K, Brunthaler R, Dán Á, Weissenböck H, Nowotny N. Usutu virus, Austria and Hungary, 2010-2016. *Emerg Microbes Infect*. 2017 Oct 11;6(10):e85.

64. Chvala S, Kolodziejek J, Nowotny N, Weissenböck H. Pathology and viral distribution in fatal Usutu virus infections of birds from the 2001 and 2002 outbreaks in Austria. *J Comp Pathol*. 2004 Oct;131(2-3):176-85.

65. Hubálek Z, Rudolf I, Čapek M, Bakonyi T, Betášová L, Nowotny N. Usutu virus in blackbirds (Turdus merula), Czech Republic, 2011-2012. *Transbound Emerg Dis*. 2014 Jun;61(3):273-6.
66. Ziegler U, Jöst H, Müller K, Fischer D, Rinder M, Tietze DT, et al. Epidemic spread of Usutu Virus in Southwest Germany in 2011 to 2013 and monitoring of wild birds for Usutu and West Nile viruses. *Vector Borne Zoonotic Dis* Larchmt N. 2015 Aug;15(8):481-8.

67. Calzolari M, Gaibani P, Bellini R, Defilippo F, Pierro A, Albieri A, et al. Mosquito, bird and human surveillance of West Nile and Usutu viruses in Emilia-Romagna Region (Italy) in 2010. *PloS One*. 2012;7(5):e38058.

68. Rouffaer LO, Steensels M, Verlinden M, Vervaeke M, Boonyarittichaikij R, Martel A, et al. Usutu Virus epizootic and Plasmodium coinfection in Eurasian blackbirds (Turdus merula) in Flanders, Belgium. *J Wildl Dis*. 2018 Oct;54(4):859-62.

69. Steinmetz HW, Bakonyi T, Weissenböck H, Hatt JM, Eulenberger U, Robert N, et al. Emergence and establishment of Usutu virus infection in wild and captive avian species in and around Zurich, Switzerland--genomic and pathologic comparison to other central European outbreaks. *Vet Microbiol*. 2011 Mar 24;148(2-4):207-12.

70. Ziegler U, Bergmann F, Fischer D, Müller K, Holicki CM, Sadeghi B, et al. Spread of West Nile virus and Usutu virus in the German bird population, 2019-2020. *Microorganisms*. 2022 Apr 12;10(4).

71. Garigliany MM, Marlier D, Tenner-Racz K, Eiden M, Cassart D, Gandar F, et al. Detection of Usutu virus in a bullfinch (Pyrrhula pyrrhula) and a great spotted woodpecker (Dendrocopos major) in north-west Europe. *Vet J Lond Engl* 1997. 2014 Jan;199(1):191-3.

72. Escribano-Romero E, Jiménez de Oya N, Camacho MC, Blázquez AB, Martín-Acebes MA, Risalde MA, et al. Previous Usutu virus exposure partially protects magpies (Pica pica) against West Nile Virus disease but does not prevent horizontal transmission. *Viruses*. 2021 Jul 20;13(7).

73. Schulze C, Ziegler U, Schlieben P, Fast C, Bock S, Schüle A, et al. Fatal systemic Usutu virus infection in two juvenile great grey owls (Strix nebulosa) from a German zoological garden. *J Comp Pathol*. 2017 Jan 31;156:113.

74. Simonin Y, Sillam O, Carles MJ, Gutierrez S, Gil P, Constant O, et al. Human Usutu virus infection with atypical neurologic presentation, Montpellier, France, 2016. *Emerg Infect Dis*. 2018 May;24(5):875-8.

75. Manarolla G, Bakonyi T, Gallazzi D, Crosta L, Weissenböck H, Dorrestein GM, et al. Usutu virus in wild birds in northern Italy. *Vet Microbiol*. 2010 Feb 24;141(1-2):159-63.

76. Lauriano A, Rossi A, Galletti G, Casadei G, Santi A, Rubini S, et al. West Nile and Usutu Viruses' surveillance in birds of the province of Ferrara, Italy, from 2015 to 2019. *Viruses*. 2021 Jul 14;13(7).

77. Störk T, de le Roi M, Haverkamp AK, Jesse ST, Peters M, Fast C, et al. Analysis of avian Usutu virus infections in Germany from 2011 to 2018 with focus on dsRNA detection to demonstrate viral infections. *Sci Rep*. 2021 Dec 17;11(1):24191.

78. Mancuso E, Cecere JG, Iapaolo F, Di Gennaro A, Sacchi M, Savini G, et al. West Nile and Usutu virus introduction via migratory birds: a retrospective analysis in Italy. *Viruses*. 2022 Feb 17;14(2).

79. Saiz JC, Blazquez AB. Usutu virus: current knowledge and future perspectives. *Virus Adapt Treat*. 2017 Oct 16;9:27-40.

80. Giglia G, Mencattelli G, Lepri E, Agliani G, Gobbi M, Gröne A, et al. West Nile Virus and Usutu virus: a post-mortem monitoring study in wild birds from rescue centers, Central Italy. *Viruses*. 2022 Sep 9;14(9).

81. Fros JJ, Miesen P, Vogels CB, Gaibani P, Sambri V, Martina BE, et al. Comparative Usutu and West Nile virus transmission potential by local Culex pipiens mosquitoes in north-western Europe. *One Health Amst Neth*. 2015 Dec;1:31-6.
82. Martinet JP, Bohers C, Vazeille M, Ferté H, Mousson L, Mathieu B, et al. Assessing vector competence of mosquitoes from northeastern France to West Nile virus and Usutu virus. *PLoS Negl Trop Dis.* 2023 Jun;17(6):e0011144.

83. Holicki CM, Scheuch DE, Ziegler U, Lettow J, Kampen H, Werner D, et al. German Culex pipiens biotype molestus and Culex torrentium are vector-competent for Usutu virus. *Parasit Vectors*. 2020 Dec 30;13(1):625.

84. Abbo SR, Visser TM, Koenraadt CJM, Pijlman GP, Wang H. Effect of blood source on vector competence of Culex pipiens biotypes for Usutu virus. *Parasit Vectors*. 2021 Apr 8;14(1):194.

85. Wang H, Abbo SR, Visser TM, Westenberg M, Geertsema C, Fros JJ, et al. Competition between Usutu virus and West Nile virus during simultaneous and sequential infection of Culex pipiens mosquitoes. *Emerg Microbes Infect.* 2020 Dec;9(1):2642-52.

86. Gaibani P, Rossini G. An overview of Usutu virus. *Microbes Infect*. 2017 Aug;19(7-8):382-7.

87. Montagnaro S, Piantedosi D, Ciarcia R, Loponte R, Veneziano V, Fusco G, et al. Serological evidence of mosquito-borne flaviviruses circulation in hunting dogs in Campania Region, Italy. *Vector-borne zoonotic dis*. 2019 Feb 1;19(2):142-7.

88. Knap N, Korva M, Ivović V, Kalan K, Jelovšek M, Sagadin M, et al. West Nile Virus in Slovenia. *Viruses*. 2020 Jul 3;12(7):720.

89. Romeo C, Lecollinet S, Caballero J, Isla J, Luzzago C, Ferrari N, et al. Are tree squirrels involved in the circulation of flaviviruses in Italy? *Transbound Emerg Dis*. 2018 Oct;65(5):1372-6.

90. Cadar D, Becker N, Campos R de M, Börstler J, Jöst H, Schmidt-Chanasit J. Usutu virus in bats, Germany, 2013. *Emerg Infect Dis*. 2014 Oct;20(10):1771-3.

91. Escribano-Romero E, Lupulović D, Merino-Ramos T, Blázquez AB, Lazić G, Lazić S, et al. West Nile virus serosurveillance in pigs, wild boars, and roe deer in Serbia. *Vet Microbiol*. 2015 Apr 17;176(3-4):365-9.

92. Garcia-Bocanegra I, Paniagua J, Gutierrez-Guzman A, Lecollinet S, Boadella M, Arenas-Montes A, et al. Spatio-temporal trends and risk factors affecting West Nile virus and related flavivirus exposure in Spanish wild ruminants. BMC *Vet Res*. 2016 Nov 9;12.

93. Bournez L, Umhang G, Faure E, Boucher JM, Boué F, Jourdain E, et al. Exposure of wild ungulates to the Usutu and tick-borne encephalitis viruses in France in 2009-2014: evidence of undetected flavivirus circulation a decade ago. *Viruses*. 2019 Dec 19;12(1).

94. Caballero-Gomez J, Terriza D, Pujols J, Martinez-Nevado E, Carbonell M, Guerra R, et al. Monitoring of bluetongue virus in zoo animals in Spain, 2007-2019. *Transbound Emerg Dis*. 2022 Jul;69(4):1739-47.

95. Čabanová V, Šikutová S, Straková P, Šebesta O, Vichová B, Zubríková D, et al. Co-circulation of West Nile and Usutu flaviviruses in mosquitoes in Slovakia, 2018. *Viruses*. 2019 Jul 12;11(7).

96. Vilibic-Cavlek T, Savic V, Petrovic T, Toplak I, Barbic L, Petric D, et al. Emerging trends in the epidemiology of West Nile and Usutu virus infections in Southern Europe. Front *Vet Sci*. 2019 Dec 6;6.

97. Brugueras S, Fernandez-Martinez B, Martinez-de la Puente J, Figuerola J, Porro T, Rius C, et al. Environmental drivers, climate change and emergent diseases transmitted by mosquitoes and their vectors in southern Europe: A systematic review. *Environ Res*. 2020 Dec;191.

98. Reiter P. Climate change and mosquito-borne disease. *Environ Health Perspect*. 2001;109(suppl 1):141-61.

99. Mohlmann T, Wennergren U, Talle M, Favia G, Damiani C, Bracchetti L, et al. Community analysis of the abundance and diversity of mosquito species (Diptera: Culicidae) in three European countries at different latitudes. *Parasit Vectors*. 2017 Oct 23;10.
100. Esser HJ, Liefting Y, Ibáñez-Justicia A, et al. Spatial risk analysis for the introduction and circulation of six arboviruses in the Netherlands. *Parasit Vectors*. 2020;13(1):464. doi:10.1186/s13071-020-04339-0

101. Gangoso L, Aragonés D, Martínez-de la Puente J, et al. Determinants of the current and future distribution of the West Nile virus mosquito vector Culex pipiens in Spain. *Environ Res*. 2020;188:109837. doi:10.1016/j.envres.2020.109837.

102. Krol L, Blom R, Dellar M, van der Beek JG, Stroo ACJ, van Bodegom PM, et al. Interactive effects of climate, land use and soil type on Culex pipiens/torrentium abundance. *One Health Amst Neth*. 2023 Dec;17:100589.

103. Becker N, Jöst A, Weitzel T. The Culex pipiens complex in Europe. *J Am Mosq Control Assoc*. 2012 Dec;28(4s):53-67.

104. Stead D. Urban planning, water management and climate change strategies: adaptation, mitigation and resilience narratives in the Netherlands. *Int J Sustain Dev World Ecol*. 2014 Jan 2;21(1):15-27.

105. Washburn JO. Regulatory factors affecting larval mosquito populations in container and pool habitats: implications for biological control. *J Am Mosq Control Assoc*. 1995 Jun;11(2 Pt 2):279-83.

106. Ramasamy R, Surendran S. Mosquito vectors developing in atypical anthropogenic habitats: Global overview of recent observations, mechanisms and impact on disease transmission. *J Vector Borne Dis*. 2016 Jun;53(2):91-8.

107. Howard C, Marjakangas EL, Morán-Ordóñez A, Milanesi P, Abuladze A, Aghababyan K, et al. Local colonisations and extinctions of European birds are poorly explained by changes in climate suitability. *Nat Commun*. 2023 Jul 20;14(1):4304.

108. Matadasmas RE, Enríquez PL, Guevara L, Navarro-Sigüenza AG. Stairway to extinction? Influence of anthropogenic climate change on distribution patterns of montane Strigiformes in Mesoamerica. *Avian Conserv Ecol*. 2022;17(2):37. https://doi.org/10.5751/ACE-02314-170237

109. Nikolay B, Diallo M, Boye CS, Sall AA. Usutu virus in Africa. *Vector Borne Zoonotic Dis*. 2011;11(11):1417-23. doi:10.1089/vbz.2011.0631.

110. Ben Hassine T, De Massis F, Calistri P, Savini G, BelHaj Mohamed B, Ranen A, et al. First detection of co-circulation of West Nile and Usutu viruses in equids in the south-west of Tunisia. *Transbound Emerg Dis*. 2014 Oct;61(5):385-9.

111. Durand B, Haskouri H, Lowenski S, Vachieri N, Beck C, Lecollinet S. Seroprevalence of West Nile and Usutu viruses in military working horses and dogs, Morocco, 2012: dog as an alternative WNV sentinel species? *Epidemiol Infect*. 2016 Jul;144(9):1857-64.

112. Mossel EC, Crabtree MB, Mutebi JP, Lutwama JJ, Borland EM, Powers AM, et al. Arboviruses isolated from mosquitoes collected in Uganda, 2008-2012. *J Med Entomol*. 2017 Sep 1;54(5):1403-9.

113. Engel D, Jost H, Wink M, Borstler J, Bosch S, Garigliany M, et al. Reconstruction of the evolutionary history and dispersal of Usutu Virus, a neglected emerging arbovirus in Europe and Africa. *MBIO*. 2016 Jan;7(1).

114. Ziegler U, Fast C, Eiden M, Bock S, Schulze C, Hoeper D, et al. Evidence for an independent third Usutu virus introduction into Germany. *Vet Microbiol*. 2016 Aug 30;192:60-6.

115. Domanović D, Gossner CM, Lieshout-Krikke R, Mayr W, Baroti-Toth K, Dobrota AM, et al. West Nile and Usutu virus infections and challenges to blood safety in the European Union. *Emerg Infect Dis*. 2019 Jun;25(6):1050-7.

116. Lustig Y, Sofer D, Bucris ED, Mendelson E. Surveillance and diagnosis of West Nile Virus in the face of flavivirus cross-reactivity. *Front Microbiol*. 2018;9:2421.

117. Cadar D, Simonin Y. Human Usutu virus infections in Europe: a new risk on horizon? *Viruses*. 2022 Dec 27;15(1).
118. Pacenti M, Sinigaglia A, Martello T, De Rui ME, Franchin E, Pagni S, et al. Clinical and virological findings in patients with Usutu virus infection, northern Italy, 2018. *Euro Surveill*. 2019 Nov;24(47).

119. Santini M, Vilibic-Cavlek T, Barsic B, Barbic L, Savic V, Stevanovic V, et al. First cases of human Usutu virus neuroinvasive infection in Croatia, August-September 2013: clinical and laboratory features. *J Neurovirol*. 2015 Feb;21(1):92-7.

120. Grottola A, Marcacci M, Tagliazucchi S, et al. Usutu virus infections in humans: a retrospective analysis in the municipality of Modena, Italy. *Clin Microbiol Infect*. 2017;23(1):33-7. doi:10.1016/j.cmi.2016.09.019

121. Cavrini F, Della Pepa ME, Gaibani P, Pierro AM, Rossini G, Landini MP, et al. A rapid and specific real-time RT-PCR assay to identify Usutu virus in human plasma, serum, and cerebrospinal fluid. *J Clin Virol*. 2011 Mar;50(3):221-3.

122. Faggioni G, De Santis R, Pomponi A, Grottola A, Serpini GF, Meacci M, et al. Prevalence of Usutu and West Nile virus antibodies in human sera, Modena, Italy, 2012. *J Med Virol*. 2018 Oct;90(10):1666-8.

123. Percivalle E, Cassaniti I, Sarasini A, Rovida F, Adzasehoun KMG, Colombini I, et al. West Nile or Usutu Virus? A Three-Year Follow-Up of Humoral and Cellular Response in a Group of Asymptomatic Blood Donors. *Viruses*. 2020 Jan 29;12(2).

124. Caracciolo I, Mora-Cardenas E, Aloise C, Carletti T, Segat L, Burali MS, et al. Comprehensive response to Usutu virus following first isolation in blood donors in the Friuli Venezia Giulia region of Italy: Development of recombinant NS1-based serology and sensitivity to antiviral drugs. *PLoS Negl Trop Dis*. 2020 Mar;14(3):e0008156.

125. Bella A, Venturi G, Riccardo F, Iapaolo F, F. Monaco FF, Calistri P. *Sorveglianza integrata del West Nile e Usutu virus. Risultati nazionali. Bollettino n. 20 del 2 novembre 2022.* Istituto Superiore di Sanità, IZS Teramo; 2022.

126. Vilibic-Cavlek T, Savic V, Sabadi D, Peric L, Barbic L, Klobucar A, et al. Prevalence and molecular epidemiology of West Nile and Usutu virus infections in Croatia in the ‘One health’ context, 2018. *Transbound Emerg Dis*. 2019 Sep;66(5):1946-57.

127. Cadar D, Maier P, Müller S, Kress J, Chudy M, Bialonski A, et al. Blood donor screening for West Nile virus (WNV) revealed acute Usutu virus (USUV) infection, Germany, September 2016. *Euro Surveill*. 2017 Apr 6;22(14):30501.

128. Agence Régionale de Santé – Nouvelle Aquitaine. Confirmation d'une infection autochtone à virus Usutu (secteurs des Landes et de Gironde). *Communiqué de presse*, 9 novembre 2022. Available from: https://www.nouvelle-aquitaine.ars.sante.fr/media/102015/download?inline; last visited July 2023.

129. Bakonyi T, Jungbauer C, Aberle SW, Kolodziejek J, Dimmel K, Stiasny K, et al. Usutu virus infections among blood donors, Austria, July and August 2017 - Raising awareness for diagnostic challenges. *Euro Surveill*. 2017 Oct;22(41).

130. Graninger M, Hubmer S, Riederer F, Kettner S, Hauk M, Auf T, et al. The first case of Usutu virus neuroinvasive disease in Austria, 2021. *Open Forum Infect Dis*. 2022 Jul;9(7):ofac255.

131. Zelená H, Kleinerová J, Šikutová S, et al. First autochthonous West Nile lineage 2 and Usutu virus infections in humans, July to October 2018, Czech Republic. *Pathogens*. 2021;10(6):651. doi:10.3390/pathogens10060651.

132. Nagy A, Mezei E, Nagy O, Bakonyi T, Csonka N, Kaposi M, et al. Extraordinary increase in West Nile virus cases and first confirmed human Usutu virus infection in Hungary, 2018. *Euro Surveill*. 2019 Jul;24(28).

133. Agliani G, Giglia G, Marshall EM, Gröne A, Rockx BHG, van den Brand JMA. Pathological features of West Nile and Usutu virus natural infections in wild and domestic animals and in humans: A comparative review. *One Health*. 2023;16:100525. doi:10.1016/j.onehlt.2023.100525
134. Giglia G, Agliani G, Munnink BBO, Sikkema RS, Mandara MT, Lepri E, et al. Pathology and Pathogenesis of Eurasian Blackbirds (Turdus merula) Naturally Infected with Usutu Virus. *Viruses*. 2021 Jul 28;13(8).

135. Wald ME, Sieg M, Schilling E, Binder M, Vahlenkamp TW, Claus C. The interferon response dampens the Usutu virus infection-associated increase in glycolysis. *Front Cell Infect Microbiol*. 2022;12:823181.

136. Segura Guerrero NA, Sharma S, Neyts J, Kaptein SJF. Favipiravir inhibits *in vitro* Usutu virus replication and delays disease progression in an infection model in mice. *Antiviral Res*. 2018 Dec;160:137-42.

137. Wald ME, Claus C, Konrath A, Nieper H, Muluneh A, Schmidt V, et al. Ivermectin inhibits the replication of Usutu virus *in vitro*. *Viruses*. 2022 Jul 27;14(8).

138. Martín-Acebes MA, Blázquez AB, Cañas-Arranz R, Vázquez-Calvo Á, Merino-Ramos T, Escribano-Romero E, et al. A recombinant DNA vaccine protects mice deficient in the alpha/beta interferon receptor against lethal challenge with Usutu virus. *Vaccine*. 2016 Apr 19;34(18):2066-73.

139. Allering L, Jöst H, Emmerich P, Günther S, Lattwein E, Schmidt M, et al. Detection of Usutu virus infection in a healthy blood donor from south-west Germany, 2012. *Euro Surveill*. 2012 Dec 13;17(50):20341.

140. Nagy A, Csonka N, Takács M, Mezei E, Barabás É. West Nile and Usutu virus seroprevalence in Hungary: A nationwide serosurvey among blood donors in 2019. *PloS One*. 2022;17(4):e0266840.

141. Europe. Directive 2002/98/EC of the European Parliament and of the Council of 27 January 2003 setting standards of quality and safety for the collection, testing, processing, storage and distribution of human blood and blood components and amending Directive 2001/83/EC. *Official Journal of the European Union* L 33, 8.2.2003

142. Berneck BS, Rockstroh A, Barzon L, Sinigaglia A, Vocale C, Landini MP, et al. Serological differentiation of West Nile virus- and Usutu virus-induced antibodies by envelope proteins with modified cross-reactive epitopes. *Transbound Emerg Dis*. 2022 Sep;69(5):2779-87.

143. Sinigaglia A, Pacenti M, Martello T, Pagni S, Franchin E, Barzon L. West Nile virus infection in individuals with pre-existing Usutu virus immunity, northern Italy, 2018. *Euro Surveill*. 2019 May;24(21).

144. Zaaijer HL, Slot E, Molier M, Reusken CBEM, Koppelman MHGM. Usutu virus infection in Dutch blood donors. *Transfusion*. 2019 Sep;59(9):2931-7.

145. Del Amo J, Sotelo E, Fernández-Pinero J, Gallardo C, Llorente F, Agüero M, et al. A novel quantitative multiplex real-time RT-PCR for the simultaneous detection and differentiation of West Nile virus lineages 1 and 2, and of Usutu virus. *J Virol Methods*. 2013 May;189(2):321-7.

146. Angeloni G, Bertola M, Lazzaro E, Morini M, Masi G, Sinigaglia A, et al. Epidemiology, surveillance and diagnosis of Usutu virus infection in the EU/EEA, 2012 to 2021. *Euro Surveill*. 2023 Aug;28(33).
# WEST NILE DISEASE

Annapaola Rizzoli (a), Giulia Mencattelli (a, b, c), Daniele Arnoldi (a), Maria Bellenghi (d), Claudia Cataldo (d), Francesca Dagostin (a), Di Luca Marco (e), Mitra Drakulovic (f), Souшиeta Jagadesh (g), Giovanni Marini (a), Beatriz Fernandez Martinez (h), Valentina Tagliapietra (a), William Wint (i), Luca Busani (d)

(a) Centro Ricerca e Innovazione, Fondazione Edmund Mach, San Michele all'Adige, Trento

(b) Centro Agricultura, Alimenti, Ambiente, Università di Trento, San Michele all'Adige, Trento

(c) Istituto Zooprofilattico Sperimentale dell'Abruzzo e del Molise, Teramo,

(d) Centro di Riferimento per la Medicina di Genere, Istituto Superiore di Sanità, Rome

(e) Dipartimento Malattie Infettive, Istituto Superiore di Sanità, Rome

(f) Department for Communicable Diseases Prevention and Control, National Public Health Institute "Dr Milan Jovanovic-Batut", Belgrade

(g) International Society of Infectious Diseases, Boston (MA), United States

(h) Centro Nacional de Epidemiología, Instituto de Salud Carlos III, Consorcio de Investigación Biomédica en Red Epidemiología y Salud Pública, Madrid

(i) Department of Biology, Environmental Research Group Oxford Ltd, Oxford

## Biological, ecological and molecular features of the causative agent

### Disease name

West Nile virus infection; West Nile virus disease; West Nile fever (WNF); West Nile virus neuroinvasive disease (WNND).

### Disease agent

#### Common, scientific and Latin name

West Nile virus (WNV).

#### Taxonomy

WNV is a mosquito-borne virus, part of the genus *Flavivirus*, family *Flaviviridae* and member of the Japanese Encephalitis antigenic serocomplex (1).

#### Disease agent characteristics

WNV is an enveloped virus of about 50 nm in diameter with an icosahedral symmetry. Like for other Flaviviruses, the genome of WNV is a single-stranded, positive-sense RNA, of about 11 kilobases (kb) enclosed in a nucleocapsid containing one long single open reading frame (ORF) confined at each side by one 5' and one 3' non-coding-region (UTR), respectively (23). The single ORF encodes for a polyprotein that is processed in three structural proteins (C, preM, and E), and seven non-structural proteins (NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5) (2). Each of the viral proteins, either structural or non-structural, plays a different and specific role in the biology and/or the pathogenesis of WNV infections. The non-structural proteins are involved in viral replication, virion assembly, and in mechanisms of host's immune response evasion (2), while
the three structural proteins make up the mature virion. In particular, the proteins M and E are
responsible for several virus properties, like host specificity, tissue tropism, replication capacity,
and cells T and B response induction (4).

WNV is part of the Japanese encephalitis virus serocomplex, sharing cross-neutralization
antibodies with viruses which are able to cause encephalitis in humans, as well as viruses which
rarely cause human disease, such as the Usutu virus (5).

Physiochemical properties

WNV is rapidly inactivated in the environment. Low temperatures preserve infectivity, with stability being greatest below -60°C. It is inactivated by heat (50 to 60°C for at least 30 minutes), ultraviolet light, and gamma irradiation.

The virus is also susceptible to disinfectants such as 3 to 8% formaldehyde, 2% glutaraldehyde, 2 to 3% hydrogen peroxide, 500 to 5,000 ppm available chlorine, alcohol, 1% iodine, and phenol iodophors. When added to Enzyme-Linked Immunosorbent Assay (ELISA) wash buffer there is a 10-fold decrease in titre per 24-hour period at 28°C. Ribavirin and interferon can inhibit WNV *in vitro*.

WNV is classified as a risk group level 3, requiring handling with biosafety level (BSL) 3 precautions (6).

Priority level for EU

WNV is considered a re-emerging public health challenge and a future health threat with significant economical implication in the European Union (EU) also in relation to the need to guarantee the quality and safety of substances of human origin intended for humans' application (SoHO) and thus to be monitored by epidemiological surveillance following a One Health integrated approach (7-14).

WNV infections became notifiable in the European Union (EU) since 2008, but surveillance with EU coverage was achieved later. The EU countries report human cases to the European Centre for Disease Prevention and Control (ECDC) which, in turn, produces annual epidemiological summaries and, since 2011, weekly surveillance updates during the WNV transmission season (May to November). EU countries also report animal outbreaks, mainly based on equine and birds' surveillance through the Animal Disease Information System (ADIS https://food.ec.europa.eu/animals/animal-diseases/animal-disease-information-system-adis_en).

Within the framework of the EU enlargement cooperation, EU enlargement countries also report human infections to the ECDC. The main objective of timely WNV surveillance at the EU level is to provide early warning to public health professionals about areas with human WNV infections thereby preventing human-to-human transmission via donation of contaminated SoHO. The EU blood safety directive obliges blood establishments to defer donors for 28 days after leaving an area where human cases were detected unless an individual donation nucleic acid test is negative (8, 12).

Distribution of the pathogen

Currently WNV is the widest distributed arbovirus globally, being found from tropical to north temperate latitudes on all the continents except Antarctica (15, 16). The virus itself has undergone
adaptive genetic changes while expanding its geographic distribution. The epidemic of 1999 in the New York City area and its expansion and establishment throughout the American continent highlighted the ability of this virus to leap from one hemisphere to another (17). WNV circulation in European Member States is known since 1958 and EU enlargement countries after 1972 (18) with an increase in the number of countries reporting local WNV transmission either in humans and animals in recent years (19, 20, 12, 21).

Since December 2022, at European scale, WNV human and animal infection have been reported in at least 15 EU countries including Austria, Bulgaria, Croatia, Cyprus, Czech Republic, France, Germany, Greece, Hungary, Italy, the Netherlands, Portugal, Romania, Slovenia and Spain, other than in 5 neighbouring EU candidate countries including Albania, Montenegro, Serbia (22), Turkey and Kosovo* (Figure 1 and Figure 2).

<figure><img src="image_69.png" /><figcaption>Figure 1. Countries of the European Union and European Economic Area that reported at least one locally acquired WNV human infection in the period 2010-2022. (Data from TESSy-ECDC and from other sources as literature and national reports)</figcaption></figure>

<footnote>* This designation is without prejudice to positions on status and is in line with United Nations Security Council Resolution 1244/99 and the International Court of Justice Opinion on the Kosovo declaration of independence.</footnote>
<figure><img src="image_70.png" /><figcaption>**Figure 2.** Distribution of WNV infections in animal hosts in the period 2010-2022, based on the data contained on the official reports (immediate notifications and follow-up reports, six-monthly reports and annual reports) submitted by the relevant Veterinary Services through the WAHIS-WOAH</figcaption></figure>

## Diversity and distribution of WNV lineages in Europe

WNV is characterized by high genetic diversity. Like other RNA viruses lacking proofreading replication, the WNV genome displays extraordinary adaptability with many variants that have evolved independently in different parts of the world. Phylogenetic analyses identified at least eight evolutionary lineages of which WNV lineages 1 (WNV-1) and 2 (WNV-2) are the most widespread and pathogenic, causing several outbreaks in humans and animals around the world (23-25). Moreover, studies have shown the high virulence of WNV-7, nowadays classified as a distinct flavivirus called Koutango virus (KOUTV), in mice and a potential risk for humans has been highlighted following a severe accidental infection in a Senegalese lab worker (26). The KOUTV is exclusively present in Africa, circulating in Senegal, Gabon, Somalia, and Niger, but its invasion into other Continents could represent a possible future threat worldwide (27).

In Europe, the maximum likelihood tree of all available WNV genome sequences showed that six lineages were detected so far in the Continent (25). Among them, WNV-1 and WNV-2 are the most widespread, responsible for an increasing number of outbreaks and deaths either in humans and animals. As the two lineages move from one area to another, principally through reservoir bird migration, strains of different origins can co-circulate in particular areas as seen in Italy (23,
24, 25). Back-and-forth exchanges between West Africa and Western-Mediterranean European countries have been recently proved for WNV-1 while WNV-2 seem to be historically characterized by two major independent one-way introductions from South Africa to Central Europe (Hungary), from where the lineage then spread, established, and co-circulated in many European countries (28).

WNV-2 accounts for 82% of all WNV sequences detected in the Continent so far, and the widest distribution since it has been found in 15 European countries (25).

The earliest WNV-2 genome (JX041631.1) was obtained from bird samples of Eastern Europe (Ukraine) in 1980 (25). The sub-lineage 2a (WNV-2a) emerged in 2004 in Hungary becoming in the past ten years the dominant lineage in Europe (29, 30). In addition, the sub-lineage 2b (WNV-2b) was reported and composed of sequences mainly of Romania, Italy, and Russia (2011-2015), Serbia (2013), and Greece (2018) (25, 29). WNV-2a has been involved in the exceptional number of infections occurred in 2018, a year characterized by an early start of the epidemic season and by the largest WNV outbreak observed so far in Central and Southern Europe, with over 2,000 symptomatic human cases, most of them reported in Italy and Serbia (24, 25, 31-33). The same year, WNV-2a emerged also in France and it was associated with the most important human WNV epidemics identified so far (34). In Spain, WNV-2a has only been detected in birds and mosquito in the Northeast (Catalonia) (35, 36).

In comparison, WNV-1 positive samples have been found in seven European countries (Austria, Italy, Spain, France, Hungary, Romania and Portugal) since 1971. Most WNV-1 sequences have been obtained in Italy (72% of total WNV-1 sequences), where the strain first appeared in Europe in 1998 (37). Most European WNV outbreaks were caused by this lineage, up to 2010, when the WNV-2 increased its circulation in the following year, causing serious epidemics in many European countries (34). Recently, WNV-1 is re-emerging in areas characterized by long unnoticed circulation as well as in new European countries and it can be associated with severe neurological forms and deaths in humans and animals (24, 23, 38). However, evidence of differences in WNV-1 and WNV-2 severity is still lacking.

WNV sequences belonging to lineages 3, 4, 8 and 9 were only sporadically reported and all of them were collected from mosquitoes. WNV-3 strains were only found in Czech Republic in 1997 and 2006; WNV-4 in Romania in 2012-13; WNV-8 in Spain in 2006, while WNV-9 genomes were obtained in Hungary in 2011 and in Austria in 2013. In addition, up to 2021, these lineages were only collected from non-human hosts (mainly birds, mosquitoes, and some equines) (25).

## Ecology and transmission routes

WNV is an ecological generalist, and it is characterized by a complex eco-epidemiology, being adaptable to different vector and host species, principally mosquitoes of the genus *Culex* and birds (Figure 3). Its circulation in Europe is usually associated with two main different cycles and habitats: the rural/sylvatic and the urban synanthropic cycles. Rural locations, including river deltas and floodplain areas, help creating the sylvatic cycle, with wild, usually nesting wetland, birds and ornithophilic mosquitoes maintaining the viral transmission. Irrigation from agriculture is also heavily linked to a greater incidence of human and veterinary WNV infections (20, 39). In urban synanthropic cycles, mosquitoes feeding preference shift can determine the enhancement of the viral transmission to humans (7). However, these two cycles can overlap, so urban areas located close to wetlands and irrigated croplands can be particularly affected by the virus circulation.

Humans and equids are highly susceptible to the infection but do not develop a sufficient level of viremia to further infect mosquitoes and are therefore considered dead-end non-competent
hosts. A minimum viremia level of $10^4-10^5$ PFU/mL has been established as necessary for infecting feeding *Culex* mosquitoes and thus allowing further virus transmission (40).

<figure><img src="image_71.png" /><figcaption>**Figure 3.** West Nile virus transmission cycle</figcaption></figure>

Mosquitoes mostly acquire infection horizontally, while feeding on an infected host, but vertical transmission (passage of virus from female parent directly to offspring) is also possible, although apparently less efficient (23, 41-43).

Several abiotic and biotic factors, such as climatic condition, habitat structure, anthropization, land cover and management, host communities' composition, host herd immunity, host behaviour, mosquito community composition, and vector host preferences, affect the virus transmission within vectors and hosts (7, 42, 44-49).

Seasonality in WNV transmission is also dependent on vector mosquitoes' annual phenology, which can vary according to the species and the local environmental condition. For example, a recent study provides new evidence that urban warming may inhibit the *Cx. pipiens* diapause initiation during autumn, thereby extending the active biting season of temperate mosquitoes (50).

Because of these features, outbreaks of WNV infection exhibit high variability in their extension across different regions and time (45).

## Vectors

Mosquito species of the genus *Culex* have a worldwide distribution and are considered among the species of greater medical importance as they can act as vectors for various zoonotic arboviruses from several virus families (51,52). Among them, *Cx. pipiens* mosquito species are considered the most important WNV vectors in Europe (53). Transmission rates vary between 0 and 60% with no intrinsic difference in vector competence between northern and Southern populations (48). Moreover, the species comprise two behaviourally different biotypes, *Cx. pipiens* (more ornithophilic and abundant in natural habitat) and *Cx. molestus* (preference for
mammals and more opportunistic feeding behaviour) which can form hybrids. Biotype *molestus* and hybrids *pipiens-molestus* are thought to play a more important role in WNV spillover from birds to humans especially in urban and periurban areas (7, 54), other than in overwintering mechanisms (23). No difference has been observed in the vector competence among the two biotypes. However, higher temperatures increase the transmission rates of biotype *pipiens* and hybrids, but not of biotype *molestus*, the latter mainly distributed in Southern and Central European countries, and considered an efficient WNV "bridge" vector, being able to transmit the pathogen between birds as well as from birds to mammals, including humans with high transmission rates of 40-55% (52).

This shows the importance of identifying *Cx. pipiens* at the biotype level and suggests that closely related mosquitoes may have different vector competence under variable climatic conditions (48, 54-56).

Among other *Culex* species, *Cx. torrentium* also exhibit high WNV transmission values observed in the laboratory, with rates of 17% (24°C) and 24% (27°C) although its role needs to be confirmed with further field studies (51). In Europe, *Cx. torrentium* usually occurs together with *Cx. pipiens*, with the first being the dominant species in northern Europe and the second prevailing in regions south of the Alps. In Central Europe such as Austria or Germany, both sister species can be found in sympatry (51, 57). It is therefore necessary to distinguish among the two species using molecular techniques (53).

*Cx. modestus*, which is distributed mainly in Southern and Central European countries, is also considered an efficient WNV "bridge" vector, being able to transmit pathogens between birds as well as from birds to mammals, including humans with high transmission rates of 40-55% (52).

Finally, *Cx. perexiguus* is established in Southwest Spain, related to rice fields. In some areas where the disease is endemic, it has been found to play an important role as the main human vector (58). In Figures 4 and 5, maps with the probability of occurrence of *Cx. torrentium*, *Cx modestus* and *Cx. pipiens* in Europe are reported.

<figure><img src="image_72.png" /><figcaption>**Figure 4.** Current 1-km probability of presence of *Cx. torrentium* and *Cx modestus* across Europe, produced using random forest and boosted regression trees analyses (source ERGO group)</figcaption></figure>
<figure><img src="image_73.png" /><figcaption>Figure 5. Current 1-km probability of presence of *Cx. pipiens* across Europe, produced using random forest and boosted regression trees analyses (source ERGO group)</figcaption></figure>

WNV vector competence and transmission rates have also been compared for various European mosquito species other than *Culex* species as in the case of *Aedes* (*Ae. albopictus*, *Ae. detritus*, *Ae. japonicus japonicus*, *Ae. vexans*) and *Ochlerotatus caspius* (43, 48, 51, 53, 57). In some experimental studies, *Ae. japonicus*, which has recently spread over considerable parts of Central Europe, has proven to be more efficient than *Cx. pipiens* (63, 76). *Ae. vexans* may also be epidemiologically important due to its generally high abundance despite its moderate competence proved in laboratory (43).

Although vector competence has been determined for a limited number of species and populations of the same mosquito species in different areas, WNV has been isolated from, or detected in, field-collected specimens of numerous mosquito species native to Europe, as shown in Table 1.

WNV has also been isolated from other arthropods other than mosquitoes including hard ticks (*Hyalomma marginatum* and *Rhipicephalus sanguineus*), soft ticks (*Ornithodoros maritimus* and *Argas hermanni*), swallow bug (*Cimex hirundinis*), and chicken mites (*Ornithonyssus sylviarum*). However, the contribution of ticks in the natural WNV transmission cycles and dispersal has not been clarified yet (16, 23).
<table><caption>Table 1. A list of mosquito species confirmed as competent vectors for West Nile virus in Europe</caption><thead><tr><td>Genus</td><td>Species</td><td>Reference</td></tr></thead><tbody><tr><td>Culex</td><td>pipiens</td><td>(59-63)</td></tr><tr><td>Culex</td><td>pipiens pipiens</td><td>(48, 64)</td></tr><tr><td>Culex</td><td>pipiens molestus</td><td>(64)</td></tr><tr><td>Culex</td><td>pipiens hybrid (pip x mol)</td><td>(64)</td></tr><tr><td>Culex</td><td>modestus</td><td>(62, 63, 65)</td></tr><tr><td>Culex</td><td>perexiguus</td><td>(66, 67)</td></tr><tr><td>Culex</td><td>quinquefasciatus</td><td>(68)</td></tr><tr><td>Culex</td><td> torrentium</td><td>(51, 69)</td></tr><tr><td>Culex</td><td>theileri</td><td>(63)</td></tr><tr><td>Culex</td><td>univittatus</td><td>(67)</td></tr><tr><td>Culex</td><td>europaeus</td><td>(70)</td></tr><tr><td>Culex</td><td>tritaeniorhynchus</td><td>(71)</td></tr><tr><td>Aedes</td><td>vexans</td><td>(55, 61, 63, 68)</td></tr><tr><td>Aedes</td><td>caspius</td><td>(62, 63)</td></tr><tr><td>Aedes</td><td>albopictus</td><td>(61, 63, 72)</td></tr><tr><td>Aedes</td><td>detritus</td><td>(63, 73)</td></tr><tr><td>Aedes</td><td>dorsalis</td><td>(63, 74)</td></tr><tr><td>Aedes</td><td>geniculatus</td><td>(63, 75)</td></tr><tr><td>Aedes</td><td>japonicus</td><td>(63, 76)</td></tr><tr><td>Aedes</td><td>punctor</td><td>(75)</td></tr><tr><td>Aedes</td><td>cinereus</td><td>(77)</td></tr><tr><td>Uranotaenia</td><td>unguiculata</td><td>(78)</td></tr><tr><td>Anopheles</td><td>plumbeus</td><td>(63, 75)</td></tr><tr><td>Anopheles</td><td>maculipennis</td><td>(77, 79, 56)</td></tr><tr><td>Coquillettidia</td><td>richiardii</td><td>(20, 79)</td></tr><tr><td>Ochlerotatus</td><td>cantans</td><td>(20, 77)</td></tr><tr><td>Ochlerotatus</td><td>sticticus</td><td>(70)</td></tr><tr><td>Culiseta</td><td>longiareolata</td><td>(20)</td></tr><tr><td>Culiseta</td><td>morsitans</td><td>(20)</td></tr></tbody></table>

## Hosts

WNV is able to infect a wide range of animal species, including birds, mammals, reptiles and amphibians, which can be infected, develop antibodies and, in some cases, clinical signs. However, several infected hosts are considered dead-end hosts since they are unable to further transmit the virus to mosquitoes (80). Competent reservoir hosts are found especially among bird species and it is generally acknowledged that *Passeriformes* (especially *Corvidae*, *Fringillidae*, and *Passeridae* families) *Charadriiformes* (*Laridae*), *Falconiformes*, and *Strigiformes* include highly competent species, although viraemic levels vary depending on species, viral lineage and strain (40, 80, 81). Domestic birds are generally less susceptible to infection; domestic chickens and turkeys do not develop clinical infection, while domestic geese, ducks and companion birds (e.g. *Psittaciformes*) can suffer encephalitis. Among mammals, high and long-lasting viremia sufficient to potentially infect vectors has been experimentally and/or naturally demonstrated in lemurs, small rodents, lagomorphs, amphibians, and reptiles (reviewed in 40, 80, 2, 82). However, most of experimental infections to assess reservoir competence have been conducted on bird species and rarely on wild mammals in Europe (40, 82-86). As such, the role of wild mammals as potential WNV amplifying hosts is uncertain, with most experimental infections resulting in detectable antibodies but no to low-level of viremia. Certain mammalian species, such as tree
squirrels, mesopredators, pigs, wild boars, and roe deer have been suggested as species of interest for WNV serological surveillance (87).

In Table 2 a list of European vertebrate species susceptible to WNV infection and with a viremia level above 4 log10 plaque-forming unit (PFU/mL) is reported. According to (40), species with viremia level ≤ 4 log10 PFU/mL (low viremia) are considered non-competent hosts. Species with medium viremia (mean peak viremia 4-6 log10 PFU/mL) and high viremia (mean peak viremia > 6 log10 PFU/mL) are considered competent hosts.

<table><caption>Table 2. List of vertebrate species with reported WNV medium and high viremia level</caption><thead><tr><td>Genera</td><td>Species</td><td>Common name</td><td>References</td><td>Note</td></tr></thead><tbody><tr><td>Alectoris</td><td>rufa</td><td>Red-legged partridge</td><td>(40, 89)</td><td></td></tr><tr><td>Anas</td><td>platyrhynchos</td><td>Mallard</td><td>(40, 90)</td><td></td></tr><tr><td>Anser</td><td>anser</td><td>Common goose</td><td>(40)</td><td></td></tr><tr><td>Anser</td><td>anser domesticus</td><td>Domestic geese</td><td>(100)</td><td>in 1- to 11-day old chicks</td></tr><tr><td>Branta</td><td>canadensis</td><td>Canada goose</td><td>(40, 90)</td><td></td></tr><tr><td>Charadrius</td><td>vociferus</td><td>Killdeer</td><td>(90)</td><td>migratory through Europe</td></tr><tr><td>Columba</td><td>livia</td><td>Rock pigeon</td><td>(40, 92)</td><td></td></tr><tr><td>Columba</td><td>livia</td><td>Rock dove</td><td>(90, 102)</td><td></td></tr><tr><td>Corvus</td><td>monedula</td><td>European jackdaws</td><td>(83)</td><td></td></tr><tr><td>Corvus</td><td>cornix</td><td>Hooded crow</td><td>(40)</td><td></td></tr><tr><td>Corvus</td><td>corone</td><td>Carrion crows</td><td>(104)</td><td></td></tr><tr><td>Coturnix</td><td>coturnix</td><td>Quails</td><td>(95)</td><td></td></tr><tr><td>Falco</td><td>rusticolus</td><td>Gyrfalcons</td><td>(38, 40)</td><td></td></tr><tr><td>Falco</td><td>rusticolus, cherrug</td><td>Hybrid falcon</td><td>(38)</td><td></td></tr><tr><td>Falco</td><td>rusticolus, peregrinus</td><td>Hybrid falcon</td><td>(38)</td><td></td></tr><tr><td>Felis</td><td>catus</td><td>Domestic cat</td><td>(99)</td><td>10<sup>3</sup> to 10<sup>4</sup> PFU/mL</td></tr><tr><td>Gallus</td><td>gallus</td><td>Chickens</td><td>(101)</td><td>young chickens - 7-week old</td></tr><tr><td>Larus</td><td>delawarensis</td><td>Ring-billed gull</td><td>(90)</td><td>occasionally migrate through Europe</td></tr><tr><td>Mesocricetus</td><td>auratus</td><td>Hamster</td><td>(96, 97)</td><td>10<sup>5</sup> TCID 50/mL</td></tr><tr><td>Mus</td><td>musculus</td><td>Lab mouse</td><td>(105)</td><td></td></tr><tr><td>Passer</td><td>domesticus</td><td>House sparrow</td><td>(40, 84, 90)</td><td></td></tr><tr><td>Pica</td><td>pica</td><td>Magpie</td><td>(103)</td><td></td></tr><tr><td>Procyon</td><td>lotor</td><td>Raccoon</td><td>(98)</td><td></td></tr><tr><td>Rana</td><td>ridibunda</td><td>Lake frog</td><td>(94, 95)</td><td></td></tr><tr><td>Sciurus</td><td>carolinensis</td><td>Grey squirrel</td><td>(88)</td><td></td></tr><tr><td>Streptopelia</td><td>decaocto</td><td>Eurasian collared-dove</td><td>(40, 93)</td><td></td></tr><tr><td>Sturnus</td><td>vulgaris</td><td>European starling</td><td>(40, 90)</td><td></td></tr></tbody></table>

TCID 50: Tissue Culture Infectious Dose (TCID 50)

In Europe, several eco-epidemiological investigations have been carried out on wild birds. In Italy, one of the most affected EU countries, the infection has been reported in 19 bird orders and 44 species, as reported by the National Reference Centre for Foreign Animal Diseases (CESME) at the Istituto Zooprofilattico Sperimentale of Abruzzo and Molise in Teramo (IZSAM-Teramo) (https://westnile.izs.it). For instance, in the Emilia Romagna region, one of the most affected areas in Europe, surveillance on wild birds, carried out from 2015 to 2019 revealed that *Galliformes* and *Strigiformes* scored the highest prevalence of infection detected by PCR (13%). They were
followed by *Columbiformes* (10%) and *Charadriformes* (7%). The order *Passeriformes* showed an average prevalence of 5%, with European blackbird (*Turdus merula*) accounting for a prevalence of 4.3%. Notable infected species were greenfinch (*Carduelis chloris*), scops owl (*Otus scops*), house martin (*Delichon urbica*), house sparrow (*Passer domesticus*), and pheasant (*Phasianus colchicus*) (106).

In Spain, sporadic WNV outbreaks observed in horses and wild birds from Extremadura (western Spain) during 2016 and 2017 seasons prompted a survey in wild birds, focused on specimens coming from two wildlife rehabilitation centres. Between October 2017 and December 2019, samples from 391 wild birds, belonging to 56 different species, were collected and analysed in search of evidence of WNV infection. The analysis of serum samples for WNV-specific antibodies by ELISA, whose specificity was subsequently confirmed by virus-neutralisation test (VNT) showed positive results in 18.23% birds belonging to 18 different species. *Pelecaniformes* (33.33%), *Accipitriformes* (25.77%) and *Strigiformes* (22.92%) orders had the higher seroprevalence. Remarkably, for the first time in Europe, WNV-specific antibodies were found in a black stork (*Ciconia nigra*). Analysis by real time RT-PCR in symptomatic birds confirmed the presence of WNV-1 RNA in griffon vulture (*Gyps fulvus*) and little owls (*Athene noctua*) (107). Moreover, following the largest epidemic outbreak ever reported in Spain, another cross-sectional study was conducted to assess the circulation and risk factors associated with WNV exposure in wild bird populations. Results showed that group species (raptors), age (>1-year old), and size (large) were the main risk factors related to WNV seropositivity in wild birds (108). Seropositivity was found in 37.8% of the 37 species analysed.

In northern Serbia, of 92 wild bird sera tested, seven (8%) were IgG ELISA positive. They belonged to three species: four Mute Swans (*Cygnus olor*); two White-tailed Eagles (*Haliaeetus albicilla*) and one Common Pheasant (*Phasianus colchicus*) (109).

Migratory birds possibly favor the WNV long-distance dispersal. Getting infected prior to or during migration, birds may carry the virus in their blood (and other tissues) over long distances and eventually infect mosquitoes and/or their predators in destination territories (110). As well as migratory birds considered to have an essential role in long distance spread, resident birds seem to have an essential role in long distance spread, resident birds have a paramount importance in local dispersal and endemization of flaviviruses at a smaller and medium spatial scale. In particular, resident bird species such as waterfowls, corvids, or birds of prey have been shown to be highly sensible to WNV (110). For instance, raptors, considered WNV reservoir and amplifying hosts of infection, are usually found to be seriously affected during WNV outbreaks, demonstrating their potential role in the virus seasonal introduction and circulation (81, 111, 108). Moreover, higher prevalence and susceptibility to WNV might point out the existence of infection through predation of infected prey in these birds. Consequently, they are considered important target species when designing cost-effective surveillance for monitoring both seasonal WNV circulation in endemic countries and its emergence into new areas (81, 112).

Corvids, such as the Eurasian magpie (*Pica pica*), the hooded crow (*Corvus corone cornix*), and the Eurasian jay (*Garrulus glandarius*), are also known to be susceptible to WNV infection. These birds are periodically shot or trapped to contain their abundance and they are screened as sentinel for WNV circulation in Italy since 2012 (113). However, the trend of infection observed in these species is indicative, but not exhaustively, of the intensity of WNV circulation. Other species, such as the small passerine birds, might be involved in WNV transmission in Europe, as previously observed in other continents such as the USA (110). For instance, the extensive capturing and sampling of wild birds in the Netherlands has allowed the recent report of the first locally acquired WNV detection in passerines, thus revealing their potential role in enzootic transmission (114).
Knowledge gaps related to bird role in circulation of WNV in Europe remain; the reservoir competence and immunological response to the infection for many European avian species is still not known (40, 7, 110, 82) and further studies in this direction are needed.

# Drivers of disease emergence and spread

## Ecological drivers

WNV displays high ecological plasticity, being able to adapt to different vector-hosts assemblages. In a recent systematic review carried out to identify the most relevant environmental factors affecting the WNV spread in Europe those which appear to exert the major effect were changes observed in temperature and precipitation patterns and the expansion of anthropized habitats (49). Environmental factors modulate the vector presence and abundance, and their Extrinsic Incubation Period (EIP) while human-derived changes in landscape may provide favorable conditions for mosquito breeding and susceptible host assemblage, especially in urban and peri-urban areas, other than in agricultural landscapes (44, 49, 115, 116).

In particular, temperature is one of the most important variables being able to affect the whole WNV transmission system. Firstly, it affects the mosquito population cycle and abundance, since it determines both the start-point and the duration of the vector's season and it is also associated with mosquitoes' population density and geographical distribution (117).

There is an optimal known range of temperatures (25-35°C) for the mosquitoes to develop and *Cx. pipiens* embryonic development cannot be completed below 7°C. Both larval development and adult survival are shorter at higher temperatures being the highest survival at 25°C. Vector abundance is a good predictor of WNV risk, also in case of *Cx. modestus* other than of *Cx. pipiens* (116).

The effect of environmental temperatures on WNV establishment in Europe via *Cx. pipiens* populations through use of a basic reproduction number (R0) model was assessed (118). WNV establishment was determined to be possible between 14°C and 34.3°C, with the optimal temperature at 23.7°C. The widespread thermal suitability for WNV establishment highlights the importance of European surveillance and the need for increased research into mosquito and bird distribution (118).

Furthermore, temperature plays a significant role in the vector competence, extrinsic incubation period, and intensity of infection of WNV within mosquito vectors. This mechanism then allows the vectors to transmit WNV earlier by shortening the gonotrophic cycle, resulting also in an increased biting rate although the effect can differ among *Cx. pipiens* biotypes (64). The average northern European summer temperatures of 18°C appear to be an important limiting factor for WNV transmission (48).

The temperature exerts an indirect effect on the intensity and duration of the outbreaks and it appears to be a reliable early warning predictor of its occurrence (44, 119, 120). Also, the mean temperature of the warmest quarter of the preceding year is considered one of the most important drivers of WNV outbreak European-wide (116). Likewise, a higher than usual spring temperature in this range (22-26°C) early in a year could also be a precursor of WNV-outbreak during the latter half of the same year (116). Similarly, temperature can be used as a predictor of vector abundance and its infection rate, as observed in Serbia where the number of collected *Cx. pipiens* mosquitoes and the monthly distribution of WNV-positive pools followed a similar profile over time as the mean temperature, between April and October 2013 in Vojvodina province (56).

In a recent study aimed at modelling the WNV Force Of Infection (FOI), spring temperatures were positively associated with FOI (121). Similarly, the timing of the peak of WNV incidence
appears to be influenced only by summer temperatures while the infection peak tends to be
demonstrated to be earlier when summer temperature is higher. Furthermore, higher temperature
is correlated with high WNV genetic diversity during the entire history of WNV-2a spread in
Europe (25). Moreover, studies carried out in France, Greece, Italy and Serbia revealed that
average temperature was a consistently good predictor across sites. In South Banat district (NUTS
level 3) in Serbia the maximum temperature of warmest part of the year and the annual
temperature range, as, well as, the presence of rivers (Danube and Tamis rivers) and its changes
in water streams were the best environmental predictors of WNV infection outbreaks in time
period 2017-2019 (122).

Regarding precipitation, ecological studies suggest that drought events can lead to outbreaks
in the following year due to changes in the mosquito food web structure. Lower precipitation in
winter implies less water availability in the region and such conditions are a likely indicator of
the aggregation of host birds and the vectors at available water bodies, which could amplify the
virus transmission rates (44, 116).

Human driven activities which determine land use and urbanization have also major effects on the vector and host abundance and community composition (49, 115, 123). The differences of WNF incidence rates among European Mediterranean countries with similar weather conditions could be explained by differences in the human interaction with suitable habitats such as wetlands or flooded fields. Areas with high levels of agricultural activities may accelerate WNV dispersal velocity as well as attract the spread direction of WNV in Europe. Meanwhile, WNV is likely to spread in urbanized areas, in line with high abundance of *Cx. pipens* which is a species able to maintain significant populations in cities (49). However, the WNV FOI is expected to be higher in anthropized semi-natural areas such as populated forests, wetlands and river basins (44, 121).

There are other local factors which could affect WNV circulation. For instance, regional
differences in mosquito species distribution, including the occurrence and rate of hybrid
population of *Cx. pipiens*, and vector competence might be crucial (25, 46). Furthermore, the
different dynamic histories of WNV variants (WNV-2a and WNV-2b) might also be associated
with the presence of different mosquito species assemblages, which needs further investigation
(25).

Among biotic factors, also bird community composition affects WNV intensity of circulation and based on available data (123), developed a structural bird borne WND risk map. However, data on presence, abundance, phenology, migration at municipal and provincial scale of the majority of the bird species of relevance for WNV ecology are usually lacking, therefore representing a significant limitation in risk modelling (123).

Bird species vary in their susceptibility to WNV infection and role in the ecology of the
disease. Migratory birds are important not only by the fact of being a reservoir but also for the
possibility of introducing new virus lineages or strains. However, resident and short-range
migratory birds play an important role in virus dispersal. For example, WNV-2a in Europe spread
at a high spread rate (88 km/year to 218 km/year) and, therefore, is more likely correlated with
short migrant bird movement than the flight range of *Culex* mosquitoes (approx. 500 m to 2
km/year) (25). Therefore, although the risk of WNV depends on both the presence of infected
birds and the presence of competent mosquitoes transmitting the disease, bird movements seed
the infections into mosquito populations occurring far away and introduce WNV into new regions.
A better understanding of bird migration pattern and phenology at European scale under a global
change scenario is essential for a better understanding of WNV temporal and spatial dynamics
(124).